• Xylanase;
  • Regulation of biosynthesis;
  • Extremophile;
  • Overexpression;
  • Protein engineering;
  • Domain structure


  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Scope of the present review
  5. 3Structure of xylan
  6. 4Xylanase production
  7. 5Regulation of xylanase synthesis
  8. 6Biochemical properties
  9. 7Xylanases of extremophilic origin
  10. 8Cloning and expression of xylanase gene(s)
  11. 9Protein engineering
  12. 10Site-directed mutagenesis (SDM)
  13. 11Mechanism of action of the xylanases
  14. 12Domain organization of xylanases
  15. 13Molecular evolution
  16. 14Biotechnological potentials of xylan and xylanases
  17. 15Future prospects
  18. Acknowledgements
  19. References

Hemicellulolytic microorganisms play a significant role in nature by recycling hemicellulose, one of the main components of plant polysaccharides. Xylanases (EC catalyze the hydrolysis of xylan, the major constituent of hemicellulose. The use of these enzymes could greatly improve the overall economics of processing lignocellulosic materials for the generation of liquid fuels and chemicals. Recently cellulase-free xylanases have received great attention in the development of environmentally friendly technologies in the paper and pulp industry. In microorganisms that produce xylanases low molecular mass fragments of xylan and their positional isomers play a key role in regulating its biosynthesis. Xylanase and cellulase production appear to be regulated separately, although the pleiotropy of mutations, which causes the elimination of both genes, suggests some linkage in the synthesis of the two enzymes. Xylanases are found in a cornucopia of organisms and the genes encoding them have been cloned in homologous and heterologous hosts with the objectives of overproducing the enzyme and altering its properties to suit commercial applications. Sequence analyses of xylanases have revealed distinct catalytic and cellulose binding domains, with a separate non-catalytic domain that has been reported to confer enhanced thermostability in some xylanases. Analyses of three-dimensional structures and the properties of mutants have revealed the involvement of specific tyrosine and tryptophan residues in the substrate binding site and of glutamate and aspartate residues in the catalytic mechanism. Many lines of evidence suggest that xylanases operate via a double displacement mechanism in which the anomeric configuration is retained, although some of the enzymes catalyze single displacement reactions with inversion of configuration. Based on a dendrogram obtained from amino acid sequence similarities the evolutionary relationship between xylanases is assessed. In addition the properties of xylanases from extremophilic organisms have been evaluated in terms of biotechnological applications.


  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Scope of the present review
  5. 3Structure of xylan
  6. 4Xylanase production
  7. 5Regulation of xylanase synthesis
  8. 6Biochemical properties
  9. 7Xylanases of extremophilic origin
  10. 8Cloning and expression of xylanase gene(s)
  11. 9Protein engineering
  12. 10Site-directed mutagenesis (SDM)
  13. 11Mechanism of action of the xylanases
  14. 12Domain organization of xylanases
  15. 13Molecular evolution
  16. 14Biotechnological potentials of xylan and xylanases
  17. 15Future prospects
  18. Acknowledgements
  19. References

The naturally occurring lignocellulosic plant biomass consists of 20–30% hemicellulosic materials which are heterogeneous polysaccharides found in association with cellulose [1]. Biomass is an alternative natural source for chemical feedstocks with a replacement cycle short enough to meet the demand in the world fuel market. Xylan is the major constituent of hemicellulose and is the second most abundant renewable resource with a high potential for degradation to useful end products. Hence the development of inexpensive technologies based on hemicellulose is called for. Eventually, if not in the near future, xylan, in combination with cellulose, will supply most of the global demand for raw materials. It is not unrealistic to foresee that coal and crude oil are likely to be substituted by biomass in another 50 years [2]. Microbial xylanases (1,4-β-d-xylan xylanohydrolase, EC are the preferred catalysts for xylan hydrolysis due to their high specificity, mild reaction conditions, negligible substrate loss and side product generation. However, the cost of enzymic hydrolysis of biomass is one of the factors limiting the economic feasibility of the process. The production of xylanases must therefore be improved by finding more potent fungal or bacterial strains, or by inducing mutant strains to excrete greater amounts of enzymes, or both. Xylanases are produced by a plethora of organisms like bacteria, algae, fungi, protozoa, gastropods, and arthropods [3]. Most of the bacteria and fungi secrete extracellular xylanases which act on the hemicellulosic material to liberate xylose as a directly assimilable end product allowing the organisms to grow heterotrophically on xylan. Ruminal microorganisms are known to be potent xylanase producers, possibly due to the high dietary hemicellulose content of the feed of ruminant animals. The use of hemicellulolytic enzymes as a substitute for chlorine chemicals in pulp bleaching has recently attracted considerable interest because of environmental concerns. The limited hydrolysis of hemicellulose in pulps by hemicellulases, mainly xylanases, increases the extractability of lignin from the kraft pulp in the subsequent bleaching sequences, reducing the chloro-organic discharges.

2Scope of the present review

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Scope of the present review
  5. 3Structure of xylan
  6. 4Xylanase production
  7. 5Regulation of xylanase synthesis
  8. 6Biochemical properties
  9. 7Xylanases of extremophilic origin
  10. 8Cloning and expression of xylanase gene(s)
  11. 9Protein engineering
  12. 10Site-directed mutagenesis (SDM)
  13. 11Mechanism of action of the xylanases
  14. 12Domain organization of xylanases
  15. 13Molecular evolution
  16. 14Biotechnological potentials of xylan and xylanases
  17. 15Future prospects
  18. Acknowledgements
  19. References

The various applications of xylanases have stimulated research on the biochemical and molecular aspects of this important enzyme of the family of glycosyl hydrolases. Considering the future prospects of xylanases for biotechnological exploitations, especially in the field of biopulping and bleaching, it is very essential to analyze the properties of xylanases with respect to regulation of biosynthesis and mechanism of action. An increasing number of publications in recent years describe numerous xylanases from new sources, their cloning, sequencing, mutagenesis and crystallographic analysis. Many of these reports are about hemicellulase-aided bleaching of kraft pulp used in the paper industry, which truly exploits the unique specificity and safety of use of biocatalysts. Since xylanases are industrially important enzymes, many reviews have been published covering the various aspects of the enzyme [4–8]. The present article is a comprehensive state-of-the-art review describing the xylanases with special emphasis on the latest developments in the area of molecular biology and biotechnology. The article also covers the xylanases from extremophilic microorganisms which have gained importance by virtue of their stability and activity at extremes of pH and temperature. Theoretical analysis of a few of the reported thermostable xylanases has also been included; it will be useful in application-oriented protein engineering studies. The recent reviews on related topics complement the present work [9,10].

3Structure of xylan

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Scope of the present review
  5. 3Structure of xylan
  6. 4Xylanase production
  7. 5Regulation of xylanase synthesis
  8. 6Biochemical properties
  9. 7Xylanases of extremophilic origin
  10. 8Cloning and expression of xylanase gene(s)
  11. 9Protein engineering
  12. 10Site-directed mutagenesis (SDM)
  13. 11Mechanism of action of the xylanases
  14. 12Domain organization of xylanases
  15. 13Molecular evolution
  16. 14Biotechnological potentials of xylan and xylanases
  17. 15Future prospects
  18. Acknowledgements
  19. References

The β-1,4-xylans are heteropolysaccharides with a homopolymeric backbone chain of 1,4-linked β-d-xylopyranose units. The backbone consists of O-acetyl, α-l-arabinofuranosyl, α-1,2-linked glucuronic or 4-O-methylglucuronic acid substituents [11]. However, unsubstituted linear xylans have been isolated from guar seed husk, esparto grass and tobacco stalks [12]. Wood xylans exist as O-acetyl-4-O-methylglucuronoxylans in hardwoods or as arabino-4-O-methylglucuronoxylans in softwoods. The degree of polymerization of hardwood xylans (150–200) is higher than that of softwoods (70–130) [13]. The cereal xylans are made up of d-glucuronic acid and/or its 4-O-methyl ether and arabinose [14]. Endospermic arabinoxylans of annual plants, also called pentosans, are more soluble in water and dilute alkali than xylans of lignocellulosic materials because of their branched structures [15].

3.1Chemical structure

Based on the common substituents found on the backbone, xylans are categorized as linear homoxylan, arabinoxylan, glucuronoxylan and glucuronoarabinoxylan. However, in each category there exists a microheterogeneity with respect to the degree and nature of branching. The basic backbone and the possible substituent groups are shown in Fig. 1A. Xylans are present in a partly acetylated form in various plants. The O-acetyl groups present at C-2 and C-3 positions of xylosyl residues inhibit xylanases from completely degrading acetylxylan, probably by steric hindrance. The synergistic action of acetylxylan esterases and xylanases is therefore essential for the complete hydrolysis of acetylxylans. The presence of small amounts of feruloyl and p-coumaroyl acids linked via l-arabinose residues has been shown on the structure of xylan in several studies. These hydroxycinnamic acids are bound to C-5 of the arabinose residue [16]. The presence of a covalent bond between lignin and hemicellulose, perhaps through xylan substituents in many cases, has been documented. Evidence for the existence of an ether linkage between arabinose and lignin [13] and an ester linkage between glucuronic acid and lignin [17] has also been shown. Feruloyl groups may also crosslink xylan and lignin [18]. 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.


Figure 1. A: Chemical structure of xylan. B: Three-dimensional structure. Projections perpendicular (top) and parallel (bottom) of (a) xylan backbone, (b) backbone and one l-arabinose side group: (c) backbone and two l-arabinose side groups. Hydrogen bonds are shown dotted. In each case the backbone is a left-handed threefold helix. (Based on [19].)

3.2Three-dimensional structure

The three-dimensional structure of the xylan molecules has been elegantly described by Atkins [19]. The xylan backbone shows a threefold left-handed conformation under crystallized conditions; the geometry of the glycosidic linkage is not affected by the side chains (Fig. 1B). Studies of the polysaccharide indicate that the backbone imposes certain minimum constraints and the interactions between the chains determine the final conformation. Electron diffraction patterns also confirm the threefold conformation and show that the chains are organized in a trigonal lattice with hexagonal morphology. The single hydrogen substituent at position 5 on the xylose ring has a dramatic effect on the intra- and inter-chain hydrogen bonding interactions. Energy calculations suggest that the d-xylose ring exists in the common 4C1 chair conformation, which indicates that in both the 1-4 and 1-3 glycosidic linked xylans the bonding is diequatorial (1e-4e). The intra-chain hydrogen bond O(2′)H?O(6) reinforces the O(3)?O(5′) hydrogen bond to support a twofold extended, helical structure, like a ribbon, and the O(6)H group hydrogen bonds to O(3) atoms in adjacent chains to form sheets [19]. The concept that the glycosidic linkage geometry maintains the basic conformation is evident from these studies. However, three-dimensional structure data on xylan, in aqueous environment, would be extremely important in understanding the xylan and xylanase interactions.

4Xylanase production

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Scope of the present review
  5. 3Structure of xylan
  6. 4Xylanase production
  7. 5Regulation of xylanase synthesis
  8. 6Biochemical properties
  9. 7Xylanases of extremophilic origin
  10. 8Cloning and expression of xylanase gene(s)
  11. 9Protein engineering
  12. 10Site-directed mutagenesis (SDM)
  13. 11Mechanism of action of the xylanases
  14. 12Domain organization of xylanases
  15. 13Molecular evolution
  16. 14Biotechnological potentials of xylan and xylanases
  17. 15Future prospects
  18. Acknowledgements
  19. References

The basic factors for efficient production of xylanolytic enzymes are the choice of an appropriate inducing substrate and an optimum medium composition. The importance of cellulase-free xylanase systems in the paper and pulp industry has initiated research into the correlation between the production of xylanases and cellulases by microorganisms. Filamentous fungi are particularly interesting producers of xylanases since they excrete the enzymes into the medium and their enzyme levels are much higher than those of yeast and bacteria. However, fungal xylanases are generally associated with cellulases [20]. Selective production of xylanase is possible in the case of Trichoderma and Aspergillus species using only xylan as the carbon source. On cellulose these strains produce both cellulase and xylanase which may be due to traces of hemicellulose present in the cellulosic substrates [6]. The mechanisms that govern the formation of extracellular enzymes with reference to 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 not contaminated by cellulose under a lower nitrogen/carbon ratio in the medium may be one of the strategies for producing xylanolytic systems free of cellulases [21]. However, cellulosic substrates were also found to be essential in the medium for maximum xylanase production by Clostridium stercorarium[22], Thermomonospora curvata[23] and Neurospora crassa[24]. Cheaper hemicellulosic substrates like corn cob, wheat bran, rice bran, rice straw, corn stalk and bagasse have also been found to be most suitable for the production of xylanases in the case of certain microorganisms such as Aspergillus awamori, Penicillium purpurogenum[25] and alkaliphilic thermophilic Bacillus sp. NCIM 59 [26]. The xylanase activity is found to be higher in fungi than in bacteria. Among fungi, the maximum activity reported is 3350 IU ml−1 in Trichoderma reesei[27]. The maximum xylanase activity in solid-state fermentation has been obtained from the fungus Schizophyllum commune (22 700 IU g−1) [28]. Trichoderma hamatum is reported to produce 7000 IU g−1 dry wt using wheat straw as substrate [29]. The production of cellulase-free xylanase has been reported in a few Bacillus sp. [26] and fungi [30,31]. Archana and Satyanarayana have reported xylanase production by the bacterial strain, Bacillus licheniformis A99, in solid state fermentation (SSF) [32]. SSF systems resemble the natural habitats of microbes and, therefore, may prove efficient in producing certain enzymes and metabolites. Although a plethora of xylanase producing strains have been described, their use for commercial production at present is restricted mainly to Trichoderma sp. and Aspergillus sp. [25]. However, the future scenario may be different since several promising strains that produce xylanases in higher yields, with increased stability at extreme conditions of pH and temperature, have been recently identified.

Actinomycetes and bacteria exhibit near-neutral pH optima for growth and enzyme production [33], in contrast to the generally acidic pH requirements of fungi. However, certain alkaliphilic Bacilli are known which have pH optima for growth and enzyme production at 9–10 [34]. In Streptomyces flavogriseus[35] and in the Bacillus sp. NCIM 59 [36] simultaneous production of glucose (xylose) isomerase, in addition to cellulases and xylanases, has been described.

4.1Factors affecting xylanase yield

The yield of xylanases in a fermentation process is governed by a few key factors in addition to the standard parameters. When xylanase fermentation is carried out on complex heterogeneous substrates, various factors have a combined effect on the level of xylanase expression. They include substrate accessibility, rate and amount of release of the xylooligosaccharides and their chemical nature and quantity of xylose released – which acts as the carbon source and as an inhibitor of xylanase synthesis in most of the cases. Generally, the slow release of the inducer molecules and the possibility of the culture filtrate converting the inducer to its non-metabolizable derivative are believed to boost the level of xylanase activity.

The xylanases bind tightly to the substrate. A part of the enzyme produced during the fermentation is often lost and discarded, as bound enzyme, along with the insoluble substrate. The metabolic enzymes of the xylanase producer such as proteases [37] and transglycosidases also affect the actual yield of the enzyme [38]. These enzymes are optimally expressed at the end of the exponential phase; and the harvesting time of the xylanases must be correlated to the production of these enzymes on the medium under consideration. Other bioprocess parameters that can affect the activity and productivity of xylanase attained in a fermentation process include the pH, temperature and agitation.

5Regulation of xylanase synthesis

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Scope of the present review
  5. 3Structure of xylan
  6. 4Xylanase production
  7. 5Regulation of xylanase synthesis
  8. 6Biochemical properties
  9. 7Xylanases of extremophilic origin
  10. 8Cloning and expression of xylanase gene(s)
  11. 9Protein engineering
  12. 10Site-directed mutagenesis (SDM)
  13. 11Mechanism of action of the xylanases
  14. 12Domain organization of xylanases
  15. 13Molecular evolution
  16. 14Biotechnological potentials of xylan and xylanases
  17. 15Future prospects
  18. Acknowledgements
  19. References

Despite the increase in knowledge of microbial xylanolytic systems in the past few years further studies on induction and secretion of xylanases are necessary to develop efficient xylanase producers for possible commercial applications. Xylanase production by various bacteria and fungi has been shown to be inducible. But rare examples of constitutive xylanase expression have also been reported [39]. In general the xylanase induction is a complex phenomenon and the level of response to an individual inducer varies with the organisms. An inducer producing maximum xylanase activity in one species may be the inhibitor of activity in an other species [40]. The substrate derivatives and the enzymatic end products may often play a key positive role in the induction of xylanases; they can also act as the end-product inhibitors, possibly at much higher concentrations.


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 by the action of small amounts of constitutively produced enzyme. Cellulose has also been shown to act as an inducer of the xylanase in a few cases, but its not clear whether the inducing effect lies with cellulose or the contaminating xylan fraction. In Streptomyces sp. xylanase activity appears to increase with the crystallinity of the cellulosic substrate [41]. Sugarcane bagasse is found to be the best inducer of xylanase and β-xylosidase in Cellulomonas flavigena[42]. A synergistic effect on the synthesis of both the enzymes was observed when cellulose and hemicellulose were used together as the carbon source.

The xylanase from Aspergillus sp. (2MI), induced by pine xylan, exhibited higher stability than that induced by other xylans [43]. In the presence of xylose higher enzyme yields were obtained from Bacillus pumilus[44], Streptomyces lividans 66 [45] and Aureobasidium pullulans[46]. However, in Cryptococcus albidus[8] xylose repressed xylanase production while in thermophilic actinomycetes [47] it had no influence on the regulation of xylanase expression. Xylobiose was found to be a specific inducer of xylanase in T. reesei[48] and Aspergillus terreus[38]. Xylanolytic systems of yeasts and fungi were also induced by positional isomers of xylobiose. In Cryptococcus albidus the positional isomers of xylobiose behave differentially from xylobiose [49] in that the response of the cells to them is slower, however, the enzyme yields are higher in the presence of isomeric xylobioses indicating that they are not direct inducers. d-Xylan fragments, as well as methyl-β-d-xylopyranoside, induce xylanase in Streptomyces sp. [50,51] and in the yeasts of the genera Cryptococcus and Trichosporon[52,53]. In Trichosporon cutaneum[54] and Trichoderma lignonum[55] thioxylobiose was found to induce xylanase activity. It is also reported that not all β-xylopyranosides serve as inducers of xylanase since their induction capacity is also dependent on the structure of aglycon.

The role of transglycosylating enzymes in the synthesis of positional isomers of these heterodisaccharides is indicated in Aspergillus terreus[38] and T. reesei[48]. The low molecular mass substances that have been identified as xylanase inducers need transferase enzymes for their translocation into the cytoplasm. Hence the level of inducers and/or the required enzymes in the culture filtrate also affect the xylanase synthesis. The possible factors affecting xylanase induction have been diagrammatically represented (Fig. 2), largely based on the figure by Thomson [56].


Figure 2. Regulation of xylanase biosynthesis (hypothetical model). Constitutive xylanases degrade xylan to xylooligosaccharides and xylobiose, which are taken up by the cell and induce other xylanase genes. The inducible xylanases degrade xylan further to xylooligosaccharides and xylobiose. The β-xylosidases, which may be produced constitutively and/or inducibly, convert xylobiose to xylose and may subsequently transglycosylate it to XylB1-2Xyl and GlcB1-2Xyl. These compounds are taken up by the cell and act as additional inducers of genes encoding xylanolytic enzymes. (Based on [56].)

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5.2Regulation at the molecular level

Xylanase biosynthesis and the phenomenon of enzyme induction at the molecular level are comparatively less investigated. This may be due to the lack of cell-free systems which are necessary for the evaluation of inducers under experimental conditions which prevent their transport into the cells [6]. A separate regulation for the formation of xylanase and cellulase has been reported in a few microorganisms [38,57]. In spite of this evidence, an additional linkage governing the synthesis of the two enzyme systems seems to be present, as evident from the pleiotropy of mutations, causing the elimination of both cellulase and xylanase genes [58].

Esteben et al. [59] have reported that xylanase was undetected in glucose grown cultures of Bacillus circulans WL-12 but xylose, mannose and cellobiose supported xylanase production. The xylanase and xylosidase in Butyrivibrio fibrisolvens GS 113 are shown to be under coordinate control, induced by xylan and repressed by glucose [60]. An analysis of DNA fragments containing β-xylanase genes from B. pumilus[61,62] indicated that the xylanase and xylosidase genes are closely associated and are linked in a 14.4-kb DNA fragment. However, they did not appear to be controlled by the same operon.

5.2.1Catabolite repression

Catabolite repression by glucose is a common phenomenon observed in xylanase biosynthesis. Relatively few reports on the relation of cAMP to xylanase induction are available. In the case of yeast C. albidus, when xylan or methyl-β-xylopyranoside were used as inducers, cAMP caused a twofold increase in xylanase production [63]. However, cAMP had no effect on the repression caused by d-xylose. It has been suggested that a 15-bp nucleotide sequence, upstream from the β-xylanase gene, may be a part of the cyclic AMP regulatory sequence. In Aspergillus tubigensis, a 158-bp region, 5′ upstream of the xylanase gene, was shown to be involved in xylan-specific induction [64]. The authors have also reported that catabolite repression of xylanase gene appeared to be controlled at two levels, directly by repression of gene transcription and indirectly by repression of transcriptional activator. The same pattern of regulation was observed in A. niger and A. nidulans. Mach et al. [65] have studied the carbon catabolite repression of xylanase gene expression and have analyzed the molecular basis for the absence of xylanase I formation by the filamentous fungus T. reesei. They have concluded on the basis of Northern blot analysis that the repression of basal xyn 1 transcription was mediated by the carbon catabolite repressor protein Cre 1. Cre 1 in vivo binds to two of four consensus sites (5′-SYG-GRG-3′) in the xyn 1 promoter, which occurred in the form of an inverted repeat.

Deletion and functional analysis of the xylanase genes from A. niger and A. tubigensis[66] led to the discovery of a triplicated sequence that appears to control the enzyme induction. Gat et al. [67] have demonstrated that, in the case of B. stearothermophilus T-6, induction of xylanase synthesis by xylose was controlled at the transcriptional level, indicating the presence of a repressor protein mediating the regulation. Enzyme synthesis was also found to be repressed when easily metabolizable carbon sources were present in the growth medium, suggesting that the synthesis of the enzyme is controlled by transition state regulators and catabolite repression. The sequence analysis of T-6 xylanase has revealed the presence of an AT-rich region preceding the promoter that has been identified as the binding site for a protein that regulates induction [68]. It has been suggested that these regions interact or facilitate interaction with regulators. A putative catabolite repression sequence was found 230 bp inside the xylanase structural gene. The catabolite repression consensus sequence was TGT/AAANC|GNTNA/TCA, where the underlined letters represent the most critical bases, N is any base, and the vertical line denotes an axis of symmetry. In B. subtilis, this consensus sequence was found inside the structural genes in several operons, including the xyl operon [69].

The biosynthesis of xylanase occurs several hours after the depletion of the inducer added to the medium, in contrast to the synthesis of β-xyloside permease and β-xylosidase which have very short induction periods. This is in agreement with the suggestion that mRNAs of secreted proteins do not undergo a rapid turnover as that of intracellular proteins [70]. Characterization of mRNA transcripts of xylanases and studies on in vivo genetics will considerably help a better understanding of the regulation of xylanase biosynthesis.

6Biochemical properties

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Scope of the present review
  5. 3Structure of xylan
  6. 4Xylanase production
  7. 5Regulation of xylanase synthesis
  8. 6Biochemical properties
  9. 7Xylanases of extremophilic origin
  10. 8Cloning and expression of xylanase gene(s)
  11. 9Protein engineering
  12. 10Site-directed mutagenesis (SDM)
  13. 11Mechanism of action of the xylanases
  14. 12Domain organization of xylanases
  15. 13Molecular evolution
  16. 14Biotechnological potentials of xylan and xylanases
  17. 15Future prospects
  18. Acknowledgements
  19. References

The available information about the properties of xylanases stems mostly from studies on bacterial and fungal enzymes. Microbial xylanases are single subunit proteins with molecular masses in the range of 8–145 kDa [10].

The optimum temperature for endoxylanase from bacterial and fungal sources varies between 40 and 60°C. Fungal xylanases are generally less thermostable than bacterial xylanases. Fungi which are mesophilic in origin but produce thermostable xylanases include Ceratocystis paradoxa, the xylanase of which is stable at 80°C for l h [71]. A low temperature active enzyme having both carboxymethyl cellulase and xylanase activities from Acremoniun alcalophilum JCM 7366 has been reported recently. The xylanase and cellulase activities at 0°C were 25 and 48.8%, respectively, of their activities at the optimum temperature of 40°C [72]. d-Xylanases from different organisms are usually stable over a wide pH range (3–10) and show optimum pH in the range of 4–7. The xylanases from fungi such as Aspergillus kawachii[73] and Penicillium herque[74] exhibit optimum pH towards the acidic side (pH 2–6). The isoelectric points for endoxylanases from various sources ranged from 3 to 10. Generally, bacteria are known to produce two xylanases – high molecular mass acidic xylanase and low molecular mass basic xylanase. However, this type of relationship is not observed in fungi; but low molecular mass basic xylanases are common. The amino acid compositions of xylanases reported from various sources indicate predominantly aspartic acid, glutamic acid, glycine, serine and threonine.

6.1Carbohydrate content

The occurrence of glycosylated enzymes is a common phenomenon among many eukaryotic xylanases [74]. The xylanases from prokaryotic sources, like Clostridium stercorarium[22], Streptomyces sp. [75] and an alkaliphilic, thermophilic Bacillus sp. [26], were found to be glycoproteins. Carbohydrate groups are covalently linked with protein or are present as dissociable complexes with xylanases. Glycosylation has been implicated in the stabilization of glycanases against extreme environments [76]. The recombinant xylanases expressed in Escherichia coli from an alkaliphilic thermophilic Bacillus sp. [77] showed lower stability at higher temperature and reduced ability to bind xylan compared to xylanases from the parent strain which is attributed to deglycosylation. In the case of xylanases from Talaromyces byssochlamydoides YH-50 the carbohydrate residues are found to be mannose, glucose and fucose [78]. It has been suggested that both differential glycosylation and proteolysis may contribute to the multiplicity of xylanases [7].

6.2Substrate specificity

Knowledge of the mechanism of action of xylan-degrading enzymes has been gained from studies on substrate specificity, the role of side chain substituents on activity, the specificity of bonds cleaved and the end products. The xylanases of fungal origin are well characterized; they are mainly of two types – non-debranching, which do not liberate arabinose, or debranching, which liberate arabinose from the side chain substituents, in addition to cleaving main chain linkages [79]. Many xylanases of fungal origin, such as Neurospora crassa[80] and Aspergillus niger[81], were found to release arabinose from arabinoxylan. An endoxylanase from Streptomyces roseiscleorticus has also been shown to be a debranching type of enzyme. However, xylanases from Trichoderma harzianum[82] and another strain of A. niger were found not to release free arabinose from arabinoxylan. The presence of debranching as well as non-debranching xylanases have been reported from T. koningii and Ceratocystis paradoxa, and various other sources. A novel enzyme has been recently isolated from A. awamori which cleaves arabinose substituents from cereal arabinoxylans, is specific to the substituent on the relevant xylose, and does not cleave the xylan main linkage [83].

It is commonly observed that substituents in the highly branched polysaccharides interfere with xylanase activity. However, enzymes having more affinity for main chain linkages, near branch points, were reported from A. niger, T. viride and other sources [3]. The xylanases also vary in their activity against various cellulosic substrates. A few of them act only on xylan whereas the non-specific xylanases from Myrothecium verrucaria, Penicillium capsulatum and P. funiculosum act against carboxymethyl cellulose and xylan. The relaxed specificity of some xylanases as against the more restricted specificity of others could be due to differences between residues involved in the catalytic groups. Generally, xylanases appear to be specific toward the intersugar linkage [3]. In C. thermocellum, endoglucanase was reported to hydrolyze β-1,3 linkages in barley, β-glucan as well as β-1,4 linkages in other substrates. Transglycosylation by endoglucanases was believed to lead to products with the same linkage as the substrate because they contain stereospecific binding sites on either side of the catalytic site [84]. The xylanase from Cryptococcus albidus was reported to form a 1,3- β-d-linkage due to transglycosylation reactions [85].

6.2.1Subsite mapping

Subsite mapping and determination of end product analysis are significantly useful in understanding the mode of action of xylanases. Based on the kinetic and end product analysis techniques the subsite maps of the endoxylanase from Aspergillus niger have been determined. The enzyme did not show any appreciable hydrolysis of xylobiose and had higher affinity towards xylooligosaccharides substrates with increasing chain length. Since the binding site of the endoxylanase is shown to consist of eight subsites with the catalytic site in the middle, xylobiose did not form a productive enzyme complex. The xylooligosaccharides of larger length showed hydrolysis of substrate with the formation of a productive enzyme complex. The presence of five main subsites and the localization of a catalytic site between the third and fourth subsites has been demonstrated in the case of xylanase from A. niger[86]. Simon et al. [87] have recently shown the occurrence of an additional subsite (subsite F, i.e. the sixth xylose binding site) in the substrate binding cleft of xylanase A from Pseudomonas fluorescens. It is found that the primary role of F subsite of xylanase A is to prevent small oligosaccharides from forming non-productive enzyme-substrate complexes. The endoxylanase from a different strain of A. niger[88] was found to have seven subsites whereas that from Cryptococcus albidus has four with the catalytic group in the middle [89]. Debeire et al. [90] mapped the subsites of a xylanase from Clostridium thermolacticum and also determined the structures of end products by nuclear magnetic resonance spectroscopy and methylation analysis. They concluded that xylanase from Clostridium thermolacticum was composed of five subsites, a–e, binding five xylosyl rings A–E. The catalytic site was located between xylosyl rings B and C and subsites d and e were able to bind substituted xylosyl residues unlike subsites a, b and c. The differences in the subsite numbers of the xylanases may be due to the different mode of action of the individual xylanases and the length of the end products released by them. The heterogeneous nature of xylan may be one of the reasons for the multiplicity of xylanases. The divergent specificities of these enzymes could play a significant role in their synergistic action towards the complex substrate.

7Xylanases of extremophilic origin

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Scope of the present review
  5. 3Structure of xylan
  6. 4Xylanase production
  7. 5Regulation of xylanase synthesis
  8. 6Biochemical properties
  9. 7Xylanases of extremophilic origin
  10. 8Cloning and expression of xylanase gene(s)
  11. 9Protein engineering
  12. 10Site-directed mutagenesis (SDM)
  13. 11Mechanism of action of the xylanases
  14. 12Domain organization of xylanases
  15. 13Molecular evolution
  16. 14Biotechnological potentials of xylan and xylanases
  17. 15Future prospects
  18. Acknowledgements
  19. References

Considerable progress has been made in the isolation of extremophilic microorganisms and their successful cultivation in the laboratory. A plethora of enzymes from diverse sources, and techniques to modify interesting candidates for basic or applied research, have laid the basis for the expansion of biocatalysts in ways not previously envisioned [91]. Commercial applications of the xylanases demand identification of highly stable enzymes active under routine handling conditions. Many advantages such as reduced contamination risk and faster reaction rates have been proposed for the use of thermophiles in biotechnology processes. In general, parameters such as temperature, pH, and chemical and enzymatic stability are important for the industrial applicability of any enzyme. Recently a simple model, based on the thermodynamic and kinetic parameters for inactivation of the enzyme, has been presented for predicting (1) thermostabilities in the presence of substrate, and (2) residual activities of enzymes in harsh processing environments. The authors have proposed that the model could be applied to a large number of industrially relevant enzymes and processing conditions where the reversible denaturation of protein is fast as compared to the rate of the subsequent irreversible inactivation step [92]. One of the ways to identify the industrially suitable xylanase preparations is to look for the enzymes from extremophilic microorganisms. In the words of M.W.W. Adams, University of Georgia, a pioneer in the study of extremophiles, “If you want highly stable enzymes, you must go to the extreme environments. Nature's bounty, it seems, abides in its black smokers, steam vents and salt lakes”[93]. The use of biocatalysts has been constrained due to their labile nature under extreme temperature and pH conditions. The study of extremophiles and their enzymes can extend the present understanding of protein chemistry in addition to expanding the potential applications of biocatalysts.

7.1Xylanases from alkaliphilic microorganisms

Studies of alkaliphiles have led to the discovery of a variety of enzymes which exhibit some unique properties. Biological detergents contain enzymes such as alkaline proteases and/or alkaline cellulases from alkaliphiles. One significant application is the use of the enzyme, cyclodextrin glycosyl transferase (CGT), to catalyze the degradation of starch to cyclodextrins. Commercial production became economical only after the discovery of alkaliphilic Bacillus producing CGT with enhanced pH stability. Alkaline xylanases have gained importance due to their application for the development of eco-friendly technologies used in paper and pulp industries. The enzymes are able to hydrolyze xylan which is soluble in alkaline solutions [94]. The first report on xylanase from alkaliphilic bacteria was published in 1973 by Horikoshi and Atsukawa [95]. The purified enzyme of Bacillus sp. C-59-2 exhibited a broad pH optimum ranging from 6.0 to 8.0. Many of the xylanases produced by alkaliphilic microorganisms such as Bacillus sp. [34] and Aeromonas sp. 212 [96] with optimum growth at pH 10.0 showed remarkable stability at pH 9–10 but were not highly active above pH 8.0. The enzymes from Bacillus sp. TAR-1 [97], C-125 [98] and alkaliphilic Bacillus sp. (NCL-86-6-10) [99] were optimally active at pH 9–10. Recently an alkali-tolerant xylanase from Aspergillus fischeri[100] was reported to exhibit remarkable (pH 9.0) stability. The xylanase from Cephalosporium was the only one reported from an alkaliphilic fungus having activity at broad pH range of 6.5–9.0 [101]. Xylanases from four alkaliphilic, thermophilic Bacillus strains (W1 JCM 2888), W2 JCM 2889, W3 and W4 with optimum pH of 6.0–7.0 and temperature of 65–70°C) are reported [102]. An alkaliphilic thermophilic Bacillus sp. (NCIM 59) producing two types of cellulase-free xylanases at pH 10 and 50°C has been documented [26].

7.2Thermophilic xylanases

The xylanases from thermophilic bacteria such as Thermonospora fusca[47], thermophilic Bacillus sp. [103] and Bacillus stearothermophilus[104] show an optimum temperature in the range of 65–80°C. The thermostable xylanase produced by a thermotolerant Aspergillus strain at 37°C showed maximum activity at 80°C [105]. Xylanase A from the thermophilic anaerobe Clostridium stercorarium has a temperature optimum of 70°C and a half-life of 90 min at 80°C, whereas the xylanase from Thermotoga sp. has been shown to have a temperature optimum of 105°C at pH 5.5 with a half-life of 90 min at 95°C [106]. Xylanase from Dictyoglomus sp.exhibited a half-life of 80 min at 90°C [107]. Among the thermophilic fungi, xylanase from Thermoascus aurantiacus has been reported to be stable at 70°C for 24 h with a half-life of 54 min at 80°C [108]. The other thermophilic fungi producing thermostable xylanases include Paecilomyces variota[109] and T. byssochlamydoides[110] which show an optimum temperature of 65–75°C at pH 5–6.5. Recently endoxylanases from the thermophilic actinomycete Microtetraspora flexuosa SIIX were reported to have an optimum temperature of 80°C, at pH 6.0 [111]. Despite the prevalence of xylan-degrading enzymes in actinomycetes, comparatively little information is available about xylanases from other groups of thermophilic actinomycetes such as Microbispora, Saccharomonospora, etc. Table 1 describes a few of the extremophilic organisms producing xylanase.

Table 1.  Xylanases from extremophilic organisms
 SourceXylanaseOptimum conditionsReference
   pHTemp. (°C)pHTemp. (°C) 
 Thermophilic bacteria      
1Bacillus acidocaldarius 3.5–4.0654.080[284]
2Bacillus licheniformis A 99 7.0607.050[32]
3Bacillus stearothermophilus T-6T-67.0–7.3609.065[104]
4Bacillus stearothermophilus No. 21A7.0557.060[285]
5Bacillus thermoalkalophilus 90606.0–7.060, 70[286]
6Clostridium acetobutylicum ATCC 824A6.0375.050[287]
  B 375.5–6.060 
7Clostridium stercorarium HX-1D6.0–7.0606.575[288]
8Clostridium stercorarium F-9A6.0–7.0656.575[22]
9Clostridium thermolacticum (TC 21) 6.0–7.0656.580[289]
10Dictyoglomus thermophilum strain B1 7.0687.080[290]
11Microtetraspora flexuosa S II X 6.0809.052[111]
12Thermophilic Bacillus Strain XE 7.0556.075[291]
13Thermophilic Bacillus sp 7.0656.5–7.078[103]
14Thermophilic bacteria ITI 283, ITI 236 7.5658.080[292]
15Thermoanaerobacterium sp. JW/SL-YS485 6.0606,280[293]
16Thermotoga sp. (Fjss3-B.1)A6.8–7.0805.3105–110[106]
17Thermotoga maritima (MSB 8)A7.0806.292[294]
  B 805.4105 
18Thermotoga thermarum16.0–7.0776.080[295]
  2 777.090–100 
19Thermomonospora curvata16.0–7.0557.875[23]
  2  7.275 
  3  6.875 
20Thermomonospora chromogena MT814 8.0505.0–8.075[47]
21Thermomonospora fusca BD 21 8.0506.0–8.065[296]
23Thermomonospora fusca YX 8.0506.0–8.070[239]
24Streptomyces thermoviolaceus OPC-520I7.0507.070[297]
  II 507.060 
 Thermophilic fungi      
1Gloephyllum trabeum  4.080[298]
2Talaromyces byssochlamydoides YH-50 6.2505.070[78]
3Thermoascus aurantiacus 6.0455.075[108]
4Thermomyces lanuginosus DSM5826 6.5506.550[299]
5Talaromyces emersonii CBS 814.7II4.5454.278[300]
 Alkaliphilic bacteria      
1Alkalophilic Bacillus 41M-1 10.3379.050[97]
2Bacillus sp. TAR-1 10.5509.070[97]
3Bacillus sp. C-59-2 8.0375.5–9.060[301]
4Bacillus sp. C-125 10.56.0–10.070[98]
5Bacillus sp. NCIM 59 10.0506.050–60[26]
6Aeromonas sp. 212 10.0377.0–8.050[118]
7Bacillus sp. NG-27 9.0–10.0277.0, 8.470[302]
8Bacillus sp 9.0–10.045–50  [102]
 W1I  6.065 
  II  7.0–9.070 
 W2I  6.065 
  II  7.0–9.570 
 W3   6.065 
 W4   6.0–7.070 
9Bacillus NCL-87-6-10 9.5288.060[99]

As pointed out by Hodgson [112], dramatic changes are predicted in the future scenario of the industrial enzyme business which is mostly based on high technology. Multinational companies like Novo, Genencor and Diversa plan to mobilize biodiversity by establishing a large culture collection of unique microorganisms and to screen for enzyme activities. Diversa (formerly Recombinant Biocatalysis) is specifically harvesting DNA from entire extremophilic communities and is using probes, based on known enzyme sequences to perform biopanning of expression libraries derived from the DNA. That the extremozymes have significant financial impact for the companies that exploit them is evident from the example of Taq polymerase which has sales of $80 million per annum [113]. The unique framework of enzymes isolated from extreme environments will be the ideal choice to engineer novel proteins with the desired functions suitable for a particular application. In spite of having interesting and novel properties, the extremophilic enzymes have not yet been commercialized. Recently, Genencor International (Rochester, NY) introduced cellulase 103, isolated from an alkaline bacterium, for application in the detergent and fabric industries. This is the first large-scale commercial application of an extremo-molecule. With the advent of recombinant DNA technology and protein engineering one can foresee a tremendous future for such enzymes.

8Cloning and expression of xylanase gene(s)

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Scope of the present review
  5. 3Structure of xylan
  6. 4Xylanase production
  7. 5Regulation of xylanase synthesis
  8. 6Biochemical properties
  9. 7Xylanases of extremophilic origin
  10. 8Cloning and expression of xylanase gene(s)
  11. 9Protein engineering
  12. 10Site-directed mutagenesis (SDM)
  13. 11Mechanism of action of the xylanases
  14. 12Domain organization of xylanases
  15. 13Molecular evolution
  16. 14Biotechnological potentials of xylan and xylanases
  17. 15Future prospects
  18. Acknowledgements
  19. References

For the commercial realization and economic viability of xylanase production, it is necessary to identify organisms which can hyperproduce the enzymes. Recombinant DNA techniques offer the means to enhance protein production. Xylanase genes have been cloned from different microbial genera into various suitable hosts. The bacterial genes encoding xylan-degrading enzymes have been found to be adjacent on the genome or in close proximity to other genes encoding cellulase-related functions. In P. fluorescens subsp. cellulosa, xylanase and arabinofuranosidase genes were transcribed in the same direction and were separated by only 148 bp [114], while the second xylanase gene mapped to within 125 bp of the endoglucanase gene. In the case of B. polymyxa, xylanase and endoglucanase genes were separated by 155 bp [115]. Similarly the close linkage between xylanase and β-xylosidase genes has been reported in B. pumilus[61] and C. saccharolyticum[116]. In spite of this frequent clustering of the genes encoding xylan-degrading enzymes, there is no evidence for polycistronic transcription [58]. The nucleotide sequence data of the 4.2-kb chromosomal segment of the genome of Bacillus stearothermophilus 21 indicated that β-xylosidase gene was located upstream of the xylanase gene. Further analysis indicated that the two genes were separated by only 2 bp and were transcribed independently [117].

8.1Heterologous cloning

The xylanase genes have been isolated from different microbial genera and expressed in E. coli. The expression in E. coli is generally found to be lower than the parent organism, and confined to the cytoplasmic or the periplasmic fractions. The absence of post-translational modifications such as glycosylation in E. coli and the intracellular accumulation of the recombinant xylanases have been suggested to be the key reasons for the low levels of activity. Extracellular activity has been reported in recombinant E. coli for the xylanases from alkaliphilic Aeromonas[118], alkaliphilic Bacillus[119], alkaliphilic, thermophilic Bacillus sp. [120] and Cellulomonas sp. [121]. The xylanase gene cloned from Bacillus stearothermophilus T-6 was of particular interest because the enzyme was optimally active at pH 9.0 and 65°C [67]. The sequence data for the gene is also available, which can be used in further studies on site-directed mutagenesis. Cloning and expression of a xylanase gene from the extreme thermophile Dictyglomus thermophilum Rt46B.1 in E. coli has been reported [122]; the recombinant enzyme was found to have an optimum temperature of 85°C. The xylanase cloned from Thermotoga maritima showed an optimum temperature of 90°C at pH 5.5 and was stable up to 100°C [123]. The recombinant xylanase from Thermotoga neapolitana was found to be stable at 90°C for 4 h with a half-life of 2 h at 100°C [124]. The latter had an optimum temperature of 102°C at pH 5.5. Studies on the cloned xylanase from Clostridium thermocellum F1 revealed that it was optimally active at 80°C and was stable up to 70°C at neutral pH and over the pH range of 4–11 at 25°C [125]. Keen et al. [126] have reported the cloning of a xylanase gene from corn strains of Erwinia chrysanthemi. Sequence analysis revealed that the leader peptide of the protein was unusual and long. The protein was found to be distinct from xylanases belonging to glycohydrolase families 10 and 11 and appeared to be intermediate between families 5 and 30. The XYN 2 gene from T. reesei QM6a [127] was expressed in yeast Saccharomyces cerevisiae under the control of alcohol dehydrogenase (ADH 2) and phosphoglycerate kinase (PGK 1) gene promoters and terminators, respectively. The recombinant strains produced 1200 and 160 nkat of xylanase activity per ml, respectively, under the control of ADH 2 and PGK 1 promoters. The recombinant xylanase had an optimum temperature of 60°C at pH 6 and retained more than 90% of its activity after 60 min at 50°C. Recently Graessle et al. [128] reported a system for regulated heterologous gene expression in the filamentous fungus Penicillium chrysogenum. The heterologous expression system was developed by placing the encoding sequences under the control of the repressible acid phosphatase gene promoter of P. chrysogenum. The cloning and expression of xylanases from various organisms is summarized in Table 2.

Table 2.  Cloning and expression of xylanase genes in E. coli
Source organismVectorMW (kDa)Expression of cloned xylanaseReference
  PRU ml−1U mg−1 
Aeromonas sp. 212 (ATCC 31085)pBR 3221451351.63[118]
Bacillus sp. C125pBR 32243430.47[119]
B. circulans NRC 9024/USDA 729pUC 1959590.04[129]
B. polymyxa NRC 2822/NRRL 8505pBR 32251480.037[304]
 pUC 13 22 
B. polymyxa NCIB 8158/ATCC 842pBR 32210.00.1[305]
B. pumilus IPOpBR 3220.002[62]
B.subtilisPAP 115220.5[303]
Bacillus sp. NCIM 59pUC 835352.0 
  15.814.5  [120]
B. ruminicola#23pUC 181.1[138]
Butyrivibrio fibrosolvens#49pUC 194546.60.01[306]
Caldocellum saccharolyticumλ1059 pBR 32242[116]
Cellulomonas sp. NCIM 2353pUC 18450.056[121]
Chainia sp. NCL 82-5-1λgt 10 pUC 860.0030.018[307]
Clostridium stercorarium F9pBR 3228.16[308]
C. acetobutylicum#P262pEcoR 2512828644.0[309]
C. thermocellum NRCCpUC 89041, 391.5[130]
F. succinogenesλgtWESλB0.5728.5[139]
Neocallimastix patriciarumλZAP II9373, 42193.75[135]
Pseudomonas fluorescens subsp. cellulosaλ 47.10.061[131]
Ruminococcus albus#SY3pBR 322560.013[310]
R. flavefaciens#17λEMBL31.6[132]
Thermomonospora fusca YXλgtWESλB0.79[311]
Trichoderma reesei C30pGEM5Z(+)1919[134]
P and R correspond to parent and recombinant protein. MW corresponds to the molecular mass in kDa.

Most of the xylanolytic organisms produce multiple isomeric forms which may result from post-translational processing of a single gene product or, more often, as the products of multiple genes. The heterogeneity of xylanases was attributed to post- translational modifications such as differential glycosylation and/or proteolysis [58]. The two xylanases from Aeromonas sp. appeared to be differentially modified products from the same gene due to their similar hydrolytic, immunological and physicochemical properties [7]. However, the presence of multiple xylanase genes has been demonstrated in the case of Bacillus circulans[129], Caldocellum saccharolyticum[116], Clostridium thermocellum[130], Pseudomonas fluorescens subsp. cellulosa[131] and Ruminococcus flavefaciens 17 [132]. Extensive cloning work has been undertaken on Clostridium sp. to track down the enzymatic specificities of the individual proteins that form the complex cellulosome and xylanosome structures. These studies have already been reviewed extensively [56].

As against the wealth of information available on the xylanase gene cloning of bacterial systems, very few reports are to be found on fungal systems. Only recently some detailed reports on the cloning and expression of xylanases from the fungi Aspergillus kawachii[133], Trichoderma reesei[134], Neocallimastix patriciarum[135] and Orpinomyces PC-2 [136] have appeared. The expression of fungal xylanase in E. coli and the rumen bacterium Butyrivibrio fibrisolvens OB 156 using the putative xylanase promoter from B. fibrisolvens strain 49 has been reported [137].

8.2Overexpression in E. coli

Enhanced xylanase production in E. coli has been achieved in the cases of alkaline Aeromonas sp. [118], Bacteroides ruminicola[138] and Fibrobacter succinogenes 135 [139]. A thermostable xylanase from C. saccharolyticum has also been cloned and overexpressed in E. coli. The enzyme expressed by the recombinant had a thermal half-life of 2–3 min at 80°C [140]. Although this enzyme cannot fit into the ideal criteria for industrial application, the isolated gene itself could prove a starting material for mutagenesis to further improve the properties of the engineered enzyme. Recently Koo et al. [141] have reported overexpression of xylanase from Clostridium thermocellum in E. coli. The recombinant enzyme had an optimum temperature of 60°C at pH 5.4. Overexpression of B. subtilis and B. circulans xylanases in E. coli has also been described. A gene encoding mature B. circulans xylanase has been designed to imitate the frequency of degenerate codons used in E. coli. Synthetic B. circulans gene is then converted to B. subtilis xylanase gene via single codon substitution (Thr147Ser). The plasmids containing both synthetic genes were further modified for direct expression in E. coli. Under the control of the lac promoter, recombinant xylanase has been produced at levels as high as 300 mg l−1 in solution form in cytoplasm. Characterization of the products indicated that the purified recombinant protein was correctly processed and enzymatically active [142]. B. subtilis xylanase gene, fused to the 5′ end of C. fimi cenA, has been overexpressed in E. coli[143]. The fusion protein exhibited strong affinity for both microcrystalline cellulose and regenerated cellulose. High level expression in E. coli has also been obtained with the modified domain II construct of xylanase cDNA from the anaerobic fungus Neocallimastix patriciarum. The modified domain II xylanase produced in E. coli had a specific activity of 1229 U mg−1 protein. The high level expression was largely attributed to the presence of a favorable N-terminal coding sequence [144]. Xue et al. [145] have reported temperature-regulated expression of recombinant xylanase from N. patriciarum in an E. coli strain containing natural lacI gene under the control of the tac promoter. The specific activites of recombinant xylanase and cellulase were 4.5 times higher at 42°C than those obtained at 23°C. The temperature-modulated Ptac system has also been used for the overproduction of xylanase in 10-l fermentation studies using a fed-batch process. Recently Lapidot et al. [146] have reported overexpression and single-step purification of a thermostable xylanase from B. stearothermophilus T-6. The xylanase gene was cloned into T-7 polymerase expression vectors. The enzyme was found to constitute over 70% of the cell protein and was efficiently purified from the host proteins by a single heating step. Over 2 g soluble and active enzyme per liter culture was achieved. Xylanase A from the extremely thermophilic eubacterial strain Rt8B.4 was overexpressed in E. coli. The xylanase activity from domain 2 was associated with a 36-kDa protein, which was stable at 70°C for at least 12 h at pH 7.0 [147].

8.3Expression in plant system

In a recent report Helbers et al. [148] have shown the high level expression of the thermostable xylanase from Clostridium thermocellum (xylanase Z-truncated protein) into transgenic tobacco plants. The xylanase gene was introduced into the tobacco plant by an integration system with Agrobacterium tumefaciens. The protein molecules were correctly targeted into the intracellular space with the help of the signal peptide from the proteinase II. Although the active enzyme was synthesized inside the tobacco cells, it did not harm the host cells due to either the high temperature optimum of the enzyme or the relative scarcity of the substrate molecules in the plant cell wall. The response of the transgenic plant to pathogenic stimuli has been shown to be mediated through the xylanase and other related enzymes secreted by the pathogen and hence the transgenic tobacco plants producing high levels of the xylanase from C. thermocellum assume a special significance. Xylanase B (Xyn B) from C. stercorarium is also expressed in tobacco suspension cells (Nicotiana tabacum L. cv BY2 cell) under the control of the CaMV 35S promoter and noparin synthetase terminator [149]. The plasmid constructed using pUC 118 was introduced into BY2 protoplasts by electroporation and the transformed cells were incubated in the medium containing kanamycin, in agarose bead-type culture. After a 4–6-week cultivation, the calli depicting clear halos on the xylan containing agar plates were selected. The amount of expressed Xyn B protein was also around 4–5% of total proteins in the soluble extracts of tobacco suspension cells. These studies have shown that the bacterial xylanases were stably expressed in the tobacco plant, as high as around 4% of total proteins, without any inhibitory effect on plant growth. In addition, the thermostable enzymes are easily isolated from other tobacco proteins by heating, which has opened an avenue of creating the bioreactors for hydrolytic enzymes.

8.4Homologous cloning

Several heterologous proteins cannot be efficiently expressed in E. coli due to any one of several reasons, such as the relatively abundant occurrence of rare codons in the cloned gene, the need for specific post-synthetic modification(s), structural complexity of the protein, toxicity of the coded protein to the host cells, and susceptibility of the foreign protein to proteases coded by E. coli. The xylanase genes cloned in E. coli may be underexpressed due to poor or complete lack of an induction mechanism. It is well known that higher expression levels are obtained in the homologous host system. Hence the cloning of xylanase genes in the homologous host systems is important, although studies on the homologous expression of cloned xylanase genes are scarce. The level of expression of the xylanases from Bacillus sp. is relatively higher in a homologous host system than that in E. coli. In the case of B. pumilus IPO, the extracellular secretion of the xylanase was achieved using the homologous host B. subtilis[150]. The homologous expression of the xylanase gene from an alkaliphilic, thermophilic Bacillus sp. has been demonstrated in a xyn- mutant B. subtilis A8 [151], and B. subtilis MI 111 [152]. The low level of expression observed has been attributed by the authors to the weak recognition of the signals from the host strain which is an unusual extremophilic species. Enhanced production of the xylanases has also been reported by chromosomal gene integration in an alkaliphilic, thermophilic Bacillus sp. [153]. This is of significance because the use of recombinants in the large-scale fermentation process has been restricted due to plasmid instability and constant requirement of antibiotic-selective pressure. An Aspergillus nidulans multicopy tranformant for the gene xylanase B encoding minor xylanase has been constructed recently [154]. The transformant was reported to secrete 114 U of xylanase per mg protein.

In the case of Streptomyces, the promoter elements may not be recognized by the E. coli sigma factor, in turn leading to the failure of the gene expression. Hence the construction of a genomic library and screening for the xylanase gene have been carried out from Streptomyces sp. #36a and S. lividans#1326 using a homologous host system. The xylanase gene of S. lividans#1326 was cloned by functional complementation of the xyn mutant of S. lividans using multicopy plasmid PIJ 702; a maximum enzyme production of 380 U ml−1 was reported [155]. Thus the cloning of a xylanase gene into a homologous system not only allowed excellent secretion but also yielded 60 times higher enzyme activity than that of the wild-type. Iwasaki et al. [156] have also reported the molecular cloning of a xylanase gene from another strain of Streptomyces; the recombinant showed maximum activity of 2830 U ml−1 of culture broth. The effects of signal peptide alterations and replacement on export of xylanase have been reported in Streptomyces[157]. The authors have suggested that the changes in the signal peptide affect the level of xylanase production. However, further experiments are necessary that will allow the combination of a suitable signal peptide with the enzyme to achieve the overproduction. The reports on homologous cloning and expression of the xylanases are summarized in Table 3.

Table 3.  Homologous expression of xylanase genes
Parent strainHostVectorXylanase activity (U ml−1)Reference
Bacillus pumilus IPOB. subtilis MI 111PUB1100.61.8[62]
Bacillus sp. (NCIM 59)Bacillus sp. NCIM-59PUC866128[153]
Streptomyces spS. lividans TK21PIJ-70238.92839[156]
Strain #36aS. kasugansis G3PSK21841 
S. lividans 1326S. lividansPIJ-7025.8380[155]
Streptomyces EC3S. lividansPIJ-702600[312]
 S. parvulusPIJ-702 
Streptomyces halstediiS. parvulusPIJ-7024.12[313]
P and R correspond to parent and recombinant protein.

8.5Cloning suitable for biotechnological application

The cloning and expression of xylanases in non-xylanolytic organisms has been largely restricted to microorganisms such as Saccharomyces, Kluyveromyces, Lactobacillus and Bacillus subtilis. These organisms have been well characterized as far as their industrial applicability is concerned. Lactobacillus plantarum can actively produce and secrete heterologous enzymes from a variety of Gram-positive bacteria and is comparable with Bacillus subtilis as a host for recombinant protein production [158]. The xylanase produced by the recombinant L. plantarum was able to release the fermentable carbohydrates from ensiled crops and thereby improve the silage quality. The cloned xylanase gene has also been shown to offer fermentative advantages to both these host organisms in the sense that the organism can directly utilize an additional substrate – hemicellulose, a cheap, renewable, abundant, fermentable, raw resource material. Saccharomyces has been considered a suitable host for the cloning and expression of the genes from eukaryotes [159]. The yeast cells do not produce endotoxins and are considered to be safe in medicine and food products. Also large-scale fermentation and downstream processing are established. Saccharomyces has been used to express the xylanases from Aspergillus and Cryptococcus. Recently cloning and expression of xylanase genes from Meripilus giganteus, Myceloipthora thermophilum and Thielavia terrestris in Aspergillus oryzae have been reported [160–162], illustrating the examples of heterologous cloning of xylanase genes. The application of recombinant xylanase in the food and paper industries has also been described. Walsh and Bergquist have reported the expression of Xyn A from an extremely thermophilic anaerobe Dictyoglomus thermophilum Rt 46 B.1 in the yeast Kluyveromyces lactis[163]. The gene was fused in frame with the secretion signal of the K. lactis killer toxin in episomal expression vectors. Xyn A was secreted predominantly as an unglycosylated protein comprising of 90% of the total extracellular proteins. Also xylanase from thermophilic bacterium Caldicellulosiruptor saccharolyticus was secreted to a level of 10μg ml−1 in the same yeast. The reports on the expression of the cloned xylanase genes in heterologous host systems with relevant application potential are summarized in Table 4.

Table 4.  Cloning of xylanase genes into hosts suitable for biotechnological application
Parent strainHostVectorReference
Aspergillus kawachiiS. cerevisiaepVT 100[133]
A. pullulansS. cerevisiaepYES 2[314]
Clostridium acetobutylicumL. plantarumpWP 37[158]
C. thermocellumL. plantarumpWP 37[158]
Cryptococcus albidusS. cerevisiaepVT 100[315]
 Pichia stipitispJHS 
C. saccharolyticumS. cerevisiaepFLAGU 2[316]

9Protein engineering

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Scope of the present review
  5. 3Structure of xylan
  6. 4Xylanase production
  7. 5Regulation of xylanase synthesis
  8. 6Biochemical properties
  9. 7Xylanases of extremophilic origin
  10. 8Cloning and expression of xylanase gene(s)
  11. 9Protein engineering
  12. 10Site-directed mutagenesis (SDM)
  13. 11Mechanism of action of the xylanases
  14. 12Domain organization of xylanases
  15. 13Molecular evolution
  16. 14Biotechnological potentials of xylan and xylanases
  17. 15Future prospects
  18. Acknowledgements
  19. References

Biotechnological applications of the xylanases require thermostable enzyme preparation with a wide pH and temperature range. Since the availability of the ideal enzyme preparation is limited, the application of protein engineering studies to xylanases has gained importance. Protein engineering is also one of the principal means of examining the active site of an enzyme to identify the roles of specific residues in catalysis; site- directed mutagenesis provides the technology required to redesign the protein. The identification of active site residues by chemical modification, X-ray crystallographic data and site-directed mutagenesis has provided basic information regarding the structure-function correlation of the xylanases. These studies have formed the basis for the protein engineering of xylanases for specific manipulation of the gene for desired enzymatic properties.

9.1Amino acid modification

Amino acid modification is carried out usually chemically or by site-directed mutagenesis, and the residues essential for substrate binding or catalysis are identified. Chemical modification, using group-specific reagents, may suggest the type of residue involved in catalysis or in substrate binding, however, it does not identify the specific residue involved. Substrates and competitive inhibitors which can bind to the active site frequently protect the enzyme against inactivation. The participation of tryptophan in the active site of xylanases from Chainia[164] and Streptomyces[165] has been reported. The fluorometric analysis of the xylanases from Chainia[166] and alkaliphilic thermophilic Bacillus[167] revealed that the tryptophan microenvironment was electronegative. Chemical modification of xylanases from the fungus Schizophyllum commune[168] and an alkaliphilic thermophilic Bacillus sp. [169] indicated the involvement of carboxyl groups in the catalysis. Evidence for the specific interaction of guanidine hydrochloride with the essential carboxyl group of xylanase from an alkaliphilic, thermophilic Bacillus sp. NCIM 59 has also been presented [170]. The presence of cysteine in the active site of a few bacterial xylanases has been reported [164,165]. However, a possible role for such residues has not so far been addressed. Cysteines may be critical for the proper folding of the enzyme and modification of the residue may result in loss in activity. It is also possible that they may participate in the formation of covalent glycosyl enzyme intermediates.

9.1.1Active site peptide

In the case of xylanases, inhibitors can be used to identify the active site residues. The characterization and sequencing of the cysteine containing active site peptide of the xylanase from Streptomyces T-7 [171] and Chainia[172] have been reported. The peptides showed the presence of a conserved aspartic acid residue consistent with the catalytic regions of other glucanases. The possibility of the cysteine residue directly participating in catalysis by forming a covalent link with the incipient reducing sugar has been postulated in the case of the variant T4 lysozyme.

9.2X-ray crystallography studies

The three-dimensional structures of low molecular mass xylanases (family 11, Mr 20) from B. pumilus[173], B. circulans, T. harzianum[174], thermophillic Bacillus sp. [175] and B. stearothermophilus T-6 [176] have been reported. These studies have helped to determine the overall structure of xylanases, in possible identification of specific residues involved in substrate binding and catalysis.

Crystallization and diffraction analysis of xylanases have been carried out at 1.5–3.0 Å. In B. pumilus IPO [177], the enzyme molecule is of ellipsoidal shape (40×35×35 Å) with a well-defined cleft down one side of the molecule. However, the crystals of two major xylanases from the fungus T. reesei[178] are reported to be monoclinic and those of T. harzianum to be orthorhombic [179]. The crystallography studies of xylanase I from A. niger indicated a characteristic fold which is unique for family 11 xylanases. It consists of a single domain composed predominantly of β-strands. Two β-sheets are twisted around a deep, long cleft which is lined with many aromatic residues and is large enough to accommodate at least four xylose residues. Two conserved glutamate residues Glu79 and Glu170 reach into the cleft from opposite sides [180]. The enzymes of family 11 are single domain proteins composed of three antiparallel β-sheets and one α-helix, with the active site lying between the second and third sheets. However, the three-dimensional structure of xylanase II from T. reesei[181] has revealed that the β-sheet structure is twisted, forming a large cleft on one side of the molecule. In the case of T. harzianum the xylanase contains two extra strands at the beginning of sheets I and II and a few insertions and deletions. The observed crystal structure of xylanase from B. circulans is in close agreement with the NMR-derived secondary structure of the protein [182]. In addition to conserved residues in the active site cleft of xylanases from B. circulans[183] there are a number of other residues conserved on either side of the cleft. The three-dimensional structures of the xylanases from T. harzianum and B. circulans were found to be very similar. Many of the conserved amino acids of xylanases are believed to be structurally important for confirming the correct folding and packing. The putative catalytic residues Glu86 and Glu177 of Xyn II from T. reesei are conserved. Also, a clear cluster of conserved residues consisting of Gln136, Tyr77 and Tyr88 is observed around Glu86. The hydrogen bond between Tyr171 and Tyr77 exists in other xylanases although the tyrosine residue is substituted by histidine in a few cases. However, the residues around Glu177 are much less conserved. In addition to these amino acids, three residues, Pro98, Asn124 and Thr133, are conserved although their role is not clear. It is speculated that they may take part in substrate binding. The flat Ser/Thr face of β-sheet A is also conserved in family 11 whose functional role may, to a certain extent, be similar to that of cellulose binding domains present in many cellulases [181].

The three-dimensional structures of the catalytic domain of a few family 10 enzymes have been solved. These are xylanase/exoglucanse (Cex) from C. fimi, xylanase A from P. fluorescens, xylanase Z from C. thermocellum and xylanase from S. lividans[184]. They all have an eight-fold α/β-barrel structure in which conserved glutamates function as catalytic nucleophile and acid/base catalytic residues. The presence of active site residues, Glu127 on strand 4 and Glu246 on strand 7, have been demonstrated in xylanase A from P. fluorescens subsp. cellulosa. The three bulge-type distortions occurring on β-strands 3, 4, and 7 seem to be functionally significant as they serve to orient important active site residues [185,186]. Xylopentose binds across the carboxy-terminal end of the α/β-barrel in an active site cleft containing the two catalytic glutamates. Recent crystal structure analysis revealed the presence of the Ca binding site (located in loop 7) in xylanase A [184]. This is the only xylanase reported to contain a Ca binding site. The authors have suggested that the occupation of Ca binding loop with its ligand protected the enzyme from thermal inactivation, thermal unfolding and proteolysis. Mutational analysis indicated that Asp256, Asn261 and Asp262 present within the Ca binding domain play a pivotal role in the affinity of Xyl A to the divalent cation. A mutant of XYLA in which the key residues of the calcium binding domain were replaced by alanine exhibited thermal stability similar to that of XYLA complexed with Ca2+ ions; however, the xylanase variant was susceptible to cleavage by chymotrypsin.

10Site-directed mutagenesis (SDM)

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Scope of the present review
  5. 3Structure of xylan
  6. 4Xylanase production
  7. 5Regulation of xylanase synthesis
  8. 6Biochemical properties
  9. 7Xylanases of extremophilic origin
  10. 8Cloning and expression of xylanase gene(s)
  11. 9Protein engineering
  12. 10Site-directed mutagenesis (SDM)
  13. 11Mechanism of action of the xylanases
  14. 12Domain organization of xylanases
  15. 13Molecular evolution
  16. 14Biotechnological potentials of xylan and xylanases
  17. 15Future prospects
  18. Acknowledgements
  19. References

Previously the identification of enzyme active site residues relied on the chemical modification of proteins. The essential reactive groups were identified by selective chemical modification. Rapid developments in molecular biology techniques have made it possible for the individual amino acids to be substituted by site-specific mutagenesis. The knowledge derived from chemical modification and crystallographic studies of the active site facilitates the understanding of the structure-function relationship of the protein. Protein engineering is also applied to alter the substrate specificity, pH optima and to increase the thermal stability of the enzymes.

10.1SDM and structure-function analysis

The highly conserved amino acid residues located at specific positions in xylanases are important in structure-function analysis and hence are targeted for SDM. Based on the sequence similarity of xylanase from Bacillus pumilus with other xylanases of known origin and the knowledge of its three-dimensional structure, the authors have proposed the residues Glu93 and Glu182 to be the most suitable candidates for the essential catalytic activity of xylanase [187]. Mutation of these residues resulted in a decrease in specific activity, suggesting that they are essential catalytical residues. No change in protein conformation was observed, as confirmed by circular dichroism (CD) spectra of the mutant enzyme. The conserved glutamate residues (Glu87 and Glu184) have also been shown to be present in the active site of xylanase A from S. commune[188]. Recently mutagenesis and analysis of 3D structure of xylanase A from S. lividans revealed Asn127 to be the important residue in maintaining the ionization states of two catalytic residues and in the stabilization of catalytic intermediates [189]. The three-dimensional structures of two bacterial xylanases from B. pumilus and B. circulans and a fungal xylanase from T. harzianum have also revealed the presence of two completely conserved glutamic acid residues corresponding to Glu87 and Glu184 of S. commune xylanase A. The mutational and crystallographic analysis of the active site residues of the B. circulans xylanase indicate the presence of six tyrosine and three tryptophan residues [183]. However, in the case of the B. circulans xylanase, a Tyr residue rather than Trp may play a significant role in substrate binding, as demonstrated by kinetic analysis of mutant enzymes. Tyrosine residues have also been implicated in the catalytic mechanism for E. coliβ-galactosidase [190]. The authors have demonstrated that two glutamic acid residues (Glu78 and Glu172) are involved in catalysis in the case of the xylanase from B. circulans. Arg112 has been implicated to play a significant role in catalysis, and Tyr69 and Tyr80 in substrate binding. Recently, NMR studies revealed His149 to be an important residue in establishing the conformation of the xylanase. His, Ser and Tyr residues are known to be completely conserved in all family 11 xylanases. Mutagenesis of H149 to Phe and Gln did not alter the active site, but the stability of the folded protein was found to decrease in irreversible thermal denaturation studies [191].

Moreau et al. [192] have identified two acidic residues, Glu128 and Glu236, as essential residues in the active site of xylanase A belonging to family 10 from Streptomyces lividans. The carboxyl group of Asp124 also contributed to the catalytic mechanism. The mutant enzymes obtained by SDM were 10–50% more active than the wild-type xylanase A. Conversely, none of the aspartic acid residues of xylanases of family 11 were completely conserved; it appears that they are not essential for activity [188]. Roberge et al. [193] showed that two histidine residues (H81 and H207) out of three were present in the active site of xylanase A from S. lividans and were found to be completely conserved in family 10 glycanases. The structural analysis revealed that they were part of an important hydrogen bond network in the vicinity of two catalytic residues (E128 and E236). SDM studies showed that they were also essential for protein stability and were probably involved in the folding of native and denatured states of protein. The mutational analysis of xylanase J from alkaliphilic Bacillus sp. strain 41M-1 indicated that Glu93, Glu183, Trp18, Trp86, Tyr84 and Tyr95 play an important role in the catalytic activity [194].

10.2SDM for altered desirable properties

The potential of hemicellulases in the biobleaching of kraft pulp has been recognized in the last decade. However, the commercial application of this technology requires highly active and thermostable enzymes. Site-directed mutagenesis of the cloned gene offers interesting research opportunities to change the properties of a protein suitable for its application. In the case of xylanase from S. lividans 1326 [195] the thermostability is increased by replacing Arg156 by glutamic acid. The modified enzyme has shown a temperature optimum 5°C higher than that of the wild-type. The half-life of Arg156Glu was 6 min longer than that of the wild-type enzyme, suggesting that although the engineered xylanase had a higher optimal temperature, the stability was not significantly affected. Previous reports suggested that the same substitution occurs naturally in the xylanases produced by Bacillus sp. C-125 and C. saccharolyticum[196]. Both xylanases have an optimum temperature of 70°C, which is the same for the modified enzyme (Arg156Glu). The studies on cellulases from T. reesei TD β-6 and Thielavia terrestris NRRL 8126 [197] have revealed that if two favorable mutations are combined, such as Arg156Glu and Asn173Asp, the resulting enzyme is twice as stable as the wild-type, with a half-life of 220 min, at 60°C. The introduction of disulfide crosslinks into proteins, to protect them from unfolding, requires the creation of cysteine residues that form disulfide bonds spontaneously in solution and do not obstruct functional domains. In B. circulans mutant xylanase proteins [198], disulfide bridges conferred thermoprotection as observed by 15°C increase in thermostability.

Site-directed mutagenesis of a xylanase gene from C. saccharolyticum has yielded six mutant xylanases with altered temperature stability and temperature optimum [196], whereas no change was observed in the pH optimum. The stabilization of xylanases by random mutagenesis of the cloned gene fragment from B. pumilus IPO has been described [199]. Four mutants, each showing a single amino acid substitution, have been selected on the basis of activity at 60°C. Based on computer graphic simulation it was confirmed that these substitutions do not change the wild-type conformation. Kinetic analysis has revealed that the mutants are stabilized by a decrease in activation entropy, except for Asn104 which was stabilized by an increase in activation enthalpy.

Even though protein engineering is the most powerful tool to redesign the protein, the reports on xylanases to date suggest that it has not been fully exploited to improve the properties of xylanases. The future scope includes alteration of the residues to improve the pH stability and shift in pH optimum of the enzyme for its commercial application at alkaline pH. SDM has also been applied to alter the substrate specificity of proteases [200] and glucose-xylose isomerases [201]. Designing a xylanase with multiple substrate specificities will also lead to efficient utilization of hemicellulosic substrates.

11Mechanism of action of the xylanases

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Scope of the present review
  5. 3Structure of xylan
  6. 4Xylanase production
  7. 5Regulation of xylanase synthesis
  8. 6Biochemical properties
  9. 7Xylanases of extremophilic origin
  10. 8Cloning and expression of xylanase gene(s)
  11. 9Protein engineering
  12. 10Site-directed mutagenesis (SDM)
  13. 11Mechanism of action of the xylanases
  14. 12Domain organization of xylanases
  15. 13Molecular evolution
  16. 14Biotechnological potentials of xylan and xylanases
  17. 15Future prospects
  18. Acknowledgements
  19. References

It has frequently been suggested that the catalytic mechanism of glycosidases resembles that of lysozyme. The hydrolysis reaction catalyzed by xylanases as well as cellulases proceeds through an acid-base mechanism involving two residues. The first residue acts as a general catalyst and protonates the oxygen of the osidic bond. The second residue acts as a nucleophile which, in the case of retaining enzymes, interacts with the oxocarbonium intermediate or promotes the formation of an OH ion from a water molecule, as observed for inverting enzymes. Reaction with retention of configuration involves a two-step mechanism in which proton transfer occurs to and from an oxygen atom in an equatorial position at the anomeric center [202] (Fig. 3). This reaction mechanism is similar to that of lysozyme [203]. Reactions leading to inversion of configuration proceed through a single substitution, as observed in the case of β-amylase [204]. It is, therefore, likely that a single amino acid residue (acid/base catalyst) is responsible in both proton transfer steps. Xylanases mainly exhibit a double displacement mechanism involving a glycosyl enzyme intermediate which is formed and hydrolyzed via oxocarbonium ion like transition state.


Figure 3. Reaction mechanism of the xylanases. (1) Single displacement reaction. Involvement of a general acid (Glu), a general base (Asp or Glu) and attack by a nucleophilic water molecule is shown. (2) Double displacement reaction (a) involving stabilization of an oxocarbonium ion by electrostatic interaction with the carboxylate of an Asp (or Glu) at the active site or (b) involving formation of a covalent intermediate by nucleophilic attack of the Asp (or Glu) on C-1 of the incipient sugar. (Based on [80].)

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11.1Nucleophile and acid-base catalyst

Wakarchuk et al. [183] have proposed Glu78 and Glu172 to be the nucleophile and acid-base catalyst, respectively, based on crystallographic studies of the enzyme-substrate complex of xylanase from B. circulans. The critical distance between two catalytic carboxylic acids (approx. 5.5 Å in B. circulans xylanase) is less in retaining enzymes as compared to that in inverting glycosidases (approx. 10–11Å). It was also observed that the precise placement of the acid/base catalyst is not critical, since both shortening and lengthening this carboxyl side chain resulted in approximately the same modest decrease in kcat/Km values [205]. This is in sharp contrast to the positional requirements of the catalytic nucleophile Glu78. Thus, as expected, the positional requirements for proton transfer are less demanding than those for carbon-oxygen bond formation. The important property of glutamic acid residue, to undergo conformational change on a rise of the pH (e.g. at pH 6.5) for xylanase II from T. reesei, could be useful in catalysis since it will initiate the reaction and consequently may assist a water molecule to attack the substrate [181]. Further studies suggested that the primary function of the distal sugar moiety in the disaccharide substrate is to increase the rate of formation of the glycosyl enzyme intermediate through improved acid catalysis and greater nucleophilic preassociation, without affecting its rate of decomposition [84]. Hence, the residues acting as nucleophiles have been identified on the basis of structural analysis and mutagenesis data. Tull et al. [206] have mapped the nucleophile (Glu274) of exoglucanase/xylanase Cex from C. fimi and Glu127 has been confirmed as the acid-base catalyst based on kinetic studies [207].

Thus, from various reports it appears that xylanases belonging to family 11 operate via a double displacement mechanism in which the anomeric configuration is retained. However, endoxylanase from A. niger[84] was found to be an inverting enzyme following the single displacement reaction mechanism (Fig. 3). Extensive NMR assignments of xylanase from B. circulans revealed the presence of potential amide-aromatic hydrogen bond between HN of Ile118 and the indole ring of Trp71. This interaction, which is conserved in all low molecular mass xylanases of known structure, may play an important role in establishing the active site conformation of these enzymes [182]. The X-ray crystallography data on xylanases from P. fluorescens[184] and kinetic studies on exoglucanase/xylanase from C. fimi[208] belonging to family 10 have also shown that the substrates are hydrolyzed with net retention of anomeric configuration.

12Domain organization of xylanases

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Scope of the present review
  5. 3Structure of xylan
  6. 4Xylanase production
  7. 5Regulation of xylanase synthesis
  8. 6Biochemical properties
  9. 7Xylanases of extremophilic origin
  10. 8Cloning and expression of xylanase gene(s)
  11. 9Protein engineering
  12. 10Site-directed mutagenesis (SDM)
  13. 11Mechanism of action of the xylanases
  14. 12Domain organization of xylanases
  15. 13Molecular evolution
  16. 14Biotechnological potentials of xylan and xylanases
  17. 15Future prospects
  18. Acknowledgements
  19. References

The understanding of the basic structure and its correlation with function has become a key step in studies dealing with the molecular aspects of the enzyme. However, detailed structural data available on the xylanases is limited. A considerable sequence variability exists among the hemicellulases making the detection of homologies difficult. Hence various means of theoretical analysis are used to complement the data. Of these, hydrophobic cluster analysis and thermostability analysis are of particular importance from the point of view of basic and industrial applications.

12.1Hydrophobic cluster analysis

Hydrophobic cluster analysis (HCA) is a sensitive method for the comparison of amino acid sequences to derive structural, functional and evolutionary information. HCA detects homologies between similar three-dimensional structures of proteins having low sequence identity. These homologies imply a structural as well as functional correlation, and hence are indispensable tools in classifying the related enzymes into families. HCA also provides information on conserved amino acids which are likely to be involved in catalysis.

On the basis of amino acid sequence homology and hydrophobic cluster analysis catalytic domains of cellulases and xylanases were first classified into six families (A–F) [209]. In a subsequent study, the families were upgraded to 11 [210] and in a third update to 45, including 482 glycosyl hydrolase amino acid sequences [211]. Currently Henrissat and Bairoch have classified all the available sequences of glycosyl hydrolases into 58 families [212]. Based on HCA the xylanases are subdivided into two families, F and G, which are also shared by other glycanases such as endoglucanase, exoglucanase, and cellobiohydrolase [211]. The families F and G, which are analogous to glycohydrolase families 10 and 11, comprise high and low molecular mass xylanases, respectively. Generally no significant homology was found between the xylanases from the two families, including the region around the catalytic residues, and they have an altogether different pattern of protein folding.

12.2Domain structure

At the molecular level the xylanase protein comprises functional or non-functional domains and linker regions. The functional domains are further dissected into the catalytic and the substrate binding domains. Classically, the domains are clusters of amino acids that represent structural homology motifs which can be well distinguished from each other on the basis of the structural and spatial identity. The domains are linked together by sequences, highly enriched in hydroxy amino acids, which are often extensively O-glycosylated. It is suggested that the serine-rich linker sequences in xylanases and cellulases may play a role analogous to that of introns (non-coding sequences) in eukaryotes. On the basis of structural similarities, it is also believed that the cellulases and the xylanases have evolved by domain shuffling, with subsequent modification of the domains [210]. Hence, it was assumed that the catalytic domains and the substrate binding domains of the xylanases, too, should be spatially separable or distinguishable. However, an analysis of the a family 10 xylanase from P. fluorescens subsp. cellulosa (Xyl A) has revealed that the spatial separation of protein domains is not necessary for substrate binding or catalytic activity [213]. The substrate binding domain was found to be a cellulose binding domain with no affinity towards xylan and therefore did not contribute to the catalytic activity. Truncated derivatives of xylanase A, lacking either the serine-rich linker sequence or cellulose binding domain, were found to be less active against xylan contained in a cellulose-hemicellulose complex as compared with the full length xylanase [214]. A 374-residue linker sequence rich in asparagine and glutamine has been reported in the xylanase from R. flavefaciens[215]. Xylanase D from C. fimi, in addition, contains two glycine-rich linker sequences that provide spatial separation of domains, imparting structural flexibility within the protein structure [216]. In the case of the xylanase from Neocallimastix patriciarum, a non-catalytic linker sequence of 455 residues has been reported [217]. The sequence unusually comprised 57 repeats of an octapeptide XSKTLPGG where X could be S, K, or N. The O-glycosylation of hydroxy amino acids present in most of the linker sequences is suggested to confer protection from proteolysis. The short linker sequence between the catalytic domain and the C-terminus of xylanase C from Clostridium thermocellum has been found to contain a docking domain which is responsible for cellulosome assembly [125].

12.2.1Catalytic domain

In general xylanases consist of a single catalytic domain. However, analysis of truncated forms of xylanase 3 from Neocallimastix frontalis has indicated that the full length protein contained two catalytic domains displaying similar substrate specificity [218]. In the case of Ruminococcus flavefaciens two catalytic domains have been reported. Catalytic activity measurements and differential scanning calorimetry of exoglucanase/xylanase Cex from C. fimi suggested that binding and catalytic domains of the protein fold independently [92]. The amino acid sequence of the endoxylanase from Cryptococcus albidus has some homology with the catalytic region of the egg white lysozyme [219]. Based on sequence homology it has been interpreted that the glycosidic bond cleavage in xylan is similar to that of lysozyme. However, West et al. [84] have shown that the catalytic domain of this enzyme is homologus to that of cellobiohydrolase from C. fimi and xylanase B from B. subtilis C 125. A xylanase from C. saccharolyticum[116] is found to be homologous with the cellobiohydrolase domain of the Caldocellum bifunctional exocellulase-endocellulase (celB) catalytic domain of cellobiohydrolase from C. fimi[220], xylanases from B. subtilis C 125, C. thermocellum[221], and Cryptococcus albidus[222]. These homology studies suggest that these enzymes may have evolved by shuffling the two catalytic domains with several substrate binding domains. When the xylanases and cellulases from such diverse groups of microbes share extensive homology in their catalytic domains, it is expected that their mechanisms of action also would be closely related. At the same time, the striking absence of homology between the two xylanases from B. circulans[129] must be noted; this could be an indication of a distinct phylogenetic route and/or catalytic mechanism. The full length xylanase C from Fibrobacter succinogenes, expressed in E. coli, is reported to be less active than the two truncated xylanase C proteins, each possessing an intact catalytic domain. It may be that the tertiary structure of the enzyme is such that the catalytic sites are partially buried, as a consequence of improper folding in E. coli. Comparison of protein sequence by HCA showed a clear similarity in the secondary structure elements for the two catalytic domains of Xyn C and the B. pumilus xylanase, indicating a common ancestry as well as related three-dimensional organization. Two tryptophan residues in each domain are found to be completely conserved [223].

12.2.2Cellulose binding domain (CBD)

Catalytic domains and CBDs are grouped into different families based on sequence similarities. Structural studies using NMR have revealed the presence of very few charged amino acids and an unusually large number of conserved aromatic residues in CBD of CbhI and Cex. The CBD of the mixed function glucanase-xylanase Cex from C. fimi contains five tryptophans, two of which are located within the α/β-barrel structure and three are exposed on the surface. NMR analysis and chemical modification studies confirmed that the exposed aromatic residues play a direct role in binding to cellulose; their interaction with cellulose appeared to be reversible [224]. The cellulose binding domains have also been detected in xylanases from Pseudomonas fluorescens, Cellulomonas fimi and Clostridium thermocellum[210]. These domains seem to play two possible roles, i.e. they either open the structure of the plant cell wall, making it more accessible to enzymic hydrolysis, or provide a general mechanism by which a consortium of hydrolases accumulate on the surface of the plant cell wall, resulting in synergistic action between the enzymes. The full length and truncated forms of xylanase D from C. fimi were found to be equally effective in hydrolyzing pulp xylans; enzyme derivatives containing a polysaccharide binding domain were marginally more efficient in decreasing kappa number [225]. Black et al. [226] have studied cellulose and xylan binding domains from xylanase D of C. fimi. The deletion of the cellulose binding domain abolished the cellulose binding capacity of the enzyme without affecting the xylan binding properties. However, the Km values of the truncated xylanase for the insoluble xylan were higher, indicating that the internal cellulose binding homologue in xylanase D constitutes a discrete xylan binding domain which influences the affinity of the enzyme for the insoluble substrate, but does not directly affect the xylanase activity. The existence of a truly functional internal CBD of the type found in C. fimi enzymes has not yet been demonstrated unambiguously [143]. Recently, domain organization and stability of xylanase A from Thermotoga maritima and its C-terminal CBD were studied by expressing the two separate proteins in E. coli. CBD was expressed as a glutathione S-transferase fusion protein. Denaturation/renaturation studies have shown that the domain folds independently [227]. As observed for all Thermotoga proteins investigated so far [228], CBD of Xyn A exhibits extremely high intrinsic stability in the case of CBD. The apparent Tm value of both proteins exceeds 100°C. CBD was found to retain residual secondary structure even at a temperature beyond Tm. CBDs encoded by xylanase 1 from Rhodothermus marinus are shown to be repeated in tandem at the N-terminus exhibiting similarity with CBD family IV [229]. It appears to be the first example of xylanase gene encoding a CBD family IV in combination with the catalytic domain of glycosyl hydrolase family 10. The molecular architecture of four new xylanases, from the aerobic soil bacteria P. fluorescens subsp. cellulosa and C. mixtus, has been studied [230]. Each of the enzymes is modular and contains a novel cellulose binding domain that is separate from the catalytic domain. Two of the enzymes, xylanase E and F from P. fluorescens subsp. cellulosa, contain a domain that is homologous with NodB from nitrogen fixing rhizobia. The authors therefore suggest that there has been a strong selective pressure for the retention of CBDs in xylanases from saprophytic soil bacteria. Recently, functional analysis of the NodB domain of xylanase D from C. fimi has shown that like other members of the family it is a deacetylase, whose function is to remove acetyl groups from acetylated xylan [231]. Characterization of the xylanase A gene from Thermoanaerobacterium thermosulfurigenes EM 1 has revealed the presence of two CBDs and a triplicated sequence at its C-terminus. The latter was found to be identical to S-layer-like domains of previously characterized pullulanase from the same organism [232]. In xylanase producing Streptomyces strains the proteolytic activity for the removal of CBD seems to be conserved [233] since a similar xylanase pattern was obtained for all of them. It is suggested that autoprocessing of protein and protease activity are the two reasons that cause the loss of CBD that might help in the selective hydrolysis of xylan.

12.2.3Thermostabilizing domain

Repeated domains unrelated to the catalytic domain are relatively common in bacterial xylanases and glucanases; however, in most cases, little is known about their functions. Recent work on the thermostable xylanase from Thermoanaerobacterium saccharolyticum[234] has correlated the intrinsic thermostability to the N-terminal domains of the enzymes. Evidence has also been presented that Xyl Y from Clostridium thermocellum[235] contained a homologue of this domain that also appeared to confer thermostability. The thermostabilizing domains have been closely associated with the xylanase catalytic domain. The 180-residue domain of xylanase Y from C. thermocellum has been shown to have 28% sequence identity with the thermostabilizing domain of xylanase A (residues 200–353) from T. saccharolyticum. Recently sequence analysis of the xylanase C gene from C. thermocellum F1 revealed the presence of a 165-amino acid region which was found to be homologous to the thermostabilizing domain [125]. We compared the sequence of Xyn A from T. saccharolyticum with the other reported xylanases from thermophilic organisms. The analysis revealed that the xylanases from B. stearothermophilus, thermophilic bacterium RT8.B4, T. maritima, Anaerocellum thermophilum and Caldicellulosiruptor saccharolyticus have a region similar to the thermostabilizing domain of Xyn A from T. saccharolyticum (Fig. 4). Multiple sequence alignment of these xylanases with Xyn A from T. saccharolyticum indicated conserved Gly and Tyr residues in the region corresponding to the thermostabilizing domain of Xyn A from T. saccharolyticum. The identity of xylanases from other thermophilic organisms with the thermostabilizing domain ranged from 5 to 32%. Xylanase C from C. thermocellum showed a maximum similarity of 32%, while xylanase from B. stearothermophilus showed very little identity. It may be possible that these domains confer thermostability to the corresponding xylanases (Fig. 4). However, more experimental evidence is necessary to validate the concept that thermostability is conferred by a specific domain. Examination of the conserved residues in the thermostabilizing domain reveals the absence of cysteine. A predominance of bulkier aliphatic residues is observed, which may increase thermostability by increasing the compactness of the folded molecule [236,237]. The occurrence of a large number of glycine residues supports the proposition that they may be located in the loop regions of thermophilic proteins [238]. However, comparatively few asparagine and glutamine residues are observed which can cause thermoinactivation, as they are more susceptible to denaturation at elevated temperatures.


Figure 4. Homology of the thermostabilizing domain of xylanase Y from Clostridium thermocellum with other thermophilic xylanases. Accession numbers are in parentheses. (1) T. saccharolyticum B6A-RI (A48490) xylanase; (2) C. thermocellum F1 xylanase C (D84188); (3) B. stearothermophilus 21 xylanase (D28122); (4) thermophilic bacterium RT8.B4 (S12745); (5) T. maritima xylanase A (Z462664); (6) Anaerocellum thermophilum xylanase A (Z69782); (7) Caldicellulosiruptor saccharolyticus xylanase I (AF005382); (8) C. thermocellum YS xylanase Y (X83269). The conserved amino acid residues are indicated by asterisks. Homology to the xylanase from T. saccharolyticum B6A-RI is shown by bold letters.

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It was observed that xylanases from thermophilic organisms exhibited more homology in their catalytic domains. The conserved residues in the catalytic domains could suggest the close evolutionary relationship between the thermophilic bacteria that might have arisen through the lateral transfer of a single ancestral gene between them. The thermophilic xylanase A of Thermomonospora fusca is the one reported to date belonging to family 11 [239]. However, non-catalytic domains conferring thermostability have not yet been detected in family 11 xylanases. One of the reasons may be that these enzymes are inherently more thermolabile, unlike family 10 enzymes in which folding is such that it has been relatively easy for the enzyme to evolve into a thermophilic enzyme [235]. The data imply that domains which confer increased thermostability may be a common phenomenon among family 10 xylanases from thermophilic organisms. Such domains are located only in xylanases and have not been observed in a number of thermophilic endoglucanases characterized to date.

13Molecular evolution

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Scope of the present review
  5. 3Structure of xylan
  6. 4Xylanase production
  7. 5Regulation of xylanase synthesis
  8. 6Biochemical properties
  9. 7Xylanases of extremophilic origin
  10. 8Cloning and expression of xylanase gene(s)
  11. 9Protein engineering
  12. 10Site-directed mutagenesis (SDM)
  13. 11Mechanism of action of the xylanases
  14. 12Domain organization of xylanases
  15. 13Molecular evolution
  16. 14Biotechnological potentials of xylan and xylanases
  17. 15Future prospects
  18. Acknowledgements
  19. References

Recent developments in the molecular genetics of xylanases have added a new dimension to the studies of evolution. Sequence alignment is one of the approaches for obtaining useful information about functional and evolutionary relationships. Residues occupying equivalent positions are believed to share common ancestors and/or to have equivalent biological roles. HCA analysis of amino acid sequences of xylanases and cellulases has revealed that isoenzyme forms of these enzymes, observed in several organisms, are a consequence of large multigene families and are not solely the result of processing of a single gene product. Cellulase and xylanase genes from different families are often present in a single organism suggesting that microorganisms have acquired multiple plant cell wall hydrolase genes not exclusively through gene duplication but via extensive horizontal gene transfer [5]. The occurrence of both fungal and bacterial enzymes in the families 5, 6, 10 and 12 and the presence of prokaryotic and plant enzymes in family 9 led to the proposition that lateral transfer of cellulase and xylanase genes has also occurred.

13.1Sequence homology

A number of reports on sequence homology of xylanases are documented. In general, sequences from the same family (10 or 11) were closely related and exhibited more homology. This criterion has been used to assign particular sequences to particular families. The amino acid sequence of Xyn II (family 11) from T. reesei showed significant homology to alkaline low molecular mass xylanases from the same family, but resembled more bacterial xylanase in several aspects [134]. Schizophyllum commune xylanase (family 11) was found to be the most similar to Trichoderma xylanases and distantly related to Bacillus xylanases [188]. Xylanase B from the fungus Penicillium purpurogenum showed significant similarity with 38 other fungal and bacterial xylanases belonging to family 11. As expected, xylanase B was more closely related to other fungal endoxylanases than to bacterial enzymes with significant homology to xylanase A from A. awamori (73%) [240]. Xylanase A from B. stearothermophilus 21 was 45–50% identical to xylanases from other thermophilic organisms such as C. saccharolyticum and C. thermocellum[117]. Xylanase C (acid xylanase) from A. kawachii showed 40–50% homology with xylanases from B. pumilus, B. circulans and C. acetobutylicum. However, Xyn C from A. kawachii showed no significant homology with xylanase A from the same organism and glucanases from other filamentous fungi [241]. The catalytic domain of STX-II (35.2 kDa) from S. thermoviolaceus OPC-520 showed extensive homology with family 11 xylanases. The two glutamic acid residues were found to be essential for catalysis in the xylanase (STX-II) [242]. Irwin et al. [239] reported that the catalytic domain of xylanase (TfxA) from T. fusca showed 40–72% identity with other xylanases. The xylanases from thermophilic organisms such as xylanase A from C. saccharolyticum[116], xylanase B from C. stercorarium[243] and xylanase T-6 from B. stearothermophilus T-6 [67] showed high homology to xylanase A from alkaliphilic Bacillus C-125. B. steararothermophilus xylanase T-6 showed comparatively less homology to other xylanases from thermophilic organisms, such as Clostidium saccharolyticum, T. saccharolyticum B6A-RI. Xylanase A from C. stercorarium had an Mr of 56 kDa; however, it consisted of a catalytic domain belonging to family 11, and showed high homology to other xylanases from family 11 such as xylanase B from C. acetobutylicum and xylanase A from B. pumilus, suggesting that these genes originate from a common ancestral gene, i.e. these genes were acquired by each bacterium through interspecies gene transfer. Xylanase A (Mr 56 kDa) was an exception as it belonged to family 11, although it had a high molecular mass. Xylanase B from C. stercorarium showed the highest homology to xylanase A irrespective of the differences in their thermal stabilities [244]. The amino-terminal domain of xylanase D (Mr 90 kDa) from R. flavefaciens showed significant sequence similarity with xylanases beloning to family 11. The C-terminal domain of xylanase D, on the other hand, showed identity with glucanases from Bacillus sp. and C. thermocellum. Surprisingly, alignment with the β(1,3-1,4)-glucanase from the rumen species F. succinogenes revealed much lower identity [132]. Xylanase C from the ruminant organism F. succinogenes S85 belonged to family 10. However, domains A and B of xylanase C shared homology with family 11 xylanases from B. circulans, B. subtilis and B. pumilus. Domain A of xylanase C from F. succinogenes showed more homology with fungal ruminal enzyme, whereas domain B exhibited similarities with the bacterial enzymes. It was suggested that these organisms were found in environmental niches totally different from the ruminal organisms, suggesting that their catalytic domains might have had a common origin but they had diverged a long time ago. On the other hand, the enzymes from the three ruminal organisms (R. flavefaciens, N. patriciarum and F. succinogenes) shared a similar multiple catalytic domain structure, which might stem from environmental selection [245]. Xylanases from two different strains of the rumen anaerobic bacterium, Prevotella ruminicola, showed similarities with the N- and C-terminal regions of other family 10 xylanases. The two glutamic acid residues that had been shown to be involved in the active site of family 10 xylanases were situated in the conserved motifs present in the carboxy-terminal regions of both enzymes, distal to the point of insertion of unrelated sequences. The structures of xylanases from the two strains of P. ruminicola represent a departure from the recognized evolutionary route for xylanases and other glycoside hydrolases, which is thought to have involved reassortment of integral domain blocks associated with catalytic, substrate binding and other functions. The relationship between the two enzymes in their N- and C-terminal family 10 sequences suggests that a common ancestral gene must have undergone two independent insertional events during evolution. A possible role for these insertions in the P. ruminicola enzymes might be in anchoring enzymes to other cell wall components, but because of their position adjacent to the catalytic center, it seems likely that they alter the substrate preferences and kinetic properties of the enzymes. It remains to be established whether these insertions are a feature unique to xylanases from the Bacteroides group of bacteria, which may have diverged early in the evolution from other eubacterial phyla [246].

The modular pattern found in the sequence of Xyn Z from C. thermocellum was similar to the structural organization of several cellulases in which similar domains are shuffled at different locations within the sequences [221,247]. Xyn Y from C. thermocellum, in common with xylanases from several thermophilic bacteria, contained a family 10 catalytic domain. This observation could reflect the close evolutionary relationship between thermophilic bacteria. Thus the evolution of xylanase activity might have arisen through the lateral transfer of a single ancestral gene between thermophilic bacteria. This scenario appeared unlikely in view of the fact that endoglucanases from C. thermocellum were derived from several distinct glycosyl hydrolase families, suggesting that the cellulase system of this bacterium arose through extensive lateral gene transfer [235].

13.2Evolutionary relationship of xylanases

Many xylanases and cellulases are known to possess a modular structure comprising a catalytic domain linked to a non-catalytic cellulose binding domain (CBD). CBDs present in cellulases are distinct from catalytic domains, and cellulases are known to bind crystalline and/or amorphous cellulose via CBDs. CBDs are also found in xylanases [216,244] and other plant cell wall hydrolases such as α-l-arabinofuranosidase and acetylxylan esterase [248] and β-mannanase [249] in addition to endoglucanases and exoglucanases. It is necessary for xylanases and other hemicellulases to act cooperatively on plant cell walls which consist of cellulose, hemicellulose and lignin. Hence the binding of hemicellulases to cellulose via CBDs should be advantageous in the hydrolysis of hemicellulose in plant cell walls. This may be the reason for the evolution of xylanases containing CBDs. Although xylan binding domains have also been reported in xylanases, such domains need to be specifically evolved for each xylan due to the heterogeneous nature of the substrate. The integration of CBD may be the efficient way to save the gene capacity exclusively for xylan.

Spurway et al. [184] have reported the presence of a Ca binding site (located in loop 7) in xylanase A from P. fluorescens subsp. cellulosa. The literature survey indicated that only five of the 28 family 10 xylanases contain an extended loop 7. The authors have suggested that the ancestral protein gave rise to family 10 xylanases containing loop 7 which, through natural selection deletions within the loop, resulted in the evolution of stable xylanases. This seems logical since loop 7 does not confer any catalytic properties on the enzyme. Phylogenetic analysis has revealed a relationship between xylanases containing loop 7 and a common ancestral sequence containing DNA insertion in the region encoding loop 7. Since similar sequences occur in taxonomically diverse groups of organisms, a considerable horizontal gene transfer between family 10 xylanases seems to have occurred.

We have compiled the amino acid sequences of xylanases from the PIR databank (Protein International Resource, Release 46.0) (Fig. 5). The sequences were aligned using Clustal V multiple sequence alignment [250]. A minimum mutation distance matrix was then constructed from the pairwise comparisons of 54 sequences. The distance matrix was then used to obtain an evolutionary tree using TAXAN (Release 2.0, Information Resources Group, University of Maryland, USA). The evolutionary tree implies that the xylanases of fungal and bacterial origin appear to be related to each other forming seven different groups. Although xylanases produced by the same organism share maximum homology as observed in the case of T. aurantiacus, T. viride and S. commune, xylanases from two different fungi, i.e. N. frontalis and F. floriforme, are also identical. Fungal xylanases from T. viride, T. aurantiacus and A. niger with bacterial xylanases from C. stercorarium and R. flavefaciens form one group. Thermophilic xylanases from C. acetobutylicum and C. thermocellum seem to be related. However, xylanases produced by two strains of P. fluorescens were found to be distantly related to each other.


Figure 5. Dendrogram showing possible evolutionary relationships among xylanases. The amino acid sequences of xylanases compiled from PIR (Release 46.0) were aligned using Clustal V multiple sequence alignment. Minimum mutation distance matrix then constructed was used to obtain the dendrogram using TAXAN (Release 2.0, Information Resources Group, University of Maryland, USA).

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Thus the xylanases that make up a family/group appear to have diverged from a common evolutionary ancestor. Such enzymes are apt to retain similar secondary and tertiary structure and have the same amino acid residues at 20–50% of the corresponding positions in their primary sequences. The folding of the polypeptide chain is essentially the same in family 11 enzymes with substantial variations occurring only in external loops, e.g. xylanases from T. reesei and T. harzianum. These enzymes are classical illustrations of diverging evolution from a common ancestor. Recently xylanase from Streptomyces viridosporus T7A has been found to be different from the reported xylanases from family 10 or 11. It does not fall into any of the two families. The large size of this protein may be explained by gene duplication of the original ancestral gene, a common occurrence in Streptomyces[251].

14Biotechnological potentials of xylan and xylanases

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Scope of the present review
  5. 3Structure of xylan
  6. 4Xylanase production
  7. 5Regulation of xylanase synthesis
  8. 6Biochemical properties
  9. 7Xylanases of extremophilic origin
  10. 8Cloning and expression of xylanase gene(s)
  11. 9Protein engineering
  12. 10Site-directed mutagenesis (SDM)
  13. 11Mechanism of action of the xylanases
  14. 12Domain organization of xylanases
  15. 13Molecular evolution
  16. 14Biotechnological potentials of xylan and xylanases
  17. 15Future prospects
  18. Acknowledgements
  19. References

During the last decade, the potential biotechnological applications of xylan and xylanases have been of particular interest to researchers. At present, the major end products of xylan, which are of considerable importance, are furfural and xylitol. Furfural production is derived mainly from agricultural residues whereas xylitol is obtained from wood residues. The hydrolysis products of xylan (xylose and oligosaccharides) have possible applications in the food industry as thickeners or as fat substitutes and as an antifreeze food additive. In the pharmaceutical industry xylan is found suitable as an agent for ‘direct tableting’ and, in combination with other components, it can be used for delayed release tablet construction. The xylan hydrolysis products can be subsequently converted to liquid fuel, single cell proteins, solvents and artificial low calorie sweeteners [252].

14.1Applications of xylanases in paper and pulp technology

Environmental regulations have put a restriction on the usage of chlorine in the bleaching process in the paper and pulp industry, especially in Western European countries and in North America [253]. Xylanases play an important role in debarking, deinking of recycled fibers, and in the purification of cellulose for the preparation of dissolving pulp [254]. Xylanase pretreatment has been reported to lower bleaching chemical consumption and to result in greater final brightness. Enzymatic bleaching results from the cleavage of bonds between lignin and carbohydrate and the the opening of the pulp structure [255]. Viikari et al. [256] first showed that treating pulps with hemicellulases can reduce subsequent chlorine bleaching requirements and other investigators [255,257,258] have subsequently confirmed these studies. Xylan and glucomannan form the basic polymers of the wood hemicellulose backbone. Xylanases and mannanases are the two main glycanases that depolymerize the hemicellulose backbone. The effects of purified or partially purified endo-acting β-mannanases from B. subtilis, A. niger and T. reesei on pulp delignification have been studied. Treatment of pine pulp with xylanase enriched with endo-β-mannanase of B. subtilis resulted in only a slight increase in delignification compared with xylanase alone [259]. Mannanase interacts synergisticaly with xylanases to improve the bleaching of kraft pulps specifically from softwoods. Xylanases promote bleaching by the hydrolysis of relocated, reprecipitated xylan on the surface of the pulp fibers, allowing for better chemical penetration and thus improving lignin extractibility [260]. The reprecipitated xylan forms a barrier for the extraction of lignin, in both hardwood and softwood pulps. Thus treatment with xylanase makes the pulp more permeable for the subsequent chemical extraction of residual brown lignin and lignin-carbohydrate molecules from the fibers [261]. Scanning electron microscope studies of xylanase-preteated pulps revealed an increase in the porosity of pulp fiber aiding in pulp accessibility to bleaching chemicals [262]. However, no transverse cracks were observed on the fibers that could decrease their mechanical strength. Alkaline-stable lignin-carbohydrate complexes present in the wood seem to be the major obstacles to solubilization of the residual lignin. During conventional bleaching, these linkages are cleaved by acidic bleaching stages, e.g. chlorine or chlorine dioxide. However, the degradation products adversely contribute to the effluent. In contrast to this, hemicellulose-degrading enzymes selectively hydrolyze polysaccharide chains attached to lignin, thereby decreasing the amounts of chemicals required for pulp bleaching. The xylanases assume special importance in the paper and pulp industry as they replace toxic chemicals such as elemental chlorine and chlorine dioxide for developing eco-friendly processes. At the same time the high molecular mass lignin molecules, which are currently gaining more importance as a valuable byproduct of the paper and pulp industry, can be recovered using enzymatic pretreatments [263].

14.1.1Enzyme-aided bleaching

Xylanases from different organisms have been evaluated for their interaction with various kinds of pulps. On the laboratory scale, xylanases from Streptomyces roseiscleroticus[264], actinomycetes [265], T. harzianum[266], and Humiola sp. [267] have been used for enzymatic pulp treatment to test their bleach boosting abilities. The biotreatment of bagasse pulp using xylanase from an alkaliphilic, thermophilic Bacillus sp. and subsequent peroxide bleaching has resulted in a decrease in kappa number by 10 units and an increase in brightness by 2.5%[268]. Recently, the thermostable xylanase from Thermotoga maritima was compared with commercial pulpzyme HC and was found to be efficient in releasing lignin from kraft pulp [123]. The cloned xylanases expressed in Bacillus cereus[269] and E. coli[255] have also been reported to improve the delignification of unbleached kraft pulps, thus reducing the chlorine required to achieve a certain degree of brightness. In the process of pulp bleaching enzymes with a high pH and temperature optima are of utmost importance. Many alkali-tolerant strains of Bacillus produce xylanases with pH optima of around 9 [270] and have been used for biobleaching. The thermostable xylanase produced by the thermophilic anaerobic bacterium Dictyoglomus sp. [271] has been evaluated for its suitability in pulp bleaching. Xylanase pretreatment at 80°C and pH 6–8 resulted in an increase in brightness by 2 ISO units in one-stage peroxide delignification. Thermostable xylanase from B. sterarothermophilus T-6 bleached the pulp effectively at 65°C and pH 9.0, and has been used successfully in an industrial-scale mill trial [146]. The first commercial xylanase preparation available for pulp bleaching was marketed by Novo Nordisk A/s, under the name ‘Pulpzyme HA’, which was produced by a strain of T. reesei. Subsequently Pulpzyme HB and HC obtained from bacterial sources were marketed. ‘Cartazyme HS’ is another xylanase preparation marketed by Sandoz Chemicals. Irgazyme 40, a commercial preparation containing xylanases from Trichoderma longibrachiatum and T. harzianum E 58, had been tested for peroxide bleaching of Douglas fir kraft pulp [272]. Recently three commercially available xylanases, viz. Ecopulp (from Alko-ICI), Cartazyme-NS-10 (from Clariant) and Pulpzyme HC (from Novo Nordisk), were tested in the bleaching of Eucalyptus kraft pulps. The results indicated a significant decrease in consumption of ClO2 and H2O2[273].

Detailed laboratory studies carried out to adapt the enzymatic treatment to existing mill conditions showed that no expensive investment is necessary for full-scale runs, except for the pH adjustment facilities. Thus, enzymatic pretreatment has been shown to be fully compatible with existing industrial equipment, which is an added advantage. Addition of an enzymatic step to any conventional bleaching sequence results in a higher final brightness value of the pulp. As world pioneers, the Finnish forest companies started mill-scale trials in 1988. Since 1991, this method has been continuously used on an industrial scale in Finland together with other low-chlorine or chlorine-free bleaching methods. The chlorine requirement in prebleaching has been shown to be reduced by 20–30%. As a result, the AOX (adsorbable organic halogen) load of the prebleaching effluent has been reduced by 15–20%. The greatest number of mill trials have been performed in Europe, mainly in Scandinavia, where most of the kraft pulp is produced.

Recently total chlorine-free (TCF) bleaching methods are being developed in which enzymes have been combined with O2, O3 and/or hydrogen peroxide [274]. In TCF bleaching sequences, the addition of enzymes increases the final brightness value, which is a key parameter in marketing the chlorine-free pulps. In addition to this, savings in the TCF bleaching chemicals are important with respect to both costs and the strength properties of the pulp. The more efficient xylanase-yielding strains and technologies will offer a low investment xylanase-aided bleaching that is both environmentally and economically advantageous.

14.2Other applications of xylanases

Xylanases also play a key role in the maceration of vegetable matter [275], protoplastation of plant cells, clarification of juices and wine [8], liquefaction of coffee mucilage for making liquid coffee, recovery of oil from subterranian mines, extraction of flavors and pigments, plant oils and starch [276] and to improve the efficiency of agricultural silage production [252]. The food processing industry is already using the commercial enzyme preparations manufactured by Novo Nordisk. The fungal β-glucanase preparation from A. niger, marketed under the tradename ‘Finizyme’, is used in the fermentation of beer to avoid the difficulties encountered in filtration and the haze caused by β-glucans. Xylanase is also one of the components of the commercial enzyme preparation ‘Ultraflo L’ produced by a selective strain of Humicola insolens. It is a heat-stable multi-active β-glucanase used in the mashing process in beer brewing to secure an efficient breakdown of β-glucans, pentosans and other gums.

The purified xylanase from Trichoderma viride was found to induce the biosynthesis of ethylene and two other pathogen-related proteins in tobacco, suggesting that xylanases may also play a role in induction of plant defence mechanisms [277]. The xylanases from the germinating plant seed primarily convert the reserve food to the assimilable end product. It is also proposed that they play a role in cell elongation, seem to be involved in fruit softening, and are believed to have yet undiscovered important physiological functions.

The xylanases find application in the bakery and the fodder industries due to the presence of substantial amounts of residual hemicellulose in the raw material. In bakeries the xylanases act on the gluten fraction of the dough and help in the even redistribution of the water content of the bread [278], thereby significantly improving the desirable texture, loaf volume and shelf life of the bread. A xylanase (Novozyme 867) has shown excellent performance in the wheat separation process [279], since it has high activity towards soluble arabinoxylan and effects a rapid decrease in the viscosity of wheat flour slurries. The dietary hemicelluloses have little nutritional significance for non- ruminant organisms as they lack the appropriate digestive enzymes. These undigested fibers increase the viscosity of the food in the gut, which interferes with penetration of digestive enzymes, absorption of the digested food and may support pathogenic conditions, especially in broiler chicks. The use of xylanases together with other hemicellulases corrects the problems and also increases the nutritive value of the feed. These biotechnological potentials of xylanases have prompted the search for suitable enzymes and technologies for large-scale economic production.

15Future prospects

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Scope of the present review
  5. 3Structure of xylan
  6. 4Xylanase production
  7. 5Regulation of xylanase synthesis
  8. 6Biochemical properties
  9. 7Xylanases of extremophilic origin
  10. 8Cloning and expression of xylanase gene(s)
  11. 9Protein engineering
  12. 10Site-directed mutagenesis (SDM)
  13. 11Mechanism of action of the xylanases
  14. 12Domain organization of xylanases
  15. 13Molecular evolution
  16. 14Biotechnological potentials of xylan and xylanases
  17. 15Future prospects
  18. Acknowledgements
  19. References

Several applications of xylanases are being developed for the food and paper industries which are based on the partial hydrolysis of xylan. The long-term application of xylanases such as conversion of renewable biomass into liquid fuels, where xylanases play a crucial role in conjunction with the cellulases, is not yet economically feasible. However, stringent environmental regulations and awareness to reduce the emission of greenhouse gases have added an incentive for future research developments in the study of xylanases. In order to make the application of xylanases realistic the improvement in enzyme yields is of utmost importance. Considerable progress has been made in the last few years in identifying the process parameters which are important for obtaining high xylanase yields and productivities, and thus influencing the economics of xylanase production. The production of xylanolytic enzymes is higher with increasing substrate concentration. However, the high concentration of solid substrate gives rise to mass transfer limitations in batch cultivations. A fed-batch mode of cultivation, where much higher substrate concentrations can be used, is an attractive alternative process. This strategy has been successfully employed for the enhanced production of cellulases by Trichoderma on both insoluble and soluble substrates. It can be assumed that similar investigations on fermentation processes for the production of xylanases will result in substantial increases in xylanase activity and productivity. In addition to the mode of operation, alternative designs and configurations of the bioreactor offer opportunities for improvement in the fermentation process. More research efforts are necessary to obtain constitutive mutants which will also eliminate the hindrance of using insoluble carbon sources for the production of xylanase in fermentors.

Developments in the field of enzyme production require strain developments as well as enzyme recovery and downstream processing [280]. Although the enzyme recovery methods are outside the scope of the present review article, their importance in the economics of enzyme production is beyond doubt and greater attention needs to be focused on this aspect. Large-scale enzyme recovery and purification methods based on two-phase separation and affinity purification may be applied to the xylanases. The laboratory-scale affinity purification method already described [272] seems to be promising. The complete bioconversion of xylans to the sugar monomers has so far not been achieved. This is the main hurdle in the commercial success of bioconversion processes from the technical as well as the economic point of view. The enzymatic cleavage of xylan to smaller oligosaccharides is itself a reversible reaction and most of the enzymes show transglycosylating activity and synthesis of higher oligosaccharides under certain reaction conditions. Hence a special process for the enzymatic cleavage of xylan needs to be designed – such as the two-phase reactor in which the product can be continuously separated from the reactants.

The specific activity of the xylanase preparations is much lower than that of commercial preparations such as amylases, proteases and glucose isomerases. There is an urgent need for identifying, otherwise developing, the strain capable of producing a high specific activity xylanase, but the task may not be simple, mainly because of the heterogeneous nature of the substrate – xylan. In addition to the chemical heterogeneity, the substrate is also divided into soluble and insoluble physical states which makes the actual catalysis an extremely complex phenomenon. This catalytic complexity may have resulted in multiplicity amongst the xylanases. Hence the xylanase multiplicity must be analyzed taking into account the microheterogeneity of the substrate. Thus it may be possible to design a mixture of xylanases that has a better specific activity than the individual components.

Cleaner biobleaching technology for the paper and pulp industry is currently concentrated in the developed countries, whereas renewable energy generation from agricultural waste has more relevance for the developing nations. A lot of work needs to be done to bring these research ideas to reality. The xylanases that are commercially available today for possible application in the paper and pulp industry, e.g. pulpzyme HA, HB, and HC from NOVO, do not meet the ideal criteria idntified for enzymatic activity, i.e. optimum activity at pH 10 and temperature >90°C. Hence it is necessary to identify the potent xylanase producer by screening for extremophiles or to design a tailor-made enzyme by the application of protein engineering. Although the creation of disulfide cross-links may prove useful for increasing the thermostability of xylanases so as to make them suitable in biotechnological processes, its use is restricted where disulfide bonds are stable. Interestingly, xylanases without S-S cross-links are known to be more thermostable than those with disulfide bonds [188]. Also, in nature, thermostability of the enzymes from hyperthermophiles appears to be the result of the reduction of water-accessible hydrophobic surfaces rather than the disulfide cross-link strategy [281]. Thus, even though protein engineering can be used to increase conformational stability and enzyme activity, much remains to be learned from natural thermophilic enzymes before precise molecular predictions can be guaranteed. Attempts have been made to recover xylanase DNA from complex soil samples by PCR amplification using degenerate primers. Recovered DNA, which is different from the known xylanases, can be used in several ways to facilitate the production of novel xylanases for industrial applications [282].

The transfer of xylanase genes to selected fermentative microbes in which the enzyme can be produced and secreted in sufficient quantities will enable the fermentative microbes to convert the xylan residue directly into liquid fuels which will have a direct implication in renewable energy conservation. Few microbes possess the metabolic pathways required for single-step conversion of xylan to ethanol [8]. Taguchi et al. [283] have reported for the first time an organism capable of producing hydrogen from xylan. The hydrogen produced from xylan was equivalent to about 73% of the xylose consumed. Thus the strain might be useful to further our understanding the basic technology involved in the biological process of hydrogen production from xylan in plant wastes. Research efforts should be focused on the improvement of such strains for the efficient utilization of biomass. Xylan being an abundant agricultural residue, the chemical energy of the molecule can be meaningfully utilized if the nitrogen fixation activity can be linked to xylan degradation by coupling the xylanase gene and the nif gene cluster.


  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Scope of the present review
  5. 3Structure of xylan
  6. 4Xylanase production
  7. 5Regulation of xylanase synthesis
  8. 6Biochemical properties
  9. 7Xylanases of extremophilic origin
  10. 8Cloning and expression of xylanase gene(s)
  11. 9Protein engineering
  12. 10Site-directed mutagenesis (SDM)
  13. 11Mechanism of action of the xylanases
  14. 12Domain organization of xylanases
  15. 13Molecular evolution
  16. 14Biotechnological potentials of xylan and xylanases
  17. 15Future prospects
  18. Acknowledgements
  19. References

The authors thank Urmila, Prashant and Sunita from the Department of Bioinformatics, University of Pune, Pune, India, for their help in the computer analysis of xylanase sequences. The authors also thank Drs. M.C. Srinivasan, V.V. Deshpande, A. Ahmad and Ms. D. Nath for valuable discussions and providing some of the literature information. We thank Mr. H.B. Sing for the critical reading of the final text. A Senior Research Fellowship to N. Kulkarni from the Council of Scientific and Industrial Research is gratefully acknowledged.


  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Scope of the present review
  5. 3Structure of xylan
  6. 4Xylanase production
  7. 5Regulation of xylanase synthesis
  8. 6Biochemical properties
  9. 7Xylanases of extremophilic origin
  10. 8Cloning and expression of xylanase gene(s)
  11. 9Protein engineering
  12. 10Site-directed mutagenesis (SDM)
  13. 11Mechanism of action of the xylanases
  14. 12Domain organization of xylanases
  15. 13Molecular evolution
  16. 14Biotechnological potentials of xylan and xylanases
  17. 15Future prospects
  18. Acknowledgements
  19. References
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