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International Biodeterioration & Biodegradation 65 (2011) 1061e1065

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International Biodeterioration & Biodegradation

journal homepage: www.elsevier .com/locate/ ibiod

Degradation of tannic acid and purification and characterizationof tannase from Enterococcus faecalis

Gunjan Goela,b,*, Ashwani Kumarc, Vikas Beniwalb, Mamta Raghava, Anil K. Puniyaa, Kishan Singha

aDairy Microbiology Division, National Dairy Research Institute, Karnal 132001, IndiabDepartment of Biotechnology, Maharishi Markandeshwar University, Mullana, Ambala 133201, IndiacDepartment of Biotechnology, JMIT Institute of Engineering and Technology, Radaur 135133, India

a r t i c l e i n f o

Article history:Received 3 May 2011Received in revised form19 July 2011Accepted 12 August 2011Available online 15 September 2011

Keywords:TanninsTannaseEnterococcus faecalisPyrogallol

* Corresponding author. Department of Biotechnolwar University, Mullana, Ambala 133201, India. Tel.1731274375.

E-mail address: gunjanmicro@gmail.com (G. Goel)

0964-8305/$ e see front matter � 2011 Elsevier Ltd.doi:10.1016/j.ibiod.2011.08.006

a b s t r a c t

Tannins, present in various foods, feeds and forages, have anti-nutritional activity; however, presence oftannase in microorganisms inhabiting rumen and gastrointestinal tract of animals results in detoxifi-cation of these tannins. The present investigation was carried out to study the degradation profile oftannins by Enterococcus faecalis and to purify tannase. E. faecalis was observed to degrade tannic acid(1.0% in minimal media) to gallic acid, pyrogallol and resorcinol. Tannase from E. faecalis was purified upto 18.7 folds, with a recovery of 41.7%, using ammonium sulphate precipitation, followed by DEAE-cellulose and Sephadex G-150. The 45 kDa protein had an optimum activity at 40 �C and pH 6.0 atsubstrate concentration of 0.25 mM methyl gallate.

� 2011 Elsevier Ltd. All rights reserved.

1. Introduction

Tannins are the most common plant secondary metabolites thatare poorly understood with respect to feeding habits, with bothbeneficial and adverse effects in ruminants (Getachew et al., 2000).Tannins (hydrolysable (HT) and condensed (CT)), being antimicro-bial; resist microbial attack and remains as recalcitrant in theenvironment (Field and Lettinga, 1992). Though destructive innature, the biological systems often have difficulty in removingtoxic polyphenols to consistently low levels. However, some rumenmicroorganisms are shown to possess the ability to grow ontannins and degrade HTs, with a few evidences showing degrada-tion of phenol ring of CTs (Bhat et al., 1998). These microorganismshave developed protective mechanisms such as secretion ofbinding polymers and synthesis of tannin resistant enzymes fortheir biodegradation (Goel et al., 2005a). A number of reviews ontannin biodegradation have appeared in the past, providinga general idea on the biodegradation of these polyphenols (Williamet al., 1986; Bhat et al., 1998). It is reported that tannase, the keyenzyme, is involved in the hydrolysis of tannins (Aguilar and

ogy, Maharishi Markandesh-: þ91 1731304148; fax: þ91

.

All rights reserved.

Gutierrez-Sanchez, 2001). The tannase catalyse the hydrolysis ofester and depside bonds presents in the hydrolysable tannins andgallic acid esters (Lekha and Lonsane, 1997). Among tannindegrading microbes, aerobic bacterial degraders and fungal specieshave been reviewed elsewhere (Bhat et al., 1998; Goel et al., 2005b).The fungi have a better activity in degrading hydrolyzable tanninshowever, a major problem in the utilization of fungal strains forindustrial applications is that degradation by fungi is relatively slow(Beniwal et al., 2010). On the other end of the spectrum, bacterialtannase can degrade and hydrolyze natural tannins and tannic acidvery efficiently (Lewis and Starkey, 1969; Deschamp et al., 1983).Therefore, the present investigation was undertaken to study thedegradation profile of an anaerobic Enterococcus faecalis isolatedfrom faeces of goat and to characterise tannase from this bacterium.

2. Materials and methods

2.1. Biodegradation of tannic acid

Tannic acid degrading E. faecalis, used in the present investiga-tion was isolated from goat faeces (Goel et al., 2007). The overnightculture was centrifuged and cell pellet was collected and mixed(at the rate 106 cfu/ml) to the minimal media supplemented with1% filter sterilized tannic acid (Nelson et al., 1995). The culturebottles were sealed under CO2 and incubated at 37 �C for 72 h. The

Fig. 1. TLC profile of degradation of tannic acid with E. faecalis.

G. Goel et al. / International Biodeterioration & Biodegradation 65 (2011) 1061e10651062

samples for analysis were removed aseptically from the bottlesafter every 12 h interval. The samples were kept at �20 �C tillfurther analysis. The tannic acid degraded products were analysedby Thin Layer Chromatography (TLC) and High Performance LiquidChromatography (HPLC). All the experiments were done intriplicates.

2.1.1. Thin Layer Chromatography (TLC)Thin Layer Chromatography was done according to the protocol

of Sharma et al. (1999). Briefly, slurry of silica (40 g in 80 ml ofdistilled water) was spread on glass plates (20 � 20 cm) toa thickness 0.5 mm. The plates were allowed to dry and wereactivated by placing in an oven at 110 �C for 1 h. The samples werecentrifuged at 2000 � g and the supernatant was transferred infresh tube. An aliquot of 10 ml was spotted on the TLC plate.Amixture of ethyl acetate: chloroform: water (50:50:1) was used asmobile phase and the detection was done using iodine vapours ina saturated chamber. Rf values were calculated and compared withthe standards of gallic acid, pyrogallol and resorcinol.

2.1.2. RP-High Performance Liquid ChromatographyTannin degraded products were estimated using RP-HPLC by the

modifiedmethod ofMakkar (2000). Briefly, the culture supernatantcontaining degraded tannic acid metabolites was filtered throughWhatman filter paper. Equal volume of ethanoleethyl acetate (95%EAþ 5% ethanol) was added to the filtered broth. The contents weremixed, vortexed and stirred for 1 h. The contents were thentransferred to separating funnel and partitioning was allowed for1 h. The upper layer was collected and aliquots of 2 ml were madein duplicates. The solvents were removed under vacuum and theextracted samples were reconstituted by adding 2 ml HPLC grademethanol. The samples were filtered through Millipore membranefilter (0.22 mm) and the filtrate was used for gradient HPLC analysisusing C18 spherisorb column (ODS 2, 4.6 � 250 mm, 5 mm particlesize, 300 Å pore size) Waters32 HPLC system. The solvent systemcomposed of degassed HPLC grade water (acidified with 0.025%phosphoric acid) and acetonitrile. Tannin degraded compoundswere eluted by a gradient of water/acetonitrile from 100 to 0% in30min. The peaks were detected at twowavelengths of 250 nm and280 nm.

2.2. Enzyme purification

E. faecalis was grown in minimal media containing 1% filtersterilised tannic acid (Sigma) for 24 h at 37 �C. The culture brothwas centrifuged at 4000 � g for 20 min and the supernatant wasused for purification of enzyme. The enzyme activity at each stepwas calculated as per Sharma et al. (1999). The enzyme activity andprotein content (Lowry et al., 1951) at each step of purification wasdetermined to calculate the specific activity.

2.2.1. Clarification and ammonium sulphate precipitationThe supernatant was treated with 1% activated charcoal to

remove the colour and phenolic compounds (Mahendran et al.,2005) for 2 h with intermittent shaking. The filtrate was clarifiedby centrifugation at 4000 � g for 20 min at 4 �C. To 950 ml ofclarified culture supernatant, ammonium sulphate was added toachieve 30e60% saturation and centrifuged at 16,000 � g for20 min at 4 �C. The supernatant was subsequently adjusted to 80and 100% saturation. The pellet in each case was dissolved in citratebuffer (0.05 M, pH 5.0). The enzyme preparation obtained aftereach ammonium sulphate fractionation step was dialysed usinga 1000 MWCO (molecular weight cut-off) cellulose acetatemembrane for about 18 h against citrate buffer (0.05 M, pH 5.0)with 3e4 intermittent changes of citrate buffer (0.05 M, pH 5.0).

2.2.2. DEAE-cellulose chromatographyA column was packed with DEAE-Cellulose to a bed size of

2.5 cm � 10 cm and was equilibrated with 0.05 M citrate buffer(pH 5.0). The elution was done in discontinuous gradient systemwith citrate buffer of increasingmolarity (0.05, 0.1, 0.2, 0.3 M citratebuffer, pH 5.0). Five, 5 ml fractions for buffer of each molarity werecollected and the fractions were monitored for the elution profile ofprotein and tannase activity as described above.

2.2.3. Gel filtration chromatographySephadex G-150 was soaked in 0.05 M citrate buffer (pH 5.0)

overnight at room temperature and the fines were decanted off.The gel suspension, after deaerating for 10e15 min, was packedinto a 55 cm � 2.5 cm glass column at the operating pressure headof 30 cm. The protein fractions were eluted from the column ata flow rate of 1ml/min. Five millilitres fractions were collected afterdraining 80% of the void volume and analysed for tannase activity.The fractions showing tannase activity were pooled and concen-trated using PEG-6000.

2.2.4. Characterization of tannaseThe molecular weight of the tannase was determined by SDS-

PAGE as per the protocol of Laemmli (1971). The molecularweight of the protein was determined by comparing with theprotein marker standard (Low range marker from Biorad). Theactivity of partially purified tannase was studied at differenttemperatures (20, 30, 40, 50, 60 and 70 �C), pH (pH 2.0e9.0) andsubstrate concentration (methyl gallate at 0.0625, 0.125, 0.25, 0.5,0.75, 1.0, 2.0, 3.0, 4.0 and 5.0 mM). The enzyme activity wasdetermined by the rhodanine method (Sharma et al., 1999).

3. Results and discussion

3.1. Degradation of tannic acid

Tannic acid degradation profile was observed using TLC andHPLC. The thin layer chromatogram showed two identificationspots, namely gallic acid and pyrogallol when compared withstandards. On TLC plate, tannic acid remained at the bottom linefollowed by gallic acid, pyrogallol and resorcinol with an Rf value of0.26, 0.59 and 0.61, respectively. Gallic acid and pyrogallol wereobserved as major degraded product of tannic acid after 72 h ofincubation (Fig. 1). However, HPLC results indicated that E. faecaliswas able to degrade tannic acid to pyrogallol and further to resor-cinol within 72 h of incubation. The retention time of 4.31 min,4.57 min and 6.05 min was observed for gallic acid, pyrogallol and

Table 1Degradation profile of tannic acid by E. faecalis at different time intervals.

IncubationTime (h)

Degraded products (mg/ml) Tannic acid (TA) % TA degradation

Gallic acid (GA) Pyrogallol (PYR) Resorcinol (Res)

12 0.16 � 0.013 0.00 0.000 0.83 � 0.101 17.4324 0.28 � 0.022 0.11 � 0.011 0.07 � 0.005 0.54 � 0.095 45.8936 0.20 � 0.012 0.16 � 0.023 0.15 � 0.010 0.49 � 0.102 51.4248 0.11 � 0.020 0.21 � 0.034 0.21 � 0.015 0.46 � 0.098 54.2860 0.08 � 0.123 0.24 � 0.077 0.40 � 0.100 0.28 � 0.111 72.1872 0.00 0.35 � 0.014 0.56 � 0.098 0.08 � 0.002 91.88

Values presented are mean � SD.

G. Goel et al. / International Biodeterioration & Biodegradation 65 (2011) 1061e1065 1063

resorcinol, respectively. The peak area of the tannic acid at reten-tion time (11.0 min) declined sharply with time of incubation withsimultaneous increase in the peak area of pyrogallol.

The degradation profile of tannic acid by E. faecalis (Table 1)shows that initially at 12 h of incubation period, only gallic acid wasformed indicating a 17% degradation of tannic acid. Further increasein incubation period upto 24 h resulted in maximum gallic acidproduction followed by pyrogallol and traces of resorcinol. Pyro-gallol was the major product after 48 h and 72 h of incubation.However, at 48 h of incubation period, all the metabolites i.e. gallicacid, pyrogallol and resorcinol were observed in significant amountresulting in 55% degradation of tannic acid and finally after 72 h ofincubation, 92% of tannic acid was mineralized into pyrogallol andresorcinol as the major end products.

Several other ruminal microorganisms were reported to possessthe ability to degrade phenolic monomers. The results are inagreement with reports from Tsai and Jones (1975) who showedthat the isolated Streptococcus strains and three Coprococcus strainsfrom the bovine rumenwere able to degrade 80% of phloroglucinol.Odenyo and Osuji (1998) have reported three strains of a tannin-tolerating bacterium (Selenomonas sp.) from the rumen micro-flora of sheep, goat and antelope that had either been fed or hadbrowsed on tanniniferous forages. One of the strains (EAT2) of thisruminal bacterium could hydrolyse tannic acid to gallic acid andsubsequently to pyrogallol, whereas the remaining two strains (ES3and EG19) were able to hydrolyse tannic acid to gallic acid only.Zeida et al. (1998) also reported a combined activity of tannase andgallate decarboxylase activity in Pantoea agglomerans T71 whichcan be used for the bioconversion of tannic acid to pyrogallol.Eubacterium oxidoreducens, a strictly anaerobe was reported todegrade gallate, phloroglucinol and pyrogallol to acetate andbutyrate in presence of hydrogen and formic acid (Krumholz andBryant, 1986). Another anaerobe, Syntrophococcus sucromutanswas reported to have ability to demethoxylate various phenolicmonomers (Krumholz and Bryant, 1988). The degradation of tannicacid by our isolate indicates that the tannineprotein complexes,which were thought to be hydrolysed only under the acidic

Table 2Summary of partial purification profile of tannase.

Purification step Volume(ml)

Tannase activity(U/ml)

Total Activity(U)

Pr(m

Culture filtrate 1000 32.58 32,580 0.Clarification 950 32.51 30,884 0.Ammonium sulphate

precipitation30 975.64 29,268 0.

DEAE-cellulose 10 898 8980 0.Sephadex G-150 15 906.81 13,602 0.

Total activity ¼ Activity (U) � Total volume (ml).Total protein ¼ Protein (mg) � Total volume (ml).Specific activity ¼ Total activity (U)/Total protein (mg).Purification fold ¼ Specific activity of the sample/Initial specific activity.Yield ¼ [Total activity of the sample/Initial total activity] � 100.

condition of the abomasum, may also be subject to microbialdegradation in rumen.

3.2. Enzyme purification

The results pertaining stepwise purification of tannase arepresented in Table 2.

3.2.1. Clarification and ammonium sulphate precipitationCulture filtrate treated with 1% activated carbon removed more

than 80% of the colour according to the visual observation. Thecolour due to oxidation of gallic acid was removed by activatedcharcoal. The clarified filtrate was having an activity of 32.51 U/mlwith a yield of 94.7%. Fractional precipitation with 50% ammoniumsulphate removed some of the nonenzymatic proteins. Tannasewasprecipitated at 70% saturation. About 89.8% of the total tannase wasrecovered by 70% saturationwith activity of 975.64 U/ml. Rajkumarand Nandy (1983) obtained a 69% of total recovery from ammo-nium sulphate at 100% saturation. Mahendran et al. (2005)obtained 78.7% of the total tannase at 70% saturation with ammo-nium sulphate. Chhokar et al. (2010a) reported maximum recoveryof tannase at 80% ammonium sulphate fractionation.

3.2.2. DEAE-cellulose chromatographyThe 70% saturated fraction was further purified through DEAE-

cellulose ion exchange chromatography. The elution profile yiel-ded a single protein peak corresponding with the tannase activity.The tannase at this stage has total activity of 8980 U with a specificactivity of 2896 U/mg. From DEAE-cellulose chromatography, theoverall purification of 25.7 fold with a yield of 27.5% was obtained.Hamdy (2008) purified tannase from Fusarium subglutinans with26.98 fold purification and 42.6% recovery by ammonium sulphate,ion exchange chromatography (DEAE-cellulose) and gel filtration(Sephedax G-150). An extracellular tannase from A. heteromorphusMTCC 8818 was also purified through ammonium sulphateprecipitation and ion exchange chromatography leading to an

oteing/ml)

Total Protein(mg)

Specific activity(U/mg)

Purificationfold

Yield (%)

29 290 112.3 1 10029 275.5 112.1 1 94.789 26.7 1096 9.7 89.8

31 3.1 2896 25.7 27.543 6.45 2109 18.7 41.7

Fig. 3. Effect of temperature on tannase activity.

G. Goel et al. / International Biodeterioration & Biodegradation 65 (2011) 1061e10651064

overall purification of 39.74 fold with a yield of 19.29% (Chhokaret al., 2010b).

3.2.3. Sephadex G-150 chromatographyThe elution profile of the enzyme from gel filtration resulted in

four protein peaks of which two were prominent. One of theprominent peak between fractions from 14 to 28 coincides with thepeak of tannase activity, thus those fractions were pooled andconcentrated. The tannase activity was not observed in other peaks.The tannase at this stage has total activity of 13,602 U witha specific activity of 2109 U/mg. At gel filtration the enzyme waspurified up to 18.7 folds with a yield of 41.7%. Rajkumar and Nandy(1983) obtained 18.5% recovery from the Sephadex G-200 columneluted with 0.02 M acetate buffer. Sharma et al. (1999) alsoobtained four peaks of the protein content with a single peak oftannase activity where the purification was obtained upto 29 foldswith 2.7% recovery of the total tannase. Mahendran et al. (2005)obtained three protein peaks when tannase was eluted fromSephadex G-200, yielding about 17.6% of total tannase.

3.3. Molecular weight (Mr) determination

The purified protein resulted in a single band of 45 kDa (Fig. 2).The tannases with a molecular weight of 59 kDa from Selenomonasruminantium (Skene and Brooker, 1995) and 45 kDa from Paecilo-myces variotii (Mahendran et al., 2005) have already been reported.However, Yamada et al. (1968) and Hatamoto et al. (1996) havereported that the multimeric tannases from Aspergillus flavus andAspergillus oryzae, respectively, with molecular weight of 186 toover 300 kDa. Bhardwaj et al. (2003) reported that the total tannaseactivity was made up of nearly equal activity of esterase and dep-sidase with a molecular weight 102 and 83 kDa.

3.4. Properties of tannase

3.4.1. Effect of temperatureThe temperature for the optimum activity of tannase from

E. faecalis was 40 �C (Fig. 3). With the further increase in temper-ature, the tannase activity was found to decline sharply with noactivity at 80 �C as increase in temperature increases the rate ofdenaturation of the enzyme, with the loss of secondary and tertiarystructures (Sabu et al., 2005). Tannase from other sources likeA. oryzae, Penicillium chrysogenum, Aspergillus niger have beenreported to have their optimal activity at 30e40 �C (Iibuchi et al.,1968; Rajkumar and Nandy, 1983; Lekha and Lonsane, 1997; Sabuet al., 2005). A lower temperature optima (30 �C) has been repor-ted for tannase from A. niger Van Tieghem MTCC 2425 (Sharmaet al., 1999) and Aspergillus awamori (Chhokar et al., 2010a).

Fig. 2. SDS profile of tannase from E. faecalis (M: Marker, low range marker Biorad).

3.4.2. Effect of pHThe tannase from E. faecalis possess an activity between 5.0 and

7.0 with an optimum activity at pH of 6.0 (Fig. 4). The same pHrange was reported for other tannase obtained from A. niger(Ramırez-Coronel et al., 2003; Srivastava and Kar, 2009), Crypho-nectria (Farias et al., 1994) and P. variotii (Mahendran et al., 2005;Battestin and Macedo, 2007). However, tannase from A. niger VanTieghemMTCC 2425 as reported by Sharma et al. (1999) resulted intwo different peaks of tannase activity at a pH of 6.0 anda secondary peak at 4.5 which were reported to be due to differentdepsidase and esterase activities (Barthomeuf et al., 1994; Lekhaand Lonsane, 1997).

3.4.3. Effect of substrate concentrationThe analysis of the graph of substrate concentration against

tannase activity yielded Km values of 0.43 mmol. This implies thattannase of E. faecalis has higher affinity for methyl gallate. The Kmvalues for tannases from A. flavus, S. ruminantium, Cryphonectria,A. niger Van Tieghem using methyl gallate as substrate have beenfound to be 0.86 mM (Yamada et al., 1968), 1.6 mM (Skene andBrooker, 1995), 7.49 mM (Farias et al., 1994) and 0.2 mM (Sharmaet al., 1999), respectively.

From the present investigation, it can be concluded that thepotential tannin degrading microorganisms reside in the gut ofsmall ruminants. Being grazers, goats are found to be moreadapted to tannins in feeds as they harbour tannin degradingmicroflora in situ. Hence, a considerable potential exists for theexploitation of such potential isolates using in vivo studies in orderto use them for trans-inoculation into other animal species toimprove the nutritional status and productivity of the livestock.The results presented in this work shows that E. faecalis tannasehas interesting and attractive properties for industrial use forproduction of pyrogallol.

Fig. 4. Effect of pH on tannase activity.

G. Goel et al. / International Biodeterioration & Biodegradation 65 (2011) 1061e1065 1065

Acknowledgements

The authors wish to thank Indian Council of AgricultureResearch, New Delhi, and National Dairy Research Institute, Karnal,India, for providing financial support and essential facilitiesrequired for the research work.

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