Keywords

Introduction

Enzymes or biocatalysts are exclusively synthesized by the living cell and no life can exist without enzymes. They are basically proteins and catalyze certain chemical reactions involving naturally occurring organic materials such as carbohydrates, protein, fats. Enzymes are important because of their extraordinary specificity and catalytic power which are greater than those of artificial catalysts. Tannin acyl hydrolase (E.C.3.1.1.20) universally known as tannase is an inducible enzyme that has been widely utilized in the biotransformation of hydrolysable tannins to simple phenolic molecules like gallic acid. Tannases precisely act upon ester and depside linkages in hydrolysable tannins particularly the gallotannins . Filamentous fungi have been documented as the prominent tannase producers amongst all tannase-producing microorganisms.

History

Van Teighem (1867) accidentally explored tannase enzyme and he reported gallic acid production after treating an aqueous solution of tannins with two fungal species. He was the pioneer who revealed that fungal activity is responsible for gallic acid production. Loraque documented the gallic acid production from tannic acid is either due an organism’s action or due to the oxidation process. In this context, he further reported numerous toxic substances inhibiting the gallic acid production from tannic acid in gallnut (Knudson 1913). Fenbach in 1901 cultivated Aspergillus niger in Raulin’s solution that consisted of tannic acid as a carbon source in place of sugar thereafter isolating tannase from the cultured organism (Knudson 1913).

Libuchi and his coworkers in (1967) proposed a spectrophotometric assay for the quantitation of tannase activity. A vast range of assay methods is available for estimation of tannase activity. Presently, two methods are predominantly used by researchers to evaluate the tannase activity. The first method as described by Mondal et al. (2001) relies on the Hydrolysis of tannic acid and measuring the leftover tannic acid using tannic acid standard curve. However, the second method as described by Sharma et al. (2000) relies on measurement of gallic acid given off post-enzymatic reaction using the gallic acid standard curve. As a matter of fact, apart from filamentous fungi tannase production by bacteria was also documented (Deschamps et al. 1983).

Tannins: Natural Substrates for Tannase

Tannins belong to phenolics are one of the major class of plant secondary metabolites that serve as a defense machinery against attack by herbivores. These phenolics are synthesized in plants as a counter response to invasion by microbes, herbivores, pathogens, cold, and UV light as well as nutrient limitation. Tannins universally exist in angiosperms, gymnosperms as well as pteridophytes. Tannins predominantly accumulate in plant parts such as bark, roots, fruits, and leaves (Scalbert 1991). Frutos et al. (2004) documented the abundance of tannins in plant leaves and flowers.

Occurrence of Tannins

Belmares et al. (2004) documented the localization and distribution of tannins in various plant parts like flowers, needles, bark, seeds, leaves. Most common sources of hydrolysable tannins used universally in commercial sector are Turkish gall (Quercus infectoria), myrobalan nuts (Terminalia chebula), sumac (Rhus coriaria), Chinese gall (Rhus semialata), chestnut (Castanea sativa, Bhat et al. 1998), tara (Caesalpinia spinosa), chestnut gum arabic tree (Acacia nilotica, Lal et al. 2012), red gram (Cajanus cajan, Kuppusamy et al. 2015), and Cashew testa (Anacardium occidentales, Lokeshwari 2016). Hydrolysable tannins can be hydrolyzed to gallic acid and glucose molecule by through chemical treatment either by acidic hydrolysis or by alkaline treatment. However, in the past few years, enzymatic hydrolysis of tannins by tannases has attained ample importance. Tannins are principally localized in vacuoles or surface wax of plants. These localization sites keep tannins in their active state to counter the invasion by herbivores and pathogens. Only after cell breakdown and death, they can act and have metabolic effects. The concentration as well as their chemical attributes vary significantly amongst different plant species. Frutos et al. (2004) documented that tannin content in plants is boosted by factors such as water stress, poor soil quality, high temperature, extreme light intensity. Such variability in tannin concentration amongst different plant species is governed by several environmental parameters such as temperature, water availability, light, CO2 as well as nutrient availability and limitation. Long-lived trees have been documented to have higher tannin content in comparison to short-lived trees.

Importance of Tannins

Bhat et al. (1998) documented tannins as the most bountiful class of polyphenolics right behind lignins. Tannins have been reported as the products of plant secondary metabolism. The term tannin refers to tanna, an “Old High German” word for oak or fir tree. The term “tannin” may also refer to “tanning” or preservation of skins to create leather. Tannins have been reported to possess molecular weights ranging from 500 to 3000 kDa (Aguilar et al. 2007). Chemical basis of tannins defense mechanism has been credited to their inherent capability of precipitating the proteins. This tannin–protein complex was documented to render the microbial attack ineffective and simultaneously limiting the accessibility of metal ions obligatory for microbial metabolism (Scalbert 1991). The defense mechanism of tannin–protein complex was documented to be because of their acrid property and masculine sensation.

Classification of Tannins

Tannins have been categorized into three subgroups: hydrolysable tannins, condensed tannins, and complex tannins. Condensed tannins differ from hydrolysable tannins in not having any sugar molecule in their structure. Hydrolysable tannins are subdivided into two types: gallotannins and ellagitannins which are esterified to a sugar molecule most commonly the glucose molecule. Hydrolysable tannins can undergo easy hydrolysis upon acidic treatment or mild alkaline treatment as well as by treatment with hot water and more efficiently with enzymatic treatment.

Gallotannins are composed of gallic acid units esterified to a glucose molecule. Examples of gallotannins are Turkish galls (Quercus infectoria), Chinese gall (Rhus semilata) and sumac tannin (Rhus coriaria), Tara pods (Caesalpinia spinosa). Gallotannins represent the simplest form of hydrolysable tannins. Tannic acid has been documented as commercial form of gallotannins. Gallotannins give off gallic acid and glucose molecule upon hydrolysis. Gallotannins can undergo easy hydrolysis to yield gallic acid and glucose molecule upon acidic treatment or mild alkaline treatment—treatment with hot water and more effectively with enzymatic treatment by tannase.

Ellagitannins are composed of ellagic acid units bonded to glucosides. Molecules carrying a quinic acid core rather than glucose also represent ellagitannins . These cannot undergo easy hydrolysis because of their complex structure including C–C bonds.

Condensed tannins represent the compounds composed of building blocks of flavonoid units usually (from 2 over 50). They predominantly exist in woods and tree bark. Examples of condensed tannins include wattle (Acacia mollisima) tannin, quebracho (Schinopsis lorentzii) tannin and from tree bark (Bhat et al. 1998).

Complex tannins are an intermediate group that shares the features of both hydrolysable tannins as well as condensed tannins. They are composed of catechin or epicatechin units linked via glycosidic linkage to a gallotannin or an ellagitannin unit. Acutissimin A, Acutissimin B, Epicutissimin A, Mongolicain, Mongolicain A, and Mongolicain B are common examples of complex tannins. They give off catechin or epicatechin and gallic acid or ellagic acid on hydrolysis (Mingshu et al. 2006). The catechin tannins principally occur in tea leaves and tropical shrub legumes (Bhat et al. 1998).

Various sources of Tannase

Plants as a Tannase Source

Tannase has been documented in hydrolysable tannin-containing plants such as Turkish gall (Quercusinfectoria), sumac (Rhus coriaria), Chinese gall (Rhus semialata), tara (Caesalpinia spinosa), chestnut (Castanea sativa, Bhat et al. 1998), gum arabic tree (Acacia nilotica, Lal et al. 2012), red gram (Cajanus cajan, Kuppusamy et al. 2015) and waste testa (Anacardium occidentales, Lenin et al. 2015), divi divi (Caesalpinia coriaria) pods, English oak (Quercus robur), Pendunculate oak (Quercus rubra), Karee tree (Rhus typhina) leaves. Plants having condensed tannins are babul (Acacia arabica), konnam (Cassia fistula), avaram (Cassia auriculata) and others. Physiological worthiness of tannase in plants has been demonstrated in the synthesis of tannins. Plants synthesize gallic acid, hexahydroxyphenic acid, and chebulinic acid in addition to significant amount of sugar. These acids possibly undergo esterification with glucose molecule during the ripening process with the aid from tannase ultimately resulting in synthesis of tannins.

Animals as a Tannase Source

Animal sources of tannase include bovine intestine and ruminal mucus. Apart from this certain insects have been reported to produce tannase in their larval state. Several gastrointestinal bacteria of animal origin have also been documented to be the effective tannase producers. Many species of these bacteria have been from explored from feces of cows, humans, goats, etc. Tannase-producing bacteria Streptococcus pneumonia and Streptococcus bovis strains isolated from fecal samples of native sheep and goats which can hydrolyse acorn tannin in rumen and reduce negative effects of tannin on animals (Mosleh et al. 2014).

Microbial Sources

Microbial route for tannase production has gained worldwide importance over other sources since the microbial enzymes offer several advantages over other sources since the microbial enzymes are much more stable in comparison to similar enzymes from other sources. Furthermore, microorganisms possess the ability to produce higher titers of tannase. Microorganisms can undergo genetic modification, thus, they are easy to manipulate genetically. This property results in a significant uplift in the enzymatic activity. Most of the reported tannase-producing organisms are fungi, only a few bacteria, and yeast. Tannases vary in their degree of specificity as well as activity toward different tannin substrates. Fungal tannases have been documented to possess higher activity titer in the degradation of hydrolysable tannins. However, as a matter of fact, yeast tannases relatively disintegrate tannic acid easily and flaunt a relatively lesser affinity on the other hand in the degradation of natural tannins (Deschamps et al. 1983).

Bacteria

Several bacterial strains have been documented for their tannase production capability. Deschamps et al. (1983) were the pioneer to document tannase production from bacteria. The main tannase-producing genera among bacteria are: Bacillus (Belur et al. 2012), Lactobacillus (Rodriguez et al. 2008), Pesudomonas (Selwal et al. 2010), Erwinia carotovora (Sahira et al. 2015), Bacillus gottheilii M2S2 Subbulaxmi and Murty (2016).

Yeast

Over the years, couple of tannin degrading yeasts have been documented viz Candida sp., Debaromyces hansenii , Pichia adeyshi, P. monospora, and P. pseudopolymer. Aoki et al. (1976a, b) documented the tannin degradation by Candida spp., which possessed the capability to synthesize both extracellular as well as intracellular tannase by utilizing tannic acid as a substrate.

Fungi

Hadi et al. (1994) documented the potential of filamentous fungi in tannin degradation. Filmenteous fungi belonging to Aspergillus genus have been documented as one of the most potent tannse producers worldwide (Banerjee et al. 2001). Aspergillus spp. possess the capability of tannase production even in the scarcity of tannic acid. However, as a matter of fact, these fungi can tolerate high tannic acid concentration up to 20% without causing any negative effect on fungal growth as well as enzyme production. Fungi have an edge in having much pronounced growth and ease of separation from fermentation broth (Belmares et al. 2004). A few examples of tannase-producing fungi are: A. fumigatus (Manjit et al. 2008), R. oryzae (Mukherjee et al. 2006), A. awamori (Beena et al. 2010), P. purpurogenum (Reddy and Rathod 2012), A. aculeatus (Bagga et al. 2015), and others.

Mode of Action of Tannase

Tannase predominantly acts on ester and depside linkages in hydrolysable tannins preferably the gallotannins. Lekha and Lonsane (1997) documented the hydrolytic action of tannase on tannic acid giving off gallic acid and glucose. Lekha and Lonsane (1997) further documented that tannases show their catalytic action on complex tannins such as in similar fashion but do not show their catalytic action on condensed tannins. Tannase also precisely acts upon the ester linkage of methyl gallate and the depside linkage of m-digallic acid. Aguilera-Carbo et al. (2008) documented a few reports on the biocatalytic action of tannse on elligitannins. However, as a matter of fact, none of the reports demonstrated a clear cut mechanism of tannase action on ellagitannins.

Induction, Synthesis and Regulation of Tannase

Tannase has been reported as an inducible enzyme. Enzyme induction, expression, and production occur to varied levels credited to the strain used and the culture conditions employed. Tannase synthesis is efficiently induced by various phenolic compounds viz tannic acid, pyrogallol, gallic acid, and methyl gallate (Costa et al. 2008). Gallic acid, the major structural component of gallotannins, has been documented to efficiently induce tannase synthesis in submerged fermentation; however, it has been reported as a repressor of tannase synthesis in solid-state fermentation (Bajpai and Patil 1997). Belmares et al. (2004) documented enhanced tannase production by Aspergillus niger upon the addition of carbon sources like sucrose, fructose, glucose to the fermentation media in the concentration range 10–30 g/L.

Production Aspects of Tannase Through Fermentation

Tannase production in fungi has been documented via liquid-surface, submerged fermentation, liquid-surface and solid-state fermentation processes, respectively. However, submerged fermentation has been the preferred method for tannase production in bacteria and yeasts (Belur and Mugeraya 2011). Submerged fermentation has been the most preferred method for tannase production worldwide; however, as a matter of fact, few studies on tannase production through solid-state fermentation have also been documented. Selection of an effective production technique is governed by several parameters such as strain to be used, nutrient availability, type and nature of substrate being used.

Tannase Production Through Submerged Fermentation (SmF)

Submerged fermentation principally involves growing the microbial culture as a suspension in the fermentation medium having all the nutrients to sustain and support its growth (Aguilar et al. 2001). In the present day scenario, submerged fermentation is the most preferred process for commercial production of several valuable enzymes including tannase credited to ease of sterilization and better process control. Bajpai and Patil (1997) documented enhanced tannase production by Aspergillus spp. at higher aeration rates. Murugan et al. (2007) documented tannase production by Aspergillus xavus, Aspergillus niger, Fusarium spp., Penicillium spp., and Trichoderma spp. in submerged fermentation under controlled conditions. A. niger was documented as the most potent tannase producer amongst all isolates with an apparent enzyme activity of 16.77 U/mL. Paranthaman et al. (2009) optimized a temperature of 35 °C, 96 h of incubation time, and 2% tannic acid substrate concentration for maximal tannase production from A. flavus through submerged fermentation. Srivastava and Kar (2009) optimized extracellular tannase and gallic acid production by A. niger isolate through submerged fermentation at 37 °C, 72 h and 4% (w/v) pomegranate rind powder as a tannin-containing substrate. (Ahmed and Rahman 2014) optimized various process parameters (pH, temperature, and incubation period) for tannase production by an Aspergillus niger strain.

Tannase Production Through Solid-State Fermentation

Solid-state fermentation (SSF) principally commences in the scarcity of free water and essentially requires a solid support which could be either natural support or an inert support material. However, as a matter of fact, the substrate being used ought to have moisture content good enough to sustain the growth and metabolism of the strain used in the process. The low moisture content renders the free water unavailable to other microorganisms thereby limiting the chances of their growth and making the fermentation process feasible only by a constricted number of microorganisms particularly yeasts and fungi and few bacteria. Most of the reports have reported fungi as highly acclimatized to SSF conditions credited to their hyphae growth on particle surfaces and colonization on solid substrate. SSF principally involves two types of process that can be differentiated from each other on the basis of characteristics of the solid phase being utilized. The first one and most widely used process principally involves a solid phase that essentially serves as a support material as well as a source of nutrition. The substrates used are predominantly heterogenous, water-insoluble, and includes major byproducts from agriculture and food industry such as tamarind seed powder, pomegranate rind powder, potato, palm kernel cake, cassava, sugar beet pulp, coffee husk, and others (Sabu et al. 2005). The second process principally consists of an inert support like polyurethane foam, sugarcane bagasse, vermiculite, resins impregnated with liquid medium consisting of all essential nutrients. Deepa et al. (2015) documented tannase production from Aspergillus niger by utilizing wood chips as a solid substrate.

Purification of Tannases

Bhardwaj et al. (2003) reported that cell-deprived fermentation broth could be directly utilized as a source of extracellular tannase. However, as a matter of fact, the intracellular enzyme needs to be released out of the cells through their lysis. This could be achieved either by grinding of cellular fraction in sand or by crushing of cells by using homogenizer and subsequently extracting the cell lysate in suitable buffer. Naidu et al. (2008) utilized ammonium sulfate precipitation as preliminary purification step which resulted in partial purification as well as concentration of the crude enzyme. Mahapatra et al. (2005) described a purification procedure of tannase produced extracellularly by A. awamori nakazawa. The properties of the purified enzyme including pH and temperature optima and effects of urea, surfactants, and chelators were investigated.

Mukherjee et al. (2006) documented tannase production by a co-culture of the two filamentous fungi, R. oryzae and A. foetidus, by modified solid-state fermentation of tannin-rich substrates, and extracellular enzyme was purified through solvent precipitation and thereafter using DEAE-Sepadex column chromatography. They also conducted studies concerned with the effects of various process parameters on the activity of enzyme. The process parameters mainly studied were pH, temperature, effect of Km and Vmax along with the thermal stability of the enzyme at different temperatures. Chhokar et al. (2009) obtained a purification fold of 19.5 with 13.5% yield upon purification of tannase from A. awamori MTCC 9299 using ammonium sulfate precipitation followed by ion-exchange chromatography.

Naidu et al. (2008) reported tannase purification from A. foetidus through aqueous two-phase extraction using Polyethylene glycol (PEG) and documented a purification fold of 2.7 with an apparent yield of 82%. Ramirez-Coronel et al. (2003) documented purification of tannase from A. niger using a different protocol involving preparative isoelectric focusing for initial purification and ion-exchange chromatography thereafter for complete purification of enzyme. SDS-PAGE analysis of purified enzyme flaunted two protein bands with apparent molecular weights of 90 and 180 kDa, respectively. Deepa et al. (2015) documented purification of tannase from A. niger using DEAE-Sephadex gel filtration chromatography followed by SDS-PAGE analysis.

Tannase Characterization

Tannase has been documented as an inducible enzyme having high molecular mass ranging from 59 to 320 kDa (Table 16.1). The molecular mass of A. niger MTCC 2425 tannase was reported to be 185 kDa which consisted of two units of apparent molecular weights 102 and 83 kDa, respectively (Bhardwaj et al. 2003). Tannase from A. niger ATCC 16620 has a single monomeric unit of 168 kDa (Sabu et al. 2005). Kasieczka-Burnecka et al. (2007) reported that tannases from Verticillium spp. were oligomeric enzymes which consisted of two kinds of subunits with molecular masses of 39.9 and 45.6 kDa, respectively. Sharma et al. (2008) documented a molecular mass of 310 kDa for tannase from Penicillium variabile . The tannase from A. niger GH1 is composed of three subunits of molecular masses of 50, 75, and 100 kDa (Mata-Gomez et al. 2009). Beena et al. (2010) documented six identical subunits of 37.8 kDa from tannase of Aspergillus awamori. Goncalves et al. (2011) reported a heteromeric tannase of Emericella nidulans with three copies of each polypeptide. This enzyme has two polypeptide bands which correspond to 45.8 and 52 kDa as observed after 12% SDS-PAGE.

Table 16.1 Molecular mass of tannase from various fungal sources
Full size table

pH Optima and Stability of Tannase

Battestin and Macedo (2007) in their research study demonstrated that pH alters enzyme activity by determining the nature of the amino acids at active site which undergo protonation and deprotonation since change in pH alters their protonation pattern. Reddy and Rathod (2012) demonstrated tannase as a protein having acidic nature with an optimum pH around 5.0–6.0 and is unstable above pH 6.0. The tannases obtained from A. aculeatus DBF9, A. niger, A. awamori nakazawa, and Erwinia carotovora had a pH optima of 5.0 (Mahapatra et al. 2005). Chhokar et al. (2010) documented pH 5.5 to be optimum for tannase of A. awamori MTCC 9299. However, an optimal pH of 6.0 was recorded for A. niger tannase (Ramirez-Coronel et al. 2003; Mata-Gomez et al. 2009). Battestin and Macedo (2007) documented more than 80% stability of tannase in the narrow pH range between 4.5 and 6.5. Mahendran et al. (2006) in their research study documented a pH optimum in the range of 5.0–7.0 for the activity of tannase of P. variotii.

Temperature Optima and Stability of Tannase

The temperature optima for fungal tannases fall between 30 and 50 °C. Battestin and Macedo (2007) demonstrated optimum temperature as that maximizes the velocity of enzymatic reaction beyond which the rate of reaction declines as an effect of thermal denaturation thus rendering the enzyme inactive. Any increase in temperature beyond optima causes decline in catalytic rate of tannase due to denaturation (Mukherjee et al. 2006). The optimal temperature for tannase from Rhizopus oryzae for the free and immobilized enzyme was 40 and 55 °C, respectively. Sharma et al. (2008) reported that temperature optima for both free and immobilized enzyme of P. variabile were similar.

pI of Tannase

The pI of tannases has been documented in the range of 3.5–8.0. Tannase from A. niger GH1 had a pI value of 3.5 (Mata-Gomez et al. 2009) and for A. niger Aa20 tannase a pI value of 3.8 (Ramirez-Coronel et al. 2003). Isoelectric point of another tannase from A. awamori is 4.4 (Beena et al. 2010).

Kinetic Constants (Km and Vmax)

Km of tannase has been documented most preferably for methyl gallate and tannic acid substrates. However, as a matter of fact, a few researchers also documented Km values for other substrates including glucose-1-gallate, propyl gallate and others. Bhardwaj et al. (2003) documented propyl gallate to have lowest affinity for tannase from A. niger with an apparent Km value of 2.05 mM. For tannic acid, highest affinity for the same enzyme with a Km value of 0.28 mM and Vmax/Km of 2.53.

Enzyme Inhibitors

Barthomeuf et al. (1994) documented inactivation of tannase from A. niger by 0-phenanthroline and phenyl methyl sulfonyl fluoride (PMSF). Kar et al. (2003) reported 1,10-O-phenathrolein as an inhibitor for R. oryzae tannase. Beena et al. (2010) observed that amongst the different inhibitors tested, PMSF showed highest enzyme inhibition (4.5% residual activity), followed by sodium deoxycholate (26.4%) and phenanthroline (61.04% R). Sharma et al. (2008) reported that N-ethylmaleimide exhibited strong inhibition of tannase activity while working with that of Penicillium variabile.

Metal Ions

The inhibitory effects of Fe3+, Cu2+, Zn2+ ions on tannase have been reported (Kar et al. 2003; Sabu et al. 2005; Kasieczka-Burnecka et al. 2007; Chhokar et al. 2010). Tannase from the strains of A. niger is strongly inhibited by Mg2+ and Mn2+ (Bhardwaj et al. 2003). However in the separate case, Mg2+, Mn2+, Ca2+, Na+, K+ were documented to boost the activity of tannase from A. awamori MTCC 9299. Cu2+, Fe3+ and Co2+ showed strong inhibition of tannase activity, whereas Zn2+ does not show any pronounced effect on tannase activity (Chhokar et al. 2010). Enzyme tannase was strongly inhibited by Fe3+, whereas Cu2+ and Zn2+ showed only a mild inhibition, while Co2+ showed stimulatory effect on the activity of tannase from A. niger GH1 (Mata-Gomez et al. 2009). Goncalves et al. (2011) documented an spike in the activity of tannase with increase in Zn2+, Hg2+, Co2+, Mg2+ (33–39%), and NH4+ (15%) and showed inhibition in presence of Fe3+, Al3+, and Ag+.

Tannase Immobilization

Immobilization of enzymes involves physical confinement on inert/insoluble polymers, such as membranes or particles, which act as carriers/support of the enzyme during a continuous catalytic process. This permits recovery or removal of enzymes from a reaction mixture and their reusage (Sanderson and Coggon 1974) as well as use in non-aqueous environment, thereby improving the economy of the process. Retention of the enzyme is favored by coating alginate matrices with high or low molecular weight chitosan (Abdel-Nabey et al. 1999) or by cross-linking with glutaraldehyde , while storage under low temperatures favors stability of activity for longer periods. Sharma et al. (2008) documented Amberlite IR 1204 as the most suitable support for immobilizing P. variabile tannase with 69% immobilization credited to its reusability up to six times without any significant decline in the activity of enzyme. Yu et al. (2004) reported microencapsulation of tannase by chitosan-alginate complex coacervate membrane. Its kinetic properties revealed a slight shift in optimum pH toward alkaline from acidic and an increase in thermal stability.

Abdel-Nabey et al. (1999) documented another method of immobilization of tannase from Aspergillus oryzae on carriers like chitosan, chitin, Dowex 50 W, DEAE-Sephadex A-25. Maximum activity was observed on chitosan with bifunctional agent glutaraldehyde. This immobilized enzyme is widely used for the production of gallic acid. Mahendran et al. (2006) and Sharma et al. (2008) recorded that immobilized tannase is widely used for gallic acid production and for gallic acid esters (Yu et al. 2004). Hota et al. (2007) and Yu et al. (2004) reported a significant increase in Km value of tannase upon immobilizing preferably with tannic acid as substrate in the fermentation process.

Applications of Tannase

Tannases have a vast range of applications in various industrial bioprocesses ranging from food, feed to chemical as well as pharma sector.

Gallic Acid (3,4,5-Trihydroxy Benzoic Acid) Production

The most significant application of tannase is the gallic acid production from hydrolysable tannins (Kar et al. 2002). Gallic acid is a versatile precursor for the manufacture of a variety of chemicals used in food and pharmaceutical industries. Gallic acid is utilized worldwide as a precursor for manufacturing of trimethoprim, a broad spectrum antibacterial agent which is bacteriostatic, since it inhibits folic acid metabolism in pathogenic bacteria (Mukherjee and Banerjee 2003). In combination with sulfonamide, trimethoprim exerts antibacterial effect at very low concentrations against Streptococci, Staphylococci, Shigella, Corynebacterium diptheriae, E. coli, Vibrio cholerae, Gonococci, Bacillus pertussis, and Clostridium welchii . Prasad et al. (2006) documented emergence of gallic acid as a highly valuable molecule credited to its worldwide use and demand as an antioxidant, antiviral, radio protective agent, anticancer, and antitumor agent. Choi et al. (2010) documented the potential of gallic acid as an antiviral agent.

Instant Tea Production

Boadi and Neufeld (2001) documented the utilization of tannase in instant tea production. The conventional process of instant tea preparation principally involves low-temperature treatment of hot water extract of tea with continuous agitation and centrifugation of tea cream thereafter. Tenco Brooke Bond Ltd. has solubilized tea cream using tannase. The product obtained at 5 °C was hazy and had an undissolved solid content of 13.5%, while an untreated sample at low temperature was visually opaque and had an undissolved solid content of 7.5%. British Patents GB-B-1, 413,351 and GB-B-1, 380,135 (Unilever) described the method of removal of tea cream and dissolution of the cold water-insoluble constituents of a hot water extract of tea by treating the tea with free or immobilized tannase.

Beer and Juices Clarification

Tannase is utlitized worldwide in juice clarification in order to remove the undesirable bitterness present in various fruits. Tannase also lowers the haze without deteriorating the quality of juices (Rout and Banerjee 2006).

Effluent Treatment

Tannery effluent wastes are ranked as high pollutants among all industrial wastes. Awareness on environmental issues among the world community especially in developing countries has reached a level which was not seen till now. Murugan and Al-Sohaibani Saleh (2010) documented eviction of tannin from tannery effluents by using biomass and enzyme from A. candidus MTCC 9628.

Future Prospectives

The industrial importance of tannase is well established. Amongst different hydrolases, tannase is gaining commercial importance due to its potential applications in food, feed, chemical, and pharmaceutical industries. Furthermore, the advancements in molecular tools and techniques have facilitated a better understanding of tannase structure, its induction, synthesis, regulation, and underlying mechanism of action. Due to surplus availability of tannin-rich agro-industrial residues and toxic tannery effluent waste, there is always an opportunity for researchers to search out novel tannases with improved activity and stability. The prospects of tannin hydrolysis by tannase from filamentous fungi are quite promising. Furthermore, tannase-based treatment of toxic tannery effluent offers a cheaper and much reliable way for bioremediation of toxic tannery effluent. In conclusion, tannase is an industrially important enzyme loaded with huge potentials for use in various bioprocessing applications. Over the years, tannase has witnessed a significant rise in its utilization in commercial sector in various industrial applications. Thus, further research related to increasing the tannin hydrolysis rate, tannin tolerance as well as to assure better process control for increased tannase production would be envisaged.