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Optimization of Tannase Production from Agro industrial wastes-Kinetics and Modeling

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CHAPTER-2

LITERATURE REVIEW

2.0 SOURCES OF TANNASE

Microbial source

A great variety of bacteria, Klebisella pneumonia (Descamps et al., 1983),

Citrobacter freundii (Kumar et al., 1999), Lactobacillus plantarum (Osawa et al.,

2000), Bacillus lichiniformis (Mondal et al., 2000), Lactobacillus sp.ASR-S1(Sabu

et al., 2006) some yeasts Candida sp (Aoki et al.,1976a,b), Debaryomyces hansenii

(Descamps et al.,1983) and Mycotorula japonica (Belmares et al., 2004) are

known as tannase enzyme producers. Mainly fungi are considered as the best

enzyme producers. Filamentous fungi of the Aspergillus and Penicillium genus

have been widely used for tannase production. Fungi like Aspergillus oryzae

(Bradoo et al., 1996) Aspergillus awamori (Beena et al., 2010), Aspergillus

fumigates (Batra and Saxena., 2005), Aspergillus ruber (Kumar et al., 2007),

Penicillium chrysogenum (Bradoo et al., 1996), Penicillium glabrum (Vande

Lagenaat and Pyle., 2005), Trichoderma viride and Trichoderma hamatum

(Bradoo et al., 1996) are reported as good tannase producers.

Plant source

Lekha and Lonsane., (1997) studied the production of tannase and reported

the presence in many tannin-rich plant materials, such as myrobolan (Terminalia

chebula) fruits, divi-divi (Caesalpinia coriaria) pods, dhawa (Anogeissus latifolia)

leaves and the bark of konnam (Cassia fistula), babul (Acacia arabica) and avaram

(Cassia auriculata) trees. Plant tannase is found to be less stable compared to

microbial sources and purification is more cumbersome.

Animal source

Sabu et al., (2006) reported many gastrointestinal bacteria of adopted

domestic and wild animals found to produce tannase and several species of these

bacteria have been isolated from faeces of koalas, goats and cows.


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2.1 TANNASE SUBSTRATES

Tannins are naturally occurring water-soluble polyphenolic compounds

with varying molecular weight that occur naturally in the plant kingdom. These

phenolic compounds differ from others by its ability to precipitate proteins from

solutions. In the plant kingdom these tannins are found in leaves, fruits, bark and

wood. They occur in many edible fruits and vegetables and are often considered

nutritionally undesirable because they form complexes with protein, starch and

digestive enzymes and cause a reduction in nutritional value of food (Chung et al.,

1998). Tannins are classified into two groups, hydrolysable tannins and condensed

tannins (Bhat et al., 1998). Current and the most accepted classification divides the

tannins into four groups namely gallo tannins, ellagi tannins, condensed tannins

and complex tannins (Fig.2.1.1). Tannins are in fact antimicrobial agents and most

of the microorganisms cannot tolerate its polyphenolic nature. Only a few

microorganisms can degrade tannic acid and utilize it as nutrient (Lekha and

Lonsane., 1997).

(a) Gallotannins are the simplest tannins that exist and are formed by units of

galloyl or di-galloyl esterified to a core of glucose or other polyhydroxy

alcohol. Upon hydrolysis, gallotannin such as Chinese gallotannin (Rhus

semilata) and sumac tannin (Rhus coriaria) yield glucose and phenolic

acids mainly gallic acid.

(b) Ellagitannins are esters of hexahydrodiphenic acid (HHDP) and during its

hydrolysis, the HHDP group dehydrates and lactonize spontaneously to

form ellagic acid. On hydrolysis, ellagitannins like myrobolan (Terminalia

chebula) tannin and divi-divi (Caesalpinia coriaria) tannin yield glucose

and ellagic acid together with gallic acid and frequently other acids

structurally related to gallic acid (Haslam et al., 1961).

(c) Condensed tannins are oligomeric and polymeric proanthocyanidins

containing flavan-3-ol (catechin) or flavan-3,4-diol linked by C–C– bonds.

In contrast to the hydrolysable tannins, they do not contain sugar residues

(Goodwin and Mercer., 1983) and are less susceptible to microbial and

chemical attack. Examples of condensed tannins include wattle (Acacia


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mollisima) tannin and quebracho (Schinopsis lorentzii) tannin (Lewis and

Starkey., 1969).

(d) The basic structure of complex tannins consists of a gallotannin or

ellagitannin unit and one of catechin (Belmares et al., 2004; Aguilera-

Carbo et al., 2008).

Fig.2.1.1 Classification of Tannins ( Khanbabaee and Van Ree., 2001)

2.1.1 Sources of Tannin

Tannins are natural polyphenolic compounds present in vascular plants.

They are characterized by their ability to form strong complexes with different

minerals and macromolecules, such as proteins, cellulose, starch, among others.

Due to their strong ability to bind with proteins, they have been used in tanning for

thousands of years (Aguilar et al., 2007). Application of agro industrial residues as

substrates is certainly economical and it also reduces environmental pollution.

Tannin are present in several naturally occurring agricultural wastes such as

redgram husk, greengram husk, blackgram husk, tamarind seed powder, tea dust,

rice bran and groundnut shell could be used in one or the other industrial

bioprocess for the production of value added products through Submerged

fermentation. Tannin content of the materials are listed in Table 2.1.


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Table 2.1 Sources of Tannin

Material Tannin content

(mg/g)

Reference

Redgram husk 2.601 Paranthaman et al., (2009)

Greengram husk 0.892 Prakasam et al., ( 2010)

Blackgram husk 0.910 Arulnathan et al., (2013)

Tamarind seed powder 1.202 Siddig et al., (2006)

Tea dust 0.102 Gowdhaman et al., (2012)

Rice bran 0.096 Paranthaman et al., (2009)

Groundnut shell 0.172 Paranthaman et al., (2009)

2.2 BIODEGRADATION OF TANNINS

Biodegradation by certain microorganism and enzymes is one of the most

efficient ways to degrade large tannin molecules into small molecules with bio-

activities of high value. The ability of microorganism to assimilate tannin differs

among yeast, bacteria and fungi. Yeast, while acting effectively against

gallotannins, loses it degradation ability against high molecular compounds.

Bacteria have the ability to degrade gallotannins as well as ellagitannins. Fungi can

efficiently degrade different types of tannins (Bhat et al., 1998).

Some of the enzymes involved in degradation of gallotannins are tannase

and gallic acid decarboxilase. Tannase is perhaps the most studied enzyme so far in

the biodegradation of tannins. It has act on ester and depside bonds of gallotannin

and may be microbial, plant, or animal in origin, of which microorganisms are the

most important source (Aguilar et al., 2007). Tannase acts on gallotannins,

ellagitannins, and complex tannins by breaking only ester bonds without affecting

the carbon–carbon bonds and hence does not affect the condensed tannins (Haslam

and Stangroom., 1966).

Gallic acid decarboxylase (E.C. 4.1.1.59) can catalyze the decarboxylation

of gallic acid to pyrogallol. This enzyme is very unstable because of its high

sensitivity to oxygen, and thus, it is difficult to isolate and purify (Zeida et al.,

1998).Some bacteria, such as selenomonas gallolyticus and Escherichia coli,

decarboxylate gallic acid to pyrogallol, but this compound is not further


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transformed. The reason is unclear but is likely to be less toxic than gallic acid or

that its production is thermodynamically more favorable and is possibly linked to

the generation of energy by pumping protons (Mingshu et al., 2006).

In the case of ellagitannin biodegradation, the release of ellagic acid has

been attributed to a new enzyme (ellagitannin acyl hydrolase). However, it is

necessary to perform a study to demonstrate the catalytic difference between tannin

acyl hydrolase and ellagitannin acyl hydrolase and to understand the

biodegradation processes of gallotannins and ellagitannins (Aguilera-Carbo et al.,

2008). On the other hand, the study of the degradation of condensed tannins and

complex is much more difficult due to their complicated structures. Therefore,

there is little progress in understanding the mechanisms of degradation of these

compounds. The biodegradation metabolic pathway of gallotannins is presented in

Fig. 2.2.1

Fig.2.2.1 Gallotannin enzymatic biodegradation pathways (Mingshu et al., 2006)


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2.2.1 Bacterial Degradation

Certain bacteria of the genera Bacillus, Staphylococcus, Klebsiella, and

Lactobacillus (Deschamps et al., 1980; Ayed and Hamdi., 2002) have the ability to

degrade tannins. Gallic acid monomers are used as a substrate after a break of

simple aliphatic acids. Gallic acid is converted to pyrogallol by the enzyme gallate

decarboxylase or gallate carboxylyase. The anaerobic decomposition of gallic acid

and hydrolyzable tannin monomer occurs by different mechanisms. The first step

is decarboxylation of gallic acid to form pyrogallol, which is converted to

phloroglucinol by pyrogallol phloroglucinol isomerase and to

dihydrophloroglucinol by phloroglucinol reductase. Later, dihydrophloroglucinol

is converted to 3-hydroxy-5- oxohexanoate (HOHN) by dihydrophloroglucinol

hydrolase. The HOHN is degraded by different pathways by rumen

microorganisms. Under anaerobic conditions, HOHN is converted to 3,5-

docosahexaenoate (triacetate) by the HOHN dehydrogenase and finally to three

molecules of acetyl-CoA by the sequential enzymatic action of triacetyl-CoA

transferase, triacetate ß-ketothiolase, acetoacetyl-CoA ß-ketothiolase,

phosphotransacetylase, and acetate kinase (Brune and Schink., 1992). HOHN can

be converted to acetate and butyrate, too, in rumen ecosystem. HOHN-CoA is

derived from the enzymatic action of HOHN-CoA transferase and is converted to

acetate and butyrate by the rumen bacterial by the sequential action of ß-

hydroxybutyryl-CoA dehydrogenase, butyryl-CoA dehydrogenase, acetyl-CoA

acetyltransferase, enoyl Co-A hydratase, phosphate acetyl transferase, and acetate

kinase (Brune and Schink., 1992; Nelson et al., 1995)

2.2.2 Fungal Degradation

The degradation of hydrolyzable tannins particularly gallotannins is best

known in fungal systems. The oxidative degradation of hydrolyzable tannins has

been studied in Aspergillus sp. (Cruz-Hernández et al., 2009; Belmares et al.,

2003). While routes of degradation of gallic acid have been determined, the

degradation pathway of hexahydroxydiphenyl and proanthocyanidins is still not

well understood.


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In Aspergillus niger, oxygenase converts gallic acid to unstable

intermediate of tricarboxylic acid. This is finally decarboxylated by an oxidative

decarboxylase to form cis-aconitic acid, which enters the citric acid cycle.

However, in Aspergillus flavus, gallic acid is degraded to oxaloacetic acid and

finally to pyruvic acid (William et al., 1986). Pyrogallol, the decarboxylated

derivative from gallic acid, is also oxidized to cis-aconitic acid by the same

mechanism.

The metabolism of hydrolyzable tannins, especially tannic acid has

received considerable interest (Ajay Kumar et al., 1999). It is well known that the

degradation of tannic acid by the action of the enzyme tannase releases gallic acid

and glucose. Attempts have been made to characterize the intermediate metabolites

of this action and found pyrogallol in addition to glucose and gallic acid. Ajay-

Kumar et al., (1999) reported the route of degradation of tannic acid by Citrobacter

freundii and observed that glucose formed by tannin hydrolysis enters glycolysis

and subsequently enters the tricarboxylic acid cycle. Gallic acid is converted to

pyrogallol by gallic acid decarboxylase. The presence of pyrogallol and resorcinol

has been observed in vitro studies with rumen systems (Singh et al., 2001).

Pyrogallol is converted to hydroxymuconic acid by pyrogallol dioxygenase

dioxygenase, which is later converted to pyruvate and it is further metabolized

through tricarboxylic acid cycle.

2.3 PHYSICOCHEMICAL PROPERTIES OF MICROBIAL TANNASE

Physicochemical properties of tannase, resumed in Table 2.3.1, are one of

the most discussed topics. Tannase properties depend on the source and culture

conditions; for example, all characterized tannases from yeast and fungi are

glycoproteins, but bacterial tannases seem not to present such post-translational

modifications. In addition, it has been found significant differences on

glycosylation between tannases produced in different culture systems by the same

microorganism (Renovato et al., 2011). Tannase glycosylations consists of neutral

sugars in the range from 5.4% to 64%, and it may be related to the resistance of the

TAH to be precipitated by tannins, as most of the proteins (Aoki et al., 1976b;

Beverini and Metche., 1990; Zhong et al., 2004; Kasieczka-Burnecka et al., 2007).


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2.3.1 Optimum temperature and temperature stability

The temperature dependence of many enzyme-catalyzed reactions can be

described by the Arrhenius equation. An increase in the temperature increases the

rate of reaction, since the atoms in the enzyme molecule have greater energies and

a greater tendency to move. However, the temperature is limited to the usual

biological range. As the temperature rises, denaturation processes progressively

destroy the activity of enzyme molecules. This is due to the unfolding of the

protein chain after the breakage of weak (hydrogen) bonds, so that the overall

reaction velocity drops. For many proteins, denaturation begins to occur at 45°C to

50°C. Some enzymes are very resistant to denaturation by high temperature,

especially the enzymes isolated from thermophilic organisms found in certain hot

environments (Rajiv Dutta., 2008).

Batra and Saxena., (2005) reported that the temperature range of the

tannase production as 30°C–70°C and the optimum temperature as 60°C for

A.flavus, A.fumigatus, A.versicolor and P. variable, whereas A. caespitosum, P.

charlesii, P. crustosum and as 40°C for P. restrictum. Tannase obtained from A.

versicolor was more stable in a broad temperature range of 30°C –80°C.

Hina Iqbal et al., (2012) studied the effect of temperature on tannase

enzyme production using T.harzianum in the temperature range from 25°C to 60°C

and the optimum temperature was found to be 40°C and was stable at 40ºC

retaining about 71% of original activity for 2 hours. Costa et al., (2008) obtained

similar results for tannase enzyme production using A.tamari in submerged culture

fermentation and the optimum temperature was found to be 35ºC and was stable at

40ºC for 2 hours.

Banerjee et al., (2007) investigated the possible use of wheat bran as

substrate for production of tannase and reported a maximum tannase activity of

8.16 U/g at 30°C by Aspergillus aculeatus DBF9. Kumar et al., (1999) used a

mineral medium with a maximum tannic acid concentration of 5% (w/v) in the

production of tannase using Citrobacter freundii and reported a maximum tannase

activity of 12.16 U/ml at 30°C. Selwal et al., (2010) studied the optimization of

process parameters for tannase production using Pseudomonas aeruginosa IIIB


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8914 and reported a maximum tannase activity of 13.16 U/ml at 37°C. Bradoo et

al., (1997) reported that the production of tannase using Aspergillus japonicas was

found to be 33.06 U/ml as maximum tannase activity with 0.2% glucose and 2%

tannic acid in Czapek- Dox's Minimal Medium at pH 6.6.

Renovato et al., (2011) studied the production of tannase using A.niger

under solid state fermentation and submerged fermentation. The optimum

temperature and pH range were found to be from 50°C to 60°C and 5–8

respectively in submerged fermentation. The optimum temperature and pH were

found to be 60°C and 6 respectively in solid state fermentation.

2.3.2 Optimum pH and Stability

All proteins are constructed from various amino acids. These biochemical

units possess basic, neutral or acidic groups. Consequently, the intact enzyme may

contain both positively or negatively charged groups at any given pH. Such

ionizable groups are often apparently part of the active site since acid- and base-

type catalytic action has been linked closely to several enzyme mechanisms. For

the appropriate acid or base catalysis to be possible, the ionizable groups in the

active site must often each possess a particular charge; i.e., the catalytically active

enzyme exists in only one particular ionization state. Thus, the catalytically active

enzyme may be a large or small fraction of the total enzyme present, depending

upon the pH (Bailey and Ollis., 1944).

Aoki et al., (1976 a) studied the production of tannase from Candida sp.

K-1 and showed an optimum activity at a pH value of 6.0. The investigation also

revealed that the enzyme was stable over a wide pH range of 3.5 to 7.5. Rajakumar

and Nandy.,(1983) reported the isolation and purification of tannase from

Penicillium chrysogenum and showed broad pH dependence in the range from 5 to

6, with the optimum enzyme activity and was stable in the pH range of 4.0 to 6.5 at

16°C.

Iibuchi et al., (1968) reported the effect of pH on tannase production using

A. oryzae and the tannase was stable in the pH range of 4.5 to 6.0 for 25 hours with

an optimum pH of 5.5. Anwar et al., (2009) studied the production and

characterization of tannase using A.niger isolated from cacao pod in a solid state


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fermentation with wheat flour as a substrate and obtained a maximum tannase

activity of 10.63 U/ml at pH 6.0. Niehaus and Gross., (1997) studied the tannase

production using penduculate oak as a substrate and the optimum pH was found to

be 5.0.

Mahendran et al., (2006) studied the purification and characterization of

tannase using P. variotii and the enzyme was stable at a pH from 5 to 7 and the

optimum pH was found to be pH 6. Mahapatra et al., (2005) reported the

production of tannase using A.awamori nakazawa and exhibited optimum activity

at 35°C and at a pH of 5.

2.3.3 Molecular Weight of Tannase

The molecular weight of tannases was found to be in the range of 50 kDa –

320 kDa (Iwamoto et al., 2008; Boer et al., 2009). It has been reported that

tannases consist of two or more subunits. Hatamoto et al., (1996) concluded that

native tannase of Aspergillus oryzae consists of four pairs of two types of subunits

(30 kDa and 34 kDa, respectively) linked together by disulfide bonds, forming a

hetero-octamer of 300 kDa. Mahendran et al., (2006) reported the molecular

weight of tannase using Pacilomyces variotii was reported to be 149.8 kDa through

native PAGE analysis with a monomeric unit of molecular mass 45kDa by SDS-

PAGE analysis. Renovato et al., (2011) studied the SDS-PAGE analysis of both

SSF and SmF purified tannases showed a single band with a molecular weight of

102 kDa and 105 kDa, respectively.

2.3.4 Kinetic Parameters of Tannase

Substrate affinity for tannase from several fungi has been found to be

different. The Km values were 0.28, 0.95, 1.05, and 0.048 for tannase from A.

niger, Cryphonectria parasitica, Verticillium sp., and Penicillium chrysogenum,

respectively, when tannic acid was used as substrate and reaction was carried out at

30°C and pH 5.0–6.0 (Bhardwaj et al., 2003; Farías et al., 1994; Kasieczka-

Burnecka et al., 2007; Rajakumar and Nandy., 1983). However, caution should be

taken when comparing these values due to the varying quality of the substrates

utilized and the different methods utilized to quantify the reaction product. Anwar

et al., (2009) reported the kinetics analysis that showed that Km and Vmax value of


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tannic acid was 0.401 mM and 10.804 U/ml respectively and Gallotannin yielded

Km and Vmax values of 6.611 mM and 12.406 U/ml respectively. Battestin and

Macedo., (2007) reported the Km and Vmax values were found to be 0.61 μmol and

0.55 U/ml with wheat bran and coffee husk as used as the substrate.

2.3.5 Effect of Carbon Sources

Selwal et al., (2011) studied the effect of carbon sources on production of

tannase using tannic acid, dextrose, glucose, glycerol, maltose, manitol, lactose and

sucrose. The strain P.atramentosum KM produced maximum tannase of 29.4 U/ml

with amla leaves and 31.1 U/ml with keekar leaves as substrates when

supplemented with 0.2% (w/v) maltose. Lokeswari et al., (2007) studied the effect

of different glucose concentrations on tannase production using Aspergillus niger.

A maximum tannase activity of 21.42 U/ml with 0.5% (w/v) glucose concentration

was reported.

Deepanjali Lal et al., (2012) studied production of tannase using Aspergillus

niger with different carbon sources namely tannic acid, mannose, galactose,

glycerol and ribose. Tannic acid (1% w/v) was the most suitable carbon source for

tannase production.

2.3.6 Effect of Additives

The effect of additives on tannase activity has been studied by several

authors. Kar et al., (2003) studied the effect of metal ions on a Rhizopus oryzae

tannase. They found that 1 mM of Mg+2

or Hg+ activated tannase activity, whereas

that Ba+2

, Ca+2

, Zn+2

, Hg+2

, and Ag+ inhibited tannase activity, and Fe

+3 and Co

+2

completely inhibited tannase activity. On the other hand, the tannase from A. niger

GH1 was highly inhibited by Fe+3

, whereas Cu+2

and Zn+2

had only a mild

inhibitory effect, and Co+2

enhanced the enzyme activity (Mata-Gómez et al.

2009). Furthermore, Mg+2

, Mn+2

, Ca+2

, Na+ , and K

+ stimulated the activity of

Aspergillus awamori tannase, while Cu+2

, Fe+3

, and Co+2

acted as inhibitors of the

enzyme (Chhokar et al., 2010a).


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Table 2.3.1 Physicochemical Properties of Tannase

Microorganism Optimum

pH and

Optimum

Temperature

pH,

Temperature

Stability

Other properties Reference

Aspergillus

niger van

Tieghem

pH 6.0, 60°C Stable at pH

3.0 –8.0,

temperature

30 °C – 60°C

Km = 0.20 mM,

Vmax = 5.0 μmol

min−1

mg−1

protein

Sharma

et al., (1999)

Aspergillus

niger ATTC

16620

pH 6.0, 40°C Enzyme was

active at pH

4.0-8.0,

30°C – 40°C

Enzyme activity

was inhibited by

Zn2+

, Mn2+

, Cu2+

,

Ca2+

, Mg2+

and

Fe2+

, and enhanced

by K+.

Sabu et al.,

(2005)

Aspergillus

niger GH1

pH 6.0, 60 °C Stable at pH

3.0–6.0,

stable at

temperatures

lower than

50°C

The enzyme was

highly inhibited by

Fe3+

, whereas Cu2+

and Zn2+

had only a

mild inhibitory

effect. Co2+

enhanced the

enzyme activity.

Mata Gomez

et al., (2009)

A.niger LCF 8 pH 6.0, 35 °C Stable at pH

3.5 –8.0,

temperature

4 °C – 45°C

Molecular weight-

186000 kDa

Inhibitor- CuSo4

(68%),

ZnCl2 (39%)

Barthomeuf et

al., (1994)

Candida sp.

K 16

pH 5.5, 50 °C Stable at pH

3.5 –7.5,

temperature

4 0°C

Molecular weight-

250000 kDa

Protein content-

35%

Aoki et

al.,(1976)

Aspergillus

aculeatus

DBF 9

pH 5.0, 60°C

for

intracellular

enzyme,

50 °C for

extracellular

enzyme

Stable at pH

4.0–6.0,

Stable for 1 h

at 50°C

Both intracellular

and extracellular

enzymes are salt

tolerant up to 3

M NaCl

Banerjee

et al., (2007)

Penicillium

variable

pH 5.0, 50°C Active at pH

3.0–8.0,

25–80°C

Mol.wt- 2 kDa

Km = 32 mM,

Vmax = 1.1 μmol

min−1

ml−1

Sharma

et al ., (2008)


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Pencillium

chrysogenum

pH 5.0-6.0

30°C -40°C

Stable at

pH 4.0–6.5,

stable

temperature

upto 30°C

Km = 0.48 x 10-4

M Rajakumar

and Nandy.,

(1983)

Bacillus cereus

KBR9

pH 4.5, 40°C Stable at

pH 4.5–5.0,

stable

temperatures

lower than

30°C

Enzyme is salt

tolerant, stable up

to 2 M of NaCl and

retains 82%

original activity in

3 M.

Mondal

et al., (2001)

Lactobacillus

plantarum

CECT 748T


22°C –37°C

Enzyme activity

was not affected by

K+, Ca

2+, Zn

2+,

Tween 80, EDTA

and urea, but

inhibited by Hg2+

Rodríguez

et al., (2008)

Lactobacillus

plantarum

ATCC 1491 T

(recombinant)

pH 8.0, 40°C Active at

pH 6.0–8.0,

stable at

temperatures

lower than

30°C

The enzymatic

activity was

increased by K+

and Ca2+

, not

significantly

affected or partially

inhibited by EDTA,

Mg2+

, Zn2+

, Triton-

X-100, Tween-80

Curiel et al.,

(2009)

Paecilomyces

variotii


pH 4.0–8.0,

30°C –50°C

Mol.wt- 45 kDa

Mahendran

et al., (2006)

A.oryzae pH 5.5,

30°C - 40°C

Stable at

pH 3.0–7.5,

30°C

Molecular weight-

200000 kDa

Inhibitor-

Cu and Zn

Iibuchi et

al.,(1972)


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2.4. STRUCTURE OF TANNASE

Enzyme name : Tannase

Enzyme Commission number: EC 3.1.1.20

Structure : Crystalline structure

Secretion : Extracellular

Reaction : digallate + H2O = 2 gallate

Other name(s) : Tannase S, Tannin acetylhydrolase

Systematic name: Tannin acylhydrolase

Comments: Also hydrolyses ester links in other tannins.

Fig.2.4.1 Structure of Tannase

2.5 PRODUCTION OF TANNASE

Studies on tannase production by bacteria, fungi and yeast have been

carried out under submerged and solid state fermentation conditions. Depending on

the strain and the culture conditions, the enzyme was induced and expressed with

different levels of activity, showing different production patterns. Tannase

synthesis was induced by phenolic compounds such as methyl gallate, gallic acid,

pyrogallol, and tannic acid (Rana and Bhat ., 2005).

2.5.1 Tannase Production in Submerged Fermentation

Submerged fermentation is always preferred for microbial tannase

production because it offers uniform fermentation conditions like substrate


22

concentration, inducer concentration, temperature, pH, dissolved oxygen

concentration, agitation and aeration. Microbial tannase production belongs to

aerobic fermentation and submerged fermentation is a better choice to provide

uniform aeration conditions. Sometimes solid state fermentation methods are

adopted for tannase production since cheaper agricultural residues based media

could be used. The disadvantages of solid state fermentation are difficulties in

scale up, control of process parameters (pH, heat, moisture, nutrient conditions

etc.,) and quality of the product.

Hina Iqbal et al., (2012) studied the production of tannase using

Trichoderma harzianum MTCC 10841 under submerged fermentation with rich

tannin materials like amla (Phyllanthus amblica, bark, leaves and fruit), amaltash

(Cassia fistula, leaves), ber (Zyzipus maurtiana, leaves), Eucalyptus (Eucalyptus

glogus, bark and leaves), jamun (Syzgium cumini, bark and leaves), guava (Psidium

guazava, bark and leaves), keekar (Acacia nilotica, leaves), mango (Magnifera

indica, leaves), mulberry (Morus macroura, leaves), tamarind (Tamarindus indica,

seed) and pomegranate (Punica granatum, rind) as carbon sources. Amla fruit,

tamarind seed, jamun leaves, mulberry leaves and keekar leaves proved to be best

substrates than tannic acid and the optimum pH and temperature for the tannase

enzyme production were found to be 5.5 and 40ºC respectively.

Das Mohapatra et al., (2006) used eight different plant extracts as the

tannin source for tannase production using Bacilus lichiniformis KBR6 under

submerged fermentation and obtained highest activity from the crude extract of

Anacardium occidentale. Darah et al., (2011) reported the production of tannase

by submerged fermentation using Aspergillus niger FETL FT3 with an initial

medium pH of 6.0 at 30ºC, agitation speed of 200 rpm and inoculums size of

6 x 106 spores/ml. Maximum tannase production of 2.81 U/ml was obtained on

the fourth day of cultivation.

Selwal et al., (2010) studied the production of tannase enzyme using

Pseudomonas aeruginosa IIIB 8914 under submerged fermentation with the leaves

of Phylanthus emblica (amla), Acacia nilotica (keekar), Eugenia cuspidate (Jamoa)

and Syzygium cumini (Jamun) as substrates and reported a maximum tannase yield


23

of 13.65 U/ml and 12.90 U/ml were obtained using amla and keekar leaves

respectively.

Wilson Peter et al., (2009) used a bacterial isolate, Citrobacter sp., from

tannery effluent has proved as a potent producer of tannase enzyme. The

production by solid state fermentation of tannase was compared with submerged

fermentation using tamarind seed as sole carbon source. Two times higher tannase

activity was observed in solid state fermentation (90 U) than submerged

fermentation (50 U) at 48 hours with 5 g of substrate.

Murugan et al., (2007) isolated 10 morphological different fungal strains

from a tannery effluent. Selected microorganisms were tested for tannase

production under submerged fermentation in a stirred tank bioreactor and reported

strain A.niger MS101 as the best tannase producer.

Sharma et al., (2007) studied the optimization of tannase production by

statistical techniques using Aspergillus niger in submerged fermentation. The

effect of concentrations of tannic acid, sodium nitrate, agitation rate and incubation

period on tannase production was studied. 5% tannic acid, 0.8% sodium nitrate,

150 rpm agitation and 48 hours were found to be optimum conditions and gave a

maximum tannase activity of 19.7 U/ml.

Beniwal et al., (2010) reported the optimization of culture conditions for

tannase production by Aspergillus awamori MTCC 9299 using the response

surface methodology. Maximum yield of tannase production was obtained at pH of

5.0, incubation temperature of 35°C, agitation speed of 125 rpm and 48 hours of

incubation period. Under the proposed optimized conditions, the tannase

experimental yield of 1.45 U/ml was closely matched the yield predicted by the

statistical model of 1.43 U/ml with R2 value of 0.99.

Kannan et al., (2011) studied the production of tannase by Lactobacillus

plantarum MTCC 1407 under submerged fermentation. Maximum tannase activity

of 5.22 U/ml was obtained at 24 hours using the following medium (g/L): tannic

acid-10: glucose-1; NH4Cl-3; MgSO4.7H2O-2; KH2PO4 - 0.5; K2HPO4 - 0.5;

CaCl2-1.


24

The summary of the production of tannase enzyme using various microorganisms

by submerged fermentation was given in Table.2.5.1

Table.2.5.1 Tannase Production by Submerged Fermentation

Strain Substrate Process conditions Enzyme

activity Reference

A.japonicus Tannic acid

pH -6.6

Temperature -35°C

Incubation time-72 hrs

Substrate conc -5%

Tannic acid conc -3%

33.06 U/ml Bradoo et al.,

(1996)

Citrobactor

species

Tamarind

seed powder

pH -5.5

Temperature- 35°C

50 U

90 U

Wilson Peter

et al., (2009)

A. niger Tannic acid pH -5.5

Temperature -35°C 120 U/ml

Lokeswari

et al., (2007)

A.oryzae 643

(NCIM)

Cashew

husk

pH -5

Temperature- 40°C

Incubation time -24hrs

32.62 U/ml Lokeswari

et al., (2010)

A. niger Pomegra

nate rind

pH -5

Temperature - 37°C


28.72 U/ml Srivastava et

al., (2009)

A.niger Tannic acid

pH -6

Temperature -50°C

Incubation time - 96hrs

65 U/ml Lokeswari

et al., (2006)

A.flavus Tannic acid

Temperature - 35°C

Incubation time - 96hrs

Tannic acid conc - 3%

30 U/g/min Paranthaman

et al., (2009)

Trichophyton

rubrum

MTCC

Tannic acid

pH-5.5

Temperature -30°C

Incubation time – 30hrs

Tannic acid conc -2.5%

36.54

U/g/min

Krishnasamy

et al., (2009)

A. awamori Tannic acid pH -5.5

Temperature - 30°C 1.43U/ml

Chhokar

et al.,

(2010a)

P. aeruginosa

Amla

Ber

Jamun

Jamoa

Keekar

pH -5

Temperature -35°C


13.65U/ml

12.90U/ml

Manjit

et al., (2010)

A.niger Tannic acid

pH -5

Temperature -30°C

Incubation time -

168hrs

101.428

U/ml

Deepanjali

Lal et al.,

(2012)


25

2.5.2 Tannase Production in Solid State Fermentation

Sabu et al., (2005) studied the production of tannase under solid-state

fermentation using A. niger ATCC 16620 with palm kernel cake and tamarind seed

powder as the substrate. A maximum enzyme yield of 13.03 IU/g dry substrate

(gds) was obtained at 30°C and 5% tannic acid as additional carbon source after 96

hours of fermentation.

Kumar et al., (2007) investigated tannase production using A. ruber by

solid state fermentation using different tannin rich agro-wastes namely ber leaves

(Zyzyphus mauritiana), jamun leaves (Syzygium cumini), amla leaves

(Phyllanthusemblica) and jawar leaves (Sorghum vulgaris). Jamun leaves were

found to be the best substrate for tannase production and a maximum production of

tannase 30.2 U/ml was recorded at 30.1ºC after 96 hours of incubation.

Bhaskar Reddy and Vandhana Rathore., (2012) studied the production of

gallic acid using tannase enzyme. Tannase enzyme was produced using acacia

pods, redgram husk, sorgum husk and spent tea powder as substrates through solid

state fermentation by the isolate P.purpurogenum BVG 7.The effect of pH and

temperature on tannase production was studied and pH 5.5 at 30 ºC were found to

be optimum conditions for maximum tannase activity and gallic acid production.

Battestin et al., (2007) studied the production of tannase using

Paecilomyces variotti with coffee husk and rice bran as the substrate for solid state

fermentation and evaluated the effects of variables namely temperature (⁰C), tannic

acid (%), residue (%) (coffee husk: wheat bran) and incubation time on the

production of tannase. The optimum conditions for tannase production by

Response Surface Methodology were found to be temperature (29⁰C –34⁰C), tannic

acid (8.5–14%); % residue (coffee husk:wheat bran 50:50) and incubation time of

5 days.

Kulkarni et al., (2012) studied the tannase production using A.oryzae with

mixture of Arjuna (Terminalia arjuna), Babul (Acacia nilotica), Jamun (Syzygium

cumini), Mulberry, Chiku stem barks as substrates by solid state fermentation. The

mixture of Jamun and Babul bark in the ratio of 4:6 was reported to be best

substrate and gave a maximum tannase activity of 116 U/g dry substrate.


26

Aguilar et al., (2001) reported the production of tannase using Aspergillus

niger Aa-20 in solid state fermentation and submerged fermentation with tannic

acid and glucose as carbon sources. The biomass yield in solid state fermentation

was found to be 2 times higher than in submerged fermentation.

Pinto et al., (2001) investigated the tannase activity of 17 wild type and 13

mutant strains of Aspergillus niger from a local culture collection in Brazil

(EMBRAPA / Food Technology stock collection) and selected the potential

tannase producers for maximum tannase production by solid state fermentation.

Paranthaman et al., (2009) did a comparative study on the suitability of

different substrates for maximum tannase production using A.oryzae in solid state

fermentation using various agricultural by-products. Rice bran was reported to be

the best substrate for the tannase enzyme production and gave a maximum tannase

activity of 14.40 U/g/min at 30⁰C, pH 5.5 in 96 hours.

Manjit et al., (2008) studied the production of tannase by solid-state

fermentation using Aspergillus fumigatus MA with different agro forest residues

such as Amla leaves (Phyllanthus emblica), Ber leaves (Zyzyphus mauritiana),

Jamun leaves (Syzygium cumini), Jamoa leaves (Syzygium sp.) and Keekar leaves

(Acacia nilotica) as substrates. The maximum tannase activity of 174.32 U/g was

obtained using Jamun leaves as substrate at 25⁰C, pH 5 in 96 hours of incubation.

Rodrigues et al., (2008) studied the effects of inoculum concentration,

temperature, and carbon sources on tannase production by solid state fermentation

with cashew apple bagasse. The maximum tannase activity of 4.63 U/g of dry

substrate was obtained at 30°C, using 107 spores/g with 1.0% (w/v) sucrose as an

additional carbon source.

Narsi reddy et al., (2011) reported tannase production using red gram husk,

green gram husk, ground nut waste, cotton seed waste, wheat bran, rice bran,

coffee husk, tamarind seed powder, cashew apple bagasse, coconut powder, corn

powder and cyceraritinum as substrates by solid-state fermentation using isolated

A.terreus. A maximum tannase activity of 41.6 U/mg was reported with wheat bran

as a substrate at pH of 3.5 and incubation period of 72 hours.


27

Kannan et al., (2012) studied production of tannase using Lactobacillus

plantarum MTCC1407 in submerged fermentation and solid state fermentation

process. They optimized the nutrients using Plackett–Burman screening and

response surface methodology. Maximum tannase activity of 9.13 U/ml was

observed at 30 hours of fermentation. Solid state fermentation was conducted with

various solid substrates and agricultural residues. Maximum tannase activity of

5.319 U/gds was obtained with coffee husk as substrate.

Boer et al., (2011) studied the large scale production of tannase using

Arxula adeninivorans strains in Fed-Batch fermentation. Transformant strains that

overexpress the ATAN1 gene from the strong A. adeninivorans TEF1 promoter

produce levels of up to 1,642 U / l when grown in glucose medium in shake flasks.

The effect of Fed-Batch fermentation on tannase productivity was found to be

51,900 U / l after 142 hours of fermentation at a dry cell weight of 162 g / l.

Zhong et al., (2004) cloned and expressed the tannase gene using A.oryzae

in the methylotrophic yeast Pichia pastoris and obtained large quantities of

extracellular tannase was found to be 7,000 U/l in a short period of 96 hours

incubation in a SmF Fed-Batch system.


28

The production of tannase using various microorganisms by solid state fermentation

was summarized in Table.2.5.2

Table.2.5.2 Tannase Production by Solid State Fermentation

Strain Substrate Process

Conditions

Enzyme

Activity Reference

A.oryzae

MTCC

Sugarcane

bagasse and

rice straw

(mixed)

pH - 5.5

Temperature -

35°C

Incubation time -

72 hrs

60.5 U/g/min Paranthaman

et al., (2008)

A.niger Redgram

husk

Temperature –

35°C

Incubation time -

96 hrs

43 U/g/min

Paranthaman

et al., (2009

b)

A.oryzae

MTCC 1122

Sugarcane

beggasse

and

ricestraw

(mixed)

pH - 5.5

Temperature -

30°C

Incubation time -

72 hrs

Tannic acid conc-

0.06%

121 U/g/min

Paranthaman

et al.,

(2009 c)

P.atramen

tosum KM Amla

pH – 5.5

Temperature -

28°C

Incubation time -

96 hrs

170.75 U/gds

Manjit

et al.,

(2011)

A.niger ATCC

16620

Tamarind

seed powder

& Palm

kernel cake

pH - 6

Temperature -

30°C

Incubation time -

96 hrs

Tannic acid

conc.-5%

13.03 IU/g Sabu et al.,

(2005)

A.oryzae

MTCC 634

Sugarcane

beggasse

and rice

straw

(mixed)

pH-5.5

Temperature -

30°C

Incubation time-

72 hrs

7.8 U/g/min Paranthaman

et al., (2010)

A. fumigates

Amla Ber

Jamun

Jamoa

Keekar

pH -5

Temperature -

25°C

Incubation time

96hrs

174.32 U/g Manjit

et al., (2008)


29

Strain Substrate Process

Conditions

Enzyme

Activity Reference

A. niger Tannic acid

pH -5.5

Temperature -

35°C

Incubation time -

36hrs

120U/ml Lokeswari

et al., (2007)

A. flavus

P.chrysogenum

T.viride

Paddy straw

pH -5.5

Temperature -

30°C

Incubation time -

96hrs

16 U/g/min

17 U/g/min

19 U/g/min

Paranthaman

et al., (2010)

A.niger Wheat flour

pH -6

Temperature -

50°C

10.8U/ml

Anwar

et al., (2009)

A.oryzae

MTCC

Paddy husk

Rice bran

Millet husk

Groundnut

shell

pH -5.5

Temperature -

35°C

Incubation time -

96hrs

4.14 U/g/min

14.4 U/g/min

7.41 U/g/min

11.4 U/g/min

Paranthaman

et al., (2009)

Lactobacillus

species

Wheat bran

Palm kernel

cake

Tamarind

seed

Coffee husk

Tannic acid conc -

0.6%

Temperature -

35°C

Moisture content -

44ml

Incubation time -

60 hrs.

0.35U/gds

0.51 U/gds

0.05 U/gds

0.21 U/gds

Sabu et al.,

(2006)

A.aculeatus

DBF9

Wheat bran

Saw dust

Rice bran

Sugarcane

pith

Rice straw

dust

pH -5.5

Temperature -

30°C

Incubation time -

72hrs

Tannic acid conc.

-5%

3.95 U/g

1.87 U/g

2.93 U/g

1.32 U/g

1.39 U/g

Banerjee

et al., (2007)

A.oryzae

Cashew

apple

baggasse

pH -5.5

Temperature -

30°C

Incubation time -

48 hrs

Tannic acid conc.

-2.5%

0.146

U/gds/hr

Rodrigues

et al., (2008)

A.oryzae

Cashew

apple

baggasse

Initial water

content -60ml

Tannic acid conc.

-2.5%

0.56U/gds Rodrigues

et al., (2007)


30

2.6 METABOLIC REGULATION OF TANNASE PRODUCTION

Depending on the strain and fermentation conditions, the tannase can be

produced either constitutively or by substrate induction. Knudson (1913) reported

that tannase production only occurs in the presence of tannic acid, resulting in the

formation of gallic acid and glucose as final products. Seiji et al., (1973) observed

tannase production when the microorganism was grown on glucose as sole carbon

source. Bradoo et al., (1997) showed that Aspergillus japonicus produced tannase

constitutively when grown in a simple culture medium with simple or complex

sugars, but the production of the enzyme was doubled when grew up with tannic

acid as sole carbon source.

Bajpai and Patil., (1997) studied the effect of different substrates such as

gallic acid, pentagalloyl glucose, methyl gallate, and pyrogallol as inducers of the

tannase activity in four species of filamentous fungi, viz., Aspergillus niger, A.

fischerii, Fusarium solani, and Trichoderma viride. They reported that only the

species that were able to grow in the presence of pyrogallol produced the enzyme.

They further observed that each species responded differently to each substrate. A.

fischerii was induced mostly by gallic acid, F. solani by gallotannin, and T. viride

by methyl gallate.

The regulation mechanism of TAH is still unclear, and there exist some

controversies about the specific role of some compounds in the induction and

repression of its expression. It is generally accepted that tannic acid cannot act

directly as an inducer; the molecule is very large and reactive to penetrate the cell

membrane of microorganisms. Gallic acid, which has been used as inducer for the

production of tannase (Bajpai and Patil., 1997), has also been linked to its

regulation (Bradoo et al., 1997). This suggests that the production of TAH is

induced by intermediate compounds produced during the hydrolysis of tannins due

to the action of constitutively produced tannase.

The regulation mechanisms of tannase are very different in SSF and SmF.

Aguilar et al., (2001a,b) studied the induction and repression patterns of production

of tannase in both systems. They found that the addition of tannic acid at

concentrations higher than 25 g/L strongly inhibited the production of tannase in


31

SmF, whereas in SSF, the enzyme was produced in tannic acid concentrations up

to 200 g/L. Furthermore, the addition of small amounts of glucose (12.5 g/L)

increased the activity titles of SSF systems, but there was very less effect in SmF.

Higher concentrations of glucose resulted in a strong catabolite repression in SmF

but had a less effect on SSF. They further reported that the tannase production was

lower than baseline levels of activity with gallic acid as sole carbon source.

2.7 TANNASE RECOVERY

Recovery of the enzyme depends on the production system used and the

time of extraction. In SSF systems, the recovery of tannase is easier due to the fact

that it is extracellular. The addition of two or three volumes of extracting agent

(distilled water or buffer) to mix and compress facilitates the extraction of the

enzyme. However, in the case of the SmF, the localization of enzyme depends on

the time of cultivation. In general, at the beginning of fermentation, the enzyme

was found to be intracellular and this implies that the recovery involves cell

disruption. The cell disruption can be achieved by enzymatic treatment or

mechanical breakdown with mortar or a homogenizer. The recovery is simple;

after cell disruption, it can be achieved by removing cell debris by filtration or

centrifugation. However, at the moment of maximal production, over 80% of the

enzyme remains bound on the mycelium, hindering its extraction ( Barthomeuf et

al., 1994; Bhardwaj et al., 2003).

2.8 TANNASE PURIFICATION

Enzyme purification is performed to increase their catalytic power, improve

stability, or prevent unwanted reactions. There are numerous studies on total and

partial tannase purification. Common protocols of two or three steps include

fractional precipitation (with salts, organic solvents, tannic acid, or pH), ion

exchange or gel permeation chromatography. Other less common steps are affinity

chromatography ( Beverini and Metche., 1990) and preparative scale isoelectric

focusing ( Ramírez-Coronel et al., 2003). The purification process is affected not

only by the type of method used but also by the order in which they are combined.


32

For example, Bhardwaj et al., (2003) increased the yield from 2.7% to 51% on the

purification protocol for the tannase from A.niger van Tieghem by simply inverting

the order of the steps previously used by Sharma et al., (1999).

Enzymes products are available as crude, dried preparations, dilute or

concentrated liquids, or purified solids. Fig.2.8.1 provides a general process

recovery scheme for enzyme derived from animal, plant, surface, or submerged

fermentations. The former source requires immediate pretreatment to release

enzyme into an extracting buffer, followed by the appropriate solids removal steps

when liquid or purified products are required. A more detailed process recovery of

a plant enzyme requires the necessity of good mixing at a low temperature to

maximize initial extraction. The two serial centrifuges perform a solids

fractionation, removing large particles first by scroll centrifugation so that the

more expensive, higher rpm bowl centrifuge is not clogged with large particles.

Subsequent acidification shift pH sufficiently to precipitate much originally soluble

protein, provided a sufficient residence time is allowed in the cooled holding coil

to form a centrifugal precipitate. A disk centrifuge removes this protein precipitate;

a second acidification and holding coil precipitate the desired protein, recovered as

wet solid from the second disc centrifuge. Thus recovery of a plant enzyme

contains two instances where similar or identical processes are placed serially to

carry out a fractionation, first of solids by centrifugation and second by

acidification/ precipitation.

As subsequent smaller scale operations occur in protein purification,

recovery steps may logically shift from continuous to batch as shown in Fig.2.8.2

for enzyme production. Note additional steps to enhance enzyme yield:

(1) repeated washing of biomass (2) two stage ultra filtration to carry out

sequential 5 fold volume reductions, (3) a switch from continuous to batch ultra

filtration to effect a further 40-fold volume reduction.


33

Fig.2.8.1 Preparation of Commercial Enzymes


34

Fig.2.8.2 Extracellular Enzyme Recovery


35

2.9 TANNASE MOLECULAR EXPRESSION

Molecular biology techniques are being employed for better understanding

of tannase and improvement of the yield production. Hatamoto et al., (1996)

cloned and sequenced the gene from tannase of A.oryzae and reported the absence

of introns in its structure. They found that the gene encodes for a sequence of 588

amino acids with a signal sequence of 18 amino acids and a molecular weight of 64

kDa, approximately. This research group hypothesized that the tannase consists of

two subunits 30 kDa and 33 kDa linked by a disulfide bond. The gene of tannase

was transcribed as a single polypeptide chain, after which the signal of 18 amino

acids was cut, and the polypeptide chain was divided in two subunits. It was

reported that the single polypeptide chain was cut by KEX-II like protease in two

amino acid chains. They concluded that native tannase consisted on four pairs of

two subunits, forming a heterooctamer with molecular weight around 300 kDa.

Cerda-Gomez et al., (2006) used conserved sequences of tannase gene from

different species of the genus Aspergillus to design a set of primers (Tan 1 and Tan

2) and used it to amplify a DNA fragment of 435 bp by PCR from four different

species of Aspergillus sp. The amino acid sequence of the tannase gene from

A. niger showed 71% of identity and 10.19% of similarity with A. oryzae. Zhong et

al., (2004) cloned and expressed the tannase gen from A.oryzae in the

methylotrophic yeast Pichia pastoris and obtained large quantities of extracellular

tannase (7,000 U/L) in a short period of incubation (96 hr) in a SmF fed-batch

system. The tannase was purified to homogeneity by a two-step protocol to achieve

a purification factor close to 3.5 with a yield of 51%. Recombinant tannase has

31% more sugar molecules than the native enzyme (22.7%). Although the native

and recombinant enzymes have different degrees of glycosylation, both presented

similar specific activity.

On the other hand, Curiel et al., (2009) reported the production and

purification of a recombinant Lactobacillus plantarum expressed in E. coli. The

tannase gene was inserted with an aminoterminal His-tag that allowed the

convenient purification of the native protein directly from the crude cell extracts.


36

In fact, using this methodology, they were able to obtain large amounts of pure

tannase (17 mg/L) by a one-step affinity procedure.

2.10 MECHANISM OF ACTION

Although the tannins are known as protein precipitants, the tannase acts

over these compounds hydrolyzing the ester bonds formed between galloyl groups

and a polyhydroxy alcohol or the depside link between two galloyl groups (Aguilar

and Gutierrez-Sánchez., 2001; Kasieczka-Burnecka et al., 2007). It has been

proposed that the tannase has a depsidase and esterase activity, and this specificity

depends on the culture conditions (Haslam and Stangroom 1966; Beverini and

Metche., 1990; Farías et al., 1994).

TAH catalyzes the complete hydrolysis of tannic acid to gallic acid and

glucose. The intermediates in the reaction are 1,2,3,4,6-pentagalloylglucose,

2,3,4,6-tetragalloylglucose, and two types of mono-galloyl glucose (Iibuchi et al.,

1972; Lekha and Lonsane., 1997). Fig. 2.10.1 depicts the hydrolysis of 1,2,3,4,6-

pentagalloylglucose catalyzed by tannase. When the substrate is formed by methyl

ester of gallic acid, the TAH produces gallic acid and methanol (Aguilar and

Gutiérrez-Sánchez., 2001).

Tannase activity is inhibited competitively by gallic acid, pyrogallol,

hydroxybenzoic acid, and di-hydroxybenzoic (Iibuchi et al., 1972). The tannase

inhibition by compounds such as diisopropyl fluorophosphate (Adachi et al., 1971;

Barthomeuf et al., 1994) and phenylmethylsulphonyl fluoride (Sharma et al., 2008)

indicates the presence of a serine residue in the active site. Studies with radioactive

isotopes suggested that the sequence of amino acids in the active site can be

threonine–serine–methionine (Adachi et al., 1971). Depending on their origin,

tannase can be inhibited or stimulated in different magnitude by metal ions. The

addition of Mg+2

increased the tannase activity, but it is still unclear whether if it

acts as a cofactor or if it has a nonspecific effect (Kar et al., 2003; Mukherjee and

Banerjee., 2006; Naidu et al., 2008; Hamdy 2008)


37

Fig.2.10.1 Hydrolysis of 1,2,3,4,6- pentagalloyl glucose catalyzed by tannase

(Aguilar and Gutierrez-Sanchez., 2001)

Fig.2.10.2 Esterase and Depsidase activities of Tannase

Toth and Barsony (1943) reported that gallotannin-decomposing tannase

contains two separate enzymes-an esterase and a depsidase with specificities for

ester linkage and m-digallic acid ester linkages, respectively shown in Fig.2.10.2


38

2.11 APPLICATIONS OF MICROBIAL TANNASES

Over the years, tannase enzyme has been used in several conventional

industrial processes, such as Pharmaceutical, Beer and wine production, cold tea

production, treatment of industrial wastewater, containing tannin material, etc

2.11.1 Pharmaceutical industry

Van de Lagemaat and Pyle., (2005) studied major applications of tannase

enzyme are in the production of gallic acid (3, 4, 5-trihydroxy benzoic acid), which

is synthesized chemically in pharmaceutical industry for production of anti

bacterial drug trimethoprim. Gallic acid is a substrate for the chemical and

enzymatic synthesis of propyl gallate, used as an anti oxidant in fats and oils,

foods, cosmetics, hair products, adhesives and lubricant industries.

2.11.2 Beer and wine production

Rout and Banerjee., (2006) reported the application of tannase enzyme in

fruit industries. In fruit industries tannase is also used for clarification and removal

of unwanted bitterness from the traditional fruit juices such as cashew,

pomegranate, cranberry, raspberry, etc. The presence of high tannin content in

these fruits is responsible for haze and sediment formation, which results from

protein–polyphenol interaction. Tannase applied to remove haze, improves color,

bitterness and astringency of the juice upon storage.

Giovanelli., (1989) showed that tannase from a certain strain of A. flavus

has been shown to dramatically reduce the haze formation in beer after storage.

This implicates tannase in the hydrolysis of wort phenolics which complex with

the other chemicals in the beer mixture and results in the haze formation. Upon

treatment of the stored beer with tannase the potential of haze formation is

dramatically reduced.

Canterelli et al., (1989) reported fifty percent of the color of the wines is

due to the presence of the tannins; however, if these compounds are oxidized to

quinines by contact with the air it could form an undesirable turbidity, presenting

severe quality problems. When the proteins of the beer are in considerably high

quantities an undesirable turbidity is presented by accomplishing with these


39

tannins. The use of tannase can be a solution to these problems. In the manufacture

of beer, the tannase could be used since the tannins are present in low quantities.

2.11.3 Cold tea products

Aguilar et al., (2001) studied the commercial applications of tannase is in

the manufacturing of instant tea. Here it is used to eliminate water – insoluble

precipitates (called “tea cream”). These precipitates are generated in a natural way

when the beverage is cooled at temperatures lower than 4°C and if these are

removed chemically (employing sulfites and molecular oxygen with an alkali), a

great amount of aromatic compounds can be eliminated. Such precipitates are

formed by polymerization of esterified poly phenols and by accomplishment of

caffeine, giving a cold water soluble tea, characterized by a high tannin content of

aromatic compounds and appropriate color.

2.11.4 Animal feed

Aguilar et al., (2001) reported the applications of tannase for animal feed

compliment. However, some cultivators of sorghum present a high content of

tannins. If this cereal is first treated with tannase, tannin content could be

decreased and this type of sorghum could then be used as compliment in animal

diet.

2.11.5 Cell wall digestion

Garcia-Conesa et al., (2001) found that tannase may contribute to plant cell

wall degradation by cleaving some of the cross-links existing between cell wall

polymers. Due to the shortage and high cost of the enzyme, the use of tannase in

large-scale applications is limited at present. It is hoped therefore that the

economic benefits of tannase production can help improve the overall viability of

the process.

2.11.6 Effluent treatment

Van de Lagemaat and Pyle., (2001) studied the applications of tannase in

effluent treatment. Tannery effluents contain high amounts of tannins, mainly

polyphenols, which are dangerous pollutants and cause serious environmental

problems. Tannase can be potentially used for the degradation of tannins present in


40

the effluents of tanneries offering a cheap treatment and removal of these

compounds.

2.12 TANNASE IMMOBILIZATION

As mentioned above, TAH has several interesting applications, but its

industrial use is limited mainly due to economical reasons. In this regard, tannase

immobilization offers several advantages over the utilization of free enzyme, such

as improvement of enzyme stability, reutilization of biocatalyst, ease of product

recovery, and continuous operation in packed bed bioreactors (Schons et al., 2011).

Several authors have reported the immobilization of tannase by different

techniques. Abdel-Naby et al., (1999) compared the immobilization of tannase on

several carriers and different methods such as physical adsorption on ASalumina

and colloidal chitin, ionic binding onto Dowex 50W and DEAE-Sephadex A-25,

covalent binding on chitosan and chitin, and entrapment on polyacrylamide and

Ca-alginate beads. They found that covalent binding to chitosan led to the highest

immobilized activity and immobilization yield. Characterization of biocatalyst

demonstrated that immobilization significantly improved the stability at low pH, as

well as thermal stability of tannase.

Mahendran et al., (2005) reported the immobilization of a Paecilomyces

variotii tannase into Ca-alginate beads and utilized them for tannic acid hydrolysis

in a batch reactor. Immobilized biocatalyst completely hydrolyzed a 1% tannic

acid solution within 6 h and retained 85% of its initial activity after eight cycles.

Hota et al., (2007) immobilized a R.oryzae by the same methodology. They

utilized the immobilized tannase for gallic acid production from several tannin rich

agro-residues (sal seed, fruit of myrobalan, and tea leaf). The biocatalyst reached a

high bio-conversion yield (about 90%) and was stable for seven cycles, since the

immobilized tannase showed an appreciable substrate bioconversion of 60–80% up

to seven cycles.

Curiel and co-workers described the immobilization of recombinant L.

plantarum tannase. The enzyme was immobilized and stabilized by one point and

multipoint covalent immobilization on highly activated glyoxyl agarose.

Biocatalyst obtained by multipoint immobilization were 500- fold and 1,000-fold


41

more stable to thermal and cosolvent inactivation than both the soluble enzyme and

the one point immobilized enzyme, respectively. In addition, the immobilized

biocatalyst preserved more than 95% of its initial activity after 1 month of

incubation under the optimal reaction conditions (Curiel et al., 2010).

Immobilized tannase has been employed for treatment of pomegranate and

myrobalan juice (Srivastava and Kar., 2009, 2010) and tea beverage (Su et al.,

2009), as well as for removal of tannin and associated color from tannery effluent

(Murugan and Al-Sohaibani., 2010). Immobilized tannase has also been utilized

for production of gallic acid esters in non-aqueous media, either by esterification of

gallic acid with the appropriate alcohol (Sharma and Gupta., 2003; Yu et al., 2004)

or by direct trans-esterification of tannic acid in presence of alcohol (Fernandez-

Lorente et al., 2011).

Other interesting approach is the utilization of self immobilized tannase.

Under certain production conditions, tannase remains strongly bound to cells; then,

whole cells or cell debris could be used as natural immobilized biocatalysts. For

example, mycelium-bound tannase from A.niger has been employed to catalyze the

synthesis of propyl gallate in organic solvents (Yu and Li., 2005). Belur et al.,

(2010 b) reported the hydrolysis of tannic acid by whole cells of Serratia ficaria

producing a cell associated tannase. This biocatalyst showed a high tolerance to a

wide range of temperature and pH. The use of naturally self-immobilized tannase

offer several technical and economical advantages for industrial applications such

as the avoidance of costly and time-consuming purification and immobilization

processes.

2.13 OPTIMIZATION OF PROCESS PARAMETERS

The production of tannase was carried out so far only in laboratory scale.

The yield of tannase was found to be less and not economical. Hence several

attempts have been made to improve the yield of tannase enzyme by using cheaper

substrates, higher yielding organisms, mixed cultures and by using mutant cultures

with higher rate of fermentation and with shorter fermentation period.

Hence optimization of process parameters for maximum production of

tannase is essential. The classical method and methods like Plackett–Burman


42

Design could not give accurate results for economical and efficient production of

tannase. Statistical optimization techniques like Central Composite Design (CCD)

through Response Surface Methodology (RSM) are widely used to optimize the

media components and process conditions due to its capacity to interpret the

variables affecting the production accurately.

2.13.1 Classical Method

The conventional method that has been used for optimization is the

“change- one-factor-at-a-time” method in which a single factor or one independent

variable is varied while fixing all others at a specific level may. This may lead to

unreliable results and less accurate conclusions.

2.13.2 Plackett–Burman Design

This study was carried out to screen the medium components with respect

to their main effects and not their interaction effects on maximum enzyme

production. Plackett–Burman, (Plackett and Burman., 1946) a two factorial design

identifies critical chemical and physical parameters required for maximum enzyme

production by screening N variables using N + 1 experiments. Minitab-16 is used

to generate experimental design with similar variables. Two values of each

variable {maximum (+) and minimum (-)} are chosen such that the difference

between the two values (+ and -) is large enough to ensure that the peak area for

the maximum enzyme production is included. The variables are analysed. The

effect of variables on enzyme production is calculated by using the following

equation:

E(xi) = 2 ( ∑M+ - ∑M- ) / N …………(2.1)

where E(xi) is the concentration effect of tested variable, M+ and M- are the tannase

production from the trials where the variable (xi) measured was present at high and

low concentrations, respectively and N is the number of experiments carried out.

Experimental error is estimated by calculating the variance among the dummy

variables as follows

Veff = ∑ (Ed)2 / n …………..(2.2)

Where Veff is the variance of the concentration effect, Ed is the

concentration effect for the dummy variable and n is the number of dummy


43

variables in Eqn (2.2). The Standard Error (SE) of the concentration effect was the

square root of the variance of an effect and the significance level (p-value ) of each

concentration effect was determined by using student’s t-test.

t (xi) = E(xi) / SE …………..(2.3)

Where E(xi) is the concentration effect of tested variable and (SE) is the

Standard Error

2.13.3 Response Surface Methodology

Response surface methodology (RSM) is an effective statistical tool and

widely used in process optimization, which includes experimental design, model

fitting, validation and optimization. An effective statistical design is the basis for

response surface optimization and the reported designs include Central Composite

design.

Central Composite Design

Central Composite Design was used to obtain a quadratic model, consisting

of factorial trails and star points to estimate quadratic effects and central points to

estimate the process variability with enzyme production.

Each factor in this design was studied at five different levels -2, -1, 0, +1,

+2 and a set of 31 experiments were carried out. All the variables were taken at a

central coded value considered as zero. The minimum and maximum ranges of

variables were used. All the experiments were carried out in triplicates and the

average value was taken as the response. The CCD experiment was designed using

the Design Expert Software package (Version 8.0.7.1). The following equation was

used for coding the actual experimental values of the factors in the range of ( −2

to +2):

Where xi is the coded value of the ith

independent variable, Xi the natural value of

the ith

independent variable, X0 the natural value of the ith

independent variable at

the center point, and Xi is the value of step change.

Analysis of the data and generation of three dimensional surface graphs

were done using Design Expert Software package (Version 8.0.7.1). After


44

conducting the experiments and measuring the tannase activity levels, a second

order polynomial equation including interactions was fitted to the response data as

given by Eqn (2.4),

Where Y is the measured response, β0 is the intercept term, βi are linear

coefficients, βii are quadratic coefficient, βij are interaction coefficient and Xi and

Xj are coded independent variables.

The significant terms in the model were found by analysis of variance

(ANOVA) for each response. The goodness of fit of the regression model obtained

was given by the coefficient of determination R2. The statistical significance of the

model was determined by F-test. Since coding of the variables enables direct

comparison of the partial regression coefficients, their significance was determined

by student ‘t’ test and associated probabilities. The interaction effects of variables

on enzyme production were studied by plotting three dimensional surface graphs

against any two independent variables while the third and fourth variables are fixed

at its central level.

The second degree polynomial equation was maximized by a constraint

search procedure using the Response optimizer of the Minitab software to obtain

the optimum levels of the independent variables and the predicted maximum

enzyme activity. The predicted enzyme activity was compared with the

experimental values. Validation of the experimental model was tested by carrying

out the batch experiments under optimal operating conditions. All the experiments

were repeated thrice, and the results were compared.

2.14 ARTIFICIAL NEURAL NETWORKS

Artificial Neural Networks (ANN) are a well-known mathematical tool

widely used and tested lately for the problems in industrial enzyme production. Its

advantages are in the ability to handle with nonlinear data, highly correlated

variables and the potential for identification of problems (Mandal et al., 2009).The

fundamental processing element of ANN is an artificial neuron. A biological

neuron receives inputs from other sources, combines them, performs generally a


45

nonlinear operation on the result, and then outputs the final result (Bas et al.,

2007). The basic advantage of ANN is that it does not need any mathematical

model since an ANN learns from examples and recognizes patterns in a series of

input and output data without any prior assumptions about their nature and

interrelations (Mandal et al., 2009).

2.14.1 Structure of ANN

An ANN consists of nodes (neurons) with weighted connections between

them. There are some nodes that receive input, some nodes that give output, and

hidden nodes in between. Each node processes all its input, for example by

summing it up and running the sum through a function, and propagates its result to

its connected nodes until an output is given at some output node(s). Fig.2.14.1

represents the topology of a simple example ANN.

Input nodes receive input data set or information and pass it along to the

hidden nodes through weighted connections. The received signal is processed in

the hidden nodes and sent along weighted connections to output node(s) which

further process the signal and produce the final output ( Jreou et al., 2012)

Fig.2.14.1 Structure of ANN

2.14.2 Properties of ANN

The main property of an ANN is its capability to learn. Learning or training

is a process by means of which a neural network adapts itself to a stimulus by

making proper parameter adjustments, resulting in the production of desired

response. Broadly, there are two kinds of learning in ANNs:

1. Parameter learning: It updates the connecting weights in a neural net.

2. Structure learning: It focuses on the change in network structure (which includes

the number of processing elements as well as their connection types).


46

The above two types of learning can be performed simultaneously or separately.

Apart from these two categories of learning, the learning in an ANN can be

generally classified into three categories as:

· Supervised Learning

· Unsupervised Learning

· Reinforcement Learning

Supervised Learning

In supervised learning, each input vector requires a corresponding target vector,

which represents the desired output. The input vector along with the target vector

is called training pair. The network here is informed precisely about what should

be emitted as output. It is assumed that the correct “target” output values are

known for each input pattern.

Unsupervised Learning

In unsupervised learning, the input vectors of similar type are grouped without the

use of training data to specify how a member of each group looks or to which

group a number belongs.

Reinforcement Learning

In this learning process, only critical information is available, not the exact

information. The learning based on this critic information is called reinforcement

learning and the feedback sent is called reinforcement signal. (Pratap et al., 2013)

2.14.3 Types of ANN

Different types of ANN are known, Kohonen, counter-propagation (CP), back-

propagation ANN, Like in the biological neural network, the artificial ANN has an

interconnection of neurons with three vital components: i) node character which

controls signals i.e. the number of inputs and outputs, the weights and activation

function associated with the node, ii) network topology defining how nodes are

organized and connected and iii) learning rules for the initialization and adjustment

of weights. There are two groups of ANN, supervised and unsupervised, which

differ in the strategy of learning. In unsupervised learning, the input data is

organised and processed without reference to the target, whereas in supervised

learning, both the input and target (output) are used.


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· Kohonen ANN is an example of unsupervised learning, where no

referential (output) data are used in training of the network, and the

algorithms used are excellent for establishing the relationship among

complex sets of data.

· Counter-propagation ANN represents an up-grade of Kohonen ANN and is

based on two-step learning procedure, unsupervised in the first step, and

supervised in the second. CP-ANN is the most suitable method for

classification of data, but can be used also as a method for developing

predictive models for new objects of unknown properties.

· Back-propagation ANN is another example of supervised learning, where

one or more target values are predicted from input data, meaning that both

inputs and outputs should be known for the training dataset. A special type

of ANN is radial basis function network which ordinarily does not involve

the training of network, but is determined using a certain transformed

function. However, the majority of algorithms work according to an

iterative principle, which is similar to training of the network (Maja

Prevolnik et al., 2012).

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