TANNIN DEGRADINGPROLOGUE

A tannin (also known as vegetable tannin, natural organic tannins, or sometimes tannoid, i.e. a type of biomolecule, as opposed to modern synthetic tannin) is an astringent, bitter plant polyphenolic compound that binds to and precipitates proteins and various other organic compounds including amino acids and alkaloids.

They are probably present in all plant materials starting fromphytoplankton to macrophytes.

Bate-Smith (1962) defined tannin as “water soluble phenolic compounds having molecular weights between 500-3000 giving usual phenolic reactions and having special propertiessuch as the ability to precipitate alkaloid, gelatin and otherproteins”.

Bate-Smith (1973) described their purpose in plants as follows: “From the biological point of view the importance of tannins in plants lies in their effectiveness as repellents topredators, whether animal or microbial. In either case the relevant property is astringency rendering the tissues unpalatable by precipitating proteins or by immobilizing enzymes, impeding the invasion of the host by the parasite”.

Structure and classes of tannins

There are three major classes of tannins: Shown below are the base unit or monomer of the tannin. Particularly in the flavone-derived tannins, the base shown must be (additionally)heavily hydroxylated and polymerized in order to give the highmolecular weight polyphenol motif that characterizes tannins. Typically, tannin molecules require at least 12 hydroxyl groups and at least five phenyl groups to function as protein binders.

Base Unit:

Gallic acid Flavone Phloroglucinol

Class/Polymer:

Hydrolyzable tannins

Non-Hydrolyzableor condensed tannins

Phlorotannins

Sources Plants Plants Brown algaeOligostilbenoids (oligo- or polystilbenes) are oligomeric forms of stilbenoids and constitute a class of tannins.[5]

Pseudo tanninsPseudo tannins are low molecular weight compounds associated with other compounds. They do not change color during the Goldbeater's skin test, unlike hydrolysable and condensed tannins, and cannot be used as tanning compounds.[4] Some examples of pseudo tannins and their sources are: Pseudo tannin Source(s)Gallic acid RhubarbFlavan-3-ols (Catechins) Tea, acacia, catechu, cocoa, guaranaChlorogenic acid Nux-vomica, coffee, mateIpecacuanhic acid Carapichea ipecacuanha

Presence in soilsThe convergent evolution of tannin-rich plant communities has occurred on nutrient-poor acidic soils throughout the world. Tannins were once believed to function as anti-herbivore defenses, but more and more ecologists now recognize them as important controllers of decomposition and nitrogen cycling processes. As concern grows about global warming, there is

great interest to better understand the role of polyphenols asregulators of carbon cycling, in particular in northern borealforests.

Leaf litter and other decaying parts of a kauri (Agathis australis), a tree species found in New Zealand, decompose much more slowly than those of most other species. Besides itsacidity, the plant also bears substances such as waxes and phenols, most notably tannins,[22] that are harmful to microorganisms.

Presence in water and woodThe leaching of highly water soluble tannins from decaying vegetation and leaves along a stream may produce what is knownas a blackwater river. Water flowing out of bogs has a characteristic brown color from dissolved peat tannins. The presence of tannins (or humic acid) in well water can make it smell bad or taste bitter, but this does not make it unsafe todrink.

Tannins leaching from an unprepared driftwood decoration in anaquarium can cause pH lowering and coloring of the water to a tea-like tinge. A way to avoid this is to boil the wood in water several times, discarding the water each time. Using peat as an aquarium substrate can have the same effect.

Many hours of boiling the driftwood may need to be followed bymany weeks or months of constant soaking and many water changes before the water will stay clear. Adding baking soda to the water to raise its pH level will accelerate the processof leaching, as the more alkaline solution can draw out tannicacid from the wood faster than the pH-neutral water.]

Softwoods, while in general much lower in tannins than hardwoods, are usually not recommended for use in an aquarium[26] so using a hardwood with a very light color, indicating a low tannin content, can be an easy way to avoid tannins. Tannic acid is brown in color, so in general white woods have a low tannin content. Woods with a lot of yellow, red, or brown coloration to them (like southern yellow pine, cedar, redwood, red oak, etc.) tend to contain a lot of tannin.

(http://www.scielo.org.mx/scielo.php?pid=S0370-59432011000200003&script=sci_arttext)

Nutritional value of Tannins

Tannins are toxic and bacteriostatic compounds (Scalbert, 1991).

Tannins have traditionally been considered antinutritional butit is now known that their beneficial or antinutritional properties depend upon their chemical structure and dosage. The new technologies used to analyze molecular and chemical structures have shown that a division into condensed and hydrolyzable tannins is far too simplistic.[Muller-Harvey I., McAllan A. B. (1992). "Tannins: Their biochemistry and nutritional properties". Adv. Plant Cell Biochem. And Biotechnol. 1: 151–217.]

Recent studies have demonstrated that products containing chestnut tannins included at low dosages (0.15–0.2%) in the diet of chickens may be beneficial.[Schiavone A., Guo K., Tassone S. et al. (March 2008). "Effects of a natural extract of chestnut wood on digestibility, performance traits, and nitrogen balance of broiler chicks". Poult. Sci. 87 (3): 521–7]

Some studies suggest that chestnut tannins have been shown to have positive effects on silage quality in the round bale silages, in particular reducing NPNs (non protein nitrogen) inthe lowest wilting level.[Tabacco E., Borreani G., Crovetto G. M., Galassi G., Colombo D., Cavallarin L. (1 December 2006). "Effect of chestnut tannin on fermentationquality, proteolysis, and protein rumen degradability of alfalfa silage". J.Dairy Sci. 89 (12): 4736–46.]

Improved fermentability of soya meal nitrogen in the rumen hasalso been reported by F. Mathieu and J. P. Jouany (1993).[Mathieu F., Jouany J. P. (1993). "Effect of chestnut tannin on the fermentability of soyabean meal nitrogen in the rumen". Ann Zootech 42 (2): 127]

Studies by S. Gonzalez et al. (2002) on in vitro ammonia release and dry matter degradation of soybean meal comparing three different types of tannins (quebracho, acacia and chestnut) demonstrated that chestnut tannins are more efficient in protecting soybean meal from in vitro degradationby rumen bacteria.

Condensed tannins inhibit herbivore digestion by binding to consumed plant proteins and making them more difficult for animals to digest, and by interfering with protein absorption

and digestive enzymes (for more on that topic, see plant defense against herbivory). Many tannin-consuming animals secrete a tannin-binding protein (mucin) in their saliva. Tannin-binding capacity of salivary mucin is directly related to its proline content. Advantages in using salivary proline-rich proteins (PRPs) to inactivate tannins are:

PRPs inactivate tannins to a greater extent than do dietary proteins; this results in reduced fecal nitrogen lossesPRPs contain non specific nitrogen and nonessential amino acids; this makes them more convenient for an animal to exploit rather than using up valuable dietary protein

Drinks with tanninsThe best-known human dietary sources of tannins are tea and wine. Other sources include fruit juices, but not citrus. Wineis a particular case. Most wines aged in charred oak wooden barrels possess tannins absorbed from the naturally occurring tannins in the wood which seep out into the wine. This concentration gives wine its signature bitterness. The tanninsdraw residual proteins from the tongue and mouth which gives wine its aftertaste. Consumption of foods with higher fat content neutralizes this effect.

Coffee pulp:has been found to contain low to trace amounts of tannins.[González S., Pabón M. L., Carulla J. (2002). "Effects of tannins on in vitro ammonia release and dry matter degradation of soybean meal". Arch. Latinoam. Prod. Anim. 10 (2): 97–101.]

Fruit juices:Although citrus fruits do not themselves contain tannins, orange-colored juices often contain food dyes with tannins. Apple juice, grape juices and berry juices are all high in tannins. Sometimes tannins are even added to juices and cidersto create a more astringent feel to the taste.

Beer:In addition to the alpha acids extracted from hops to provide bitterness in beer, condensed tannins are also present. These originate both from the malt and hops.[citation needed] Especially in Germany, trained brewmasters consider the presence of tannins as a flaw.[citation needed] In some

styles, the presence of this astringency is acceptable or evendesired, as, for example, in a Flanders red ale.

In lager type beers, the tannins can form a precipitate with specific haze-forming proteins in the beer resulting in turbidity at low temperature. This chill haze can be preventedby removing part of the tannins or part of the haze-forming proteins. Tannins are removed using PVPP, haze-forming proteins by using silica or tannic acid.

Food items with tanninsFruitsPomegranatesPersimmons:Some persimmons are highly astringent and therefore inedible when they are not extremely ripe (to be specific, the Korean, American, and Hachiya or Japanese).Berries:Most berries, such as cranberries, strawberries and blueberries, contain both hydrolyzable and condensed tannins.Nuts:Nuts that can be consumed raw, such as hazelnuts, walnuts, andpecans, contain high amounts of tannins. Almonds have a lower content. Tannin concentration in the crude extract of these nuts did not directly translate to the same relationships for the condensed fraction. Acorns contain such high concentrations of tannins that they must be processed before they can be consumed safely.The areca nut also contains tannin.

Smoked foods:Tannins from the wood of mesquite, cherry, oak, and other woods used in smoking are present on the surface of smoked fish and meat.

Herbs and spices:Cloves, tarragon, cumin, thyme, vanilla, and cinnamon all contain tannins.[Navia, Jeanette. "Could Tannins Explain Classic Migraine Triggers?" 1988]

Legumes:

Most legumes contain tannins. Red-colored beans contain the most tannins, and white-colored beans have the least. Peanuts without shells have a very low tannin content. Chickpeas (garbanzo beans) have a smaller amount of tannins.[Reed J. D. (1 May 1995). "Nutritional toxicology of tannins and related polyphenols in forage legumes". J. Anim. Sci. 73 (5): 1516–28]

Chocolate:Chocolate liquor contains about 6% tannins.[Robert L. Wolke; Marlene Parrish (29 March 2005). What Einstein told his cook 2: the sequel: further adventures in kitchen science. W. W. Norton & Company. p. 433]

INTRODUCTION

Tannic acid is used worldwide as clarifying agent in alcoholic drinks and as aroma ingredient in both alcoholic andsoft drinks or juices. Tannins from different botanical origins also find extensive uses in the wine industry.

Tannins are an important ingredient in the process of tanning leather. Tanbark from oak, mimosa, chestnut and quebracho treehas traditionally been the primary source of tannery tannin, though inorganic tanning agents are also in use today and account for 90% of the world's leather production.[Marion Kite; Roy Thomson (2006). Conservation of leather and related materials. Butterworth-Heinemann. p. 23]

Tannins produce different colors with ferric chloride (either blue, blue black, or green to greenish-black) according to thetype of tannin. Iron gall ink is produced by treating a solution of tannins with iron(II) sulfate.

Condensed tannins, e.g., quebracho tannin, and Hydrolyzable tannins, e.g., chestnut tannin, appear to be able to substitute a high proportion of synthetic phenol in phenol-formaldehyde resins for wood particleboard.

Tannins can be used for production of anti-corrosive primer, sold under brand name-Nox Primer for treatment of rusted steelsurfaces prior to painting, rust converter to transform oxidized steel into a smooth sealed surface and rust inhibitor.

The use of resins made of tannins has been investigated to remove mercury and methylmercury from solution.(Torres J., Olivares S., De La Rosa D., Lima L., Martínez F., Munita C. S.,Favaro D. I. T. (1999). "Removal of mercury(II) and methylmercury from solution by tannin adsorbents". Journal of Radioanalytical and Nuclear Chemistry 240 (1): 361–5.)

Immobilized tannins have been tested to recover uranium from seawater.[Takashi Sakaguchia, Akira Nakajimaa (June 1987). "Recovery of Uranium fromSeawater by Immobilized Tannin". Separation Science and Technology 22 (6): 1609–23]

Some of the tannins are used due to their antioxidant properties for protect metals of corrosion.

Binding ability of the tannins to proteins facilitates the protein to bypass the rumen fermentation and allows better utilization of dietary protein in ruminant animals. High urinary N output into the environment is an implication of sub-optimal protein utilization.

The process of tannin-protein binding is complex and may depend on quantity, structure and size of both tannin and protein. Tannins have a high degree of hydroxylation that offers many possible sites for binding to proteins. Complexes are formed mainly via hydrogen bonding. Proline-rich proteins have been described as particularly reactive with tannins (Hagerman & Butler, 1981). Precipitation increases with size of the complex. This occurs as a result of more tannins binding to the protein, cross linking with other proteins, causing decreased hydrophilicity. The polymer polyethylene glycol (PEG) has been shown to have high binding affinity to tannins and may even replace already tannin bound proteins (Jones & Mangan, 1977). This effect has been used to study tannin-protein binding (Silanikove et al., 1996; Makkar et al., 1993) or to alleviate detrimental nutritional effects caused by tannins (Priolo et al., 2000).

The stability of tannin-protein bindings has been shown to be pH dependant (Oh & Hoff, 1987; Hagerman & Butler, 1981). They seem to be more stable close to the isoelectric point of the protein, i.e. for the most abundant plant protein Ribulose-1,5-bisphosphate carboxylase oxygenase (Rubisco) in a pH range from 3.5 to 7.0. The complexes are susceptible to release, particularly at low pH (Jones & Mangan, 1977).

(http://www.slideshare.net/CarlaSouza1/tannins-33124697)

ROLE OF TANNINS IN ANIMAL FEEDSWide use and acceptability of plant derived protein sources asalternatives to FM have been constrained by the imbalanced amino acid profiles, and the presence of some anti-nutritionalfactors (ANFs), such as tannin, phytic acid, trypsin inhibitor, lectins, saponins and many more (Francis et al., 2001). Tannins are probably the most common among the plant derived ANFs (Francis et al., 2001), which are considered as secondary compounds of various chemical structures widely

occurring in the plant kingdom starting from phytoplankton to the macrophytes (Hernes and Hedges, 2000).

Experimental evidence has shown to induce feed rejection by hydrolysable tannin when included in the feed of the common carp, Cyprinus carpio (Becker and Makkar, 1999). Poor growth performance has also been recorded in the same species as a result of feedingtannic acid incorporated diet (Hossain and Jauncey, 1989). In addition, it has beenhypothesized that tannins interfere with protein and dry matter digestibility byinhibiting protease and also by forming indigestible complexeswith dietary protein(Krogdahl, 1989).

Vohra et al. (1966) reported that tannins caused growth depression in chickens at levels as low as 0.5% of the diet. Similarly, Hossain and Jauncy (1989) observed poor growth response in common carp (Cyprinus carpio) fed diets containing 0.57% and 1.14% tannins. In addition, high levels of tannins in feed have been shown to have adverse effect on herbivorous and omnivorous fish (Olvera et al., 1988; Al-Owafeir, 1999; Becker and Makkar, 1999).

Tannins may reduce macro nutrient utilization by forming tannin protein complexes with various digestive enzymes precluding the formation of the product absorbable by the small intestine (Carmona et al., 1996). In addition, it may inhibit transport systems concerning carbohydrate assimilationsuch as glucosidase/ maltase (Bjorck and Nyman, 1987), sucrase(Welsch et al., 1989) and the intestinal sodium dependent glucoseuptake system (Karasov et al., 1992). Bean tannins were shown to strongly inhibit pancreatic trypsin, chymotrypsin and α-amylase (Sing, 1984; Carmona et al., 1991).

Maitra and Ray (2003) demonstrated the inhibition of the intestinal enzymes in L. rohita fingerlings to a significant level by the tannin extracted from Acacia, though, the effect of the hydrolysable tannin on the digestive enzymes of the other carp species has not been worked out.

Tannin not only affected the protease activity, but also inhibited comparatively lipaseactivities to a greater extent and amylase activities to a lesser extent.

Interference by tannin with the activities of metabolic enzymes at the tissuelevel may serve as a basis to explain the poor growth responseof fish fed tannincontaining feed (Olvera et al., 1988; Al-Owafeir, 1999; Becker and Makker, 1999).(http://www.google.co.in/url?sa=t&rct=j&q=&esrc=s&source=web&cd=1&cad=rja&uact=8&ved=0CCIQFjAA&url=http%3A%2F%2Fshodhganga.inflibnet.ac.in%3A8080%2Fjspui%2Fbitstream%2F10603%2F21336%2F2%2Fthesis%2520complete.pdf&ei=wLFEVcvyJ4GNuASzmoDoCg&usg=AFQjCNFutp8Y97EvBHCODKuQI9oFjadwzQ&bvm=bv.92291466,d.c2E)

Concentration of tannin (mean ± S.E. of three observations) in natural and potential fish food items / ingredients.

Potential fish food Amount of tannin(mg g-1 dry weight)

Phytoplankton 34.3±0.67iZooplankton 6.58±0.19cLemna minor 5.83±0.21bLemna major 7.5±0.27dAzolla sp. 19.58±0.38fPistia sp. 26.67±0.41hIpomoea sp. 5.81±0.32bGroundnut oil cake 27.05±0.43hSunflower oil cake 26.25±.38hCoconut oil cake 5.38±0.18 aMustard oil cake 21.25±0.35 gSoybean seed 10.42±0.26eValues with the same superscript in the same column are not significantly different (P<0.05) from each other.

Hence the need arises for degrading the tannins.

TANNIN DEGRADATIONHydrolysable and condensed tannins are the two major classes of tannins. These compounds have a range of effects on variousorganisms – from toxic effects on animals to growth inhibitionof microorganisms. Some microbes are, however, resistant to tannins, and have developed various mechanisms and pathways for tannin degradation in their natural milieu. The microbial degradation of condensed tannins is, however, less than hydrolysable tannins in both aerobic and anaerobic environments. A number of microbes have also been isolated from the gastrointestinal tract of animals, which have the ability to break tannin-protein complexes and degrade tannins,especially hydrolysable tannins. Tannase, a key enzyme in the degradation of hydrolysable tannins, is present in a diverse group of microorganisms, including rumen bacteria. This enzymeis being increasingly used in a number of processes. Presently, there is a need for increased understanding of the biodegradation of condensed tannins, particularly in ruminants.(http://link.springer.com/article/10.1023%2FA%3A1008397506963)

Tannins in aquatic environment cause harmful effects to aquatic life. They are inhibitory to microbial growth, respiration and metabolism.

In animals, especially ruminants, these effects include decreased feed intake, lowernutrient utilization and toxicity at higher intake levels (VanSoest 1994). Tannins have long been considered toxic to micro-organisms, and this activity is mainly due to enzymeinhibition and substrate deprivation, action on membranes, andmetal ion deprivation (Reed 1995). Nevertheless, some fungi, yeasts and bacteria are quite resistant to tannins and can also degrade them (Scalbert 1991 ; Saxena et al . 1996). A strain of Aspergillus niger van Tieghem has been reported to possess tannin –protein complex degrading activity (Bhat et al. 1996)

Let us take examples of some conventional and non conventionalfeeding stuff for use in Animal Feeds and their usable levels.

Groundnut Oil Cake is an excellent source of arginine but deficient in lysine, cysteine and methionine. It contains manyANFs including trypsin inhibitor, tannin and phytic acid that lowers the quality of protein(Nyina-wamwiza et al., 2010).

Mohanta et al. (2007) reported poor performance of silver barb, Puntius gonionotus fingerlings fed diet containing 28% Sunflower Oil Cake which may be due to the presence of some anti-nutritional factors like protease inhibitors and polyphenolic compounds like tannin and chlorogenic acid as reported by Liener (1980).

The limitation of using mustard oil cake as feed ingredient for fish is the presence of glucosinolates, a toxic factor causing reduced growth rate in fish (Jackson et al., 1982).

Jatropha curcas cake also contains several anti-nutritional factors such as phorbol esters, tannin, phytic acid, trypsin inhibitors, lectins and saponins.

Edwards et al. (1985) observed better growth and feed utilizationefficiency in tilapia, O. niloticus fed pelleted diets formulated with 75% composted water hyacinth. On the contrary, Hasan et al. (1990) observed less acceptability of water hyacinth incorporated diet by rohu fry. Keshavanath et al. (1990) reportedbetter body weight gains in rohu, grass carp and common carp fed diets incorporating the leaf meal powder at 50% level. Kumar et al. (1991) also reported positive utility of water hyacinth for carp spawns.

Hajra and Tripathi (1985) concluded that although duckweed incorporated diets resulted in a little less growth of fish ascompared to the control diet with fish meal, there are possibilities for replacement of the control diet up to 60% level with dried Lemna leaf meal without any significant reduction in fish growth. According to Bairagi et al. (2002b) thenutritive value of Lemna leaf meal was enhanced by fermentation.

Soya Bean Meal contains some anti-nutritional factors (ANFs), such as protease (trypsin) inhibitors, phytohaemagglutinin (lectins), anti-vitamins, phytic acid, saponins, andphytoestrogens (El-Sayed, 1999; Francis et al., 2001). Though phytic acid, protease inhibitors, lectins and anti-vitamins can be removed from SBM by thermal processing, phytoestrogens and saponins are heat-stable (Francis et al., 2001).

The major ANFs of canola meal include fibre, oligosaccharides,phenoliccompounds, phytic acid and glucosinolates (Bell, 1993; Higgs etal., 1995). Thephytic acid content of canola meal ranges from 3.1 to 3.7 g kg-1 (Higgs et al., 1995)and significantly reduces phosphorus digestibility, protein utilization and growth(Spinelli et al., 1983; Forster et al., 1999).

Polyphenolic compounds like tannins and gossypol impair nutritional quality, decrease digestibility, thereby reducing feed consumption. Reaction of these compounds with protein reduces protein quality, especially by reducing the bioavailability of lysine. Their antinutritional effects

include interference with the digestive processes either by binding with the enzymes or by binding to feed components likeproteins or minerals. Tannins also reduce the absorption of vitamin B12 (Liener, 1989).

TANNIN DEGRADATION BY MICROORGANISMSCondensed tannins are more resistant to microbial decomposition, while hydrolysable tannins are more easily degraded by some microorganisms (Lewis and Starkey, 1968; Lekha and Lonsane, 1997; Aguilar et al., 2000, 2001; Aguilar and Gutierrez-Sanchez, 2001). Condensed tannins are polymers of catechin or similar flavans and hydrolysable tannins are gallotannins (gallic acid and glucose) or ellagitannins (ellagic acid and glucose).(http://www.znaturforsch.com/ac/v60c/s60c0844.pdf)

Lactobacillus plantarum is frequently isolated from the fermentation of plant material where tannins are abundant. L. plantarum strains possess tannase activity to degrade plant tannins. A L. plantarum tannase (TanBLp, formerly called TanLp1) has been previously identified and biochemically characterized. In this study, we report the identification andcharacterization of a novel tannase (TanALp). While all the 29L. plantarum strains analyzed in this study possess the tanBLpgene, the gene tanALp was only present in four strains. Upon

methyl gallate exposure, the expression of tanBLp was induced,whereas tanALp expression was not affected. TanALp showed only27% sequence identity to TanBLp but the residues involved in tannase activity are conserved. Optimum activity for TanALp was observed at 30 °C and pH 6 in presence of Ca(2+) ions. TanALp was able to hydrolyze gallate and protocatechuate esters having a short aliphatic alcohol substituent. Moreover,TanALp was able to fully hydrolyze complex gallotannins such as tannic acid. The presence of the extracellular TanALp tannase in some L. plantarum strains will provide them an advantage for the initial degradation of complex tannins present in plant environments. (Tannin Degradation by a Novel Tannase Enzyme Present in Some Lactobacillus plantarum Strains. https://www.researchgate.net/publication/260647188_Tannin_degradation_by_a_novel_tannase_enzyme_present_in_some_Lactobacillus_plantarum_strains )

Aspergillus niger PSH and Penicillium commune EH2 exhibited their biotechnological potential because they can utilize condensed tannins while Aspergillus niger PSH and GH1 degradedhigh tannic acid levels used as carbon source. They could be used to produce tannase and degraded tannins directly in residues of coffee pulp, being a good alternative to solve thebig problem that represents the accumulations of this agroindustrial byproduct in some countries like Mexico, Colombia or Brazil.(http://www.znaturforsch.com/ac/v60c/s60c0844.pdf)

Microbial Biotechnology (Fermentation)Ingredient(s)processed

Flour of Finger Millet (Eleusinecoracana)

Duckweed (Lemna polyrhiza) leaf meal

Soybean (Glycine max), cowpea(Vigna unguiculata),Ground bean (Macrotylomageocarpa) seed meal

Leucaena leucocephala leafmeal

Grass pea (Lathyrus sativus) seed meal

Methodology

Fermentation of finger millet flour using endogenousgrain microflora

Pretreatment with Bacillus sp., fish gut isolate

Pretreated with conventionalprocesses (soaking–dehulling–washing cooking) andfermentation with Rhizopusoligosporus

Leaf meal inoculated with fishintestinal bacteria, Bacillus subtilis and Bacillus circulansisolated from Common carp, C.carpio and Mozambique tilapia,O. mossambicus, respectively

Brief results and application

Showed a significant ↓ oftannins by 52% (and alsoofother ANFs at varying levels) at the end of 24h. and asimultaneous ↑ in in vitroprotein digestibility.

↓ ANFs and ↑ total free fattyacids and amino acids. ↑growthand body composition in L.rohita fingerlings up to 30%incorporation level.

Conventional processessignificantly ↓ trypsin inhibitor.Phytic acid ↓ significantlyduring. Tannins ↓ mainlybyconventional processes andfermentation. No tannindetected in dehulled andcookedand in fermented soybean,cowpea and groundbean.

The crude fibre, cellulose andhemicellulose contents and theANF, tannin, phytic acidandmimosine in the leaf meal ↓;free amino acids and fatty acids↑; the group fed inoculated leaf

Reference

Antony andChandra, 1998

Bairagi et al.,2002b

Egounlety andAworh, 2003

Bairagi et al.,2004

Ramachandran

Grass pea, Lathyrus sativus L., seed meal

Pretreatment with Bacillus sp. Isolated from gut of adultcommon Carp, Cyprinus carpio

Processed through fermentation, extrusion,autoclaving, and germination.

meal performed better intermsof growth response, feedconversion ratio, proteinefficiency ratio and apparent netprotein utilization in L. rohitafingerlings

↓ in ANFs (including tannin)and ↑ of nutritive valueof grasspea seed meal. Apparent protein digestibility ↑ in L. rohita fingerlings

Fermentation, extrusion andgermination significantly ↓ theANFs (including tannin).Autoclaving ↓ tannins but wasnot effective for other ANFs.30% fermented, extruded,andgerminated grass pea seed mealincorporated diets resulted insignificantly (P < 0.05)bettergrowth performance and feedutilization in L. rohitafingerlings.

et al., 2005

Ramachandranand Ray, 2008

Symbols represent an increase/ improvement (↑) or decrease (↓).#Adapted from Ghosh and Ray (2012).

TANNIN DEGRADATION BY ENZYMESMicrobial tannases, tannin acyl hydrolases (E.C.3.1.1.20), areamong st important industrial enzymes with immense applications in food, feed, leather and pharmaceutical industries (Lekha and Lonsane, 1997).

Tannase (Tannin Acyl Hydrolase, E.C. 3.1.1.20) is an importantinducible enzyme which catalyses decomposition of hydrolysable

tannins especially gallo-tannins to glucose and gallic acid (Van de Lagemaat and Pyle, 2005). Gallic acid, the product of tannin hydrolysis finds application in many fields including dye-making, serves as a precursor for the commercial production of an anti-microbial drug trimethoprim, a food –preservative propyl gallate(Hota et al., 2007).

Conventionally gallic acid is produced by acid hydrolysis of tannic acid but it has cost, yield and low purity disadvantages.

Alternatively, gallic acid can be produced by the microbial hydrolysis of tannic acid by tannase. The use of tannin-rich agricultural and forest residues as a substitute of costly tannic acid substrate is an economic alternative for gallic acid production which also suggest the beneficial utilization of agro wastes.

Tannases (Tannin Acyl Hydrolase, E.C. 3.1.1.20) are industrially important enzymes which find applications mainly in production of Gallic acid, in beer, wine and fruit juice clarification and detannification of poultry feed. Hence, the present investigation was carried out with a view to obtain industrially potent extracellular tannase from the fungal source. Among the total eighty four fungal strains isolated, the isolate Trichoderma harzianum (MTCC no. 10841) was selected for tannase productionand optimization studies.

The crude tannase had maximum activity of 31.36 U/ml with 1.0%tannic acid as substrate. The industrial potential of tannase from Trichoderma harzianum in fruit juice clarification and gallic acid production was explored. On treatment of varying amount of tannase with pomegranate juice at 40oC tannin reduction of 57% was observed without the loss of its biochemical attributes such as pH, viscosity and sugar contentand protein content. The natural tannin degradation efficiencyof the enzyme was also investigated for gallic acid productionusing different tannin-rich agro residues as substrate. The

enzyme showed maximum activity with amla fruits as substrate which was higher than tannic acid. The tannase from Trichoderma harzianum can therefore be employed for gallic acid production using cheaper agro residues(http://www.ripublication.com/ijbbr_spl/ijbbrv4n6spl_25.pdf)

Microbial source of tannase*Microorganisms ReferencesBacteriaAchromobacter sp. Lewis and Starkey (1969)Bacillus spp. Deschamps et al. (1983), Mondal et al. (2001c),Corynebacterium sp. Deschamps et al, (1983)Klebisella spp. Deschamps et al. (1983)Pseudomonas solanaceanum Deschamps et al. (1983)Streptococcus spp. Belmares et al. (2004), Sasaki et al. (2005)Lactobacillus spp. Ayed and Hamdi (2002), Kostinek et al. (2007),Nishitani and Osawa (2003), Nishitani et al. (2004), Sabu et al. (2006)Enterococcus faecalis Goel et al. (2005)Citrobacter freundii Belmares et al. (2004)Selenomonas ruminantium Belmares et al. (2004)

YeastsCandida sp. Aoki et al. (1976)Pichia spp. Deschamps et al. (1983)Debaryomyces hansenii Deschamps et al. (1983)Saccharomyces cerevisiae Zhong et al. (2004)Mycotorula japonica Belmares et al. (2004)

FungiAspergillus spp. Bradoo et al. (1996; 1997), Belmares et al. (2004),Mahapatra et al. (2005), Murugan et al. (2007), Treviño-Cueto et al. (2007)Penicillium spp. Bradoo et al. (1996), Lekha and Lonsane (1997),

Batra and Saxena (2005), Van de Lagemaat and Pyle (2005)

Neurospora crassa Bradoo et al. (1996)Fusarium spp. Bradoo et al. (1996)

Rhizopus oryzae Hadi et al. (1994), Purohit et al. (2006)*Adapted from Aguilar et al. (2007)

Mode of actionTannase catalyses the breakdown of hydrolysable tannin such astannic acid,methylgallate, ethylgallate, n-propylgallate, and isoamylgallate (Fig.2.4). It is well known that tannase hydrolyses the ester bonds of tannic acid, although tannic acid is known to denature proteins (Aguilar et al., 2007). Tannase hydrolyses tannic acid completely to gallic acid and glucose through 2,3,4,6,-tetragalloyl glucose and two kinds ofmonogalloyl glucose.

Regression equations of the activities of enzymes from the Indian major carps against graded levels of

tanninEnzymes Fish species Regression Equation

(y = enzyme activity, x = Dose of tannin)Protease L. rohita y=13.36-0.064x; F=181.98, df=1,19; p<0.001; R2=0.905

C. catla y=11.37-0.023x; F=834.63, df=1,19; p<0.001; R2=0.978

C. cirrhosus y=10.12-0.04x; F=848.1, df=1,19; p<0.001; R2=0.978

Amylase L. rohita y=7.08-0.013x; F=254.35, df=1,19; p<0.001; R2=0.93

C. catla y=8.3-0.009x; F=614.45, df=1,19; p<0.001; R2=0.97

C. cirrhosus y=6.58-0.01x; F=304.04, df=1,19; p<0.001; R2=0.941

Lipase L. rohita y=11.98-0.05x; F=2687.66, df=1,19; p<0.001; R2=0.993

C. catla y=8.84-0.02x; F=348.86, df=1,19; p<0.001; R2=0.948

C. cirrhosus y=10.49-0.04x; F=2099.89, df=1,19; p<0.001; R2=0.991

Percent reduction of in vitro activities of the intestinalenzymes

from the three Indian major carps by gradedlevels of tannin extracted from Pistia leaf meal

% reduction in enzyme activity (control, without tannin at 100%activity)Tannin Protease Amylase Lipaseadded(μg) L. rohita C. catla C. cirrhosus L. rohita C. catla C. cirrhosus L. rohita C. catla C. cirrhosus12.5 9.65±0.13 4.37±0.06 6.43±0.17 3.22±0.07 3.12±0.03 3.67±0.14

7.93±0.2 4.43±0.04 6.41±0.0325 17.87±0.35 6.74±0.14 10.66±0.19 9.31±0.23 3.68±0.08 5.75±0.14

12.95±0.12 8.9±0.1 12.17±0.1650 47.17±0.51 13.23±0.21 28.2±0.19 18.53±0.31 7.79±0.16 14.42±0.29 24.69±0.13 17.62±0.36 25.79±0.16100 67.35±0.56 20.75±0.11 38.61±0.48 22.35±0.37 12.65±0.23 17.29±0.15 49.28±0.67 29.87±1.04 43.1±0.67150 71.42±0.6 27.47±0.27 54.12±0.42 26.52±0.28 16.81±0.3 21.84±0.53 62.81±0.41 37.28±0.75 59.09±0.15200 88.28±0.68 43.96±0.09 82.53±0.99 37.88±0.77 21.63±0.33 32.74±0.39 87.07±0.59 41.35±0.4 75.35±0.62

The study clearly indicated that tannin (extracted from Pistia) even at very low concentration (12.5μg to 200μg) can inhibit or, lower the digestive enzyme activity in the Indian major carps in a dose dependent manner as evident from the regression equations.

Among the fish species, digestive enzymes from L. rohita were found tobe the most sensitive to tannin followed by C. cirrhosus, while enzymes from C. catla were comparatively least affected

(http://www.google.co.in/url?sa=t&rct=j&q=&esrc=s&source=web&cd=1&cad=rja&uact=8&ved=0CCIQFjAA&url=http%3A%2F%2Fshodhganga.inflibnet.ac.in%3A8080%2Fjspui%2Fbitstream%2F10603%2F21336%2F2%2Fthesis%2520complete.pdf&ei=wLFEVcvyJ4GNuASzmoDoCg&usg=AFQjCNFutp8Y97EvBHCODKuQI9oFjadwzQ&bvm=bv.92291466,d.c2E)

ANALYTICAL PROTOCOLSTests for tanninsThere are three groups of methods for the analysis of tannins:precipitation of proteins or alkaloids, reaction with phenolicrings, and depolymerization.[30]

Goldbeater's skin test

When goldbeater's skin or ox skin is dipped in HCl, rinsed in water, soaked in the tannin solution for 5 minutes, washed in water, and then treated with 1% FeSO4 solution, it gives a blue black color if tannin was present.

Ferric chloride (FeCl3) testIt is rather a test for phenolics in general. Powdered plant leaves of the test plant (1.0 g) are weighed into a beaker and10 ml of distilled water are added. The mixture is boiled for five minutes. Two drops of 5% FeCl3 are then added. Productionof a greenish precipitate was an indication of the presence oftannins.[31] Alternatively, a portion of the water extract is diluted with distilled water in a ratio of 1:4 and few drops of 10% ferric chloride solution is added. A blue or green color indicates the presence of tannins (Evans, 1989).[32]

Other methodsThe hide-powder method is used in tannin analysis for leather tannin and the Stiasny method for wood adhesives.[33][34] Statistical analysis reveals that there is no significant relationship between the results from the hide-powder and the Stiasny methods.[35][36]

Hide-powder method400 mg of sample tannins are dissolved in 100 ml of distilled water. 3 g of slightly chromated hide-powder previously dried

in vacuum for 24h over CaCl2 are added and the mixture stirredfor 1 h at ambient temperature. The suspension is filtered without vacuum through a sintered glass filter. The weight gain of the hide-powder expressed as a percentage of the weight of the starting material is equated to the percentage of tannin in the sample.

Stiasny's method100 mg of sample tannins are dissolved in 10 ml distilled water. 1 ml of 10M HCl and 2 ml of 37% formaldehyde are added and the mixture heated under reflux for 30 min. The reaction mixture is filtered while hot through a sintered glass filter.The precipitate is washed with hot water (5x 10 ml) and dried over CaCl2. The yield of tannin is expressed as a percentage of the weight of the starting material.

Tannase assayTannase activity was determined spectrophotometrically following Mondal et al. (2001b) with minor modifications, based on the measurement of residual tannic acid content after enzymatic reaction. The reaction mixture consisting of 0.3 ml tannic acid [0.5% (w/v), in phosphate buffer, pH 6.5] and 0.1 ml crude enzyme was incubated at 35°C for 1 h. The enzymatic reaction was stopped by the addition of 3 ml bovine serum albumin (1 mg ml-1), which also leads to the precipitation of the remaining tannic acid. A reference tube was prepared in the same way with heat-denatured enzyme. The tubes were then centrifuged (8,000×g, for 20 min) and the precipitate dissolved in 3 ml SDS-triethanolamine [1% (w/v) of SDS in 5% (v/v) triethanolamine] solution mixed with 1 ml FeCl3 and absorbance was measured at 530 nm using a spectrophotometer (Shimadzu UV-1800). One unit of tannaseactivity was defined as the amount of enzyme that can hydrolyze 1 μmol of tannic acid in 1 min under assay

conditions. Enzyme yield was expressed as U gds-1 (gram dry substrate).

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Becker K and Makkar HPS (1999). Effects of dietary tannic acid and quebracho tannin on growth performance and metabolic rates of common carp (Cyprinus carpio L.). Aquaculture, 175: 327–335.

Bhat TK, Singh B and Sharma OP (1998). Microbial degradation of tannins – Acurrent perspective. Biodegradation, 9: 343-357.

Carmona A, Borgudd L, Borges G and Levy-Benshimol A (1996). Effect of blackbean tannins on in vitro carbohydrate digestion and absorption. Journal of Nutritional Biochemistry, 7: 445-450.

Deschamps AM, Otuk G and Lebeault JM (1983). Production of tannase and degradation of chestnut tannin by bacteria. Journal of Fermentation Technology, 61: 55–59.

Ephraim E, Odenyo A and Ashenafi M (2005). Isolation and characterization of tannin degrading bacteria from faecal samples of some wild ruminants in Ethiopia. Animal Feed Science and Technology, 118: 243–253.

Ghosh K and Ray AK (2011).Tannins in Plant Feed Ingredients: Facts and Probable Consequences in Fish Nutrition. In: Tannins: Types, Foods Containing, and Nutrition (Ed. G. K. Petridis). Nova Publishers, Hauppauge,New York, pp. 265-280.

Goel G, Puniya AK, Aguilar CN and Sing K (2005). Interaction of gut microflora with tannins in feeds. Naturwissenschaften, 92: 497–503.

Hernes PJ and Hedges JI (2000). Determination of condensed tannin monomers in plant tissues, soils, and sediments by capillary gas chromatography of acid hydrolysis extracts. Analytical Chemistry, 72: 5115-5124.

Lewis JA and Starkey RL (1969). Decomposition of plant tannins by some soilmicroorganisms. Soil Science, 107: 235–241

Maitra S and Ray AK (2003). Inhibition of digestive enzymes in rohu, Labeo rohita (Hamilton) fingerlings by tannin, an in vitro study. Aquaculture Research, 34: 93–95.

Mondal KC, Banerjee D, Jana M and Pati BR (2001b). Colorimetric assay method for determination of the tannin acyl hydrolase (EC3.1.1.20) activity. Analytical Biochemistry, 295: 168–171.

Murugan K, Saravanababu S and Arunachalam M (2007). Screening of tannin acyl hydrolase (E.C.3.1.1.20) producing tannery effluent fungal isolates using simple agar plate and SmF process. Bioresource Technology, 98: 946–949.

Nelson KE, Pell AN, Schofield P and Zinder SH (1995). Isolation and characterization of anaerobic ruminal bacterium capable of degrading hydrolyzable tannins. Applied and Environmental Microbiology, 61: 3293–3298.

Prusty AK, Sahu NP, Pal AK, Reddy AK and Kumar S (2007). Effect of dietary tannin on growth and haemato-immunological parameters of Labeo rohita (Hamilton) fingerlings. Animal Feed Science and Technology, 136: 96-108.

Vohra P, Kratzer FH and Joslyn MA (1966). The growth depressing and toxic effects of tannins to chicks. Poultry Science, 46: 135-142.