By Dhawan, Samriti Kaur, Jagdeep
ABSTRACT Microbial mannanases have become biotechnologically important since they target the hydrolysis of complex polysaccharides of plant tissues into simple molecules like manno- oligosaccharides and mannoses. The role of mannanases in the paper and pulp industry is well established and recently they have found application in the food and feed technology, coffee extraction, oil drilling and detergent industry. Mannanses are enzymes produced mainly from microorganisms but mannanases produced from plants and animals have also been reported. Bacterial mannanases are mostly extracellular and can act in a wide range of pH and temperature, though acidic and neutral mannanases are more common. This review will focus on complex mannan structure and the microbial enzyme complex involved in its complete breakdown, mannanase sources, production conditions and their applications in the commercial sector. The reference to plant and animal mannanases has been made to complete the overview. However, the major emphasis of the review is on the microbial mannanases. KEYWORDS hemicellulases, mannanase, mannooligosaccharides, mannose, locust bean gum, konjac glucomannan
Enzymes are the catalytic keystone of metabolism and are the hub of concentrated worldwide research. The advent of enzymology represents an important breakthrough in the biotechnology industry, with the worldwide increase in usage of enzymes. The latter half of the twentieth century saw an unparalleled expansion in our knowledge of the use of microorganisms, their metabolic products, and enzymes in a broad area of basic research and their potential industrial applications. Hydrolases such as proteases, amylases, amidases, esterases and carbohydrolases such as cellulases, hemicellulases and pectinases occupy the major share of the industrial enzyme market. Cellulases and hemicellulases have numerous applications and biotechnological potential (Bhat, 2000; Sun and Cheng, 2002; Wong and Saddler, 1992; Beauchemin et al, 2001). It is estimated that approximately 20% of the more than one billion US dollars of the world’s sale of industrial enzymes consist of cellulases, hemicellulases and pectinases (Bhat, 2000).
Mannan and heteromannans are a part of the hemicellulose fraction in plant cell walls. Structural analysis of plant cell wall polysaccharides has revealed that the cell wall of a dicot sp. contains three major classes of polysaccharides viz. cellulose, hemicellulose and lignin (Dekker et al., 1985). Hemicelluloses are defined as those plant cell wall polysaccharides that are not solubilized by water or chelating agents but are solubilized by aqueous alkali (Selvendran and O’Neill, 1985). According to this definition, hemicelluloses include mannan, xylan, galactan and arabinan. Hemicelluloses are also defined chemically as cell wall polysaccharides that are structurally homologous to cellulose because they have a backbone composed of 1,4-linked beta-D- pyranosyl residues (O’Neill and York, 2003). Hemicellulose is abundant in primary walls but is also found in secondary walls (Puls, 1997).
In recent years, hemicellulases have emerged as key enzymes in the rapidly growing biotechnology industry, owing to their multifaceted properties, which find usage in a wide array of industrial applications (Hongpattarkere et al., 2002). Most studies on hemicellulases have focused until now on enzymes that hydrolyse xylan, the primary constituent of hemicellulose in grasses. Enzymes that hydrolyse mannan have been largely neglected, even though it is an abundant hemicellulose, therefore the application of mannanase for catalyzing the random hydrolysis of beta-D-1,4 mannopyranoside linkages in beta-1,4 mannans is as important as the application of xylanases.
The use of mannanases in the paper and pulp industry increased significantly with the discovery of Gubitz et al. (1997). Since then researchers worldwide have focused their attention toward newer microbial isolates for mannanases. Despite the availability of several reports on mannolytic microorganisms, there are only a few reviews available on mannanases. These reviews emphasize mainly production and properties of mannanases and not their biotechnological potential. Thus there is a need to assemble information available on these enzymes from different sources in order to bring to light their importance and direct research into the probable application of these enzymes. The present review will encompass the microbial sources, production, properties and potential industrial applications of mannanases.
MANNAN: OCCURRENCE AND STRUCTURE
Mannans and heteromannans are widely distributed in nature as part of the hemicellulose fraction in hardwoods and softwoods (Capoe et al., 2000), seeds of leguminous plants (Handford et al., 2003; Buckeridge et al, 2000) and in beans (Lundqvist et al., 2002). Hemicelluloses are copolymers of both hexose and pentose sugars. The branched structure allows hemicellulose to exist in an amorphous form that is more susceptible to hydrolysis. Within biomass, mannans or the hemicelluloses are situated between the lignin and the collection of cellulose fibres underneath. Consistent with their structure and side group substitutions, mannans seem to be interspersed and covalently linked with lignins at various points while producing a coat around underlying cellulose strands via hydrogen bonds, but as few H-bonds are involved they are much more easily broken down than cellulose. The mannan layer with its covalent linkage to lignin and its non-covalent interaction with cellulose may be important in maintaining the integrity of the cellulose in situ and in helping protect the fibers against degradation to cellulases (Puls and Schuseil, 1993).
Mannan is the predominant hemicellulosic polysaccharide in softwoods from gymnosperms, but is the minor hemicellulose in hardwood from angiosperms (Puls, 1997). Unsubstituted beta-1,4- mannan, composed of a main chain of beta-mannopyranose residues, is an important structural component of some marine algae (Yamasaki etal., 1998) and terrestrial plants such as ivory nut (Chanzy etal., 2004) and coffee bean (Nunes etal., 2006). It resembles cellulose in the conformation of the individual polysaccharide chains, and is water insoluble.
Hardwood mannans are composed of beta-1,4-linked mannopyranose and glucopyranose units, whereas softwood contains two different types of acetylated galactoglucomannans. These consist of glucose, mannose and galactose in the ratio 1:3:1 and 1:4:0.1 respectively (Lundqvist etal., 2002). In leguminous seeds, water soluble galactomannan is the main storage carbohydrate, comprising up to 20% of the total dry weight (McCleary, 1988). It has a a-galactose linked at the O-6 position of some mannose residues and may also have some beta-D-glucose residues incorporated in the backbone. Furthermore, the mannose residues can be acetylated to various degrees at the C-2 and C-3 positions (Figure 1). The galactomannans from different leguminous taxonomic groupings differ in their degrees of galactose substitution and M:G ratios between 1.1:1 (high galactose) and 3.5:1 (low galactose) are encountered. The high degree of galactose substitution of the (1-4) beta-D-mannan in galactomannans is clearly sufficient to prevent the chain aggregation that leads to insolubility and crystalline order in the mannans and glucomannans. The mannose: galactose ratio (M:G) is key in determining the amount of intermolecular association called hyperentanglements. Without any galactose side chains, mannan backbone will aggregate due to intermolecular interaction between the unbranched parts of heteromannans.
FIGURE 1 General structure of mannan and heteromannans. A) A typical mannan structure, a main chain of beta-1,4 linked mannose (Man) residues; B) A typical galactomannan structure, a main chain of beta-1,4 linked mannose residues with alpha-1,6 linked galactose (Gal) residues attached to some (Man) residues; C) A typical glucomannan structure, a main chain of beta-1,4 linked mannose (Man) and glucose (Glc) residues; D) A typical galactoglucomannan structure, a main chain of beta-1,4 linked mannose (Man) and glucose (Glc) residues, with alpha-1,6 linked galactose (Gal) residues attached to some (Man) residues.
The substrate used routinely for the study of mannanases is galactomannan from locust bean gum (Ceratonia siliqua) with a mannose: galactose ratio of 4:1 (De Nicolas-Santiago et al., 2006). In addition, ivory nut (Phytelephas macrocarpa) mannan, an unbranched beta-1,4-linked mannan homopolymer and manno- oligosaccharides (mannobiose, mannotriose, mannotetraose and mannopentoase), can also be used as substrates in the hydrolysis (Stoll et al., 2005).
TABLE 1 The major mannanases and their classification
ENZYMATIC HYDROLYSIS OF MANNAN
The main component of mannan is D-mannose, a six carbon sugar, but due to the heterogeneity and complex chemical nature of plant mannans, its complete breakdown into simple sugars that can be readily used as energy sources by particular microorganisms, the synergistic action of endo-1,4-beta-mannanases (E.C 126.96.36.199, mannan endo-1,4-beta-mannosidase) and exoacting beta-mannosidases (E.C 188.8.131.52) is required (Table 1). Additional enzymes, such as beta- glucosidases (EC 184.108.40.206), alpha-galactosidases (EC 220.127.116.11) and acetyl mannan esterases (Tenkanen, 1998) are required to remove side chain sugars that are attached at various points on mannans. A galactomannan structure whose glycosidic bonds are hydrolyzed by two enzymes viz. endo-1,4-beta-mannanases and alpha-galactosidases is shown in Figure 2. FIGURE 2 Scheme of Enzymatic Action on Galactomannan. The beta-(1-4) linked polymannose chain is substituted with alpha-(1-6) linked galactose (GAL) residues. The arrows represents the glycoside links recognized by beta-mannanase and alpha-galactosidase.
beta-1,4-mannanases are endohydrolases that cleave randomly within the 1,4-beta-D mannan main chain of galactomannan, glucomannan, galactoglucomannan, and mannan (McCleary and Matheson, 1986). Apart from their ability to hydrolyze different mannans, some beta-D-mannanases have also been reported to transglycosylate manno- oligosaccharide substrates (Harjunpaa etal., 1995; Schroder etal., 2004).
Hydrolysis of these polysaccharides is affected by the degree and pattern of substitution of the main chain by beta-D-galactosyl residues in galactomannan and galactoglucomannan (McCleary, 1979) and by the pattern of distribution of D-glucosyl residues within the main chain in glucomannan and galactoglucomannan. In glucomannan, the pattern of distribution of O-acetyl groups may also affect the susceptibility of the polysaccharide to hydrolysis. Like beta- glucosidases in the cellulase system, beta-mannosidases are essential for the complete hydrolysis of plant heteromannans. They convert the manno-oligosaccharides produced by beta-mannanases to mannose (Franco etal., 2004). The galactose release from softwood pulp is enhanced by the presence of mannanase in combination with alpha-galactosidase (Clark etal., 2000). The main products obtained during the hydrolysis of mannan by beta-mannanases are mannobiose and mannotriose. beta-mannanases from Aspergillus tamarii(Civas etal., 1984), Trichoderma reesei (Stalbrand et al., 1993) and Aspergillus niger (Ademark et al., 1998) all produced mainly mannobiose and mannotriose and traces of higher oligosaccharides.
The pattern of galactomannan hydrolysis and the apparent subsite- binding requirements have been interpreted from structural analysis by NMR of isolated hydrolysis products from glucomannan and galactomannan incubated with Aspergillus niger beta-mannanase. It is suggested that the enzyme has five substrate binding subsites- alpha, beta, gamma, delta and epsilon, which is equivalent to -3, – 2, -1, +1, and +2, according to the nomenclature given by Davies et al. (1998). Binding to at least four subsites is required for efficient hydrolysis. Substitution of the substrate monomers at two of the subsite positions restricted hydrolysis, most likely by preventing binding (McCleary and Matheson, 1986). The hydrolysis of the pine craft pulp by the Trichoderma reesei beta-mannanase was also studied (Tenkanen etal, 1997). Judging from this study, Trichoderma reesei enzyme also appears to be restricted by galactosyl residues in a similar way to the Aspergillus niger enzyme.
SOURCES OF MANNANASES
Mannanases are ubiquitous in nature and are elaborated by a compendia of microorganisms largely isolated from natural environments. A vast variety of bacteria, actinomycetes, yeasts and fungi are known to be mannan degraders (Talbot and Sygusch, 1990; Puchart et al., 2004). The important degraders are listed in Table 2.
Among bacteria, degradation is mostly confined to gram- positives, including various Bacillus species (Yanhe et al, 2004; Sun et al., 2003) and Clostridia species (Kataoka et al., 1998; Perret et al., 2004; Nakajima and Matsuura, 1997). However, a few strains of gramnegative bacteria, viz. Vibrio (Tamaru etal., 1997, Pseudomonas (Braithwaite et al., 1995) and Bacteroides (Gherardini and Salyers, 1987) have also been reported. In addition, a few thermophiles and extremophiles belonging to genera Bacillus, Caldocellum, Caldibacillus, Rhodothermus, have also been described (Hatada et al., 2005; Morris et al., 1995; Sunna et al., 2000; Politz et al., 2000). Besides these, actinomycetes from the streptomycetes group, viz. Streptomyces galbus (Kansoh and Nagieb, 2004), Streptomyces lividans (Arcand et al., 1993), actinobacteria group, viz. Cellulomonas fimi (Stoll et al., 1999) and the actinoplanetes group viz. Thermomonospora fusca KW3 (Hilge et al., 1996) are described as mannan degraders with an ability to act on a wide variety of mannan substrates. The most mannolytic group among fungi belongs to genera Aspergillus, Agaricus, Trichoderma, Sclerotium (Huang et al., 2007; Chen et al., 2007; Tang et al., 2001; Franco et al., 2004; Sachslehner et al., 2000). Thus the property of mannolysis is widespread in the microbial world. Mannanases of microbial origin have been reported to be both induced as well as constitutive enzymes and are usually being secreted extracellularly into the medium in which the microorganism is cultured. The bacterial mannanases produced by Sporocytophaga coccoids and Aerobacter mannolyticus were found to be intracellular (Dekker and Richards, 1976). Extracellular mannanases are of considerable commercial importance, as their bulk production is much easier. Mannanases are also produced in higher plants (Shimahara et al., 1975; Marraccini et al., 2001) and animals (Yamaura et al., 1993, 1996) Although a number of mannanase-producing bacterial sources are available, only a few are commercially exploited as wild or recombinant strains, of these, the important ones are: Bacillus sp., Streptomyces, Caldibacillus cellulovorans, Caldicellulosiruptor Rt8B, Caldocellum saccharolyticum. (Zhang et al., 2006; Hatada et al., 2005; Morris et al., 1995; Sunna et al., 2000).
TABLE 2 Sources of mannanases
Numerous screening methods exist for detecting mannanase activity in microorganisms. A solid medium providing rapid assays is useful for the direct measurement and isolation of mannanase-producing organisms from natural substrates and for the isolation of mutants. Common screening techniques devised for the detection of mannan degrading enzymes by microorganisms involve plate assays where the respective polymer, or its derivative, is incorporated into the basal growth medium. The production of corresponding hydrolases is indicated by the clearing of the opaque medium as the substrate is dissolved by the enzymes produced by the growing colonies. In such procedures, the results are often difficult to interpret in case of fungal isolates, because the zones of clearing are not always easy to distinguish from the unaffected medium and activity might not be detected because of the masking effect of the mycelial mass when the rate of fungal growth exceeds enzyme diffusion. A gel-diffusion assay for the quantification of endo-beta-mannanase activity has been developed (Downie et al., 1994). The assay is specific and detects activity as low as 0.07 pkatal. The assay was equally effective at acidic and neutral pH. One per cent (w/v) Congo Red dye resulted in fast staining times (15 min) and good contrast. The diameter of the clearing zones decreased linearly with increasing substrate concentration in the range of 0.1-1.0% w/v galactomannan. The assay was specific for this endoenzyme, with no zone of clearing developing for alpha-galactosidase, and only a slight decrease in dye intensity with beta-mannosidase. The clearing zone diameter was greater in the gel-diffusion assay using Congo Red than using Remazol Brilliant Blue colored Carob substrate at identical concentrations, enzyme activities and assay conditions. This problem was further overcome by inoculating a polycarbonate membrane 0.2 [mu]m, 90 mm diameter, Nucleopore Co (Chang et al., 1992), or a circle of sweet cellophane 400 (De Nicolas-Santiago et al., 2006) with the test fungus. After incubation, the membrane is removed and the enzyme activity recorded. The membranes, which can resist penetration by fungi, are placed between moistened filter papers and sterilized by autoclaving before use.
Solid media are also used to screen cell-free extracts from microorganisms for mannan degrading enzyme activity. Generally, the enzyme solution is placed in a well (cup) cut in an agar medium containing a mannan/heteromannan substrate. The agar surface usually needs to be flooded with a reagent to develop the zones of enzyme activity around the cup. The diameter of the cleared zone is measured with calipers and recorded. Diameters are converted to enzyme activity by running a dilution of the enzyme and plotting zone diameter against the log of the enzyme concentration (Dingle et al., 1953).
An assay based on the quantification of hydrolysis products has been described by McCleary (1988). Dyed polysaccharide substrates (AZO-carob galactomannan, Megazyme) can also be used as a substrate and incubated with enzyme for ten minutes; ethanol is added to stop the reaction and to precipitate high molecular-weight galactomannan. Low molecular-weight fragments stay soluble and can be spectrophotometrically quantified at 590 nm, due to the attached dye (Stalbrand, 2003). To relate to enzyme activity units, a standard curve (supplied by Megazyme for each batch of substrate) with known enzyme amounts can be prepared.
Microbial mannanases are mostly extracellular and are greatly influenced by nutritional and physicochemical factors, such as temperature, pH, nitrogen and carbon sources, inorganic salts, agitation and dissolved oxygen concentration (Mabyalwa and Setati, 2006; Li et al., 2006; Lin et al., 2007).
However their production is significantly influenced by using different mannan substrates such as glucomannan, galactomannan, galactoglucomannan. Galactomannans of guar gum (mannose:galactose) ratio of 2:1 and locust bean gum (mannose:galactose) ratio of 4:1 and glucomannan of konjac (mannose:glucose) ratio of 1.6:1 induced their production (Dong et al., 1991; Yague et al., 1997). Besides carbon source, the type of nitrogen source in the medium also influences the mannanase titers in production broth (Kataoka and Tokiwa, 1998). Generally, organic nitrogen is preferred, such as peptone and yeast autolysate (Puchart et al., 2004), which have been used as a nitrogen source for mannanase production by various Bacillus strain viz. Bacillus polymyxa, Bacillus subtilis, Bacillus brevis (Zhang et al., 2006; Cui et al., 1999) while beef extract and peptone have been used in the case of Clostridium tertium (Kataoka and Tokiwa, 1998). Inorganic nitrogen sources such as ammonium sulfate, diammonium hydrogen phosphate, ammonium dihydrogen phosphate and sodium nitrate have also been reported to be effective in some microbes (Zakaria et al., 1998; Perret et al., 2004; Araujo and Ward, 1990). Divalent cations stimulate or inhibit enzyme production in microorganisms. Nakajima and Matsuura (1997) observed stimulation in mannanase production from Clostridium sp. in the presence of Ca^sup 2+^ and Mg^sup 2+^. However, most other metal ion salts Fe^sup 3+^, Al^sup 3+^, EDTA, Hg^sup 2+^ were inhibitory to mannanase production (Chen et al., 2000; Sun et al., 2003). Iron was found to play a critical role in the production of mannanase by most of the microorganisms. In addition to the various chemical constituents of a production medium, physiological parameters such as pH, temperature, agitation, aeration and incubation period also play an important role in influencing production by different microorganisms. The initial pH of the growth medium is important for mannanase production. Largely, bacteria prefer pH around 7.0 for best growth and mannanase production, such as in the case of Bacillus sp., Clostridium sp. (Helow and Khattab, 1997; Talbot and Sygusch, 1990) However, maximum activity at higher pH (>7.0) has been observed in some cases (Hatada et al, 2005). The optimum temperature for mannanase production corresponds with the growth temperature of the respective microorganism, for example, the best temperature for growth and mannanase production in the case of Bacillus sp. M50 was 50[degrees]C (Chen et al., 2000). It has been observed that, in general, mannanases are produced in the temperature range 4-85[degrees]C (Politz and Borriss, 2000; Kansoh and Nagieb, 2004). Incubation periods ranging from a few hours to several days have been found to be best suited for maximum mannanase production by bacteria. An incubation period of 24 h was optimum for mannanase production by Bacillus subtilis 168 (Helow and Khattab, 1996), and 36 h for Bacillus subtilis BM 9602 (Cui etal., 1999), maximum mannanase was produced after 72 h and 96 h of incubation. Mannanases are often secreted into the culture liquid as multiple enzyme forms as observed for some bacteria (Stoll etal., 1999; Akino et al., 1989), and several fungi including Schkrotium rolfsii (Gubitz etal., 1996), Aspergillus fumigatus (Puchart et al., 2004) and Trichoderma reesei (Stalbrand etal., 1993). The multiplicity of these mannanases is thought to be due to their ability to bind and degrade different substrates (Johnson et al., 1990). These beta-1,4- mannanase isoforms may be secreted as products of the same gene differing only in their post-translational modification, such as with A. fumigatus (Puchart et al., 2004) or their production may be regulated differently such as Sclerotium rolfii isoforms that exhibited different functions on substrates of varying sizes (Grosswindhager etal., 1999). Kinetics of mannan depolymerization, i.e. Michaelis-Menten constant (K^sub m^) and the maximal reaction velocity (V^sub max^) values, are reported for different fungal and bacterial beta-mannanases. K^sub m^, and V^sub max^ values reported for Aspergillus Jumigatus are 3.07 mg/ml and 1935 [mu]mol min^sup – 1^ mg^sup -1^ for Man I and 3.12 mg/ml and 2139 [mu]mol min^sup -1^ mg ^sup -1^ for Man II on locust bean gum (Puchart et al., 2004). For Aspergillus tamarii V^sub max^ and K^sub m^, reported are 43.5 [mu]mol min^sup -1^ mg^sup -1^ and 2 x 10^sup -2^ g/100 ml (Civas et al., 1984) on locust bean gum. Franco et al (2004) reported V^sub max^ and K^sub m^ values as 20.1 units /ml and 6.0 mg/ml, respectively, for Trichoderma barzanium strain T4 on wheat bran. Among bacterial beta-mannanases (Talbot and Sygusch, 1990) reported for Bacillus stearothermophilus a V^sub max^ value of 455 +- 60 U/ mg and the estimated Km value of 1.5 +- 0.3 mg/ml on locust bean gum. In Bacillus subtilis SA-22 the Michaelis constants (K^sub m^) were measured as 11.30 mg/ml for locust bean gum and 4.76 mg/ml for konjac powder, while V^sub max^ for these two polysaccharides were 188.68 [mu]mol min^sup -1^ ml^sup -1^ and 114.94 [mu]mol min-1 ml^sup -1^ respectively (Sun etal., 2003).
Thus, microbial mannanases are generally produced in the presence of different mannan substrates, viz. guar gum (Cyamopsis tetragonobola), locust bean gum (Ceratonia siliqua), and konjac glucomannan (Amorphophallus konjac) in the presence of any complex nitrogen source. Dye-based substrate ostazin brilliant red mannan and azo-carob galactomannan can also be used for easy screening of mannanase producing microorganisms. The requirement for metal ions varies with the organism. However, physical parameters such as pH, temperature, agitation and aeration influence mannanase production via modulating the growth of the bacterium. Mannanases are produced throughout bacterial growth, with peak production being obtained by the late logarithimic phase. The production period for mannanases varies from a few hours to a few days. A list of various production conditions used with different microorganisms is presented in Table 3.
FAMILY CLASSIFICATION AND MODULARITY OF MANNANASES
Mannanases are glycoside hydrolases that degrade mannans and heteromannans. Glycoside hydrolases (EC 3.2.1.-) are a widespread group of enzymes that hydrolyze the glycosidic bonds in oligo- and polysaccharides. Due to the complicated structures of the carbohydrates with different combinations in nature, a large number of enzymes with different substrate specificities are required. Glycoside hydrolases from various sources were classified into different families based on their amino acid sequence similarities (Henrissat and Bairoch, 1993, 1996). The basic principle behind the family classification is that the family membership of a particular enzyme can be defined from its sequence alone and there is a direct relationship between sequence and folding similarities. Structure conservation was shown to be much stronger than amino acid conservation. Families can thus be grouped into “clans” according to their three-dimensional structure (Henrissat, 1990). So far there are a total of 110 families and some of them are well studied. The beta-mannanase sequence comparison studies permit assignment of these enzymes to either glycoside hydrolase family 5 or 26. Family 5 comprises several bacterial mannanases including Caldocellum saccharolyticum, Caldibacillus, Vibrio species, fungal mannanases from Aspergillus aculeatus, Trichoderma reesei and Agaricus bisporus and eukaryotic mannanase from Lycopersicon esculentum and Mytilus edulis. Family 26 comprises mannanases from Bacilli sp., Cellvibrio japonicus, Pseudomonas fluorescens and Rhodothermus marinus. Thus both bacterial and eukaryotic mannanases have been annotated to family 5. With the exception of a few anaerobic fungi, the mannanases in family 26 are of bacterial origin. In some cases, mannanases from the same genus have been classified in different families; beta-mannanases from different strains of Caldocellulosiruptor saccharolyticus have been classified in both families 5 and 26 (Gibbs et al., 1996), and multiple beta- mannanases in Cellvibrio japonicus have been classified in both families 5 and 26 (Hogg et al., 2003). Besides this, beta- mannanases from different Bacilli species are also found in both families (Sygusch et al., 1998; Hatada et al., 2005). Some glycoside hydrolases are multifunctional enzymes that contain catalytic domains that belong to different GH families like Paenibacillus polymyxa. A cel44C-man26A gene was cloned from this endophytic strain. This gene encodes for a protein that contains a glycoside hydrolase family 44 (GH44) catalytic domain and a glycoside hydrolase family 26 (GH26) catalytic domain. The multifunctional enzyme domain GH44 possesses cellulase, xylanase, and lichenase activities, while the enzyme domain GH26 possesses mannanase activity (Han et al., 2006).
TABLE 3 Overview of various production conditions (molecular weight, optimum pH, optimum temperature, carbon source, and pl values) of mannanase producing microorganisms
Most mannanases often display a modular organization and usually consist of two-domain proteins (Henrissat etal., 1995; Perret et al., 2004). These proteins generally contain structurally discrete catalytic and non-catalytic modules (Warren et al., 1996).
beta-mannanases in both families belong to the 3-D structure group (beta/alpha)z fold catalytic module characteristic of clan Glycoside hydrolase-A (Stoll et al., 2005). The most important non- catalytic module consist of carbohydrate binding module (CBM) which facilitate the targeting of the enzyme to the polysaccharide, By analogy to the glycoside hydrolases, CBMs are classified into families based on sequence similarities and three dimensional structures (Boraston et al., 2004). Presently, 49 families of CBMs have been reported and a number of these families are closely related and consequently grouped into superfamilies or clans. A comprehensive classification of mannanases and other hydrolases is available at http://afmb.cnrs-mrs.fr/CAZY. Amongst beta-mannanases from aerobic fungi, the enzymes from T. reesei and A. bisporus are composed of a family 5 catalytic module linked to a family 1 CBM (Tang et al., 2001; Yague etal., 1997). Some bacterial beta- mannanases from families 5 and 26 have more complex structures: the C. fimi beta-mannanase Man26A contains both a mannan binding family 23 CBM, a putative SLH-module and a module of unknown function (Stoll et al., 1999, 2000; Stalbrand et al., 2006). The family 5 beta-mannanase from Thermoanaerobacteriumpolysaccharolyticum also contain a SLH-module and, in addition, two internal family 16 CBMs (Cann et al., 1999). Several beta-mannanases from anaerobic bacteria and fungi contain dockerin modules, which attach the beta- mannanases either to the microbial cell surface or to multienzyme complexes such as the cellulosome (Shallom and Shoham, 2003; Halstead et al., 1999; Tamaru and Doi, 2000; Perret et al., 2004). The family 26 beta-mannanases from Caldocellulosiruptor saccharolyticus contains two family 27 mannan-binding modules (Sunna et al., 2001) and from Paenibacillus pofymyxa contain a fibronectin domain type 3, CBM family 3 in addition to two catalytic domains (Han et al., 2006). A comparison of the modular structure of different mannanases reveals CBMs have been predicted for several beta-mannanases from family 5 and family 26. However, only in a few cases has the binding been experimentally confirmed. In addition to Man5A, family 1 CBMs have been indicated in die fungal beta- mannanases from Agaricus bisporus (Tang et al., 2001; Yague et al., 1997). However, the beta-mannanases from Aspergillus aculeatus and Aspergillus niger appear to lack CBMs (Ademark et al., 1998; Christgau et al., 1994). Some other family 5 beta-mannanases have CBMs that have been classified into family 3, 16 and family 35. Moreover, some family 26 beta-mannanases have family 23, 27 and 35 CBMs, which have been shown to bind to mannan (Stoll et al., 2000; Sunna et al., 2001). However, many beta-mannanases from both families 5 and 26 appear to be sole catalytic modules. Thus, at present, it is difficult to establish any obvious patterns in modularity among beta-mannanases. It clearly appears as if several different strategies in mannan-hydrolysis subsist. Henrissat et al. (1998) proposed a scheme for designating enzymes that hydrolyze the polysaccharides in cell walls of plants. A written designation of the module organization of recendy studied beta-mannanases is depicted in Table 4. CLONING AND EXPRESSION OF THE MANNANASE GENE
Gene cloning is a rapidly progressing technology that has been instrumental in improving our understanding of the structure- function relationship of genetic systems. It provides an excellent method for the manipulation and control of genes. More than 50% of the industrially important enzymes are now produced from genetically engineered microorganisms. Recombinant DNA techniques offer a plethora of opportunities for construction of genetically modified microbial strains with selected enzyme machinery. To ensure the commercial utilization of hemicellulosic residues in paper and pulp industries, the production of higher mannanase yields at low capital cost is required. Microbial genes encoding proteins associated with sugar metabolism are clustered which indicates that their expression is coordinated. The sequence of some of the genes encoding beta-1,4- mannanases have been determined and analyzed to deduce the primary structure of mannanase enzymes. In this respect, isolation and cloning of the mannanse gene represents an essential step in the engineering of the most efficient microorganism. Several reports have been published in the past decade (Table 4) on the isolation and manipulation of microbial mannanase genes with the aim of enzyme overproduction, studying the primary structure of the protein, its role in the secreting microorganism, and protein engineering to locate the activesite residues and/or to alter the enzyme properties to suit its commercial applications. Henrissat (1990) recognized two families encoding beta-mannanases on the basis of amino acid sequence similarities. Only two beta-mannanases genes were available for comparison at the time his enzyme classification was made. ManA from Caldicellulosiruptor saccharolyticus (Luthi et al., 1991) and ManA from a Bacillus sp. (Akino et al., 1989) were compared with 301 glycosyl hydrolases and classified respectively as type 5 and type 26 glycosyl hydrolases. A number of mannanase genes had been sequenced since the classification of mannanases by Henrissat (1990) and all share homology with either manA from Caldicellulosiruptor saccharolyticus or the Bacillus mannanase gene (Gibbs et al., 1999). Attempts have been made to clone and express mannanase from bacteria such as Bacillus stearothermophilus, Caldibacillus cellulovorans, Caldicellulosiruptor Rt8B, Caldocellum saccharolyticum (Sygusch et al., 1998; Morris et al., 1995; Sunna et al., 2000) into a non mannanase producing strain of Escherichia coli. As alkaline beta- mannanases provide obvious advantage for the application in the manufacture of kraft pulp and in the detergent industry, where high pH processes are common. Few reports describe the molecular cloning, sequencing and expression of mannanase genes from alkaliphilic Bacillus sp. (Takeda et al., 2004a; Takeda et al., 2004b; Hatada et al., 2005), Bacillus subtilis B36 (Li et al., 2006), Bacillus N16-5 (Yanhe Ma et al., 2004), Bacillus agaradhaerens (Bettiol and Showell, 2002), Bacillus sp. 1633 (Kauppinen et al., 2003) and Thermoanaerobacterium polysaccbarolyticum (Cann et al., 1999). The sizes of mannanase genes and their protein products are highly divergent (Table 4). This indicates that mannanases form a group of proteins that possess similar enzyme activities even though they have highly different primary protein structure. Amino acid sequence alignments of members of family 5 rarely reveal levels of sequence identity greater than 20%, As the amino acid sequence of members of family 5 were divergent, members of family 5 have been classified in eight subfamilies, subfamilies A1-A8 (Hilge et al., 1998). Subfamilies A7 (eukaryotic mannanases) and A8 (bacterial mannanases) have recendy been included. Members of subfamily A8 exhibit levels of sequence identity greater than 43%, while the levels of sequence identity for members of subfamily A7 and members of subfamily A8 are less than 20%. Three dimensional structures of two /5-1,4- mannanases from family 5 in complex with oligosaccharide products have been determined by Xray crystallography (Hilge etal, 1998; Sabini etal., 2000). Recendy, the preliminary structure determination of a bacterial family 5 /3-l,4-mannanase (Akita et al., 2004) has been reported. Eight conserved residues of all family 5 enzymes were demonstrated to be responsible for the catalytic activity of the enzyme including two catalytic glutamates (Sakon etal., 1996).
Yanhe et al. (2004) reported the eight essential conserved active site residues of /3-mannanase in strain Bacillus N16-5 as Arg-83, His-119, Asn-157, Glu-158, His-224, Tyr-226, Glu-254, and Trp-283. Computer analysis of the deduced amino acid sequence of ManA from Bacillus sp. N16-5 with sequences in the GenBank and UniProt databases showed that ManA consists of a single catalytic domain. The amino acid sequence of ManA showed high homology to family 5 mannanases: 96.8% sequence similarity to ManA of B. agaradhaerens (GenBank accession number: AAN27517), 58.2% to ManG of B. circulans (BAA25878), 52.3% to ManA of Bacillus sp. I633 (AAQ31835), 39.4% to ManA of Thermobifida fusca (CAA06924), 34.4% to ManA of Vibrio sp. MA-138 (BAA25188), and 33.1% to ManA of Streptomyces lividans 66 (S30386).
TABLE 4 Occurrence of the main mannanase genes and their products
Yanhe et al. (2004) compared the sequences of mannanases from Bacillus sp. N16-5 and Bacillus sp. AM001, the only two mannanases extensively characterized among alkaline mannanases reported to date. A higher level of similarity (48%) was observed for the C terminal region (about 150 amino acid residues long) of these two enzymes. Hatada et al. (2005) reported a novel alkaline mannanase Man26A in the culture of an alkaliphilic Bacillus sp. strain JAMB- 750. The N-terminal half (Glu27-Val486) of the enzyme exhibited moderate similarities to other mannanases belonging to glycoside hydrolase family 26, such as the enzymes from Cellvibrio japonicus (37% identity), Cellulomonas fimi (33% identity), and Bacillus sp. strain AM-001 (28% identity).
Another family 26 beta-mannanase Man A of D. thermophilum Rt46B.1 on alignment with related enzymes shows that it shares considerable sequence similarity (58% sequence identity) with the catalytic domain of ManA from Caldicellulosiruptor sp. strain Rt8B.4 (Gibbs et al., 1996), and at least 25% sequence identity with the other family 26 mannanases listed in Table 2. Matsushita et al. (1991) published a sequence coding for a beta-1,4-endoglucanase from Bacteroides ruminicola. However, comparison with the mannanase domains from D. thermophilum Rt46B.1 and Caldicellulosiruptor Rt8B.4 ManA suggests that the B. ruminicola enzyme is a multidomain enzyme possessing both a mannanase and an endoglucanase domain. Currently, the reported three-dimensional structures for GH26 members are for the Cellvibrio japonicus mannanase CjMan26A, and for the Cellulomonas fimi mannanase CfMan26A. These enzymes possess a conserved motif in which two glutamates have been identified as the putative catalytic residues; at least two additional residues are conserved (Bolam et al., 1996; Le Nours et al., 2005). Notably, the location of the catalytic apparatus is completely conserved, with the catalytic residues positioned on strands beta-4 and beta-7 (Jenkins et al., 1995).
Amongst the fungal mannanases, The T. reesei man 1 gene was the first fungal mannanase gene characterized (Stalbrand et al., 1995). Tang et al. (2001) demonstrated that the catalytic domain of CEL4 from Agaricus bisporus had the most amino acid sequence similarity with Ascomycete mannanases from Aspergillus aculeatus and Trichoderma reesei, which belong to glycosyl hydrolase family 5 (43 and 42%, respectively). The availability of a number of family 26 mannanase sequences provides the opportunity for the design of PCR primers to highly conserved regions. It has allowed the identification of strains that carry mannanase genes from this family as well as providing the necessary sequence information for the isolation of the entire genes by genomic walking. These consensus primers have been successfully tested against the genes known to be present in genomic DNA from D. thermophilum Rt46B.1 and Caldicellulosiruptor Rt8B.4 and have revealed several other family 26 mannanases present in culturable and unculturable hemicellulolytic thermophilic bacteria (Bergquist et al., 1996). Further sequence information on family 5 mannanases is required before consensus primers can be designed to allow the identification and isolation of these genes.
There are many applications for mannanases in the industrial processes. The worldwide requirement for enzymes for individual applications varies considerably. Mannanases are used mainly for improving the quality of food, feed and aiding in enzymatic bleaching of softwood pulps in the paper and pulp industries. There is a paucity of knowledge about the roles that govern the diverse specificity of these enzymes. Deciphering these secrets would enable us to exploit mannanases for their applications in biotechnology. The following section will discuss details of some of the most promising applications of mannanases (Howard et al., 2003; Galbe et al., 2002; Comfort et al., 2004).
Paper and Pulp Industry
The most potent application of mannanase is consistent with its potential use in enzymatic bleaching of softwood pulps. The extraction of lignin from wood fibers is an essential step in bleaching of dissolving pulps. Pulp pretreatment under alkaline conditions hydrolyzes hemicelluloses covalently bound to lignin and thus facilitates subsequent removal of lignin. There is a drawback to alkaline treatment of wood pulps, however, in that it creates an environmental pollution problem (Hongpattarakere, 2002). The alternate use of mannanases equally facilitates lignin removal in pulp bleaching and yields results comparable to alkaline pretreatment (Cuevas et al., 1996). Consequently, to substitute within a pulp-bleaching sequence an enzymatic pretreatment for the ultrahot alkaline extraction stage (Clarke et al., 2000) offers the possibility of significant reduction in environmental pollution and thus is of considerable interest to the pulp and paper industry. To be feasible, however, enzymatic bleaching requires that hemicellulase treatment not impair pulp quality by attacking cellulose fibers. Softwoods from which the majority of pulps are derived contain as much as 15 to 20% hemicellulose in the form of galactomannan (Suurnakki et al., 1996). Mannanases having substrate specificities for galactomannan constituents would make excellent candidates for use in enzymatic bleaching of softwood pulps (Gubitz et al., 1997). Moreover, pulping is best carried out at elevated temperatures, thermophilic mannanases could offer significant advantages over mesophilic mannanases in terms of their higher intrinsic stability and catalytic efficiencies at such elevated temperatures (Yanhe et al., 2004). Mannanase is useful in chlorine- free bleaching processes for paper pulp (chemical pulps, semichemical pulps, mechanical pulps or kraft pulps) in order to increase the brightness, thus decreasing or eliminating the need for hydrogen peroxide in the bleaching process (Tenkanen et al., 1997).
Hydrolysis of Coffee Extract
Different mannanase preparations are used for the hydrolysis of coffee mannan, thus reducing significantly the viscosity of coffee extracts. Mannan is the main polysaccharide component of these extracts and is responsible for their high viscosity, which negatively affects the technological processing of instant coffee. Mannanase may also be used for hydrolyzing galactomannans present in a liquid coffee extract, preferably in order to inhibit gel formation during freeze drying of the (instant) coffee. Coffee mannan is isolated from green defatted beans by delignification, acid wash and subsequent alkali extraction with a yield of 12.8%. Additionally, coffee extract polysaccharides are separated by alcohol precipitation and are found to form nearly half of the coffee extract dry weight. These isolated mannans as well as the mannans in the coffee extract are efficiently hydrolyzed by the mannanase, which resulted in significant viscosity reductions (Sachslehner et al., 2000; Nicolas et al., 1998). Concurrently, the reducing sugar content increased continuously due to the release of various mannooligosaccharides including mannotetraose, mannotriose, and mannobiose. Both a partially purified, immobilized and a soluble, crude mannanase preparation are successfully employed for the degradation of coffee mannan (Nunes et al., 1998, 2006).
Use in Detergent Industry
Application of carbohydrases in detergents is well known. Amylases and cellulases being the most common enzymes. Recently, alkaline mannanases stable to constituents of detergents have found application in certain laundry segments as stain removal boosters. Mannanases hydrolyze mannan containing materials like gums (galactomannans, glucomannans and guar gum). These gums are used worldwide as a thickener or stabiliser in many types of household products and foods including ice-creams, BBQ sauces, hair styling gels, shampoos, conditioners and toothpaste (Wong and Saddler, 1992). Stains containing mannan are generally difficult to remove because mannans have a tendency to adsorb to cellulose fibers and therefore bind to cotton textiles. Further, mannan also has a “glue effect.” This gluing effect means that particulate soils released during the wash cycle or from subsequent wear can bind to invisible residual mannan. In other words, not only many mannan stains reappear, but mannan can also be transferred to otherwise clean fabrics during washing and result in fabric greying. Mannanases cleave the beta-1,4-linkages of mannans through hydrolysis, thus breaking down the gum polymer into smaller carbohydrate fragments. These smaller, more water-soluble polysaccharide fragments remain free from the fabric and are siphoned out of the wash. Mannanases can thus prevent redeposition of soil released during washing.
The cleaning compositions must contain at least one additional detergent component. These compositions can also be formulated as sanitization products, contact lens cleansers, dishwashing, hard surface cleaner, and health and beauty care products (Bettiol et al., 2000). Such additive products are intended to supplement the performance of conventional compositions and can be added at any stage of the cleaning process (McCoy, 2001). Treatment with cleaning or detergent compositions comprising the mannanase can improve whiteness as well as prevent binding of certain soils to the cellulosic material. Accordingly, mannanase are used in cleaning compositions, including laundry, personal cleansing and oral/dental compositions, Such cleaning compositions comprising a mannanase and an enzyme selected from cellulases, proteases, lipases, amylases, pectin degrading enzymes and xyloglucanases, are prepared to provide superior cleaning performance.
To Improve the Nutritional Value of Poultry Feeds
Reduction in weight gain and feed conversion efficiency in poultry birds has been associated with intestinal viscosity due to different feeds. beta-Mannan is a polysaccharide commonly found in feed ingredients such as soybean meal (SBM), palm kernel meal (PKM), copra meal (CM), guar gum meal and sesame meal. There is almost universal use of soybean meal and full fat soy as protein sources in poultry feeds, PKM and CM as ruminant feed (Moss and Givens, 1994; Chandrasekariah et al., 2001) pig diets (Pettey et al., 2002) and rabbit diets (Aganga et al., 1991). All these meals have some common properties: high fiber content, low palatability, lack of several essential amino acids and high viscosity coupled with several anti- nutritional properties such as mannan, galactomannan, xylan and arabinoxylan, their utilization in the intestine is very limited. beta-Mannans have been found to be highly deleterious to animal performance, severely compromising weight gain and feed conversion as well as glucose and water absorption. Incorporation of beta- mannanase into these diets results in decreased intestinal viscosity, thus improving both the weight gain of chicks and their feed conversion efficiency. More recently, the beneficial effect of enzymatic degradation of beta-mannan by addition of beta-mannanase to diets containing SBM has been documented in broilers (Jackson et al., 2004; Daskiran et al., 2004; Lee et al., 2003), layers (Wu et al., 2005), turkeys (Odetallah et al., 2002) and swine (Pettey et al., 2002). Using endo-mannanase alone may only produce a small proportion of mannose and thus only a small amount of mannan is likely to be absorbed in the intestine of broilers (Saki et al., 2005). A combination of endo-mannanase from different strains may have a complementary ability to cleave sugars, the use of a combination of endo-mannanase from different strains of organisms may be helpful and that is more possible in the poultry industry. A study of endo-mannanases has been done by Tamaru et al. (1997), who found that endo-mannanase from Vibrio sp. could not hydrolyse mannotriose but produced mannotriose, while endo-mannanase from Streptomyces could hydrolyse mannotriose to form mannose and mannobiose (Kusakabe and Takashi, 1988). Due to the fact that most of mannanase in the market is in form of endo beta-mannanase (e.g. gamanase and hemicell mannanase), it can be speculated that manno- oligosaccharides, mannotriose and mannobiose as well as a small amount of mannose are generated when this type of enzyme is included in the diet. Since only mannose can be absorbed in the intestine, the production of mannobiose and mannooligosaccharides are, to some extent, useless. So these components of carbohydrates may not be absorbed and are therefore unable to supply energy to the host. However, in the sense of the health of the poultry, the production of mannooligosaccharides can improve a chicken’s health (Laere et al., 1999), either by increasing the population of specific bacteria such as Bifidiobacteria because the supply of these types of carbohydrates are a source of feed for bacteria in the caeca and thus suppressing the pathogenic ones, or by flushing out the pathogenic bacteria which attach to the mannooligosaccharides. In fact, mannooligosaccharides are added to the poultry diets for this effect (Lyons and Jacques, 2002). The use of mannanase in poultry research has been widespread and proven to be successful (Sundu et al., 2006). Use in Oil Drilling
Mannanases can also be used to enhance the flow of oil or gas in drilling operations. Practically, product flow to the well bore is stimulated by forcing out open crevices in the surrounding bedrock, which is done by flooding the well with a natural polymer (guar gum) solution and sand particles, capping the well and then pressurizing the bedrock until it fractures. The viscous polymer solution carries the sand through the fractures, propping open cracks for oil and gas flow. To facilitate product flow, the polymer solution is thinned (Adams et al., 1995; Christoffersen, 2004). As mannanases are able to hydrolyze guar gum at elevated temperatures (Politz etal., 2000), so implementation of oil production by these enzymes could be feasible.
In oil Extraction of Coconut Meats
Mannanases can be used in enzymatic oil extraction of coconut meat. The main components of the structural cell wall of coconut meat are mannan and galactomannan (Saittagaroon et al., 1983). A high oil yield is achieved in this process. In the traditional method of oil extraction by expeller, unsanitary handling, drastic processing of copra for oil extraction, refining of the product, the presence of polycylic aromatic hydrocarbons (PAH) retained in the coconut oil not only lessen the quality of the by-product like protein but also render the product susceptible to aflatoxin contamination and oxidative rancidity. The enzymatic oil extraction process not only eliminates such problems but also improves the sensory qualities of the products. In addition, the improvement of the quality of coconut oil minimizes the need for refining the oil as it is already comparable to the semi-refined coconut oil. This enzymatic process gives three valuable products: coconut oil, an aromatic protein containing liquid that may be used as a beverage base and a coconut protein isolate (Chen et al., 2003).
In the Textile and Cellulosic Fiber Processing Industries
The mannanase can be used for preparation of fibers or for cleaning of fibers in combination with detergents in the textile and cellulosic processing industry. The processing of cellulosic material for the textile industry into a material ready for garment manufacture involves several steps: spinning of the fiber into a yarn, construction of woven or knit fabric from the yarn and subsequent preparation, dyeing and finishing operations. Woven goods are constructed by weaving a filling yam between a series of warp yarns; the yarns could be of two different types. Mannanase is useful in an enzymatic scouring process and in desizing (removal of mannan size) during the preparation of cellulosic material for proper response in subsequent dyeing operations (Palackal et al., 2006).
For Degradation of Thickening Agents
Galactomannans such as guar gum and locust bean gum are widely used as thickening agents, e.g. in food and print paste for textile printing such as prints on T-shirts. Mannanase can be used for reducing the viscosity of residual food in processing equipment and thereby facilitate cleaning after processing. The enzyme or enzyme preparation is useful for reducing viscosity of print paste, thereby facilitating wash out of surplus print paste after textile printings (Ademark et al., 1998).
As Non-Nutritional Food Additives
Mannanases also contribute to the human health as they degrade mannans, which otherwise are resistant to mammalian digestive enzymes in the small intestine but are readily fermented in the large intestine, particularly by probiotic bacteria belonging to the genera Bifidobacteria and Lactobacillus. Prebiotic oligosaccharides including mannooligosaccharides, i.e. hydrolysis products of mannan degradation, are believed to promote the selective growth and proliferation of human beneficial intestinal microflora (Kobayashi et al., 1984, 1987).
Thus, mannanases only recendy attracted increased scientific and commercial attention due to potential applications in several industries. A few commercial products have been launched successfully worldwide in the past few years (Table 5).
The benefit of employing novel enzymes for specific industrial processes is well recognized with the discovery of beta-mannanases. beta-Mannanases (18.104.22.168) hydrolyze mannan based hemicelluloses and liberate short beta-1,4 manno-oligomers, which can be further hydrolyzed to mannose by beta-mannosidases (EC 22.214.171.124). There are currently about 50 beta-mannanase gene sequences in GH families 5 and 26. The increasing number of new microbial genomes is revealing new mannolytic systems. Major challenges in this field include the design of efficient enzyme system for commercial applications. Mannanases occur ubiquitously in animals, plants, and microbes. However, microbes are most potent producers of mannanases and represent the preferred source of enzymes in view of their rapid growth, limited space required for cultivation, and ready accessibility to genetic manipulation. Microbial mannanases have been used recently in the food, feed and detergent industries. Advances in genetic manipulation of microorganisms have opened new possibilities for the introduction of predesigned changes, resulting in the production of tailor-made mannanases with novel and desirable properties. The development of recombinant mannanases and their commercialization by P&G, ChemGen and Genencor is an excellent example of the successful application of modem biology to biotechnology.
TABLE 5 List of commercial mannanases and their suppliers
Industrial applications of mannanases have posed several problems and challenges for their further improvements. The biodiversity represents an invaluable resource for biotechnological innovations and plays an important role in the search for improved strains of microorganisms used in the industry. A recent trend has involved conducting industrial reactions with enzymes reaped from exotic microorganisms that inhabit hot waters, freezing Arctic waters, saline waters, or extremely acidic or alkaline habitats. The mannanases isolated from extremophilic organisms are likely to mimic some of the unnatural properties of the enzymes that are desirable for their commercial applications. Exploitation of biodiversity to provide microorganisms that produce mannanases well suited for their diverse applications is considered to be one of the most promising future alternatives. The existing knowledge about the structure- function relationship of mannanases, coupled with novel techniques, promises a fair chance of success, in the near future, in evolving mannanases that were never made in nature and that would meet the requirements of the multitude of mannanase applications.
Adams, M. W., and Kelly, R. M. 1995. Enzymes from microorganisms in extreme environment. Chem. Eng. News 18: 32-42.
Adernark, P., Varga, A., Medve, J., Harjunpaa, V., Drakenberg, T., Tjerneld, F., and Stalbrand, H. 1998. Softwood hemicellulose- degrading enzymes from Aspergillus niger. purification and properties of a beta-mannanase. J. Biotechnol. 63: 199-210.
Aganga, A. A., Aduku, A. O., Abdulmalik, M., and Sekoni, A. 1991. The effect of different protein sources and their levels on the production of breeding rabbits. J. Appl. Rabbit Research 14: 30-33.
Akino, T., Kato, C., and Horikoshi, K. 1989. The cloned beta- mannanase gene from alkalophilic Bacillus sp. AM-001 produces two beta-mannanases in Escherichia coli. Arch. Microbiol. 152: 10-15.
Akita, M., Takeda, N., Hirasawa, K., Sakai, H., Kawamoto, M., Yamamoto, M., Grant, W. D., Hatada, Y., Ito, S., and Horikoshi, K. 2004. Crystallization and preliminary X-ray study of alkaline mannanase from an alkaliphilic Bacillus isolate. Acta Crystallogr. 60: 1490-1492.
Araujo, A., and Ward, O. P. 1990. Hemicellulases of Bacillus species: preliminary comparative studies on production and properties of mannanases and galactanases. J. Appl. Bacteriol. 68: 253-261.
Arcand, N., Kluepfel, D., Paradis, F. W., Morosoli, R., and Sharek, F. 1993. beta-Mannanase of Streptomyces lividans 66: cloning and DNA sequencing of the manA gene and characterization of the enzyme. J. Biochem. 290: 857-863.
Beauchemin, K. A., Morgavi, D. P., Mcallister, T. A., Yang, W. Z., and Rode, L. M. 2001. The use of enzymes in ruminant diets. pp 296-322. In Wiseman, J., Garnsworthy, P.C. (eds.), Recent Advances in Animal Nutrition. Nottingham University Press, Nottingham, UK.
Bettiol, J. L. P., Boutique, J. P., Gualco, L. M. P., and Johnston, J. P. 2000. Nonaqueous liquid detergent compositions comprising a borate releasing compound and a mannanase. EP1059351.
Bettiol, J. P., and Showell, M. S. 2002. Detergent compositions comprising a mannanase and a protease. US Patent 6376445.
Bhat, M. K. 2000. Cellulases and related enzymes in biotechnology. Biotechnol. Adv. 18: 355-383. Bicho, P. A., Clark, T. A., Mackie, K., Morgan, H. W., and Daniel, R. M. 1991. The characterisation of a thermostable endo-beta-mannanase cloned from Caldocellum saccharolyticum. Appl. Microbiol. Biotechnol. 36: 337- 343.
Bolam, D. N., Hughes, N., Virden, R., Lakey, J. H., Hazlewood, G. P., Henrissat, B., Braithwaite, K. L., and Gilbert, H. J. 1996. Mannanase A from Pseudomonas fluorescens sp. Cellulosa is a retaining glycosyl hydrolase in which E212 and E230 are the putative catalytic residues. Biochem. 35: 16195-16204.
Boraston, A. B., Bolam, D. N., Gilbert, H. J., and G. J. Davies. 2004. Carbohydrate-binding modules: fine tuning polysaccharide recognition. Biochem. J. 382: 769-781.
Braithwaite, K. L., Black, G. W., Hazlewood, G. P., Ali, B. R., Gilbert, and H. J. 1995. A non-modular endo-beta-1,4-mannanase from Pseudomonas fluorescens subspecies cellulosa. Biochem J. 305: 1005- 1010.
Buckeridge, M. S., Pessoa, dos Santos, H., and Tine, M. A. S. 2000. Mobilisation of storage cell wall polysaccharides in seeds. Plant Physiol. Biochem. 38: 141-156.
Cann, I. K., Kocherginskaya, S., King, M. R., White, B. A., and Mackie, R. I. 1999. Molecular cloning, sequencing, and expression of a novel multidomain mannanase gene from Thermoanaerobacterium polysaccharolyticum. J. Bacteriol. 181: 1643-1651.
Capoe, P., Kubackova, M., Alfoldi, J., et al. 2000. Galactoglucomannan from the secondary cell wall of Picea abies.L.Krast. Carbohydr. Res. 329: 635-645.
Chandrasekariah, M., Sampath, K. T., Thulasi, A., and Anandan, S. 2001. Insitu protein degradability of certain feedstuffs in cattle. Ind. J. Ani. Sci. 71: 261-264.
Chang, T. T., Yang, X. Y., and Ko, W. H. 1992. A sensitive method for detecting producion of extracellular enzymes by fungi on solid media. Mycologia. 84: 923-925.
Chanzy, H., Dube, M., Marchessault, R. H., et al. 2004. Single crystals and oriented crystallization of ivory nut mannan. Biopoly. 18: 887-898.
Charrier, M., and Rouland, C. 2001. Mannan-degrading enzymes purified from the crop of the brown garden snail Helix aspersa Muller (Gastropoda Pulmonata). J. Expt. Zool. 290: 125-135.
Chen, B. K., and Diosady, L. L. 2003. Enzymatic aqueous processing of coconuts. Int. J. Appl. Sci. Engg. 1: 55-61.
Chen, X., Cao, Y., Ding, Y., Lu, W., and Li, D. 2007. Cloning, functional expression and characterization of Aspergillus sulphureus beta-mannanase in Pichia pastoris. J. Biotechnol. 128(3): 452-461.
Chen, Y., Long, J., Liao, L., et al. 2000. Study on the production of betamannanase by Bacillus M50. Wei Sheng Wu Xue Bao. 40: 62-68.
Chhabra, S. R., Shockley, K. R., Ward, D. E., and Kelly, R. M. 2002. Regulation of endo-acting glycosyl hydrolases in the hyperthermophilic bacterium Thermotoga maritima grown on glucan and mannan based polysaccharides. Appl. Environ. Microbiol. 68: 545- 554.
Christgau, S., Kauppinen, S., Vind, J., Kofod, L. V., and Dalboge, H. 1994. Expression, cloning, purification and characterization of a beta-1,4-mannanase from Aspergillus aculeatus. Biochem. Mol. Biol. Int. 33: 917-925.
Christoffersen, L. 2004. Products and applications that promote sustainability. In REBIO 2004 (DIVERSA), Boca Chica, Dominican Republic.
Civas, A., Eberhard, R., le Dizet, P., and Petek, F. 1984. Glycosidases induced in Aspergillus tamarii secreted alpha-D- galactosidase and beta-D-mannanase. Biochem. J. 219: 857-863.
Clarke, J. H., Davidson, K., Rixon, J. E., Halstead, J. R., Fransen, M. P., and Gilbert, H. J. 2000. A comparison of enzyme aided bleaching of softwood paper pulp using combinations of xylanase, mannanase and alpha-galactosidase. Appl. Microbiol. Biotechnol. 53: 661-667.
Comfort, D. A., Chhabra, S. R., Conners, S. B., Chou, C.-J., Epting, K. L., Johnson, M.R., Jones, K.L., Sehgal, A.C., and Kelly, R.M. 2004. Strategic biocatalysis with hyperthermophilic enzymes. Green Chemistry 6: 459-465.
Cuevas, W. A., Kantelinen, A., Tanner, P., Bodie, B., and Leskinen, S. 1996. Purification and characterization of novel mannanases used in pulp bleaching, pp. 123-126. In: Srebotnik E., Messner K., Srebotnik, E., Messner, K., eds. Biotechnology in the Pulp and Paper Industry. Vienna, Austria: Facultas- Universitatsverlag.
Cui, F., Shi, J., and Lu, Z. 1999. Production of neutral beta- mannanase by Bacillus subtilis and its properties. Wei Sheng Wu Xue Bao. 39(1): 60-63.
Daskiran, M., Teeter, R. G., Fodge, D., and Hsiao, H. Y. 2004. An evaluation of endo-beta-D-mannanase (Hemicell) effects on broiler performance and energy use in diets varying in beta-mannan content. Poultry Sci. 83: 662-668.
Davies, G., Sinnott, M. L., and Withers, S. G. 1998. Glycosyl Transfer. Comp. Biol. Catal. 1: 119-209.
Dekker, R. F. H. 1985. Biodegradation of hemicelluloses. pp. 505- 533. In J. Higuchi (ed.), Biosynthesis and Biodegradation of Wood Components, Academic Press Inc., Orlando, FL.
Dekker, R. F. H., and Richards, G. N. 1976. Hemicelluases: their occurrence, purification, properties and mode of actions. Adv. Carbohydr. Chem. Biochem. 32: 277-352.
De-Nicolas-Santiago, S., Regalado-Gonzalez, C., Garcia- Almendarez, B., Fernandez, F. J., Tellez-Jurado, A., and Huerta- Ochoa, S. 2006. Physiological, morphological, and mannanase production studies on Aspergillus niger uam-gs1 mutants. Electron. J. Biotechnol. 9 Epub Jan. 2006.
Dingle, J., Reid, W. W., and Solamons, G. L. 1953. The enzymatic degradation of Pectin and other polysaccahrides II, Application of the “Cup Plate” Assay to the estimation of enzymes. J. Sci. Food Agric. 4: 149.
Dong, X. Z., Schyns, P. J., and Stams, A. J. 1991. Degradation of galactomannan by a Clostridium butyricum strain. Antonie Van Leeuwenhoek. 60: 109-114.
Downie, B., Hilhorst, W. M., and Bewley, J. D. 1994. A new assay for quantifying endo-beta-mannanase activity using congo red dye. Phytochemistry 36: 829-835.
Duffaud, G. D., McCutchen, C. M., Leduc, P., Parker, K. N., and Kelly, R. M. 1997. Purification and characterization of extremely ther