Economic Production of Tannase by Aspergillus niger ... - Rombio.eu

Romanian Biotechnological Letters Vol.17, No.4, 2012 

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ORIGINAL PAPER 

Economic Production of Tannase by Aspergillus niger van Tiegh Adopting 

Different Fermentation Protocols and Possible Applications 

Abstract 

Romanian Biotechnological Letters, Vol. 17, No. 4, 2012 

Received for publication, February 7, 2011 

Accepted, October 31, 2011 

H. S. HAMDY 1 AND E. M. FAWZY 2 

Biological Sciences & Geology Department, Faculty of Education, Ain Shams 

University, Roxy, Heliopolis, Cairo, 11757, Egypt 

1 hosamhamdy@yahoo.com 2 emanfawzy@hotmail.com 

Here, we demonstrated the ability of Aspergillus niger to utilize Ficus nitida' leaves and crude 

tannic acid extract to produce tannase. The kinetics of tannase synthesis using free and immobilised 

cells of A. niger were studied by adopting submerged (LSF), solid-state (SSF) and slurry-state (SlSF) 

cultures. The optimum parameters for maximum production were recorded at 30°C, initial pH of 5 

after 120 hours of shaking incubation at 175 rpm for SSF and SlSF and 200 for LSF. The maximum 

yield of tannase was produced in the following descending order: SSF (457.2 U 50ml -1 ) > LSF (451 U 

50ml -1 ) > SlSF (324.3 U 50ml -1 ). The concentration and balance of trace elements had a critical role 

on tannase synthesis where a dramatic beneficial effect was observed when a mixture of Ca 2+ , Mg 2+ , 

Mn 2+ and Zn 2+ was added to the fermentation media. Applicability of the research findings was 

illustrated through suggesting the possibility of using the fermentation residue of F. nitida obtained 

from SSF in animal feed. Moreover, the ability of the produced tannase to remove up to 55% and 45% 

of tannin after 120 minute from tea extract and pomegranate juice respectively was confirmed. 

Keywords: Aspergillus niger, Ficus nitida, tannase, LSF, SSF, SlSF, detannification, 

partial purification. 

Introduction 

Tannase (tannin acylhydrolase, E.C 3.1.1.20) is an important inducible enzyme 

responsible for the breakdown of ester linakages in tannins and gallic acid esters to produce 

glucose and gallic acid [31]. Tannase has a wide range of application in industrial, 

biotechnological, environmental, pharmaceutical and other important fields, such as instant 

tea production, clarifying coffee-flavoured soft drinks, removing haze, bitterness, astringency 

and improving the colour of fruit juices, stabilising wine color, [7, 8, 35], enzymatic treatment 

for nutritive use of protein and carbohydrates from peas [64], enhancing the antioxidant 

activity of green tea [37], and improving the quality of forage used in animal and bird feeds 

[50]. It is also applied in the synthesis of pharmaceutically important drugs, i.e., 

trimethoprim, gallic acid, propyl gallate, antifolic agents, antioxidants, antibacterial and 

antiviral agents, analgaesics, anti-inflammatories, and active anticancer agents that do not 

harm normal cells [11, 20, 26, 69]. Tannase is also used in the production of semiconductors, 

ink, dye, photo developers, [27], manufacture of sensitive analytical probes to determine 

gallic acid esters [24], high grade leather [6], plant cell wall degradation [49] and cleaning 

hard acidic industrial effluent containing tannin materials [4]. 

In spite of the varied fields for tannase applications, the large-scale use of tannase is 

still limited at present, partly due to the shortage and high cost of the enzyme production [65]. 

This could partly be due to the high price of the enzymes produced by submerged 

fermentation and the relatively high cost of tannic acid used as the substrate. On the other 

hand, there is often a serious problem in disposing agricultural residues and wastes, which 

7441

H. S. HAMDY AND E. M. FAWZY 

may cause serious environmental contamination and propagation of flies and rats if not 

correctly dealt with. Meanwhile, agricultural wastes can represent large renewable resources 

for enzyme production by filamentous fungi and has been practiced worldwide as waste-based 

cultures adopting solid state fermentation [67]. To bridge this gap, alternative techniques 

such as solid/slurry state fermentations, use of whole-cell-immobilization, and/or use of less 

expensive substrates such as agricultural residues may be used in obtaining relevant costeffective 

products such as enzymes and forage among others. In Egypt, Ficus nitida L. 

(Moraceae) is a widely cultivated ornamental plant that annually yields a vast amount of waste 

in the form of leaves. Although Ficus wastes are highly nutritious for microorganisms [29], 

there is still no study that examines its potential use as a bioresource for tannase production. 

Solid state fermentation (SSF) is a batch process used for its many advantages such as 

the low cost of raw materials, simple equipments and facilities, labour and technology. 

Moreover, the secreted enzymes in the culture filtrate are more concentrated and of higher 

quality, with a less effluent generated and lower a lower cost of recovery than in the liquid 

state cultures [43]. It is also a natural environment that provides easy aeration, high surface 

exchange and serves as a support for the cells [44]. Along with the SSF, there is a growing 

interest in the whole-immobilized cells to produce enzymes as a promising technique 

proposed for improving the fermentation process where it offers several advantages, i.e., easy 

to handle, reusable, continuous operation over a prolonged period with operational stability, 

reducing the overall cost and delays associated with the time needed for sterilisation, 

inoculation and mycelium growth, lower susceptibility to microbial contamination, lower 

vulnerability to inhibitory compounds and nutrient depletion, and increased rates of 

generating of microbial products [17, 51]. Meanwhile, very little information can be found in 

the literature about slurry state fermentation and their applications are mostly restricted to 

anaerobic fermentations [18]. 

Many fungal species have been reported to produce tannase, including Aspergillus 

aculeatus, A. aureus, A. flavus, A. foetidus, A. japonicas, A. niger, A. oryzae Aureobasidium 

pullulans, Fusarium solani F. subglutinans, Paecilomyces variotii, Penicillium 

atramentosum; P.chrysogenum, P. variable, and R. oryzae as reviewed by Belur and 

Mugeraya [10] and Chavez-Gonzalez [16]. The vast majority produce in submerged cultures, 

while Aspergillus and Penicillium are the most active microorganisms capable of producing 

tannase through both submerged and solid state fermentations [45, 46]. Different agricultural 

residues such as cassava, carob bean, wine-grape, tea and coffee wastes, [65]; Acacia nilotica, 

A. auriculiformis, Casuarina equisetifolia, Cassia fistula, Ficus benghalensis, [39] and 

gobernadora [62], have been associated with microbial production of tannase on SSF, while 

many studies have investigated tannase production with pure tannic acid as the soul carbon 

source [39]. 

Thus, the aim of this study was to investigate, compare and optimise production of 

tannase utilising crude tannic acid extract and leaves of F. nitida adopting LSF, SSF and SlSF 

protocols inoculated with free and immobilised cells of A. niger. Possible applications of the 

produced tannase as well as the fermentation residue are also suggested. 

Materials and Methods 

Preparation of materials and equipments 

Plant leaves were collected in May from mature trees of Ficus nitida, cultivated in 

different Egyptian localities, cut into small pieces, and then oven-dried at 55°C for 24 h until 

they reached a constant weight. The material was then finely ground to pass through a 50μm 

sieve. The powder was stored in dry flasks under dark conditions at room temperature. 

7442 Romanian Biotechnological Letters, Vol. 17, No. 4, 2012

Economic Production of Tannase by Aspergillus niger van Tiegh 

Adopting Different Fermentation Protocols and Possible Applications 

Crude tannin extract was prepared according to the method of Schanderi [55] where 

50 g of dry plant materials were mixed with 200 ml of distilled water and kept at room 

temperature overnight. After soaking, the mixture was boiled for 10 min and filtered. The 

filtrate served as a crude tannin extract from which different dilutions were prepared using 

distilled water. 

Cleaning of equipments used in studying the requirements of metallic nutrition was 

performed by washing in hot detergent solution, rinsing in tap water, drying and submerging 

in a bath of hot, concentrated nitric-sulphuric acid (2:1) for 15 minutes. The glassware was 

removed from this bath and rinsed 5 times with distilled water and finally 2 times with glassdistilled 

water. 

Microorganism 

Aspergillus niger van Tiegh CBC 122722 used throughout this work was previously 

isolated from an Egyptian soil sample, identified by the CBS-KNAW (Fungal Biodiversity 

Centre, Uppsalalaan, Netherlands) and kept on malt extract agar at 4 °C and routinely 

cultured. 

Spore suspension (containing about 2 x 10 6 spores ml -1 ) was freshly prepared from 7day-old 

cultures of A. niger grown on malt extract agar slants at 30°C using deionised double 

distilled water. 

Entrapped spores of A. niger was prepared according to the method described by 

Yalcinkaya [71] where 10 ml of 5% Na-alginate was mixed with 1 ml of the fungal spore 

suspension. The mixture was introduced into a solution containing 0.1 M CaCl2 with a 

syringe with a constant stirring to prevent aggregation of the beads. The fungal sporeentrapped 

beads (about 2.5 mm in diameter to overcome the anaerobic conditions inside the 

large beads [60]) were cured in this solution for 1 h and then washed twice with 200 ml of 

sterile distilled water. The beads with immobilised spores (about 20 ml volume) were then 

transferred to the fermentation medium. 

Screening 

The potential of different fungal strains for tannase production was screened in tannic 

acid medium composed of (g/l): freshly prepared filter-sterilised tannic acid (30g), (NH4)2SO4 

(2g), KH2PO4 (0.5g), MgSO4 (2g), and agar agar (30g) [4]. The tannase-producing organisms 

were grown according to one of the following fermentation protocols. 

Enzyme production and fermentation media 

(A) Batch cultures 

Flasks containing one of the following fermentation media were prepared in 

triplicates, the initial pH value adjusted to 5.5, and incubated for 96 h at 30°C under static and 

shaking conditions (GFL shaking incubator, 150 rpm). 

Liquid state fermentation (LSF) medium 

Liquid state fermentation (LSF) medium was composed of (g/l): freshly prepared 

filter-sterilised crude tannin extract (30g), (NH4)2SO4 (2.0g), K2HPO4 (0.5g) and 

MgSO4.7H2O (1.0g) in a final volume of 50 ml per flask [4]. Medium was inoculated with 1 

ml of free A. niger spore suspension, or with about 20 ml of immobilised cells. 

Solid state fermentation (SSF) 

Five ml of LSF medium without any of the carbon (crude tannin extract) and nitrogen 

((NH4)2SO4) sources were added to 10 g of powdered Ficus nitida leaves and inoculated with 

1 ml of free spore suspension. 

Romanian Biotechnological Letters, Vol. 17, No. 4, 2012 7443


Slurry state fermentation (SlSF) 

Similar sets of flasks used in SSF were prepared, while an extra amount of 45 ml of 

distilled water was added to the medium to set the slurry state condition (SlSF) and then were 

inoculated with 1 ml of free spore suspension. 

(B) Repeated batch cultures 

Flasks (LSF) inoculated with immobilised spores were incubated for 120 hours at the 

first cycle to allow the mycelium to develop and then the duration of each cycle was 48 hour. 

At the end of each run the beads were aseptically filtered, washed thoroughly with 25 ml 

saline, acetate buffer (0.2 M, pH 5) and distilled water and then recultivated into 50 ml 

aliquots of fresh medium. 

Enzyme extraction 

At the end of incubation, the cell-free filtrate in LSF was obtained by filtering through 

Whatman No. 1 filter paper in a Buchner funnel. In SSF and SlSF, a suitable amount of the 

fermented matter was thoroughly mixed with 10 ml of cold distilled water by keeping the 

flasks on a rotary shaker for 1 h at 200 rpm. The mixture was filtered through muslin cloth 

and the filtrate was centrifuged at 10000 rpm for 20 min at 4°C and served as crude enzyme 

preparation. The volume of all the enzyme extracts obtained from the different protocols was 

restored to 50 ml by cold distilled water and served as the crude enzyme preparation. 

Tannase and protein assays 

The activity of tannase was estimated according to the method of Mondal [41] where 

0.5 ml of enzyme solution was incubated with 3 ml of 1.0% (w/v) tannic acid, in 0.2 M acetate 

buffer (pH 5.0) at 40 °C for 30 min. The reaction was terminated by transferring the tubes 

containing the reaction mixture to an ice bath followed by the addition of 2.0 ml (1%, w/v) of 

bovine serum albumin to precipitate the remaining tannic acid. The precipitate was collected by 

centrifugation (12 x 10 3 g for 15 min) and dissolved in 2 ml sodium dodecyl sulphate (SDS)triethanolamine 

(1.0%, w/v, SDS in 5% v/v, triethanolamine) solution. Absorbency was 

measured at 550 nm 15 min after the addition of 1.0 ml of FeCl3 (0.13 M dissolved in 0.01 N 

HCl). Control treatments were performed using heat-killed enzyme. One unit of tannase is the 

amount of enzyme capable of hydrolysing 1.0 mM min -1 of tannic acid under the assay 

conditions by reference to a standard curve constructed with pure tannic acid [41]. 

Protein content was determined using bovine serum albumin dissolved in 0.17 M 

NaCl as a standard [14]. 

Chemical analyses 

Cellulose, hemicellulose and pectin contents of F. nitida' leaves were analysed as 

described by Jermyn [25] and the total nitrogen was estimated by the conventional micro- 

Kjeldahl method [33, 48]. 

Tannin was extracted as described by Schanderi [55] and estimated as described by 

Lokeswari [35] adopting the Folin–Denis method where a 50 ml aliquot of tannin-containing 

solution was mixed with 5 ml of Folin–Denis reagent, and then 10 ml of 15% (w/v) Na2CO3 

solution was added and then diluted to the 100 ml mark with distilled water. After thorough 

mixing, the flasks were kept in the dark for 30 minutes at 25°C. The absorbency of tannin was 

measured spectrophotometrically at 700 nm and its concentration calculated using pure tannic 

acid as a standard. Distilled water was used as blank regarding the calibration curve. From 

this curve, the concentrations for each sample was obtained and used for the tannin content 

calculation. 




Optimising process parameters for tannase production 

Optimising tannase biosynthesis was performed by optimizing various physicochemical 

process parameters where the optimum condition found for each parameter was 

settled for the subsequent experiments. The effect of incubation temperature (25 to 45°C) for 

different incubation periods (96 to 168 h) at different agitation speeds (125 to 225 rpm) was 

explored. Initial pH value of the fermentation medium (4 to 7) was adjusted by 0.1 M HCl or 

NaOH. The effect of various initial tannic acid concentrations (2 to 9 g %, w/v) and the 

addition of various mineral salts i.e. Mg, Ba, Ca, Co, Fe, K, Mn, Na and Zn (added as 

chlorides), were also optimised. 

Partial purification of tannase 

Partially purified tannase was prepared as previously described by Hamdy [23] where 

protein content of 500 ml of crude tannase of A. niger was precipitated overnight with 65% 

ammonium sulphate, collected by centrifugation at 12,000 ×g for 15 min, desalted by passing 

through a column of Sephadex G-25 and then fractionated by applying 2 ml to a column 

(2.5×82 cm) of diethylaminoethyl cellulose (DEAE cellulose, fast flow, fibrous form, Sigma 

Chemicals, Germany) for ion-exchange chromatography. 5 ml fractions were eluted with a 

linearly increasing molarity of NaCl solution (0.0 to 0.5 M) prepared in 0.2-M acetate buffer, 

pH 5.0. Tannase activity and the protein content of each fraction were determined and used to 

calculate the corresponding specific activities. Fractions possessing the highest specific 

activities were pooled, desalted, lyophilised and used as partially purified tannase in the 

experiments of reducing tannin. 

Application of tannase in reducing tannin from tea extract and pomegranate juice 

50 g of fresh pomegranate seeds were washed with distilled water and extracted in 100 

ml distilled water using an electric blender. The extract was filtered through muslin cloth and 

kept cooled at 4°C. 

Tea extract was prepared by adding 2 g of locally purchased black tea leaves to 100 ml 

of boiled distilled water, left to stand for 20 minutes and then kept cold at 4°C. 

15 ml of the tea or pomegranate juice was incubated with 1 ml of partially purified 

tannase containing 17.32 U/ mg protein at 30°C for different incubation periods (30 to 150 

min) under shaking conditions (150 rpm). Tannin content was determined pre and post 

incubation with the enzyme. 

Statistical validation of treatment effects 

The mean, standard deviation, Tukey's test “T” and probability "P" values of three 

replicates of the investigated parameters and the control were calculated according to the 

mathematical principles described by Glantz [21]. Results were considered highly significant, 

significant or non-significant where P < 0.01, > 0.01 and < 0.05, and > 0.05, respectively. 

Results and Discussion 

Cellulose, hemicelluloses, lignin, pectin, tannin, nitrogen and protein contents of Ficus 

nitida’ leaves used in the fermentation medium were determined and the results are given in 

Table 1. It is known that the detected amounts are subject to dramatic changes according to 

genotype, age of plant and leaves, soil fertility and acidity, growing season and methods of 

extraction, drying and assay [3]. 



Table 1: Chemical composition of Ficus nitida’ leaves. 

Chemical composition (%) 

Cellulose 1 

Hemicellulose 1 

Lignin 1 

Pectin 1 

Tannin 2 

Ash 

Nitrogen (N) and protein content 3 

32.7 21.93 0.71 11.03 15.01 7.5 0.19 0.83 1.02 6.37 

Analyses carried out adopting the methods of 1 Jermyn, [25] 2 Lokeswari [35], 3 Pirie, [48] and Lexander [33]. 

Aspergillus niger was selected in this study for being classified as GRAS (generally 

regarded as safe) and is officially approved in France for enzyme production for the food 

industry [6] where some applications of tannase, especially in detanification processes, are 

accepted by partially purified enzymes [12], which requires a safe source of enzyme. 

Entrapment in alginate was selected in this study for its highest effectiveness factor (i.e. 1.12 

= amounts of tannase produced by immobilized cells / corresponding amounts produced by 

free cells under the same conditions) versus the other investigated methods, i.e., adsorption to 

kieselguhr (0.68), covalent bonding to silica gel beads (0.53) and cross-linking with 

glutaraldehyde (0.43). Moreover, alginate gel is characterised by its high biodegradability, 

hydrophilicity, mechanical stability, rigidity, chemical inertness, ability to bind cells firmly 

and high loading capacity [71]. 

Tannase production by free or immobilized cells of A. niger under static or shaking 

conditions, adopting LSF, SSF or SlSF was monitored at different temperatures (25 to 45°C) 

for different incubation periods (96 to 168 h) in a bifacatorial design. It was found that 30°C 

enabled maximum tannase biosynthesis in all cases and the results of the time course 

production at 30°C are graphically represented in Fig. 1. The optimum temperature for 

tannase production by most fungal species is around 30°C [10], while tannase production by 

Trichoderma viride [35] has been maximally recorded at 45°C. Decrease in tannase 

biosynthesis by A. niger above 30°C is in agreement with other findings, such as the sharp 

decrease in tannase production by A. oryzae and almost no tannase production was detected at 

40°C [32]. 

The results of time course of tannase production by A. niger in LSF (Fig. 1 a), SSF or 

SlSF (Fig. 1 b) at 30°C showed that the maximum biosynthesis achieved after 120 h of 

shaking incubation, and 144 h of static incubation. Production under shaking conditions was 

significantly higher than the static one within the same fermentation protocol. The increased 

order of enzyme production in relation to different fermentation protocols is as per the 

following sequence: SSF (81.5 U 50 ml 

7446 Romanian Biotechnological Letters, Vol. 17, No. 4, 2012 

-1 ) > Immobilised LSF (77.5 U 50 ml -1 ) > SlF (75.5 U 

50 ml -1 ) > Free LSF (69.5 U 50 ml -1 ) (Fig. 1). Therefore, the next stage of experiments 

focused on the first three for their significant level of production as well as representing the 

various fermentation protocols. It has been previously found that tannase production by A. 

niger on solid substrates is higher than in LSF (2), where tannase in LSF is partially 

intracellular and subsequently secreted into the medium during fermentation, while the 

enzyme in SSF is completely secreted out of the cells [31]. On the other hand, Srivastava and 

Kar [59] stated that extracellular tannase of A. niger was optimally produced in submerged 

cultures. Our results (Fig. 1 a) also showed that tannase biosynthesis by immobilized cells of 

A. niger in either shaking (77.5 U 50ml -1 ) or static (66 U 50 ml -1 ) conditions was significantly 

Soluble N 

Insoluble N 

Total N 

Total protein



higher (P < 0.01) than that by free cells (69.5 & 52.5 U 50 ml -1 , in order). Higher amounts of 

tannase production by immobilised cells of A. niger and Bacillus licheniformis KBR6 

compared to free cells have also been reported [17, 40]. 

a- LSF 

b- SSF and SlSF 

Fig. 1. Time course of tannase production by A. niger at different incubation periods adopting (a) liquid state 

(LSF), (b) solid state (SSF) and slurry state (SlSF) fermentation protocols using crude tannic acid extract and F. 

nitida leaves, respectively. 

Cultures were incubated for the indicated incubation periods while pH was initially 

Adjusted to 5.5 and the velocity of shaking, if applied, was 150 rpm. 

The data represent the mean of 3 different readings and the error bars represent the standard deviation of means. 

Tannase biosynthesis in LSF by A. flavus has been maximally recorded after 96 h 

[45]; while in SSF P. atramentosum has been maximally recorded after 96 h [56]; A. niger 



after 96 h [32], A. niger [19], and Paecilomyces variotii, [7] after 120 h; and after 144 h for A. 

niger ATCC 16620, [54]. Comparison of the incubation periods should be undertaken 

cautiously even for the same fungal species due to the variation in state of the fermentation 

and substrate utilised. The recorded decreases in tannase biosynthesis with prolonged 

incubation (Fig 1 a & b) could be explained by substrate scarcity in the medium, a shift in the 

reaction equilibrium due to accumulation of gallic acid [27] or an accumulation of toxic 

metabolities in the fermentation medium. 

The effect of shaking velocity on tannase production by A. niger was evaluated and 

the maximum yield was obtained at 175 rpm for SSF (89.93 U / 50 ml) and SlSF (88.76 U / 

50 ml), while 200 rpm was the optimum (86.76 U/50 ml) for LSF inoculated with 

immobilized cells (Fig. 2). A concurrent increase in tannase biosynthesis with an increase in 

shaking velocity could be due to the enhancement effect of shaking on heat, mass and oxygen 

transfer capabilities in the fermentation media that minimizes moisture loss and drying in SSF 

[27] and overcomes a serious constraint to high enzyme productivity by immobilized aerobic 

cells where most of the interiors of large beads may be anaerobic [60]. On the other hand, a 

decline in tannase biosynthesis as the velocity of shaking increases above the optimum level 

could be ascribed to increasing the frequency of particle tumbling that disrupts the mycelium 

in the early growth stages [36], thus increasing vaculoation of older hyphal compartments and 

leading to autolysis or weakened hyphae [47] and/or making the mycelium easier to peel off 

from the substrate surface [61]. A decrease in enzyme biosynthesis at higher rotation speeds 

was more pronounced in SSF and SlSF than in LSF using immobilised cells where the curve 

was broader (Fig. 2), most probably, due to the known protective effect of the immobilisation 

on cells. 

Fig. 2. Effect of shaking velocity on tannase production by A. niger after 120 h of incubation at 30°C, in the 

different fermentation protocols (LSF __ ♦ __ , inoculated with immobilized cells of A. niger; SSF __ ■ __ and 

SlSF __ ▲ __ inoculated with free cells of A. niger and grown on leaves of F. nitida) at the indicated velocity of 

shaking. Initial pH of the medium was adjusted to 5.5. 




Maximum tannase production by A. niger (96.11, 114.76, 106.8 U 50 ml -1 for LSF, 

SSF, SlSF in order) was optimally recorded at an initial pH of 5.0 of the fermentation medium 

(Fig. 3). Applying different fermentation protocols have so far not been found to shift the 

optimum pH value, where a range of 4.5 to 6.5 enabled maximum tannase production by A. 

flavus, A. foetidus, A. japonicus, A. niger, A. oryzae, Paecilomyces variotti, Penicillium 

atramentosum, P. chrysogenum, grown on tannery solid substrates or in submerged media [1, 

6, 7, 27, 50, 56]. It is known that biological surfaces consist of different functional groups 

that become dissociated if the pH is above a certain value [71]. As shown in Fig. 3, the curve 

of tannase synthesis in LSF is broader, where a decline in tannase synthesis is lower and the 

active pH range is wider than that in SSF and SlSF. 

Fig. 3. pH-dependency of tannase production by A. niger after 120 h of shaking incubation at 30°C in different 

fermentation protocols (LSF __ ♦ __ , 200 rpm, inoculated with immobilised cells of A. niger;SSF __ ■ __ , and 

SlSF __ ▲ __ , 175 rpm, inoculated with free cells of A. niger and grown on leaves of F. nitida). Initial pH of the 

medium was adjusted using 0.1 M HCl or NaOH. 

The effect of initial tannin concentration on production of tannase by A. niger was 

studied and the results showed that maximum productivity was attained at 7% (w/v) in both 

LSF (146.10 U 50 ml -1 ) and SSF (190.50 U 50 ml -1 ) and at 6% in SlSF (153.72 U 50 ml -1 ), 

representing an increase in production of 1.88, 2.34 and 2.04-fold, respectively (Table 2). 

Tannin concentration is an important detrimental factor in microbial production of tannase 

and its optimum concentration for A. niger has been recorded at 4%, 5% and 10% by Lekha 

and Lonsane [31, 32], Aguilar [2] and Sharma [58], respectively, while various concentrations 

ranging from 0.5 to 10% have been recorded for other fungal species [10]. A reduction in the 

tannase yield at higher concentrations of tannin could be due to its toxic effect where it forms 

non-reversible bonds with surface proteins of the organism [5], deposition of gallic acid on 

cell surfaces [57], as well as depriving metal ions and substrates on membranes [13, 52] that 

impair metabolism, inhibit the organism and decrease tannase production. The ability of 

several fungi to grow and tolerate relatively high tannin concentrations of up to 20%, as in A. 

niger [22], is an advantage for fungi as tannase producers. 



Table 2: The effect of varying initial crude tannin concentration on tannase production by A. niger in different 

fermentation protocols. 

Crude 

Tannase productivity 

tannin 

conc. (%) 

* 

LSF 1 SSF 2 SlSF 3 

U 50 ml -1 

% U 50 ml -1 

% U 50 ml -1 % 

3 (control) 96.11 ± 2.21 100 114.76 * ± 4.36 100 109.80 ± 2.74 100 

4 123.00 * ± 3.93 128 161.81 * ± 5.17 141 132.86 * ± 4.51 121 

5 131.67 * ± 5.13 137 174.43 * ± 6.27 152 133.95 * ± 4.28 122 

6 138.39 * ± 2.90 144 182.47 * ± 4.01 159 153.72 * ± 4.45 140 

7 146.10 * ± 6.28 152 190.50 * ± 5.90 166 142.74 * ± 5.42 130 

8 143.20 * ± 2.72 149 183.62 * ± 7.52 160 131.76 * ± 5.13 120 

9 131.67 * ± 3.42 137 161.81 * ± 4.20 141 117.49 * ± 2.70 107 

Initial pH of the fermentation medium was adjusted to 5.0, and incubated at 30ºC for 120 h with shacking 

incubation at: 200 rpm for LSF and 175 rpm for SSF and SlSF. 

1 Liquid state fermentation using crude tannic acid extract and inoculated with immobilised A. niger cell. 

2 , 3 Solid-state and slurry-state fermentations using F. nitida and inoculated with free cells of A. niger. 10 g of 

dried F. nitida’ leaves (15% tannin on dry weight base –Table 1- that equals 3% w/v in the fermentation 

medium) were used and higher tannin concentrations were applied to the liquid part of the medium. 

Significance of results was statistically validated compared to the control (3% crude tannin concentration) 

where * indicates highly significant difference. 

The effect of adding different metal ions at concentrations of 0.1, 0.2 and 0.3 g % (w/v) 

to the fermentation media was investigated and the results showed that tannase production 

was stimulated in the following descending order: Ca 


2+ , Mg 2+ , Mn 2+ , Zn 2+ in LSF and SSF, 

and Ca 2+ , Mn 2+ , Mg 2+ , Zn 2+ in SlSF (Table 3). It has been established that metal ions 

significantly affect the progress and efficiency of fermentation, secretion of active enzymes 

and synthesis of secondary metabolites by microorganisms and their ability to attach to their 

various substrates [38] by affecting the dynamics of cell membranes, cell viability, 

permeability, membrane fluidity, stability and signalling systems [30]. Tannase synthesis by 

A. foetidus, A. japonicas A. niger A. oryzae, A. pullulans and R. oryzae are also stimulated by 

Ca 2+ , Mg 2+ , Mn 2+ and Zn 2+ [1, 4, 8, 28, 62], while tannase synthesis by A. niger [6] and P. 

chrysogenum [50] is inhibited by Zn +2 . On the other hand, in the present investigation, 

tannase synthesis by A. niger was inhibited in the following descending order: Ba 2+ , Co 2+ , 

Fe 2+ in LSF and Co 2+ , Ba 2+ and Fe 2+ in SSF and SlSF. Ba 2+ and Co 2+ also inhibit tannase 

synthesis by A. niger and A. pullulans [4]. Inhibition mediated by Fe 2+ is in agreement with 

the results of Rajakumar and Nandy [50] and Barthomeuf [6] on tannase synthesis by A. 

niger, A. pullulans and P. chrysogenum but disagree with those of Banerjee [4] and Trevino- 

Cueto [62] on tannase synthesised by A. aculeatus and A. niger, respectively. The presence of 

some metal ions in the fermentation medium may inhibit tannase synthesis due to their 

binding with tannic acid to form insoluble complexes, hence, limiting its bioavailability as a 

carbon source and inducer for enzyme synthesis [22]. This assumption was excluded in the 

present study where raising the level of tannin concentration in the fermentation medium 

containing the inhibitory metal salts (i.e. Ba 2+ , Co 2+ , or Fe 2+ ) did not show a significant 

enhancement effect on tannase synthesis (P > 0.05, data not shown). On the other hand, 

possible inhibition due to the binding of tannic acid to metal ions affecting the metal 

bioavailability, which in turn affects efficiencies of metabolic processes, secretion of active 

enzymes and the ability of the microorganisms to attach to their various substrates [38], was 

also ruled out since inhibition of tannase synthesis by A. niger was more pronounced at higher 

concentrations of Ba 2+ , Co 2+ or Fe 2+ (Table 3). The previous discussion suggests that the 

effect of metals is, most probably, on the activity of the secreted tannase rather than on 

tannase synthesis itself, consequently affecting the ability of A. niger to utilize the available 

tannin in the medium. Moreover, reduction in tannase productivity at higher concentrations



of metal ions could be due to the partial denaturation of some other important enzymes 

because of the presence of excessive free ions in the media [63], nonspecific binding or 

aggregation of the enzyme [28] or possible osmotic stress on the cells. 

Table 3: Effect of different metal salts at different concentrations on tannase production by A. niger in different 

fermentation protocols. 

Metal 

salt 

BaCl2 

CaCl2 

CoCl2 

FeCl2 

KCl 

MnCl 

MgCl2 

NaCl 

ZnCl2 

2 

Conc. 

(g %, 

w/v) 

Tannase productivity (U 50 ml -1 ) 

LSF 1 SSF 2 SlSF 3 

0.1 60.8 ± 2.37 * 167.64 ± 5.69 * 107.60 ± 4.41 * 

0.2 53.2 ± 1.17 * 146.68 ± 3.81 * 83.00 ± 2.15 * 

0.3 41.04 ± 0.98 * 131.44 ± 4.86 * 56.87 ± 1.42 * 

0.1 296.4 ± 9.18 * 321.94 ± 13.19 * 265.93 ± 9.04 * 

0.2 278.16 ± 5.28 * 280.03 ± 8.68 * 232.11 ± 7.42 * 

0.3 264.48 ± 11.37 * 253.36 ± 7.34 * 222.89 ± 6.46 * 

0.1 136.8 ± 3.83 * 62.865 ± 2.01 * 75.32 ± 2.86 * 

0.2 63.84 ± 2.17 * 59.05 ± 2.24 * 58.41 ± 2.27 * 

0.3 57.76 ± 2.02 * 36.19 ± 0.83 * 39.96 ± 0.91 * 

0.1 141.36 ± 4.09 * 173.35 ± 5.37 * 115.29 ± 3.22 * 

0.2 133.76 ± 4.14 * 152.4 ± 5.18 * 107.60 ± 4.19 * 

0.3 110.96 ± 4.32 * 133.35 ± 4.26 * 93.77 ± 3.46 * 

0.1 151.24 ± 5.44 n 190.5 ± 5.52 n 152.18 ± 5.02 n 

0.2 152 ± 4.25 n 192.40 ± 7.50 n 158.33 ± 3.64 n 

0.3 155.04 ± 3.25 n 203.84 ± 7.94 156.79 ± 3.29 n 

0.1 121.6 ± 4.37 * 169.54 ± 6.61 * 138.35 ± 4.98 * 

0.2• 152 ± 5.77 190.5 ± 4.95 153.72 ± 3.38 

0.3 232.56 ± 6.27 * 276.23 ± 9.11 * 182.93 ± 5.67 * 

0.1 200.64 ± 7.42 * 234.31 ± 5.38 * 201.37 ± 6.24 * 

0.2 205.2 ± 3.89 * 262.89 ± 8.41 * 204.45 ± 4.49 * 

0.3 212.8 ± 5.32 * 266.7 ± 10.40 * 216.74 ± 9.31 * 

0.1 152 ± 4.86 n 190.5 ± 4.00 n 146.03 ± 4.23 ** 

0.2 165.68 ± 6.79 * 190.5 ± 8.19 n 139.88 ± 4.89 * 

0.3 167.2 ± 6.35 * 190.5 ± 3.61 n 138.35 ± 4.70 * 

0.1 183.92 ± 5.70 * 211.45 ± 5.49 * 178.32 ± 4.27 * 

0.2 196.08 ± 4.50 * 190.5 ± 7.23 n 184.46 ± 7.93 * 

0.3 183.92 ± 5.88 * 188.95 ± 6.04 n 179.85 ± 3.95 * 

* = highly significant (P < 0.01), ** = significant (P > 0.01 and < 0.05) and n = non significant (P > 0.05) in 

comparison to • Control (i.e., the original fermentation medium). 

1 Liquid-state fermentation inoculated with immobilised cells of A. niger and grown on 7 % crude tannic acid 

extract at 30°C, 200 rpm, and 2 , 3 Solid-state and slurry-state fermentations using F. nitida and inoculated with 

free cells of A. niger. 

The investigated metal salts were separately added at the indicated concentrations to the fermentation media. No 

growth was determined in the presence of HgCl2 or PbCl2. Initial pH was adjusted to 5 with ammonium 

hydroxide instead of NaOH to prevent the interference from sodium ions. 

The effect of adding all the possible combinations of Mg 2+ , Ca 2+ , Mn 2+ and Zn 2+ , 

considered as stimulators of tannase biosynthesis by A. niger, to the fermentation media was 

investigated and the results showed that any such combination enhanced the levels of tannase 

synthesis at variable significant levels (Table 4). The maximum stimulatory effect was 



attained in the presence of Mg 2+ , Ca 2+ , Mn 2+ and Zn 2+ with a 2.9 (451 U 50 ml -1 ), 2.4 (457.2 

U 50 ml -1 ) and 2.1 (324.3 U 50 ml -1 ) fold increment in tannase synthesis in LSF, SSF and 

SlSF, respectively. Optimising the mineral nutrition requirements in the fermentation media 

for tannase synthesis by different microorganisms enhanced the yield at different levels, i.e., 

by 2.8 (A. aculeatus), and 1.9 (A. pullulans) fold [4] and 4 fold in the case of B. licheniformis 

[39]. The exerted stimulatory effect of the combinations of metallic salts on tannase synthesis 

by A. niger is the net result of the interactions between the available metals either in a 

synergetic or antagonistic manner. For example, a balanced mixture of K 2+ and Mg 2+ salts 

enhances tannase synthesis by B. licheniformis [39] while the stimulatory effect of Ca 2+ on 

tannase production by yeasts declines at higher concentrations due to the known antagonistic 

effect of calcium on magnesium uptake and its consequent effect on some essential magnesiumdependent 

metabolic processes [66]. Moreover, the known depressive effect of Mn 2+ on 

respiration, nucleic acid synthesis and proteins synthesis can be antagonised by Ca 2+ [70]. 

Repeated batch culture 

The efficiency of the immobilised A. niger for tannase production in a repeated batch 

process was evaluated in 250 ml Erlenmeyer flasks and the productivity over 7 cycles is given 

in Fig. 4. Significant amounts of tannase were produced up to the 5 th cycle (266.54 U 50 ml -1 , 

representing 59% of the production at the 1 st cycle), while almost stable amounts were 

produced during the first 3 cycles. In this respect, immobilised cultures of A. niger have 

exhibited significant tannase production stability of two repeated runs [17], while tannase 

production by immobilised cells of B. licheniformis have been demonstrated to be successful 

for up to 13 cycles, with its maximum level being reached at the 3 rd cycle [40]. 

Table 4: The effect of adding different mixtures of metal salts on tannase production by A. niger in different 

fermentation protocols 

Minerals 1 

Tannase productivity (U 50 ml -1 LSF 

) 

SSF SlSF 

(Original Medium) Control 152 190.5 153.7 (100) 

Ca 2+ 296.4 ± 8.00 * 321.95 ± 12.55 * 265.90 ± 5.84 * 

Mg 2+ 232.56 ± 6.97 * 276.23 ± 9.94 * 182.90 ± 6.95 * 

Mn 2+ 212.8 ± 5.10 * 266.7 ± 8.26 * 216.72 ± 4.11 * 

Zn 2+ 196.1 ± 8.43 * 211.46 ± 5.92 * 184.44 ± 6.27 * 

Ca 2+ + Mg 2+ 375.44 ± 13.14 * 329.56 ± 9.55 * 276.66 ± 8.57 * 

Ca 2+ + Mn 2+ 320.72 ± 8.02 * 346.71 ± 11.09 * 290.49 ± 11.91 * 

Ca 2+ + Zn 2+ 314.64 ± 11.95 * 323.85 ± 10.03 * 272.05 ± 6.25 * 

Mg 2+ + Mn 2+ 264.48 ± 10.31 * 304.8 ± 10.97 * 233.62 ± 6.54 * 

Mg 2+ + Zn 2+ 255.36 ± 5.36 * 297.18 ± 10.99 * 210.57 ± 4.00 * 

Mn 2+ + Zn 2+ 258.4 ± 9.30 * 291.47 ± 11.07 * 230.55 ± 6.22 * 

Ca 2+ + Mg 2+ + Mn 2+ 351.12 ± 13.69 * 379.10 ± 8.34 * 282.81 ± 6.78 * 

Ca 2+ + Mg 2+ + Zn 2+ 319.2 ± 9.89 * 342.9 ± 6.51 * 276.66 ± 11.89 * 

Ca 2+ + Mn 2+ + Zn 2+ 349.6 ± 10.13 * 375.29 ± 11.63 * 305.86 ± 11.92 * 

Mg 2+ + Mn 2+ + Zn 2+ 270.56 ± 7.57 * 302.90 ± 10.29 * 233.62 ± 8.17 * 

Ca 2+ + Mg 2+ + Mn 2+ + Zn 2+ 451.44 ± 16.25 * 457.2 ± 12.80 * 324.31 ± 10.54 * 

1 

Added at the optimum concentration enabled maximum tannase synthesis shown in Table 3 

* 

highly significant (P < 0.01) in comparison to control 

Significant production over repeated cycles could be due to long term cell viability, 

reducing the damage of fungal mycelia and continuous metabolic activities that are known to 

be advantages for immobilised cells [17]. The observed decrease in enzyme productivity after 

several cycles could be attributable to the high growth of biomass, increasing cell density 




and/or the presence of dead cells that cause diffusional limitation of oxygen, substrate and 

product inside and out of the beads, in addition to the mass transfer limitation that is still the 

major drawback in the application of an entrapment technique [68]. 

Total activity (U / 50 ml) 

500 

400 

300 

200 

100 

0 

1 2 3 4 5 6 7 

Number of cycle 

Fig. 4. Repeated batch culture for tannase production using immobilised cells of A. niger 

The presence of large amounts of tannins in human food and animal forages reduces 

the nutritive value [22] and has serious health implications [15]. Biological processes reduce 

the antinutritional effects of tannin and improve digestibility by adopting tannase preparations 

or various tannase-producing fungal strains [42]. Meanwhile, the fermentation residue 

resulting from SSF for tannase production is an agricultural waste that has already been 

detannified and may represent an extra added value if used in animal feed. To this end, tannin 

and protein contents of the leaves of F. nitida were monitored throughout fermentation and 

the results are given in Fig. 5 which show that tannin levels reduced by 27.33 % after 120 h of 

fermentation. A reduction in tannin content due to fermentation by A. oryzae and B. 

licheniforms utilising different substrates range from 0.1 %, (Psidium guazava), 20 to 35% 

(Acacia auriculiformis, Casuarina equisetifolia and Delonix regia); and up to 53 to 75% 

(Anacardium occidentale, Cassia fistula, Eucalyptus tereticornis, and Ficus benghalensis) 

[34, 39]. Meanwhile, 90% of the tannin content of creosote bush plant materials is degraded 

by A. niger [9]. Such variations in tannin biodegradation and digestibility depend on the 

nature of leaves, fungi and fermentation times. The residual amount of tannin in the 

fermentation residue of F. nitida (10.90 %) might benefit ruminants by protecting proteins 

from microbial deamination and preventing bloats [22]. Meanwhile, in the present study, the 

protein content of F. nitida leaves increased by 24.33 % after 120 h of incubation before 

decreasing (Fig 5). This decrease in tannase biosynthesis could be because of changes in the 

prevailing conditions and physiology of the fungus where protein serves as a substrate. The 

protein content of creosote bush (Larrea tridentata) and tar bush (Flourensia cernua) used in 

SSF for tannase production by A. niger PSH and GH1 has been shown to increase by 30% and 

11 %, and 0.62%, 0.76%, respectively [9]. 


Total protein (%) 

10 

8 

Total tannin (%) 

6 

4 

2 

0 


Total Protein (%) Tannin (%) 

0 24 48 72 96 120 144 

Incubation time (h) 

Fig. 5: Changes in tannin and protein contents of the fermented F. nitida' leaves used for the production of 

tannase by A. niger. 

Possible applications of the partially purified tannase produced by A. niger was 

investigated by reducing tannin levels in tea extract and pomegranate juice. 55% of tannin 

was hydrolysed after 30 minutes of treating tea extract with A. niger tannase, while 45 % was 

removed from pomegranate juice after 120 minutes (Table 5). Tannase from P. 

atramentosum has been shown to reduce 38% of jamum wine; 43.5% of grape wine and 74% 

of tea extract after 3 h at 35° C [56] while 25% of tannin content of pomegranate juice has 

been reported to be reduced [53]. 

Table (5): Hydrolysis of tannin content of tea extract and pomegranate juice with treatment of partially purified 

tannase from A. niger 

Incubation time 

(Minutes) 


20 

15 

10 

% Tannin hydrolysis 

Tea extract Pomegranate 

0 100 % (175.12 µg ml -1 ) 100 % (975 µg ml -1 ) 

30 22 17 

60 29 21 

90 38 32 

120 55 45 

150 55 44 

1 ml of the partially purified tannase containing 17.32 U/ mg protein was incubated at 30° C in a rotary 

shaker (150 rpm) with 15 ml of tea extract or pome granate juice at pH 5 for the indicated time periods. 

Concluding remarks 

In the present work, we report for the first time the production of tannase by A. niger 

using F. nitida leaves and crude tannic acid extract as a cost-effective alternative to largescale 

production where pure tannic acid is a very costly substrate. The optimum temperature 

for tannase production was recorded at 30°C, indicating A. niger as a suitable candidate for 

5 

0



tannase production at room temperature using F. nitida leaves in tropical countries such as 

Egypt with minor control processes where much of the necessary energy cost can be reduced. 

Parameters of tannase production (incubation period, incubation temperature, shaking 

velocity, initial pH, initial tannic acid concentration and mineral nutritional requirements) 

were optimised and increased production by 5.82, 5.61, and 4.29 fold in LSF, SSF, SlSF, 

respectively. Moreover, suitability of the SSF residue as a candidate for animal feed is 

suggested for further research. We also demonstrated the potentiality of the partially purified 

tannase for application in the removal of tannin from tea extract and pomegranate juice. 

References 

1. M. A. ABDEL-NABEY, A. A. SHERIEF, A. B. EL-TANASH, Tannin biodegradation and some 

factors affecting Tannase production by two Aspergillus sp. Biotechnology. 10(2): 149-158 (2011). 

2. C. N.AGUILAR, C. AUGUR, E. FAVELA, Z. VINIEGRA-GONZALEZ, Production of tannase by 

Aspergillus niger Aa-20 in submerged and solid-state fermentation: influence of glucose and tannic 

acid. J. Ind. Microbiol. Biotechnol. 26: 296-302 (2001). 

3. I. T. BALDWIN, J. C. SCHULTZ, D. WARD, Patterns and sources of leaf tannin variation in yellow 

birch (Betula allegheniensis) and sugar maple (Acer saccharum). J. Chem. Ecol. 13, 1069-1078 (1987). 

4. S. D. BANERJEE, Microbial production of tannase and gallic acid. Ph. D. Thesis, Vidyasagar 

University, India. (2005). 

5. D. Banerjee, K.C. Mondal, B.R. Pati Tannase production by Aspergillus aculeatus DBF9 through solidstate 

fermentation. Acta Microbiol Immunol Hung; 54(2):159-66 (2007). 

6. C. BARTHOMEUF, F. Regerat, H. Pourrat, Production, purification and characterization of a tannase 

from A. niger LCF8. J. Ferment. Bioeng. 77(3): 320-323 (1994). 

7. V. BATTESTIN, G.A. MACEDO, Effects of temperature, pH and additives on the activity of tannase 

produced by Paecilomyces variotii. Elec. J. Biotechnol. Vol. 10(2): 191 -199 (2007). 

8. R. BELMARES, J.C. CONTRERAS-ESQUIVEL, R. RODRÍGUEZ- HERRERA, A. RAMÍREZ- 

CORONEL, C.N. AGUILAR, Microbial production of tannase: an enzyme with potential use in food 

industry. Lebensm. Wiss. U.- Technol. 37: pp. 857 – 864 (2004). 

9. R. BELMARES, Y. GARZA, R. RODRÍGUEZ, J. CONTRERAS-ESQUIVEL, C. Aguilar, 

Composition and fungal degradation of tannins present in semiarid plants. Electronic J. Environ. Agric. 

Food Chem. 8(7), 512-518 (2009). 

10. P. D. BELUR, G. MUGERAYA, Microbial production of tannase: State of the art. Res. J. Microbiol. 6, 

25-40 (2011). 

11. V. BENIWAL, V. CHHOKAR, Statistical optimization of culture conditions for Tannase production by 

Aspergillus awamori MTCC 9299 under submerged fermentation. Asian J. Biotechnol. 2(1), 46-52 

(2010). 

12. R. BHARDWAJ, B. SINGH, T.K. BHAT, Purification and characterization of tannin acyl hydrolase 

from Aspergillus niger MTCC 2425. J. Basic Microbiol. 43, 449-461 (2003). 

13. T.K. BHAT, H.P. MAKKAR, B. SINGH, Preliminary studies on tannin degradation by Aspergillus 

niger van Tieghem MTCC 2425. Lett. Appl. Microbiol. 25, 22-23 (1997). 

14. M. M. BRADFORD, A Rapid and Sensitive Method for the Quantitation of Microgram Quantities of 

Protein Utilizing the Principle of Protein-Dye Binding. Anal. Biochem. 72. pp. 248 – 254 (1976). 

15. K.T. CHANG, T.Y. WONG, C.I. WEI, Y.W. HUANG, Y. LIN, Tannins and human health: a review. 

Crit. Rev. Food Sci. Nutr. 38(6), 421-464 (1998). 

16. M. CHAVEZ-GONZALEZ, L.V. RODRIGUEZ-DURAN, N. BALAGURUSAMY, A. PRADO- 

BARRAGAN, R. RODRIGUEZ, J. CONTRERAS, C.N. AGUILAR, Biotechnological advances and 

challenges of tannase: An overview. Food Bioprocess Technol. Published online first. DOI 

1001007/s11947-011-0608-5 (2011). 

17. I. DARAH, G. SUMATHI, K. JAIN, S.H. LIM, Tannase enzyme production by entrapped cells of 

Aspergillus niger FELT FT3 in submerged culture system. Bioprocess Biosyst. Eng. 19(13), 1-7 (2011). 

18. A. DE-GREGORIO, G. MANDALARI, N. ARENA, F. NUCITA, M.M. TRIPODO, R.B. Lo CURTO, 

SCP and crude pectinase production by slurry-state fermentation of lemon pulps. Biores. Technol. 83, 

89-94 (2002). 

19. M.Z. EL-FOULY, Z. EL-AWAMRY, A.M. SHAHIN, H. EL-BIALY, E. NAEEM, G. EL-SAEED, 

Biosynthesis and characterization of Aspergillus niger AUMC 4301 tannase. J. American Sci., 6(12), 709- 

721 (2010). 



20. A. GATHON, Z. GROSS, M. ROZHANSKI, Propyl gallate: enzymatic synthesis in a reverse micelle 

system. Enzyme Microb Technol. 11, 604-9 (1989). 

21. AS. Glantz, Primer of biostatistics. McGraw Hill, Inc., USA. pp. 2-18 (1992). 

22. G. Goel, A. Puniya, C. Aguilar, K. Singh, Interaction of gut microflora with tannins in feeds. 

Naturwissenschaften, 92(11), 497-503 (2005). 

23. H. S. Hamdy, Purification and characterization of a newly isolated stable long-life tannase produced by F. 

subglutinans (Wollenweber and Reinking) Nelson et al. J. Pharm. Innov. 3(3), 142-151 (2008). 

24. E. Haslam, J.N. Tanner, Spectrophotometric assay of tannase. Phytochem; 90, 2305-2309 (1970). 

25. M. A. Jermyn, Cellulose and hemicellulose. In: Peach K., Tracey M. V., Eds., Modern Methods of Plant 

Analysis. 2, 197-224 (1955). 

26. C.S. Jun, M.J. Yoo, W.Y. Lee, K.C. Kwak, M.S. Bae, W.T. Hwang, D. H. SON, K.Y. Chai, Ester derivatives from 

tannase-treated prunioside A and their anti-inflammatory activities. Bull Korean Chem. Soc. 28, 73-76 (2007). 

27. B. Kar, R. Banerjee, B.C. Bhattacharya, Microbial production of gallic acid by modified solid state 

fermentation., J Ind Microbiol Biotechnol. 23,173-7. (1999). 

28. B. Kar, R. Banerjee, BC. Bhattacharyya, Effect of additives on the behavioural properties of tannin acyl 

hydrolase. Process Biochem. 38,1285-93 (2003). 

29. J. Kitajima, K. Kimizuka, Constituents of Ficus pumila leaves. Chem. Pharm. Bull. Tokyo, 46(10), 1647- 

1649 (1998). 

30. R.P. Learmonth, E. Gratton, Assessment of membrane fluidity in individual yeast cells by Laurdan 

Generalised polarization and multi-photon scanning fluorescence microscopy, In: Fluorescence 

Spectroscopy, Imaging and probes- New tools in Chemical, Physical and life science, Springer Series on 

Fluorescence: Methods and Applications, Springer, Heidelberg, 2, 241-252. (2002). 

31. P.K. Lekha, B.K. Lonsane, Comparative titers, location and properties of tannin acyl hydrolase produced by 

Aspergillus niger PKL 104 in solid state, liquid surface and submerged fermentation. Process Biochem. 29, 

497-503. (1994). 

32. P.K. LEKHA, B.K. LONSANE, Production and application of tannin acyl hydrolase. State of the art. Adv. 

Appl. Microbiol. 44, 215-260 (1997). 

33. K. LEXANDER, E. CARLSSON, V. SCHALEN, A. SINONSSON, T. LUNDBORG, Quantities and 

qulaities of leaf protein concentrates from wild species and crop species grown under controlled conditions. 

Ann. App. Biol. 66(2), 193-216 (1970). 

34. N. LOKESWARI, Production of tannase through submerged fermentation of tannin-containing cashew husk 

by Aspergillus oryzae. Rasayan J. Chem. 3(1), 32-37 (2010). 

35. N. LOKESWARI, K. JAYA-RAJU, S. POLA, V. BOBBARALA, Tannin acyl hydrolase from 

Trichoderma viride. Int. J. Chemical and Analytical Science. 1(5), 106-109 (2010). 

36. B. K. LONSANE, N.P. GHILDYAL, S. BUDIATMAN, S.V. RAMAKRISHNA, Engineering aspects of 

solid-state fermentation. Enzyme Microbiol. Technol. 7, 258-265 (1985). 

37. M.J. LU, C. CHEN, Enzymatic modification by tannase increases the antioxidant activity of green tea. Food 

Res. Int. 41, 130-137 (2008). 

38. M. MCDONALD, I. MILA, A. SCALBERT, Precipitation of metal ions by plant polyphenols: optimal 

conditions and origin of precipitation. J. Agric. Food Chem. 44, 599-606 (1996). 

39. P.K. MOHAPATRA, K.C. MONDAL, B.K. PATI, Production of tannase through submerged fermentation 

of tannin-containing plant extracts by Bacillus lichiniformis KBR6. Polish J. Microbiol. 55(4), 297- 

301(2006). 

40. P.K.D. MOHAPATRA, K.C. MONDAL, B.R. PATI, Production of tannase by the immobilized cells of 

Bacillus licheniformis KBR6 in Ca-alginate beads. J. Appl. Microbiol. 102(6), 1462-1467 (2007). 

41. K.C. MONDAL, D. BANERJEE, R. BANERJEE, B.R. Pati, Production and characterization of tannase 

from Bacillus cereus KBR9. J. Gen. Appl. Microbiol., 47, pp. 263 – 267 (2001). 

42. O.M. NUERO, F. REYES, Enzymes for animal feeding from Penicillium chrysogenum mycelial wastes 

from penicillin manufacture. Lett. Appl. Microbiol. 34(6), 413-416 (2002). 

43. A. PANDEY, Solid state fermentation: An overview. In: Solid-state fermentation, A. Pandey (Ed.), Wiley 

Eastern Ltd., New Delhi, India, PP, 3-10(1994). 

44. A. PANDEY, P. SELVAKUMAR, C.R. SOCCOL, P. NIGAM, Solid-state fermentation for the production 

of industrial enzymes. Curr. Sci. 77: 146-162 (1999). 

45. R. PARANTHAMAN, R.R. VIDYALAKSHMI, S. MURUGESH, K. SINGARAVADIV, Optimization of 

fermentation conditions for production of tannase enzyme by Aspergillus oryzae using sugarcane bagasse 

and rice straw. Global J. Biotechnol. Biochemist., 3, 105-110 (2008). 

46. R. PARANTHAMAN, R. VIDYALAKSHMI, S. MURUGESH, K. SINGARAVADIVE, Effects of fungal 

co-culture for the biosynthesis of tannase and gallic acid from grape wastes under solid state fermentation. 

Glob. J Biotechnol. Biochem. 4(1), 29-36 (2009). 




47. G.C. PAUL, C.A. KENT, C.R. THOMAS, Hyphal vaculoation and fragmentation in Penicillium 

chrysogenum. Biotechnol. Bioeng. 44 655-660 (1994). 

48. N.W. PIRIE, Proteins in modern methods of plant analysis. FIZCL. Biokhim. Kul't Rast., 8(6) 619-625. [C. 

F. Chemical Abstract., 86(9), 52761 (1955). 

49. J.S. PUROHIT, J.R. DUTTA, R.K. NANDA, R. BANERJEE, Strain improvement for tannase production 

from co-culture of Aspergillus foetidus and Rhizopus oryzae. Biores Technol; 97,795-801 (2006). 

50. G.S. RAJAKUMAR, S.C. NANDY, Isolation, purification, and some properties of Penicillium chrysogenum 

tannase. Appl. Environ. Microbiol. 46, 525-527 (1983). 

51. S.V. RAMAKRISHNA, R.S. PRAKASHAM, Microbial fermentatioins with immobilized cells. Curr. Sci. 

77, 87-100 (1999). 

52. J.D. REED, Nutritional toxicology of tannins and related polyphenols in forage legumes. J. Animal Sci. 

73(5), 1516-28 (1995). 

53. S. ROUT, R. BANERJEE, Production of Tannase under mSSF and its application in fruit juice debittering. 

Ind. J. Biotechnol. 5, 346-350 (2006). 

54. A. SABU, A. PANDEY, M. JAAFAR-DAUD, G. SZAKACS, Tamarind seed powder and palm kernel 

cake: two novel agro residues for the production of tannase under solid state fermentation by Aspergillus 

niger ATCC 16620. Bioresource Technology. 96, 1223–1228 (2005). 

55. S.H. SCHANDERI, Methods in Food Analysis. Academic Press. New York. P, 709 (1970). 

56. M. SELWAL, A. YADAV, K. SELWAL, N.K. AGGARWAL, K. GUPTA, S.K. GAUTAM, Tannase 

production by Penicillium atramentosum KM under SSF and its applications in wine clarification and tea 

cream solubilization. Braz. J. Microbiol. 42, 374-387 (2011). 

57. M. SETH, S. CHAND, Biosynthesis of Tannase and hydrolysis of tannins to gallic acid by Aspergillus 

awamori- optimization of process parameters. Proc. Biochem. 36, 39-44 (2000). 

58. S. SHARMA, L. AGARWAL, R.K. SAXENA, Statistical optimization for tannase production from 

Aspergillus niger under submerged fermentation. Indian J. Microbiol. 47, pp. 132 – 138 (2007). 

59. A. SRIVASTAVA, R. KAR, Characterization and application of tannase produced by Aspergillus niger 

ITCC 6514.07 on pomegranate rind. Braz. J. Microbiol. 40, 782-789 (2009) 

60. K. STORMO, R. CRAWFORD, Preparation of encapsulated microbial cells for environmental applications. 

App. Environ. Microbiol. 58 (2), 727-730 (1992). 

61. D.M. STUART, D.A. MITCHELL, J.D. JOHNS, LITSTER, Solid-state fermentation in rotating drum 

bioreactors: operating variables affect performance through their effects on transport phenomena. 

Biotechnol. Bioeng. 63, 383-391 (1999). 

62. B. TREVINO-CUETO, M. LUIS, J.C. CONTRERAS-ESQUIVEL, R. RODRIGUEZ, A. AGUILERA, 

C.N. AGUILAR, Gallic acid and tannase accumulation during fungal solid state culture of a tannin-rich 

desert plant (Larrea tridentata Cov.). Biores. Technol. 98, 721-724 (2007). 

63. R. TUNGA, R. BANERJEE, B.C. BHATTACHARYYA, Optimization of n variable biological experiments 

by evolutionary operation-factorial design technique. J. Biosci. Bioeng. 87(2), 125-131. (1999). 

64. G. URBANO, M. LOPEZ-JURADO, J.M. PORRES, S. FREJNAGEL, E. GOMEZ-VILLALVA, J. FRIAS, 

C. VIDALVALVERDE, P.ARANDA, Effect of treatment with α-galactosidase, Tannase or a cell-walldegrading 

enzyme complex on the nutritive utilization of protein and carbohydrates from pea (Pisum sativum 

L.) flour. J. Sci. Food Agric. 87, 1356-1363. (2007). 

65. J. VAN DE LAGEMAAT, D.L. PYLE, Solid state fermentation and bioremediation: development of a 

continuous process for the production of fungal tannase. Chem. Engin. J. 84, 115-123. (2001). 

66. G. WALKER, R. DE NICOLA, S. ANTHONY, R. Learmonth, Yeast-metal interactions: impact on brewing 

and distilling Asia Pacific Section. 19-24 March, Hobart, Tasmania. (2006) 

67. H.H. WANG, A review: Development and/or reclamation of bioresources with solid state fermentation. 

Proc. Natl. Sci. Counc. Roc (B). 23(2), 45-61. (1999). 

68. X. WANG, Z GAI, B. YU, J. FENG, C. XU, Y. YUAN, Z. LIN, P. XU, Degradation of carbazole by 

microbial cells immobilized in magnetic gellan gum gel beads. Appl. Environ. Microbiol. 73(20), 6421-6428 

(2007). 

69. H.H. WEETAL, Enzymatic gallic acid esterification. Biotechnol. Bioeng. 27, 124-127 (1985). 

70. E.D. WEINBERG, Roles of metallic ions in host-parasite interactions. Bacteriol. Reviews. 30(1), 136-151 

(1966). 

71. Y. YALCINKAYA, M.Y. ARICA, L. SOYSAL, A. DENIZLI, O. GENC, S. BEKTAS, Cadmium and 

mercury uptake by immobilized Pleurotus sapidus. Turk J. Chem. 26, 441-452 (2002).

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