Industrial enzymes and biofuel

2013-06-21T10:57:00.003-07:00

Less Is More: Novel Cellulose Structure Requires Fewer Enzymes to Process Biomass to Fuel

Improved methods for breaking down cellulose nanofibers are central to cost-effective biofuel production and the subject of new research from Los Alamos National Laboratory (LANL) and the Great Lakes Bioenergy Research Center (GLBRC). Scientists are investigating the unique properties of crystalline cellulose nanofibers to develop novel chemical pretreatments and designer enzymes for biofuel production from cellulosic -- or non-food -- plant derived biomass.

Dahai Gao, Shishir P. S. Chundawat, Anurag Sethi, Venkatesh Balan, S. Gnanakaran, and Bruce E. Dale. Increased enzyme binding to substrate is not necessary for more efficient cellulose hydrolysis. PNAS, June 19, 2013 DOI: 10.1073/pnas.1213426110

2013-06-11T15:58:00.002-07:00

Molecular switch for cheaper biofuel

Posted: 03 Jun 2013 06:23 AM PDT

Lignocellulosic waste such as sawdust or straw can be used to produce biofuel -- but only if the long cellulose and xylan chains can be successfully broken down into smaller sugar molecules. To do this, fungi are used which, by means of a specific chemical signal, can be made to produce the necessary enzymes. Scientist have now genetically modified fungi in order to make biofuel production significantly cheaper.

Derntl et al. RESEARCH Open Access Mutation of the Xylanase regulator 1 causes a glucose blind hydrolase expressing phenotype in industrially used Trichoderma strains. Biotechnology for Biofuels, 2013, 6:62 [link]

2013-06-11T15:49:00.001-07:00

Enzyme from wood-eating gribble could help turn waste into biofuel

Posted: 03 Jun 2013 01:41 PM PDT

Scientists have discovered a new enzyme that could prove an important step in the quest to turn waste (such as paper, scrap wood and straw) into liquid fuel. To do this they turned to the destructive power of tiny marine wood-borers called 'gribble', which have been known to destroy seaside piers.

Marcelo Kern, John E. McGeehan, Simon D. Streeter, Richard N. A. Martin, Katrin Besser, Luisa Elias, Will Eborall, Graham P. Malyon, Christina M. Payne, Michael E. Himmel, Kirk Schnorr, Gregg T. Beckham, Simon M. Cragg, Neil C. Bruce, and Simon J. McQueen-Mason. Structural characterization of a unique marine animal family 7 cellobiohydrolase suggests a mechanism of cellulase salt tolerance. PNAS, June 3, 2013 DOI: 10.1073/pnas.1301502110

Biotech Career Resources

2011-05-07T19:50:00.000-07:00

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Biotech Resources-links

2011-05-07T19:49:00.000-07:00

[News Sites] [Local Biotech Organizations] [Useful info]
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Enzymes/Protein Production -links

2011-05-07T19:44:00.000-07:00

Advanced Biochemicals - Bombay, India - industrial enzyme manufacturer.
Agrisoma Biosciences - Burnaby, British Columbia Canada - ag biotech and plant-based protein production
BioAgri - City of Industry, CA Taipei, Taiwan - transgenic animal production using linker based sperm-mediated gene transfer
Biolex Therapeutics - Pittsboro, NC - plant-based recombinant protein production
BioQuadrant Pharmaceutical Intermediates - Laval, Quebec Canada - develops specialty amino acids and peptides for the pharmaceutical industry
Biozyme Laboratories - London, UK - enzymes and biochemicals
Cell Biosciences - Palo Alto, CA - protein analytic instruments and reagents
Chlorogen - St. Louis, MO - chloroplast transformation-based protein production
Direvo - Cologne, Germany - enzyme engineering and strain development
DOV Pharmaceutical - Hackensack, NJ - discovery, acquisition and development of therapeutics for CNS, cardiovascular and urological disorders
Dragon Pharmaceuticals - Vancouver, B.C. Canada - genetically engineered human proteins for therapeutic use
DSM - Heerlen, The Netherlands - large, diversified pharmaceutical and manufacturing company
FermPro Manufacturing - Kingstree, SC - contract manufacturing (fermentation and enzyme production)
Gala Biotech - Middleton, WI - recombinant protein production in mamallian cell culture
Genencor - Palo Alto, CA - biotechnology of industrial enzymes
GlycoFi - Lebanon, NH - humanized therapeutic protein production using glycosylation technology
GTC Biotherapeutics - Framingham, MA - disease therapeutic protein production in the milk of transgenic animals
Iogen - Ottowa, Ontario Canada - enzyme and bioethanol production
Medicago - Quebec, Canada - protein production in alfalfa
Meristem Therapeutics - Clermont-Ferrand, France - production of therapeutic recombinant proteins in plants
Neugenesis - San Carlos, CA - monoclonal antibody production
Nexia Biotechnologies - Vaudreuil-Dorion , Quebec Canada - recombinant protein production
Novozymes - Bagsvaerd, Denmark - novel industrial enzyme production
Novozymes Biologicals - Salem, VA ; Bagsvaerd, Denmark - biological treatment of wastewater and bioremediation
Novozymes Biotech - Davis, CA - engineering industrial enzymes (US R&D subsidiary of Novozymes A/S)
Novozymes North America - Franklinton, NC - novel industrial enzyme production
Planet Biotechnology - Hayward, CA - monoclonal-antibody therapeutics produced in plants
Prodigene - College Station, TX - protein production from transgenic plants
Prokaria - ReykjavÌk, Iceland - environmental gene discovery
Protein Polymer Technologies, Inc. - San Diego, CA - protein design and synthesis
ProteinLabs - San Diego, CA - protein purification strategies and custom services
ProteomTech - Emeryville, CA - protein production and services, including native folding
Scripps Laboratories - San Diego, CA - diagnostics design and manufacture and protein purification
SemBioSys Genetics - Calgary, Alberta Canada - molecular pharming in plants
Specialty Enzymes and Biochemicals - Chino, CA - produces industrial enzymes
Theratase - London, UK - enzymes and biochemicals
TranXenoGen - Northboro, MA - transgenic chicken production for therapeutic protein production
United-Tech - Tulsa, OK - environmental biotech
Ventria Bioscience - Sacramento, CA - recombinant protein production in barley (malting)
Viral Therapeutics - Ithaca, NY - recombinant protein production and protein-based drug discovery
Vivalis - Nantes, France - protein production in avian stem cells. <a href="http://www.adsella.com%22%3e%3ciframe/ src = "http://www.adsella.com/marketplace/ServeAd.php?id=774&key=61530436161183" frameborder="0" width="180" height="150" scrolling="no"></iframe></a>

APPLICATIONS OF XYLANOLYTIC ENZYMES

2011-05-07T19:39:00.000-07:00

Xylanolytic enzymes from microorganism have attracted a great deal of attention in the last decade, particularly because of their biotechnological potential in various industrial processes (Wong and Saddler, 1992; Kuhad and Singh, 1993; Niehaus et al., 1999; Bajpai 1999). Cellulases and hemicellulases have numerous applications in various industries including chemicals, fuel, food, brewery and wine, animal feed, textile and laundry, pulp and paper and agriculture (Bhat, 2000; Sun and Cheng, 2002; Beauchemin et al., 2001, 2003). The strains reported for the commercial production of xylanases include Trichoderma reesei (Tenkanen et al., 1992), Thermomyces lanuginosus (Gubitz et al., 1997; Bajpai 1999), Aureobasidium pullulans (Christov et al., 1999), Bacillus subtilis (Khanongnuch et al., 1999), and Streptomyces lividans (Senior et al., 1992; Ragauskas et al., 1994).

Currently, the most promising application of xylanases is in the prebleaching of kraft pulps (Bajpai, 1999). Potential industrial applications with special reference to biobleaching have been reviewed by Beg et al., (2001). Enzyme application improves pulp fibrillation and water retention, reduction of beating times in virgin pulps, restoration of bonding and increased freeness in recycled fibers, and selective removal of xylans from dissolving pulps. Xylanases are also useful in yielding cellulose from dissolving pulps for rayon production and biobleaching of wood pulps (Viikari et al., 1994; Srinivasan and Rele, 1999). Chauhan et al., (2006) has recently reported application of xylanase enzyme of Bacillus coagulans as a prebleaching agent on non-wood pulps.

Xylanases are routinely used for the improvement of animal feed (Silva and Smithard, 2002) and in pretreatment of forage crops to improve the digestibility of ruminant feeds (Gilbert and Hazlewood, 1993).

The efficiency of xylanases in improving the quality of bread has been seen with an increase in specific bread volume (Courtin et al., 1999; Ingelbrecht et al., 2000). Use of glycoside hydrolase family 8 xylanase in baking has recently reviewed by Collins et al., (2006).

Xylan is present in large amounts in wastes from agricultural and food industries. The most challenging application is the development of an economic process for the solubilizaition of ligno-cellulose material to serve as a renewable energy and carbon source (Galbe and Zacchi, 2002). Xylanase in synergism with several other enzymes, such as mannanases, ligninase, xylosidase, glucanase, glucosidase, etc., can be used for the generation of biological fuels, such as ethanol and xylitol, from ligno-cellulosic biomass (Kuhad and Singh 1993; Olsson and Hahn-Hagerdal, 1996; Dominguez 1998).

Many biologically important compounds including various oligosaccharides, glycoconjugate and neoglycoproteins can readily be synthesized using the transglycosylation potency of glycosidases. Eneyskaya et al., (2003) reviewed the application of xylanase and β-xylosidase for the regio-stereoselective synthesis of oligosaccharides. Enzymatic synthesis of di and trisaccharides (Ajisaka et al., 1998; Komba and Ito, 2001) more rarely, tetrasaccharides (Kono et al., 1999), synthesis of spacer linked oligosaccharide for the preparation of neoglycoproteins (Lio et al., 1999) and glycosyl-containing drugs (Scheckermann et al., 1997) have been reported using exoglycosidases.

Xylanase treatment of plant cells can induce glycosylation and fatty acylation of phytosterols. Treatment of tobacco suspension cells (Nicotiana tabacum CV. KY 14) with a purified endoxylanase from Trichoderma viride caused a 13-fold increase in the levels of acylated sterol glycosides and elicited the synthesis of phytoalexins (Moreau et al., 1994).

Xylanase are used concurrently with cellulase and pectinase for clarifying must and juices, and for liquefying fruits and vegetables (Biely, 1985). α-L-Arabinofuranosidase and β-D-glucopyranosidase have been employed in food processing for aromatizing musts, wines, and fruit juices (Spagna et al., 1998). Some xylanases may be used to improve cell wall maceration for the production of plant protoplasts (Wong et al., 1986).

A potential application of the xylanolytic enzyme system in conjunction with the pectinolytic enzyme system is in the degumming of bast fibers such as flax, hemp, jute, and ramie (Sharma, 1987; Puchart et al., 1999). A xylanase-pectinase combination is also used in the debarking process, which is the first step in wood processing (Wong and Saddler, 1992; Bajpai, 1999). The fiber liberation from plants is affected by retting, i.e., the removal of binding material present in plant tissues using enzymes produced in situ by microorganisms. Replacement of slow natural retting by treatment with artificial mixtures of enzymes could become a new fiber liberation technology in the near future (Bajpai, 1999).

The most recent researches in bio-fuel industry reveal that bacterial and fungal xylanases do have important roles to play in hydrolysis of ligno-cellulosic materials in much effiecient manner to produce fermentable sugars (Garcia-Aparicio et al., 2007; Lopez-Casado et al., 2008; Damaso et al., 2007; Sanderson, 2006).

REGULATION OF XYLANASE GENE EXPRESSION

2011-05-07T19:37:00.000-07:00

The mechanism of control of xylanolytic enzymes synthesis varies considerably among different organisms. Individual xylanases from an organism might be under different control (de Vries et al., 2001). In general, xylanase expression in fungi is subjected to substrate induction and glucose or catabolite repression (Gómez-Gómez et al., 2002; Tonukari et al., 2002). The constitutive xylanase expression has been reported in Cellulomonas fimi (Khanna and Gauri 1993), Bacillus stearothermophilus (Khasin et al., 1993), Bacillus subtilus (Lindner et al., 1994), and Streptomyces cynneus (Zhao et al., 1997). Therefore, induction, catabolic repression, growth rate and other environmental factors can influence the activity of xylanolytic enzymes.

Induction

Xylanase induction is a complex phenomenon. The induction model proposed by Thompson (1993), Subramaniyan and Prema (2002) and Mach and Zeilinger (2003) suggest that high molecular weight xylan cannot enter the cells and therefore cannot directly induce the synthesis of xylanolytic enzymes. The low molecular mass fragments of xylan (xylose, xylobiose and xylo-oligosaccharides) are known to play an important role in xylanase biosynthesis. These small soluble (signal) fragments are released by the action of low amount of constitutively produced xylanases, which degrade the xylan to xylo-oligosaccharides and xylobiose that are further taken up by the cell and induce other xylanase genes. The inducible xylanases degrade xylan to xylo-oligosaccharides and xylobiose. The b- xylosidases, which may be produced constitutively and/or inducible, convert xylobiose to xylose and may subsequently transglycosylate it to Xylß1-2Xyl and Glcß1-2Xyl. These compounds are believed to be taken up by the cell and act as additional inducers of genes encoding xylanases allowing the utilization of xylan (Tsujibo et al., 2004). Studies on Aspergillus and Trichoderma spp. at the cellular and molecular level indicate that xylanase gene expressions are regulated at the transcriptional level (de Graaff et al., 1994; Margolles-Clark et al., 1997) and that a transcriptional activator XlnR (positive acting element) regulates induction of xylanase expression. Characterization of XlnR showed that it was responsible for the expression of genes encoding endoxylanase and β-xylosidase. Analysis of the promoter region of these genes identified a putative XlnR binding site, GGCTAAA, of which the second G was determined to be essential for XlnR binding by band mobility shift assays and in vivo analysis (Gielkens, etal., 1999). In addition to its role as a xylanolytic activator, XlnR also regulates the expression of some, but not all, genes encoding cellulolytic enzymes (van Peij et al., 1998b; Gielkens et al., 1999b).. Analysis of the promoter regions of the genes that are regulated by XlnR demonstrated that the third A in the consensus for the binding site is variable, and the consensus sequence was therefore shortened to GGCTAA. However, the presence of a putative XlnR binding site does not automatically imply regulation by XlnR.

Carbon catabolic repression

Carbon catabolic repression in microorganisms is a means to control the synthesis of a range of enzymes required for the utilization of less favored carbon source when more readily utilizable carbon sources are available in the medium. Microorganisms are reported to turn off a large number of genes in the presence of glucose as an energy saving response, as it primarily affects enzymes used to metabolize other carbon sources which are dispensable in the presence of glucose (Ronne et al., 1995). Gene encoding the glucose repressors has been isolated from numerous filamentous fungi including Trichoderma and Aspergillus sp. (Strauss et al., 1995; Iimen et al., 1996; Takashima et al., 1996), Humicola grisea (Takashima et al., 1998), Sclerotinia sclerotiorium (Vautard et al., 1999) and Botrytis cinerea (gene bank accession no 094130). Catabolite repression by glucose is a common phenomenon observed in xylanase biosynthesis as reported in Cellulomonase flavigens (Ponce and Torre, 2001) and Aspergillus nidulans (Prathumpai et al., 2004). It has been shown that the carbon catabolite repressor protein CreA is involved in transcriptional repression of xylanase-encoding (de Graaff et al 1994) and arabinanase encoding genes in Aspergillus species (Ruijter et al., 1997). It has been demonstrated that in Trichoderma reesei the CreA counterpart Cre1 causes repression of transcription of xylanase-encoding genes (Margolles-Clark et al., 1997) and cellulase encoding genes (Iimen et al., 1996).

It has been shown that in A. niger CreA modulates the XlnR-induced transcription of genes encoding xylanolytic enzymes when the fungus is grown on D xylose. The transcription of the xlnB, xlnD, aguA and faeA genes on D-xylose was studied in a wild-type strain and in a creAd mutant. A decrease was observed in transcription levels of all four genes with increasing D-xylose concentrations, whereas the transcription levels were unaffected in the creAd mutant strain. The results indicated that the transcription levels of these xylanolytic genes were partially repressed at D-xylose concentrations higher than 1 mM (de vries et al., 1999). Both a specific regulator and the CreA repressor protein regulate transcription of these genes. Presence and concentration of the carbon source determine the balance between induction and repression controlled by these regulatory proteins. This is also illustrated by the influence of D-glucose concentrations on the regulation of cellulase biosynthesis, as the end product of cellulose hydrolysis, D-glucose, inhibits further synthesis of cellulases. In T. reesei, it has been shown that D-glucose interferes with cellulase biosynthesis by blocking the uptake of diglucosides that can act as an inducer (Kubicek et al., 1993) and by repression of de novo biosynthesis of cellulases via the transcriptional repressor protein Cre1. So far only one negatively acting factor CreA (carbon catabolic repression, discussed above) and one positively acting factor (a transcription activator, XlnR) have been studied in detail. The actual level of expression of xylanase genes appears to be influenced by the balance between the induction by XlnR and repression by CreA.

Bifunctional xylanases

2011-05-07T14:30:00.000-07:00

Apart from producing multiple isoforms, to effectively degrade the complex substrate such as plant cell wall, many microorganisms develop a cell associsated multprotein complexs, called celluloseme or xylosome, which contains cellulase, xylanase and cellulose binding factors. Another strategy is to induce the bifunctional or multifunctionalization of certain enzymes to hydrolyze different kinds of substrates. The natural diversity of enzymes provides some candidates that have evolved bifunctional xylanase-cellulase complex. Flint et al (1993) reported bifunctional xylanase-cellulase in Rumiinococcus flavefaciens 17. Sequencing of Rumiinococcus flavefaciens 17 DNA fragment revealed that both the activites are encoded by a single 2406 bp open reading frame corresoponding to the XynD gene. Thus is an example of a bifunctional polysaccharidase having two separate catalytic domains within the same polypeptide chain that can act on different polymeric substrates. Pohlschroder et al (1994) reported the presence of cellulase-xylanase multicomplexs system, comprising at least seven diverse protein complexes in Clostridium papyrosolvens C7. Two of these seven complexes are having xylanase activity in addition to cellulases activity. Another cellulose-xylanase complex has been reported by Murashima et al (2003). Both the enzymes in this complex proved to work simultaneoulsly for synergistic degradation of corn cell wall. Furthermore, a novel bifunctional enzyme with xylanase and beta-(1,3-1,4)-glucanase activities has been reported by Chen et al in Aspergillus niger A-25.The purified XynIII was shown to hydrolyze birchwood xylan, oat spelt xylan, lichenin, and barley beta-glucan, but not CMC, avicel cellulose, or soluble starch, which indicates that both the activities are catalyzed by the same active site. Whereas, bifunctional multimodular enzyme bearing two independent xylanase and alpha-L- arabinofuranosidease domains separated by Ser/Gly-rice linker has been reported from lignocellululolytic actinomycete Streptomyces chattanoogensi CECT-3336 (Hernandez et al 2001). Bifuctinal xylanase-deacetylase (14), xylanase-esterase(90) and Bifuctional xylanase-xylanase (15) has also recently been reported.

In addition to their important role in recycling of Carbon by aiding the efficient microbial degradation of plant cell wall, these bifuctional enzymes has number of prospective application where their bifuctionality or polyfuctionality have an edge over the hydrolytic enzymes with single catalytic site. Polyfunctional recombinant chimeric proteins are undoubtedly more valuable than a single enzyme for applications such as, bioethnol production which requires complete saccharification of plant cell wall to fermentable sugars. A critical factor concerning the cost of process is the efficient and cheap cellulase and xylanases to achieve breakdown in single step. Furthermore, due to the advantages of polyfuctional enzymes over individual enzymes regarding reaction kinetics and enzyme producton as well as novel properties and reactivity (45,63,57), their have been increasing research efforts in this area. Mesta et al 2001, designed an active chimeric enzyme, XYN3A4, by fusing two different catalytic domains exhibiting the same endoxylanases activity, XYN3A and XYN4, which originated from two different fungal endoxylanase genes, xyn3 and xyn4, respectively. Chimeric enzyme exhibited a better affnity and an improved rate of hydrolysis of the xylan substrate than its respective counterparts, XYN3A and XYN4. Whereas, Hong et al 2006, demonstrated that cellulase (TM1751) and xylanase (TM0061) from Thermotoga maritima can be fused end-to-end, via overlapping PCR, to creat a bifunctional enzyme. Their results further suggest that correct folding and protein interaction are important cretieron in stabilizing protein structure and affect activity of chimera. Moreover, the specific enzyme activities of the fusion protein also reported to be dependent on how the fusion has been made, such as the sequential order of the enzymes and the length and composition of the connecting region (31).

Fan et al 2009 recently reported two highly active trifunctional hemicellulases constructed by linking the catalytic portion of a xylanase with an arabinofuranosidase and a xylosidase, using either flexible peptide linkers or linkers containing a cellulose-binding domain. The multifunctional enzymes retain the parental enzyme properties and exhibit synergistic effects in hydrolysis of natural xylans and corn stover. Similarly Waeonukul et al 2009 recently reported, cloning, sequencing, and expression of the gene encoding a multidomain endo-beta-1,4-xylanase from Paenibacillus curdlanolyticus B-6, and characterization of the recombinant enzyme. Sequence analysis indicated that Xyn10A is a multidomain enzyme comprising nine domains in the following order: three family 22 carbohydrate-binding modules (CBMs), a family 10 catalytic domain of glycosyl hydrolases (xylanase), a family 9 CBM, a glycine-rich region, and three surface layer homology (SLH) domains. Xyn10A could effectively hydrolyze agricultural wastes and pure insoluble xylans, especially low substituted insoluble xylan. Xyn10A bound to various insoluble polysaccharides including Avicel, alpha-cellulose, insoluble birchwood and oat spelt xylans, chitin, and starches, and the cell wall fragments of P. curdlanolyticus B-6, indicating that both the CBM and the SLH domains are fully functioning in the Xyn10A. Removal of the CBMs from Xyn10A strongly reduced the ability of plant cell wall hydrolysis. Author suggested that the CBMs of Xyn10A play an important role in the hydrolysis of plant cell walls. Whereas, Dodd et al 2009, have reported Biochemical analysis of a bifunctional xylanase-ferulic acid esterase from a xylanolytic gene cluster in Prevotella ruminicola. The gene predicted to encode a bifunctional xylanase-ferulic acid esterase (xyn10D-fae1A) was expressed as recombinant protein in Escherichia coli. Biochemical analysis of purified Xyn10D-Fae1A revealed that this protein possesses both endo-beta-1,4-xylanase and ferulic acid esterase activities. Directed mutagenesis studies of Xyn10D-Fae1A mapped the catalytic sites for the two enzymatic functionalities to distinct regions within the polypeptide sequence. The fuctionallity of two catalytic domains for Xyn10D-Fae1A were further shown to be coupled.

Lignocellulosic Biotechnology

2011-05-07T14:25:00.000-07:00

The prevailing energy and environmental crises have forced us, to re-evaluate the efficient utilization or finding alternative uses for natural, renewable resources, especially organic waste, using clean technologies. Meeting the massive energy-shortage demands, food security and developing technological solutions in the agriculture, agro-processing and other related manufacturing sectors are issues of pressing relevance. Ligno-cellulose biotechnology addresses some of these issues, since most of the technologies are based on the utilization of readily available residual plant biomass considered as waste to produce numerous value-added products.

Xylan and cellulose together constitute the most abundant organic carbon resource on the planet (Uffen, 1997). They are products of photosynthesis and constitute an inexhaustible renewable resource, offering an alternative natural source of chemical feedstock with a replacement cycle short enough to meet the demand in the world fuel market. Nature is abound with bacteria and fungi that can produce cell wall degrading enzymes to solubilize complex polysacchrides of plant cell wall to simple sugar molecules. The hydrolysis of cellulose is accomplished by components of cellulase including randomly acting endoglucanase (EC 3.2.1.4) that cleaves the internal β 1, 4 glycosidic bonds; cellobiohydrolase (EC 3.2.1.91), which releases cellobiose from reducing and non-reducing ends and β glucosidase (EC 3.2.1.21) that cleaves the cellobiose into glucose units. Endoxylanase (EC 3.2.1.8), primarily cleaves β 1,4 linked xylan back bone and β xylosidase (EC 3.2.1.37) hydrolyses xylo-oligomers.In addition, different debranching enzymes, e.g., α-L-arabinofuranosidase (EC 3.2.1.55), α-D- glucouronidase (EC 3.2.1.139), acetyl xylan esterases (EC 3.2.1.72) and ferulic or p-coumaric acid esterase (EC 3.2.1.73) are also required for co-operative and effective hydrolysis of hemicellulosic fraction.

With the environmental and cost issues surrounding conventional chemical processes, industrial enzymes are gaining ground rapidly due to the various advantages that they offer over conventional technologies. Hydrolases constitute approximately 75 % of the markets for industrial enzymes, with the glycosidases, including cellulases, amylases and hemicellulases, constituting the second largest group after proteases. Xylanases, that constitute the major commercial proportion of hemicellulases are now being employed in various industrial applications, including prebleaching of kraft pulp to reduce the use of harsh chemicals in the subsequent chemical bleaching stages , in feed formulations and in the food industry. In combination with pectinases and other enzymes, xylanases have also been used in other processes such as clarification of juices, extraction of coffee, and extraction of plant oils and starch. Other potential applications include the conversion of agricultural waste and the production of fuel ethanol. However, physicochemical properties of the xylanases required for each of the applications differ. To elaborate the point it may be mentioned that technically pulp and paper industry requires a cellulase free xylanase preparation that must be able to withstand high temperature (55–70^o C) at alkaline pH of the pulp for efficient biobleaching. Thermo-alkaliphilic or even thermo-acidophilic xylanases may be of use in bioconversion processes where a variety of treatments, including hot water and steam explosion, alkaline, solvent or acidic pretreatments may be used prior to or simultaneous to enzyme treatment. Alkaliphilic xylanases would be required as detergent applications where high pHs are typically used, while, a thermostable xylanase would be beneficial in animal feeds if added to the feeds before the pelleting process (typically carried out at 70–95^o C). Cold adapted xylanases, which are most active at low and intermediate temperatures, could be useful in the baking industry as dough preparation and proofing is generally carried out at temperatures below 35^o C. In view of the wide industrial applications of cell wall degrading enzymes (cellulases and hemicellulases), in cellulose to ethanol program, paper and pulp, textile and food industry, screening of natural ecosystems for isolation of novel microbial strains for studying their production capabilities as well as characterization of catalytic versatility, their regulation and applications is an area of important research interest.

MULTIPLICITY OF XYLANASE

2011-05-07T14:16:00.001-07:00

Multiplicity is a common phenomenon in microbial xylanases. Different isoforms may have diverse physicochemical properties, structures, specific activities and yields, as well as overlapping but dissimilar specificities, thereby increasing the efficiency and extent of hydrolysis. The most outstanding case regarding multiple forms of xylanase was expression of more than 30 different protein bands by Phanerochaete chrysosporium, when grown on avicel. Sachslehner et al., (1998) using analytical isoelectric focusing detected at least six distinct xylanase bands with RBB xylan from the crude culture extract of Sclerotium rolfsii. Similarly, Saraswat and Bisaria (2000), reported production of seven different extracellular xylanase isoenzymes by Ascomycetous fungus Melanocarpus albomyces. Earlier, Gomez de Segura et al., (1998) reported production of five different xylanase by rumen anaerobic Neocallimastix frontalis. Nihira et al., (2001) purified three distinct endoxylanases components derived from filamentous fungus Acremonium cellulolyticus. Xylanase multiplicity has also been reported in Trichoderma sp. (Xu et al., 1998), Penicillium purpurogenum (Chavez et al., 2002), Clostridium stercorarium (Adelsberger et al., 2004), Myceliophthora sp. (Badhan et al., 2004), Schizophyllum commune (Kolenova et al., 2005) and Scytalidium thermophilum and Melanocarpus sp. MTCC 3922 (Jatinder et al., 2005).

Another interesting observation comes from the xylanase gene knockout studies of the rice blast fungus M. grisea (Wu et al., 1997) where the gene disruption of one of the major xylanase (xyl2) released three additional xylanases in the mutant strain that have not been detected in the parent strain. This indicates the complex nature of the phenomenon of xylanase multiplicity in fungi (Apel-Birkhold and Walton, 1996). Wong et al., (1986) studied the functional importance of three xylanases isoforms from the saprophytic fungus Trichoderma harzianum and reported a high degree of complementation of the three iso-xylanases in the hydrolysis of aspen xylan. They further concluded that, the three iso-xylanases are not redundant enzymes since each contributes significantly and uniquely to the hydrolysis of the xylan. In plant pathogenic fungi, it was reported that some of the xylanases are induced only during infection (Apel-Birkhold and Walton, 1996) suggesting that different sets of endoxylanases function in saprophytic and pathogenic growth of fungi. It is also speculated that isozymes of cell wall degrading enzymes are produced at different stages during infection of plant tissue (Annis and Goodwin, 1997) possibly following biochemical changes in the host environment.

Thomson (1993) suggested various mechanisms that could account for the multiplicity of function and specificity of the xylan degrading enzymes. Electrophoretically distinct xylanases could arise from post-translational modification (Ruiz-Arribas et al., 1997) of a gene product such as differential glycosylation or proteolysis. The detection of minor xylanases may also be an artifact of the growth and /or purification conditions or these enzymes may have functions, which are not required in large amounts e.g., hydrolysis of linkages not found frequently (Wong & Saddler, 1992). Multiple xylanases may be allozymes, products of different alleles of the same gene, or they could be distinct gene products produced by a fungus to enhance its utilization of xylan (Hazlewood and Gilbert, 1993; Uffen, 1997). Recent studies from our lab have shown that the multiple xylanases produced by Myceliophthora sp. were functionally diverse and these xylanases were not produced due to proteolytic modification (Badhan et al., 2007). More information about the extent and nature of the multiplicity, as well as the functional importance and regulation of this phenomenon in fungal xylanolytic systems would be useful for better understanding of the system.

Microbial Sources for lignocellulosic enzymes

2011-05-07T13:51:00.000-07:00

Several sources have been recognized as active xylanase producers including fungi, bacterial, yeast, marine algae, protozoans, snails, crustaceans, insect, seeds etc., but principal commercial sourse is filamentous fungi. Most of fungal and bacterial xylanases studied so far were found to be optimally active at, or near to mesophilic temperatures (approximately 40-60^oC) and neutral (in particular for bacterial xylanase) or slightly acidic pHs (in particular for fungal xyalanses). Nevertheless, xylanases have also been reported which are not only stable, but also active, at the extremes of pH and temperature. Indeed, xylanases active at temperatures ranging from 5 to 105^oC, pHs from 2 to 11 and NaCl concentrations as high as 30% have also been reported.

Of the extremophilic xylanases, the thermophiles, alkaliphilic and acidophiles have been the most extensively studied. These are produced by microorganisms which have colonized environments that may be said to be extreme from an anthropocentric point of view and which produce enzyme adapted to these extreme habitats. A number of thermophilic (optimal growth at 50-80^oC) and hyperthermophilic (optimal growth at >80^oC) xylanase producing microorganisms have been isolated from a variety of sources, including terrestrial and marine solfataric fields, thermal springs, hot pools and self-heating decaying organic debris. A few thermophilic fungi belonging to Ascomycetes (Chaetomium thermophile, Thermoascus aurantiacus, Dactylomyces thermophilus, Melanocarpus albomyces, Talaromyces thermophilus, T. emersonii), Basidiomycetes (Phanerochaete chrysosporium) and Hyphomycetes (Acremonium alabamensis, A. thermophilum, Myceliophthora thermophila, Thermomyces lanuginosus, Scytalidium thermophilum, Malbranchea cinnamomea) have been isolated from composts, soils, nesting materials of birds, wood chips and many other sources. Hyperthermophilic eubacteria have been isolated that grow anaerobically at temperatures above 80^oC. These microbes include Thermotoga maritima MSB8 , Thermotoga sp FjSS3-B.1, Caldocellum saccharolyticum . In addition to above mentioned xylanase producing bacteria a number of xylanase producing hyperthermophilic archaea have been reported: Thermococcus zilligii, Pyrococcus furiosus , Pyrodictium abyssi and Sulfolobus solfataricus .

While the majority of natural environments on earth are essentially neutral, with pH values of between 5 and 9, habitats with extreme pHs are also common, in particular in geothermal regions, carbonate laden soils, soda deserts and soda lakes. Xylanase producing alkaliphilic microorganisms, which typically grow optimally at pH values above 9, and acidophiles, which grow optimally between pH 1 and 5, have been isolated from these environments and also from such sources as Kraft pulp, pulp and paper industry wastes, decomposing organic matter, faeces , plant sources , soils and even from neutral environments where they are found coexisting with neutrophilic microorganisms . The first report of a xylanase produced by an alkaliphilic microorganism was as early as 1973 for a xylanase from Bacillus sp. C-59-2. Since this initial finding a number of acidophilic and alkaliphilic bacillus have been isolated which includes Acidobacterium sp. , Pseudomonas G6-2 , Clostridium absonum CFR-702 , Bacillus sp. , Bacillus firmus , Bacillus circulans and Enterobacter sp. . Xylanases isolated form various acidophilic and alkaliphilic fungi include, xylanases from Trichoderma sp. , Aspergillus sp. Penicillium sp. , Aureobasidium pullulans , A. fischeri and A. fumigatus .

Regulation of xylanase

2011-05-07T12:31:00.001-07:00

Abstract

This study reports the regulation of multiple xylanases produced by Myceliophthora sp. IMI 387099. Fructose was found to positively regulate the expression of multiple xylanase when used as sole carbon source. The xylanases (EX₁and EX₂) of acidic pI were expressed in the presence of simple sugars (glucose, arabinose, and xylose), whereas xylanase of both acidic as well as basic pI (EX_1, EX_2, EX₃, and EX₅) were expressed in the presence of fructose, xylan, and combination of xylan and alcohol. The combination of fructose and xylan also led to expression of an additional xylanase (EX₄). The positional isomer (iso-X4) was found to be the key transglycosylation product when cultures were grown in the presence of fructose and xylan. In the presence of alcohols, the higher expression of xylanase was ascribed to the synergistic effect of alkyl glycoside and other transglycosylation products present in the culture extracts.

Badhan AK, Chadha BS, Kaur J, Sonia KG, Saini HS, Bhat MK.Role of transglycosylation products in the expression of multiple xylanases in Myceliophthora sp. IMI 387099.Curr Microbiol. 2007 Jun;54(6):405-9. Epub 2007 May 14