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Romanian Biotechnological Letters Copyright © 2012 University of Bucharest

Vol.17, No.4, 2012 Printed in Romania. All rights reserved ORIGINAL PAPER

Economic Production of Tannase by Aspergillus niger van Tiegh Adopting Different Fermentation Protocols and Possible Applications Received for publication, February 7, 2011 Accepted, October 31, 2011 H. S. HAMDY1 AND E. M. FAWZY2

Biological Sciences & Geology Department, Faculty of Education, Ain Shams University, Roxy, Heliopolis, Cairo, 11757, Egypt 1 [email protected] [email protected]

Abstract 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 Ca2+, Mg2+, Mn2+ and Zn2+ 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 Romanian Biotechnological Letters, Vol. 17, No. 4, 2012

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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

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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 106 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.

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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 103 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 microKjeldahl 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. 7444

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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].

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H. S. HAMDY AND E. M. FAWZY Table 1: Chemical composition of Ficus nitida’ leaves.

Chemical composition (%) Cellulose1

Hemicellulose1

Lignin1

Pectin1

Tannin2

Ash

Soluble N

Insoluble N

Total N

Total protein

Nitrogen (N) and protein content3

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

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

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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.

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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.

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H. S. HAMDY AND E. M. FAWZY Table 2: The effect of varying initial crude tannin concentration on tannase production by A. niger in different fermentation protocols.

Crude tannin conc. (%) 3 (control) 4 5 6 7 8 9

LSF1 U 50 ml-1 96.11 ± 2.21 123.00* ± 3.93 131.67* ± 5.13 138.39* ± 2.90 146.10* ± 6.28 143.20* ± 2.72 131.67* ± 3.42

% 100 128 137 144 152 149 137

Tannase productivity* SSF2 U 50 ml-1 % 114.76* ± 4.36 100 161.81* ± 5.17 141 174.43* ± 6.27 152 182.47* ± 4.01 159 190.50* ± 5.90 166 183.62* ± 7.52 160 161.81* ± 4.20 141

SlSF3 U 50 ml-1 109.80 ± 2.74 132.86* ± 4.51 133.95* ± 4.28 153.72* ± 4.45 142.74* ± 5.42 131.76* ± 5.13 117.49* ± 2.70

% 100 121 122 140 130 120 107

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: Ca2+, Mg2+, Mn2+, Zn2+ in LSF and SSF, and Ca2+, Mn2+, Mg2+, Zn2+ 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 Ca2+, Mg2+, Mn2+ and Zn2+ [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: Ba2+, Co2+, Fe2+ in LSF and Co2+, Ba2+ and Fe2+ in SSF and SlSF. Ba2+ and Co2+ also inhibit tannase synthesis by A. niger and A. pullulans [4]. Inhibition mediated by Fe2+ 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 TrevinoCueto [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. Ba2+, Co2+, or Fe2+) 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 Ba2+, Co2+ or Fe2+ (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 7450 Romanian Biotechnological Letters, Vol. 17, No. 4, 2012

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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.

2

ZnCl2

NaCl

MnCl

MgCl2

KCl

FeCl2

CoCl2

CaCl2

BaCl2

Metal salt

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

Tannase productivity (U 50 ml-1)

Conc. (g %, w/v)

LSF1

SSF2

SlSF3

0.1 0.2 0.3 0.1 0.2 0.3 0.1 0.2 0.3 0.1 0.2 0.3 0.1 0.2 0.3 0.1 0.2• 0.3 0.1 0.2 0.3 0.1 0.2 0.3 0.1 0.2 0.3

60.8 ± 2.37* 53.2 ± 1.17* 41.04 ± 0.98* 296.4 ± 9.18* 278.16 ± 5.28* 264.48 ± 11.37* 136.8 ± 3.83* 63.84 ± 2.17* 57.76 ± 2.02* 141.36 ± 4.09* 133.76 ± 4.14* 110.96 ± 4.32* 151.24 ± 5.44n 152 ± 4.25n 155.04 ± 3.25n 121.6 ± 4.37* 152 ± 5.77 232.56 ± 6.27* 200.64 ± 7.42* 205.2 ± 3.89* 212.8 ± 5.32* 152 ± 4.86n 165.68 ± 6.79* 167.2 ± 6.35* 183.92 ± 5.70* 196.08 ± 4.50* 183.92 ± 5.88*

167.64 ± 5.69* 146.68 ± 3.81* 131.44 ± 4.86* 321.94 ± 13.19* 280.03 ± 8.68* 253.36 ± 7.34* 62.865 ± 2.01* 59.05 ± 2.24* 36.19 ± 0.83* 173.35 ± 5.37* 152.4 ± 5.18* 133.35 ± 4.26* 190.5 ± 5.52n 192.40 ± 7.50n 203.84 ± 7.94 169.54 ± 6.61* 190.5 ± 4.95 276.23 ± 9.11* 234.31 ± 5.38* 262.89 ± 8.41* 266.7 ± 10.40* 190.5 ± 4.00n 190.5 ± 8.19n 190.5 ± 3.61n 211.45 ± 5.49* 190.5 ± 7.23n 188.95 ± 6.04n

107.60 ± 4.41* 83.00 ± 2.15* 56.87 ± 1.42* 265.93 ± 9.04* 232.11 ± 7.42* 222.89 ± 6.46* 75.32 ± 2.86* 58.41 ± 2.27* 39.96 ± 0.91* 115.29 ± 3.22* 107.60 ± 4.19* 93.77 ± 3.46* 152.18 ± 5.02n 158.33 ± 3.64n 156.79 ± 3.29n 138.35 ± 4.98* 153.72 ± 3.38 182.93 ± 5.67* 201.37 ± 6.24* 204.45 ± 4.49* 216.74 ± 9.31* 146.03 ± 4.23** 139.88 ± 4.89* 138.35 ± 4.70* 178.32 ± 4.27* 184.46 ± 7.93* 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 Mg2+, Ca2+, Mn2+ and Zn2+, 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 Romanian Biotechnological Letters, Vol. 17, No. 4, 2012

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attained in the presence of Mg2+, Ca2+, Mn2+ and Zn2+ 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 K2+ and Mg2+ salts enhances tannase synthesis by B. licheniformis [39] while the stimulatory effect of Ca2+ 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 Mn2+ on respiration, nucleic acid synthesis and proteins synthesis can be antagonised by Ca2+ [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 5th cycle (266.54 U 50 ml-1, representing 59% of the production at the 1st 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 3rd cycle [40]. Table 4: The effect of adding different mixtures of metal salts on tannase production by A. niger in different fermentation protocols Tannase productivity (U 50 ml-1) 1

Minerals

1 *

(Original Medium) Control Ca2+ Mg2+ Mn2+ Zn2+ Ca2+ + Mg2+ Ca2+ + Mn2+ Ca2+ + Zn2+ Mg2+ + Mn2+ Mg2+ + Zn2+ Mn2+ + Zn2+ Ca2+ + Mg2+ + Mn2+ Ca2+ + Mg2+ + Zn2+ Ca2+ + Mn2+ + Zn2+ Mg2+ + Mn2+ + Zn2+ Ca2+ + Mg2+ + Mn2+ + Zn2+

LSF 152 296.4 ± 8.00* 232.56 ± 6.97* 212.8 ± 5.10* 196.1 ± 8.43* 375.44 ± 13.14* 320.72 ± 8.02* 314.64 ± 11.95* 264.48 ± 10.31* 255.36 ± 5.36* 258.4 ± 9.30* 351.12 ± 13.69* 319.2 ± 9.89* 349.6 ± 10.13* 270.56 ± 7.57* 451.44 ± 16.25*

SSF 190.5 321.95 ± 12.55* 276.23 ± 9.94* 266.7 ± 8.26* 211.46 ± 5.92* 329.56 ± 9.55* 346.71 ± 11.09* 323.85 ± 10.03* 304.8 ± 10.97* 297.18 ± 10.99* 291.47 ± 11.07* 379.10 ± 8.34* 342.9 ± 6.51* 375.29 ± 11.63* 302.90 ± 10.29* 457.2 ± 12.80*

SlSF 153.7 (100) 265.90 ± 5.84* 182.90 ± 6.95* 216.72 ± 4.11* 184.44 ± 6.27* 276.66 ± 8.57* 290.49 ± 11.91* 272.05 ± 6.25* 233.62 ± 6.54* 210.57 ± 4.00* 230.55 ± 6.22* 282.81 ± 6.78* 276.66 ± 11.89* 305.86 ± 11.92* 233.62 ± 8.17* 324.31 ± 10.54*

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 7452

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Economic Production of Tannase by Aspergillus niger van Tiegh Adopting Different Fermentation Protocols and Possible Applications

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

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Total Protein (%)

Tannin (%)

20

Total tannin (%)

Total protein (%)

10 8

15

6

10

4 2

5

0

0 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) 0 30 60 90 120 150

% Tannin hydrolysis Tea extract Pomegranate 100 % (975 µg ml-1) 100 % (175.12 µg ml-1) 22 17 29 21 38 32 55 45 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 7454

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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.

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