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1 2 3 1 4 5 2 Running title: L. PLANTARUM TANNASE 6 7 F 8 3 9 o 10 4 11 r 12 13 5 ProductionR and physicochemical properties of recombinant 14 15 16 6 e Lactobacillus plantarum tannase 17 v 18 19 7 i 20 e 21 8 22 w 23 † † § 24 9 JOSÉ ANTONIO CURIEL , H.ÉCTOR RODRÍGUEZ , IVÁN ACEBRÓN , JOSÉ MIGUEL 25 § † , † 26 10 MANCHEÑO , BLANCAC DE LAS RIVAS , AND ROSARIO MUÑOZ * 27 28 11 29 o 30 31 12 n 32 f 33 13 Departamento de Microbiología, Instituto de Ferimentaciones Industriales CSIC, Juan de la 34 d 35 14 Cierva 3, and Grupo de Cristalografía Molecular y Biología Estructural, Instituto de 36 e 37 38 15 Química-Física “Rocasolano” CSIC, Serrano 119n, Madrid 28006, Madrid. 39 40 16 t 41 i 42 a 43 17 44 l 45 18 * Corresponding author: 46 - 47 19 Address: Instituto de Fermentaciones Industriales, CSIC, Juan de la Cierva 3, 28006 48 A 49 50 20 Madrid, Spain. Phone: +34 91 5622900; Fax: +34 91 564 4853. C 51 52 21 E-mail: [email protected] S 53 54 22 † Departamento de Microbiología 55 56 § 57 23 Grupo de Cristalografía Molecular y Biología Estructural 58 59 60 1 ACS Paragon Plus Environment Submitted to Journal of Agricultural and Food Chemistry Page 2 of 27

1 2 3 1 Abstract 4 5 2 Tannase is an enzyme with important biotechnological applications in food industry. 6 7 F 8 3 Previous studies have identified the tannase encoding gene in Lactobacillus plantarum and 9 o 10 4 also have reported the description of the purification of recombinant L. plantarum tannase 11 r 12 13 5 through a Rprotocol involving several chromatographic steps. Here, we describe the high- 14 15 6 yield production of pure recombinant tannase (17 mg/L) by a one-step affinity procedure. 16 e 17 7 The purified recomvbinant tannase exhibits optimal activity at pH 7 and 40 ºC. Addition of 18 19 2+ i 20 8 Ca to the reaction mixeture greatly increased tannase activity. The enzymatic activity of 21 22 9 tannase was assayed againwst 18 simple esters. Only esters derived from gallic 23 24 10 acid and we.re hydrolyzed. In addition, tannase activity was also 25 26 C 27 11 assayed against the tannins tannic acid, gallocatechin gallate and epigallocatechin gallate. 28 o 29 12 Despite L. plantarum tannase representing a novel family of tannases which shows no 30 n 31 13 significant similarity to tannases from fungal sources, both families of enzymes shared 32 f 33 i 34 14 similar substrate specificity range. The physicochemicald characteristics exhibited by L. 35 36 15 plantarum recombinant tannase make it an adequate aleternative to the currently used fungal 37 38 n 39 16 tannases. 40 t 41 17 i 42 a 43 18 44 l 45 46 19 - 47 48 20 KEYWORDS: Tannin acyl hydrolase; tannase; hydrolyzable tannins; propylA gallate; 49 50 21 . C 51 52 S 53 22 54 55 56 57 58 59 60 2 ACS Paragon Plus Environment Page 3 of 27 Submitted to Journal of Agricultural and Food Chemistry

1 2 3 1 INTRODUCTION 4 5 2 6 7 F 8 3 The enzyme tannase (or tannin acyl hydrolase EC 3.1.1.20) catalyzes the hydrolysis 9 o 10 4 of ester bonds in hydrolyzable tannins such as tannic acid, thereby releasing glucose and 11 r 12 13 5 gallic acidR ( 1-3 ). Currently, tannase plays an important role in the industries of drinks and 14 15 6 foods, chemical-pharmaceutics, brewing and production of aminal feed. Thus, tannase is 16 e 17 7 used in the elaboravtion of acorn liquor and also in the production of gallic acid, which is 18 19 i 20 8 subsequently employede for the synthesis of propylgallate, a potent antioxidant. Also, 21 22 9 tannase is used as clarifyinwg agent in the manufacturing of instant tea, some wines, beers, 23 24 10 juices or fruits and in coffee-flavored. soft drinks ( 1-4 ). 25 26 C 27 11 It has long been known that several fungal species, such as Aspergillus spp., are 28 o 29 12 capable of producing large amounts of tannase ( 5). However, the use of tannase on a large 30 n 31 13 scale was limited by a variety of factors, including production costs and insufficient 32 f 33 i 34 14 knowledge of the enzyme. The advent of recombindant DNA technology has revolutionized 35 36 15 research in the field of enzymology. Hatamoto et al. (1996)e cloned and sequenced the gene- 37 38 n 39 16 encoding tannase from Aspergillus oryzae (6). Later, the Aspergillus tannase gene was 40 t 41 17 heterologously expressed in Saccharomyces cerevisiae , althoughi with a low yield of protein 42 a 43 18 production ( 3). Conversely, large quantities of enzyme were obtained when produced in 44 l 45 46 19 Pichia pastoris ( 7). Obviously, the high-yield production of fully active r-ecombinant 47 48 20 tannases is an attractive goal both for basic research and for industrial purposAes. 49 50 21 In addition to fungal tannases, many bacterial species have been reported Cto produce 51 52 S 53 22 tannase ( 3), and in fact several putative bacterial tannase sequences have been included in 54 55 23 the databases. To our knowledge, the only bacterial tannases that have been analyzed both 56 57 58 24 genetically and biochemically are those from Staphylococcus lugdunensis ( 8) and 59 60 3 ACS Paragon Plus Environment Submitted to Journal of Agricultural and Food Chemistry Page 4 of 27

1 2 3 1 Lactobacillus plantarum ( 9). As S. lugdunensis strains have been associated with human 4 5 2 disease ( 10-13 ) and microorganisms used in the industrial production of food processing 6 7 F 8 3 enzymes should be listed as GRAS (generally recognized as safe), so far, the L. plantarum 9 o 10 4 protein is the only bacterial tannase alternative for its use in the food industry. 11 r 12 13 5 SinRce i) a low-yield protocol has been described for its production, and ii) the 14 15 6 substrate specificity of the L. plantarum tannase remains unknown, the aims of this study 16 e 17 7 were to design a pvrotocol for a high-yield tannase production which could facilitate its 18 19 i 20 8 biochemical characterizeation, as well as to make it highly attractive for their use in the 21 22 9 industrial sector. w 23 24 10 . 25 26 C 27 11 28 o 29 12 MATERIALS AND METHODS 30 31 n 32 13 33 f i T 34 14 Bacterial strains and materials. L. plantarumd CECT 748 strain was purchased 35 36 15 from the Spanish Culture Type Collection (CECT). E. ecoli DH5 α was used for all DNA 37 38 n 39 16 manipulations. E. coli JM109 (DE3) was used for expression in pURI3 vector ( 14 ). L. 40 t 41 17 plantarum strain was grown in MRS medium at 30 ºC without shaking.i E. coli strains were 42 a 43 18 cultured in Luria-Bertani (LB) medium at 37 ºC and 200 rpm. When required, ampicillin 44 l 45 46 19 was added to the medium at a concentration of 100 g/mL. - 47 48 20 The compounds assayed in this study were (Fluka 486A90), ethyl 49 50 21 gallate (Aldrich 48640), propyl gallate (Sigma P3130), lauryl gallate (Aldrich 48C660), 51 52 S 53 22 methyl benzoate (Fluka 18344), ethyl benzoate (Sigma E12907), methyl 4- 54 55 23 hydroxybenzoate (Sigma H5501), ethyl 4-hydroxybenzoate (Aldrich 111988), propyl 4- 56 57 58 24 hydroxybenzoate (Scharlau Hi200), butyl 4-hydroxybenzoate (Scharlau Hi180), methyl 59 60 4 ACS Paragon Plus Environment Page 5 of 27 Submitted to Journal of Agricultural and Food Chemistry

1 2 3 1 vanillate (Aldrich 138126), methyl 2,4-dihydroxybenzoate (Aldrich M42505), methyl 4 5 2 gentisate (Aldrich 426091), methyl salicylate (Sigma M6752), ethyl 3,4-dihydroxybenzoate 6 7 F 8 3 (protocatechuic acid ethyl ester) (Aldrich E24859), ethyl 3,5-dihydroxybenzoate (Aldrich 9 o 10 4 541087), ferulic methyl ester (Extrasynthèse 6267), ferulic ethyl ester (Extrasynthèse 11 r 12 13 5 6275), tanRnic acid (Sigma T0125), ellagic acid (Sigma, E2250), chlorogenic acid (Aldrich 14 15 6 C3878), quercetin (Sigma Q4951), catechin (Sigma C1251), epicatechin (Sigma E1753), 16 e 17 7 gallocatechin (Fluvka G6657), gallocatechin gallate (Sigma G6782), epigallocatechin (Fluka 18 19 i 20 8 08108), epigallocatechine gallate (Fluka 50299) and 4-nitrophenyl β-D-glucopyranoside 21 22 9 (Sigma N7006). w 23 24 10 . 25 26 C 27 11 DNA manipulations . Bacterial DNA was isolated from overnight cultures using a 28 o 29 12 protocol previously described ( 15 ). DNA sequencing was carried out by using an Abi Prism 30 n 31 13 377 DNA™ sequencer (Applied Biosystems, Inc.). Sequence similarity searches were 32 f 33 i 34 14 carried out using FASTA on EBI site ( http://www.ebi.ac.ukd ). Signatures, pI/MW, etc. were 35 36 15 analysed on EXPASY site (http//www.expasy.ch). Mueltiple alignments were done using 37 38 n 39 16 CLUSTAL W on EBI site after retrieval of sequences from GenBank and Swiss-Prot. 40 t 41 17 i 42 a 43 18 Expression, and purification of recombinant tannase . The gene coding for the L. 44 l 45 T 46 19 plantarum CECT 748 tannase, tanLp1 (named lp_2956 in the L. plantarum- WCFS1 strain) 47 48 20 was cloned and overexpressed following a strategy previously described ( 14 A). Briefly, the 49 50 21 gene was PCR-amplified with Hot-start Turbo Pfu DNA polymerase by using theC primers 51 52 S 53 22 513 (5´- CATCATGGTGACGATGACGATAAG atgagtaaccgattgatttttgatg) and 514 (5´- 54 55 23 AAGCTTAGTTAGCTATTATGCGTA tcattggcacaagccatcaatccag) (the nucleotides pairing 56 57 58 24 the expression vector sequence are indicated in italics, and the nucleotides pairing the 59 60 5 ACS Paragon Plus Environment Submitted to Journal of Agricultural and Food Chemistry Page 6 of 27

1 2 3 1 tanLp1 gene sequence is written in lowercase letters). The corresponding 1.4-kb purified 4 5 2 PCR product was inserted into pURI3 vector by using the restriction enzyme- and ligation- 6 7 F 8 3 free cloning strategy described previously ( 14 ). Briefly, to clone any target protein into the 9 o 10 4 pURI3 vector, PCR products of the gene of interest need to be generated with specific 11 r 12 13 5 overhangsR that are complementary to the integration site sequence of the vector. After the 14 15 6 first PCR reaction, the fragment is added to the methylated recipient template plasmid. The 16 e 17 7 elongated and modifiedv strands are not methylated. Dpn I digestion is used to eliminate the 18 19 i 20 8 methylated plasmid DNeA. Expression vector pURI3 was constructed based on the 21 22 9 commercial expression vecwtor pT7-7 (USB) but expressing a protein containing the 23 24 10 following leader sequence MGG.SHHHHHHGDDDDKM consisting of a N-terminal 25 26 C 27 11 methionine followed by three spacer amino acids, a six-histidine affinity tag, a spacer 28 o 29 12 glycine residue, and the five-amino acid enterokinase recognition site ( 14 ). E. coli DH5 α 30 n 31 13 cells were transformed, recombinant plasmids were isolated and those containing the 32 f 33 i 34 14 correct insert were identified by restriction-enzymed analysis, verified by DNA sequencing 35 36 15 and then transformed into E. coli JM109(DE3) cells. e 37 38 n 39 16 Cells carrying the recombinant plasmid, pURI3-TanLp1, were grown at 37 ºC in 40 t 41 17 Luria-Bertani media containing ampicillin (100 g/mL), and indiuced by adding 0.4 mM 42 a 43 18 IPTG. After induction, the cells were grown at 22 ºC during 20 h and collected by 44 l 45 46 19 centrifugation. Cells were resuspended in 20 mM Tris-HCl, pH 8.0, 100 -mM NaCl. Crude 47 48 20 extracts were prepared by French Press lysis of cell suspensions (three cycleAs at 1100 49 50 21 p.s.i.). The insoluble fraction of the lysate was removed by centrifugation at 47 000C g for 30 51 52 S 53 22 min at 4 ºC. 54 55 23 The supernatant was filtered through a 0.45 m filter and applied to a His-Trap TM - 56 57 58 24 FF crude chelating affinity column (Amersham Biosciences) equilibrated with 20 mM Tris- 59 60 6 ACS Paragon Plus Environment Page 7 of 27 Submitted to Journal of Agricultural and Food Chemistry

1 2 3 1 HCl, pH 8.0, 100 mM NaCl containing 10 mM imidazole, to improve the interaction 4 5 2 specificity in the affinity chromatograpy step. The bound enzyme was eluted by applying a 6 7 F 8 3 stepwise gradient of imidazole concentration, from 20 mM Tris-HCl, pH 8.0, 100 mM 9 o 10 4 NaCl containing 10 mM imidazole to 20 mM Tris-HCl, pH 8.0, 100 mM NaCl containing 11 r 12 13 5 500 mM iRmidazole. Fractions containing the eluted proteins were pooled and the protein 14 15 6 was then dialyzed overnight at 4 ºC in a membrane (3500 cut-off) against 25 mM sodium 16 e 17 7 phosphate buffer, vpH 6.5. The purity of the enzymes was determined by 12% sodium 18 19 i 20 8 dodecyl sulphate polyaecrylamide-gel electrophoresis (SDS-PAGE) in Tris-glycine buffer. 21 22 9 w 23 24 10 SDS-PAGE and determination. of protein concentration . Samples were analyzed 25 26 C 27 11 by 12 % SDS-PAGE under reducing conditions. Protein bands were visualized by 28 o 29 12 Coomassie blue staining. The gels were calibrated using molecular weight markers. Protein 30 n 31 13 concentration was measured according to the method of Bradford using a protein assay kit 32 f 33 i 34 14 (Bio-Rad) with bovine serum albumin as standard. d 35 36 15 e 37 38 n 39 16 Enzyme activity assay . Tannase activity was determined using a rhodanine assay 40 t 41 17 specific for gallic acid ( 16 ). Rhodanine reacts only with gallic aciid and not with galloyl 42 a 43 18 esters or other phenolics. Gallic acid analysis in the reactions was determined using the 44 l 45 46 19 following assay. Tannase enzyme (10 µg) in 700 µL of 50 mM phosphat-e buffer pH 6.5 47 48 20 was incubated with 40 µL of 25 mM methyl gallate (1 mM final concentratioAn), during 5 49 50 21 min at 37 ºC. After this incubation, 150 µL of a methanolic rhodanine solution (0C.667% w/v 51 52 S 53 22 rhodanine in 100% methanol) was added to the mixture. After 5 min incubation at 30 ºC, 54 55 23 100 µL of 500 mM KOH was added. After an additional incubation of 5-10 min, the 56 57 58 24 absorbance at 520 nm was measured on a spectrophotometer. A standard curve using gallic 59 60 7 ACS Paragon Plus Environment Submitted to Journal of Agricultural and Food Chemistry Page 8 of 27

1 2 3 1 acid concentrations ranging from 0.125 to 1 mM was prepared. One unit of tannase activity 4 5 2 was defined as the amount of enzyme required to release 1 µmol of gallic acid per minute 6 7 F 8 3 under standard reaction conditions. 9 o 10 4 To study the effect of substrate concentration, the standard activity assay was done 11 r 12 13 5 by using sRolutions of methyl gallate at different concentrations (0.1 to 100 mM). To study 14 15 6 the effect of pH and temperature, standard activity assays were done using the conditions 16 e 17 7 described below. v 18 19 i 20 8 e 21 22 9 Optimum temperaturew and pH-dependence . Activities of recombinant L. 23 24 10 plantarum tannase were measur.ed at 20, 30, 40, 50, 60, and 70 ºC to determine the optimal 25 26 C 27 11 temperature for enzymatic activity. The optimum pH value for tannase activity was 28 o 29 12 determined by studying its pH-dependence within the pH range between 3 and 10. Acetic 30 n 31 13 acid-sodium acetate buffer was used for pH 3-5, citric acid-sodium citrate buffer for pH 6, 32 f 33 i 34 14 sodium phosphate buffer for pH 7, Tris-HCl bufferd for pH 8, glycine-NaOH buffer for pH 35 36 15 9, and sodium carbonate-bicarbonate for pH 10. A 100e mM concentration was used in all 37 38 n 39 16 the buffers. The rhodanine assay was used for the optimal pH characterization of tannase. 40 t 41 17 Since the rhodanine-gallic acid complex forms only in basic coniditions, after the 42 a 43 18 completion of the enzymatic degradation of methyl gallate, KOH was added to the reaction 44 l 45 46 19 mixture to ensure that the same pH value (pH 11) was achieved in all sam-ples assayed. 47 48 20 For temperature stability measurements, the recombinant tannase wasA incubated in 49 50 21 50 mM phosphate buffer pH 6.5 at 25, 30, 37, and 45 ºC for 15 min, 30 min, and C1, 2, 3, 4, 51 52 S 53 22 and 20 h. After incubation, the residual activity was measured as above described. 54 55 23 56 57 58 59 60 8 ACS Paragon Plus Environment Page 9 of 27 Submitted to Journal of Agricultural and Food Chemistry

1 2 3 1 Effect of additives on tannase activity. To test the effect of metals and ions on the 4 5 2 activity of the recombinant tannase, the enzymatic activity was measured in the presence of 6 7 F 8 3 different additives at a final concentration of 1mM. The additives analyzed were MgCl 2, 9 o 10 4 KCl, CaCl 2, HgCl 2, ZnCl 2, Triton-X-100, Urea, Tween 80, EDTA, DMSO, and β- 11 r 12 13 5 mercaptoeRthanol. The potential substrate was also assayed. 14 15 6 16 e 17 7 HPLC analysisv of L. plantarum tannase activity on several substrates. The 18 19 i 20 8 activity of recombinante tannase against 28 potential substrates has been analyzed. The 21 22 9 standard enzyme activity awssay was modified by using 100 µg of tannase, 1 mM substrate, 23 24 10 and 1mM CaCl in the reaction .mixture, and incubated at 37 ºC during 10 min. As controls, 25 2 26 C 27 11 phosphate buffer containing the reagents but the enzyme were incubated in the same 28 o 29 12 conditions. The reaction products were extracted twice with ethyl acetate (Lab-Scan, 30 n 31 13 Ireland) and analyzed by HPLC-DAD. 32 f 33 i 34 14 A Thermo (Thermo Electron Corporation, Wdaltham, Massachussetts, USA) 35 36 15 chromatograph equipped with a P400 SpectraSystem peump, and AS3000 autosampler, and 37 38 n 39 16 a UV6000LP photodiode array detector were used. A gradient of solvent A (water/acetic 40 t 41 17 acid, 98:2, v/v) and solvent B (water/acetonitrile/acetic acid, 78:i20:2, v/v/v) was applied to 42 a 43 18 a reversed-phase Nova-pack C (25 cm x 4.0 mm i.d.) 4.6µm particle size, cartridge at 44 18 l 45 46 19 room temperature as follows: 0-55 min, 80% B linear, 1.1 mL/min; 55-5-7 min, 90% B 47 48 20 linear, 1.2 mL/min; 57-70 min, 90% B isocratic, 1.2 mL/min; 70-80 min, 95%A B linear, 1.2 49 50 21 mL/min; 80-90 min, 100% linear, 1.2 mL/min; 100-120 min, washing 1.0 mL/miCn, and 51 52 S 53 22 reequilibration of the column under initial gradient conditions. Detection was performed by 54 55 23 scanning from 220 to 380 nm. Samples were injected in duplicate onto the cartridge after 56 57 58 24 being filtered through a 0.45 m PVDF filter. The identification of degradation compounds 59 60 9 ACS Paragon Plus Environment Submitted to Journal of Agricultural and Food Chemistry Page 10 of 27

1 2 3 1 was carried out by comparing the retention times and spectral data of each peak with those 4 5 2 of standards from commercial suppliers. 6 7 F 8 3 9 o 10 4 11 r 12 13 5 RESULTSR AND DISCUSSION 14 15 6 16 e 17 7 Productionv and enzymatic activity of recombinant L. plantarum tannase. 18 19 i 20 8 Tannases are enzymes extensively used in the food industry. In previous studies we showed 21 22 9 the existence of tannase activityw in cell-free extracts from L. plantarum CECT 748 T ( 17 ). 23 24 10 Later, it was reported the identification. of a gene encoding a putative tannase ( 9), which 25 26 C 27 11 was similar to the sequence of the S. lugdunensis tannase. The gene encoding the putative 28 o 29 12 tannase from L. plantarum CECT748 T, tanLp1 , has been cloned and hyperexpressed in E. 30 n 31 13 coli ; the recombinant tannase was further purified and biochemically characterized. The 32 f 33 i 34 14 purification protocol comprised several chromatogdraphic steps (Q-Sepharose Fast Flow, 35 36 15 hydroxylapatite, and Mono-Q GL5/5 columns). With tehe aim of improving the purification 37 38 T n 39 16 of tannase from L. plantarum CECT48 , we decided to use the expression vector pURI3 40 t 41 17 constructed in our laboratory ( 14 ). The pURI3 vector was createid using the pT7-7 vector as 42 a 43 18 template and contains an amino-terminal His-tag that allows convenient purification of the 44 l 45 46 19 native protein directly from the crude cell extracts. In fact, we have previ-ously employed 47 48 20 this methodology to successfully overproduce other proteins from L. plantarumA ( 18, 19 ). 49 50 21 The tanLp1 gene from L. plantarum CECT 748 T (ATCC 14917 T) was PCCR- 51 52 S 53 22 amplified by using 513 and 514 primers and cloned into pURI3 vector. When the sequence 54 55 23 of the recombinant plasmid pURI3-TanLp1 was analyzed, it could be observed that, in 56 57 T 58 24 relation to the previously deposited L. plantarum ATCC 14917 sequence, it contains two 59 60 10 ACS Paragon Plus Environment Page 11 of 27 Submitted to Journal of Agricultural and Food Chemistry

1 2 3 1 nucleotide changes at positions 801 (C to T) and 962 (T to C). This last nucleotide change 4 5 2 produces a conservative amino acid substitution at position 321 (Val-321 to Ala). The 6 7 F 8 3 nucleotide changes were confirmed by the sequencing of the gene from chromosomal 9 o 10 4 DNA. This was a surprising result, as both sequences proceed from the same strain. It is 11 r 12 13 5 possible thRat the different evolution of both strain stocks could be responsible of these 14 15 6 changes. In this sense, Iwamoto et al. (2008) described that the tanLp1 sequence was 99.6% 16 e 17 7 identical to that derivedv from the L. plantarum WCFS1 strain, lp_2956. These differences 18 19 i 20 8 determine protein sequencese differing in two amino acid residues ( 9). Therefore in relation 21 22 9 to tannase from L. plantarumw WCFS1, the sequence reported in this study, showed three 23 24 10 amino acid substitutions, two no.n conservative changes Thr-154 to Ala, and Arg-406 to 25 26 C 27 11 Gln, and the conservative Val-321 to Ala change. 28 o 29 12 To confirm that L. plantarum CECT 748 T tanLp1 gene encodes a functional 30 n 31 13 tannase, we expressed this gene in E. coli under the control of the T7 RNA polymerase- 32 f 33 i 34 14 inducible ф10 promoter. Cell extracts were used tod detect the presence of hyperproduced 35 36 15 proteins by SDS-PAGE analysis. Control cells containeing the pURI3 vector plasmid alone 37 38 n 39 16 did not show overexpression, whereas expression of additional 50 kDa protein was 40 t 41 17 apparent with cells harbouring pURI3-TanLp1 ( Figure 1 ). As thie protein was cloned 42 a 43 18 containing an affinity hexa-His tag, recombinant L. plantarum tannase was purified on a 44 l 45 46 19 His-Trap-FF chelating column and eluted with a stepwise gradient of imi-dazole. Highly 47 48 20 purified recombinant tannase protein was obtained from pURI3-TanLp1 ( FigureA 1 ). The 49 50 21 eluted protein was dialyzed to eliminate the imidazole and checked for its tannaseC activity 51 52 S 53 22 on methyl gallate. The purification protocol previously described resulted in a production 54 55 23 yield of 1.48 mg/L, with a specific activity of 84.3 U/mg ( 9). The one-step protocol herein 56 57 58 59 60 11 ACS Paragon Plus Environment Submitted to Journal of Agricultural and Food Chemistry Page 12 of 27

1 2 3 1 described, 17 mg of recombinant protein with specific activity of 2.35 x 10 6 U/mg were 4 5 2 obtained per 1 L culture. 6 7 F 8 3 The biochemical characterization of tannase from L. plantarum has been previously 9 o 10 4 reported from cell-free extracts ( 17 ) and also the pure recombinant enzyme ( 9). As some 11 r 12 13 5 contradictRory results were found among both studies, we decided to confirm the 14 15 6 biochemical data on the pure recombinant L. plantarum CECT 748 T tannase by the 16 e 17 7 colorimetric assayv using methyl gallate as substrate. Figure 2A shows that recombinant 18 19 i 20 8 tannase presented an opetimal pH around 7, being also highly active at pH 6-8. This pH 21 22 9 value is similar to that previouslyw reported for the purified tannase (pH 8) ( 9), and clear 23 24 10 differs from the value reported .from cell-free extracts (pH 5). As recombinant tannases 25 26 C 27 11 showed less than 20% activity at pH 5, the presence of a factor affecting tannase activity in 28 o 29 12 the cell free extract at this pH must be considered. Although the dissimilar observed pH- 30 n 31 13 dependences may result from the above described amino acid substitutions, the effect of 32 f 33 i 34 14 experimental factors such as the different buffer sydstems or the standard assay used for the 35 36 15 determination cannot be discarded. e 37 38 n 39 16 The neutral optimum pH here reported for the recombinant L. plantarum tannase is 40 t 41 17 in contrast to the pH dependence of fungal tannases which are aicidic proteins with an 42 a 43 18 optimum pH around 5.5 ( 1). Figure 2B shows the temperature dependence for recombinant 44 l 45 46 19 tannase activity. The optimum temperature was found to be ~40 ºC, in ag-reement with 47 48 20 previous results ( 9); similarly to cell-free extracts, tannase activity at 50 ºC wAas 75% of the 49 50 21 maximal activity ( 17 ). Tannase from Bacillus cereus KBR9 also showed optimumC activity 51 52 S 53 22 at ~40 ºC ( 20 ). This temperature range represents an advantage since some processes 54 55 23 assisted by tannases are performed at increased temperatures. Figure 2C shows that L. 56 57 58 24 plantarum tannase activity was markedly decreased after incubation at a temperature of 37 59 60 12 ACS Paragon Plus Environment Page 13 of 27 Submitted to Journal of Agricultural and Food Chemistry

1 2 3 1 ºC or higher. The pure recombinant enzyme appears to be more heat-labile that when 4 5 2 present in cell-free extracts ( 17 ). 6 7 F 8 3 As a low substrate level for the maximal activity of the enzyme was considered a 9 o 10 4 positive factor for industrial applications, the effect of substrate concentrations on tannase 11 r 12 13 5 activity wRas studied. The assay was performed at substrate concentrations ranging from 0.1 14 15 6 to 100 mM. Enzyme activity was maximal a 0.7-0.9 mM methyl gallate, with further 16 e 17 7 increases in substrvate concentration resulting in reduced tannase activity (data not shown). 18 19 i 20 8 This substrate concentreation is lower that the previously reported from cell extracts (6.25 21 22 9 mM); however it should bew considered that, in the previous work lower enzyme units were 23 24 10 used in the assay. . 25 26 C 27 11 Table 1 shows the effects of various additives (1 mM final concentration) on L. 28 o 29 12 plantarum CECT 748 T recombinant tannase activity. Compared to the enzyme incubated in 30 n 31 13 50 mM phosphate pH 6.5, the enzymatic activity was increased by KCl and CaCl (relative 32 f 2 33 i 34 14 activity 126-181%), not significantly affected or partiallyd inhibited by EDTA, MgCl 2, 35 36 15 ZnCl 2, Triton-X-100, Tween-80, and DMSO (relative aectivity 80-107%), significantly 37 38 n 39 16 inhibited by urea (relative activity 48%), and was greatly inhibited by HgCl 2 and β- 40 t 41 17 mercaptoethanol (relative activity 15-22%). Whereas the great inhibitoryi effect of HgCl 2 42 a 43 18 agrees with the results derived from cells extracts ( 17 ), the positive effect of CaCl was not 44 l 2 45 46 19 identified. As cell extracts contained all the soluble cell components, the -effect of a 47 48 20 particular additive could be masked. Cell extracts could contain the Ca 2+ ionAs needed for 49 50 21 the optimal activity of L. plantarum tannase. As Ca 2+ ions greatly increased tannaCse 51 52 S 53 22 activity, the substrate specificity assays were done in the presence of 1 mM CaCl 2. 54 55 23 The addition of 1 mM ethyl gallate leads to a 52% relative activity suggesting that 56 57 58 24 ethyl gallate acts as an operative competitive inhibitor of methyl gallate. Taking into 59 60 13 ACS Paragon Plus Environment Submitted to Journal of Agricultural and Food Chemistry Page 14 of 27

1 2 3 1 account that both compounds, methyl and ethyl gallate, are present at the same 4 5 2 concentration on the reaction, this result must indicate that recombinant L. plantarum 6 7 F 8 3 tannase showed similar affinity for both compounds. 9 o 10 4 11 r 12 13 5 SubstrateR specificity of recombinant L. plantarum tannase . When Osawa et al. 14 15 6 (2000) reported for the first time tannase activity in L. plantarum strains by a colorimetric 16 e 17 7 method, they usedv methyl gallate, a simple galloyl ester of methanol, as a substrate of 18 19 i T 20 8 tannase ( 21 ). When tanLp1e was cloned from L. plantarum ATCC 14917 only methyl 21 22 9 gallate was assayed as subswtrate in the colorimetric assay. In order to analyze the substrate 23 24 10 specificity of pure recombinant. L. plantarum CECT 748 T tannase, 18 different simple 25 26 C 27 11 phenolic acid esters were assayed. Among these compounds gallic esters (methyl gallate, 28 o 29 12 ethyl gallate, propyl gallate, and lauryl gallate), benzoic esters (methyl benzoate, and ethyl 30 n 31 13 benzoate), hydroxybenzoic esters (methyl 4-hydroxybenzoate, ethyl 4-hydroxybenzoate, 32 f 33 i 34 14 propyl 4-hydroxybenzoate, and butyl 4-hydroxybendzoate), vanillic ester (methyl vanillate), 35 36 15 dyhydroxybenzoic esters (methyl 2,4-dihydroxybenzoaete, ethyl 3,4-dihydroxybenzoate or 37 38 n 39 16 protocatechuic acid ethyl ester, and ethyl 3,5-dihydroxybenzoate), gentisic ester (methyl 40 t 41 17 gentisate), salicylic ester (methyl salicylate), and ferulic esters (fierulic methyl ester, and 42 a 43 18 ferulic ethyl ester) are found. Only all the esters derived form the gallic acid and the 44 l 45 46 19 derived from the protocatechuic acid (ethyl 3,4-dihydroxybenzoate) were- hydrolyzed 47 48 20 (Figure 3 ). This result seems to indicate that other cinnamic acids without hyAdroxyl groups, 49 50 21 and with substituents other than –H, or –OH at position 2 were not metabolized bCy 51 52 S 53 22 recombinant tannase. It can be also concluded that there are not significant esteric 54 55 23 hindrances regarding the aliphatic alcohol constituent of the ester bond, as ethyl as well as 56 57 58 24 lauryl substituent could be effectively hydrolyzed by the L. plantarum tannase. 59 60 14 ACS Paragon Plus Environment Page 15 of 27 Submitted to Journal of Agricultural and Food Chemistry

1 2 3 1 In vegetable foods and agricultural wastes, together with simple phenolic acid 4 5 2 esters, complex tannins can be found, such as tannic acid or some , which can be 6 7 F 8 3 also potential substrates for the tannase enzyme ( 22 ). In this regard, it has been described 9 o 10 4 that recombinant E. coli colonies producing L. plantarum tannase showed a clear zone on 11 r 12 13 5 tannic acidR-treated plates, which had an opaque surface due to the formation of a tannin- 14 15 6 protein complex. This clear zone indicated apparent bacterial degradation of tannic acid. 16 e 17 7 The effect of L. plantarumv tannase on tannins was clearly demonstrated by incubating cell- 18 19 i 20 8 free extracts on a tannice acid solution ( 17, 23 ). The tannic acid used is a gallotannin mainly 21 22 9 composed of monomers tow tetramers of galloyl glucose ( 23 ). L. plantarum extract degrades 23 24 10 this gallotannin by depolymeriz.ation of high molecular weight tannins and a reduction of 25 26 C 27 11 low molecular weight tannins, being gallic acid and pyrogallol the final metabolic products 28 o 29 12 (23 ). 30 31 n 32 13 To corroborate these previous results, assays were performed to determine the 33 f T i 34 14 activity of pure L. plantarum CECT 748 recombindant tannase on tannic, ellagic, and 35 36 15 chlorogenic acids, and on quercetin, catechin, epicatecehin, gallocatechin, gallocatechin 37 38 n 39 16 gallate, epigallocatechin, and epigallocatechin gallate. From these compounds, only tannic 40 t 41 17 acid, gallocatechin gallate and epigallocatechin gallate were metiabolized by tannase 42 a 43 18 (Figure 4 ). Free gallic acid and pyrogallol were identified as the final products resulting 44 l 45 46 19 from the degradation of hydrolyzable tannins by L. plantarum extracts ( 18,- 23 ). As in our 47 48 20 experimental conditions no gallate decarboxylase is present, the gallic acid thAus formed 49 50 21 cannot be decarboxylated becoming the only final product from tannic acid degraCdation. 51 52 S 53 22 This is notable due to the antioxidant properties of gallic acid. In fact, among 54 55 23 hydroxybenzoic acids, gallic acid is the most potent antioxidant, being 1.6 and 3.4-fold 56 57 58 24 more active than protocatechuic and syringic acids, respectively ( 24 ). Therefore, the use of 59 60 15 ACS Paragon Plus Environment Submitted to Journal of Agricultural and Food Chemistry Page 16 of 27

1 2 3 1 L. plantarum tannase may provide an efficient means for obtaining molecules with valuable 4 5 2 activities from the degradation of complex tannins present in food and agricultural wastes. 6 7 F 8 3 It has been reported that some fungal tannases exhibited also β-glucosidase activity 9 o 10 4 (25 ). In order to know whether L. plantarum CECT 748 T tannase exhibits such an activity, 11 r 12 13 5 the standaRrd assay was performed by incubating recombinant tannase in the presence of 14 15 6 1mM 4-nitrophenyl β-D-glucopyranoside. No activity was detected against this substrate by 16 e 17 7 using the colorimevtric assay as well as by the HPLC analysis of the samples (data not 18 19 i 20 8 shown). e 21 22 9 Conversely, it has bween reported that tannase enzymes cleave ester linkages in 23 24 10 tannic acid ( 26 ), also acting on .the ester linkages in methyl gallate and m-digallic acid, 25 26 C 27 11 respectively. Tannases hydrolyze only those substrates that contain at least two phenolic 28 o 29 12 OH groups in the acid component ( Figure 5 ). The esterified COOH group must be on the 30 n 31 13 oxidized benzene ring and must not be ortho to one of the OH groups. However, 32 f 33 i 34 14 chlorogenic acid, originated from caffeic and quinidc acid, is resistant to tannase activity. 35 36 15 This is attributed to the presence of a double bond in thee side-chain carrying the esterified 37 38 n 39 16 COOH group ( 1). Despite that L. plantarum tannase represents a novel family of tannases 40 t 41 17 showing no significant sequence similarity to fungal tannases, thie reported substrate 42 a 43 18 spectrum of fungal tannases and L. plantarum tannase seems to be very similar ( 27 ). 44 l 45 46 19 - 47 48 20 In conclusion we have described here an improved method for the higAh production 49 50 21 of L. plantarum tannase. Although this protein is not similar to fungal tannases, tChe analysis 51 52 S 53 22 of a broad range of potential substrates reveals that they shared common substrate 54 55 23 specificity. Therefore the use of L. plantarum tannase is an adequate alternative to the 56 57 58 24 fungal tannases currently used in the food industry. 59 60 16 ACS Paragon Plus Environment Page 17 of 27 Submitted to Journal of Agricultural and Food Chemistry

1 2 3 1 4 5 2 6 7 F 8 3 ACKNOWLEDGMENTS 9 o 10 4 11 r 12 13 5 The help oRf J. M. Barcenilla and M. V. Santamaría is greatly appreciated. 14 15 6 16 e 17 7 LITERATURE CITEDv 18 19 i 20 8 e 21 9 (1) Lekha, P. K.; Lonsane, B. K. Production and application of tannin acyl 22 w 23 10 hydrolase: state of the art. Adv. Appl. Microbiol. 1997 , 44, 212-260. 24 . 25 26 11 (2) Aguilar, C. N., GutiérrezC-Sánchez, G. Review: sources, properties, applications 27 28 12 and potential uses of tannin acyl hydrolase. Food Sci. Tech. Int. 2001 , 7, 373- 29 o 30 13 382. 31 n 32 f 33 14 (3) Aguilar, . N.; Rodríguez, R.; Gutiérrezi-Sánchez, G., Augur, C.; Favela-Torres, 34 d 35 15 E.; Prado-Barragán, L. A.; Ramírez-Coronel, A.; Contreras-Esquivel, J. C. 36 e 37 16 Microbial tannases: advances and perspectives. Appl. Microbiol. Biotechnol . 38 n 39 40 17 2007 , 76:47-49. t 41 i 42 18 (4) Belmares, R.; Contreras-Esquivel, J. C.; Rodríguez, R.;a Ramirez-Coronel, A.; 43 44 l 45 19 Aguilar C. N. Microbial production of tannase: an enzyme w ith potential use in 46 - 47 20 food industry. Lebensm.-Wiss. u.-Technol. 2004 , 37, 857-864. 48 A 49 21 (5) Banerjee, R.; Mondal, K.; Bikas, R. Production and characterization of 50 C 51 52 22 extracellular and intracellular tannase from newly isolated Aspergillus aculeatusS 53 54 23 DBF9. J. Basic Microbiol. 2001 ; 6, 313, 318. 55 56 57 58 59 60 17 ACS Paragon Plus Environment Submitted to Journal of Agricultural and Food Chemistry Page 18 of 27

1 2 3 1 (6) Hatamoto, O.; Watarai, T.; Kikuchi, M.; Mizusawa, K.; Sekine, H. Cloning and 4 5 2 sequencing of the gene encoding tannase and a structural study of the tannase 6 7 F 8 3 subunit from Aspergillus oryzae . Gene 1996, 175, 215-221. 9 o 10 4 (7) Zhong, X:, Peng, L.; Zheng, S.; Sun, Z.; Ren, Y.; Dong, M.; Xu, A. Secretion, 11 r 12 13 5 Rpurification, and characterization of a recombinant Aspergillus oryzae tannase in 14 15 6 Pichia pastoris . Protein Expr. Purif. 2004 , 36, 165-169. 16 e 17 7 (8) Noguchvi, N.; Ohashi, T.; Shiratori, T.; Narui, K.; Hagiwara, T.; Ko, M.; 18 19 i 20 8 Watanabe, Ke.; Miyahara, T.; Taira, S.; Moriyasu, F.; Sasatsu, M. Association of 21 22 9 tannase-producingw Staphylococcus lugdunensis with colon cancer and 23 24 10 characterization of a .novel tannase gene. J Gastroenterol . 2007 , 42, 346-351. 25 26 C 27 11 (9) Iwamoto, K.; Tsuruta, H.; Nishitaini, Y.; Osawa, R. Identification and cloning of 28 o 29 12 a gene encoding tannase (tannin acylhydrolase) from Lactobacillus plantarum 30 n 31 13 ATCC 14917 T. Syst. Appl. Microbiol . 2008 , 31, 269-277. 32 f 33 i 34 14 (10) Freney, J.; Brun, Y.; Bes, M.; Meugnier,d H.; Grimont, F.; Grimont, P. A. D. ; 35 36 15 Nervi, C. ; Fleurette, J. Staphylococcus lugdunensise sp. nov. and Staphylococcus 37 38 n 39 16 schleiferi sp. nov., two species from human clinical specimens. Int. J. Syst. 40 t 41 17 Bacteriol. 1988 , 38, 168-172. i 42 a 43 18 (11) Bellamy, R.; Barhkam, T. Staphylococcus lugdunensis infection sites: 44 l 45 46 19 predominance of abscesses in the pelvic girdle region. Clin. Infect.- Dis . 2002 ; 35, 47 48 20 E32-E34. A 49 50 21 (12) Greig, J. M., Wood, M. J. Staphylococcus lugdunensis vertebral osteomCyelitis. 51 52 S 53 22 Clin. Microbiol. Infect . 2003 , 9, 1139-1141. 54 55 23 (13) Patel, R.; Piper, K. E.; Rouse, M. S.; Uhl, J. R.; Cockerill, F. R.; Steckelberg, J. 56 57 58 24 M. Frequency of isolation of Staphylococcus lugdunensis among staphylococcal 59 60 18 ACS Paragon Plus Environment Page 19 of 27 Submitted to Journal of Agricultural and Food Chemistry

1 2 3 1 isolated causing endocarditis: a 20-year experience. J. Clin. Microbiol . 2000 , 38, 4 5 2 4262-4263. 6 7 F 8 3 (14) De las Rivas, B.; Curiel, J. A.; Mancheño, J. M.; Muñoz, R. Expression vectors 9 o 10 4 for enzyme restriction- and ligation-independent cloning for producing 11 r 12 13 5 Rrecombinant His-fusion proteins. Biotech. Progress 2007 , 23, 680-686. 14 15 6 (15) Sambrook, J.; Fritsch, E. F.; Maniatis, T. Molecular Cloning: A Laboratory 16 e 17 7 Manual,v 2 nd ed.; Cold Spring Harbor Laboratory Press: Cold Spring Harbor, NY, 18 19 i 20 8 1989. e 21 22 9 (16) Inoue, K. H.; Hawgerman, A. E. Determination of gallotannin with rhodanine. 23 24 10 Anal. Biochem . 1988 ., 169, 363-369. 25 26 C 27 11 (17) Rodríguez, H.; de las Rivas, B.; Gómez-Cordovés, C:, Muñoz, R. 28 o 29 12 Characterization of tannase activity in cell-free extracts of Lactobacillus 30 n 31 13 plantarum CECT 748 T. Int. J. Food Microbiol. 2008 , 121, 92-98. 32 f 33 i 34 14 (18) Rodríguez, H.; Landete, J. M.; Curiel, J.d A.; de las Rivas, B.; Mancheño, J. M.; 35 36 15 Muñoz, R. Characterization of the p-coumareic acid decarboxylase from 37 38 T n 39 16 Lactobacillus plantarum CECT 748 . J. Agric. Food Chem . 2008 , 56, 3068- 40 t 41 17 3072. i 42 a 43 18 (19) Landete, J. M.; Rodríguez, H.; de las Rivas, B.; Muñoz, R. Characterization of a 44 l 45 46 19 benzyl alcohol dehydrogenase from Lactobacillus plantarum WCFS1.- J. Agric. 47 48 20 Food Chem. 2008 , 56, 4497-4503. A 49 50 21 (20) Mondal, K. C.; Banerjee, D.; Banerjee, R.; Pati, B. R. Production and C 51 52 S 53 22 characterization of tannase from Bacillus cereus KBR9. J. Gen. Appl. Microbiol . 54 55 23 2001 , 47, 263-267. 56 57 58 59 60 19 ACS Paragon Plus Environment Submitted to Journal of Agricultural and Food Chemistry Page 20 of 27

1 2 3 1 (21) Osawa, R.; Kuroiso, K.; Goto, S.; Shimizu, A. Isolation of tannin-degrading 4 5 2 lactobacilli from humans and fermented foods. Appl. Environ. Microbiol. 2000 , 6 7 F 8 3 66, 3039-3097. 9 o 10 4 (22) Battestin, V.; Macedo, G. A.; De Freitas, V. A. P. Hydrolysis of epigallocatechin 11 r 12 13 5 Rgallate using a tannase from Paecilomyces variotii . Food Chem . 2008 , 108, 228- 14 15 6 233. 16 e 17 7 (23) Rodríguvez, H.; de las Rivas, B.; Gómez-Cordovés, C.; Muñoz, R. Degradation of 18 19 i 20 8 tannic acid bye cell-free extracts of Lactobacillus plantarum . Food Chem . 2008 , 21 22 9 107, 664-670. w 23 24 10 (24) Ordoudi, S., A.; Tsim.idou, M. Z. Crocin bleaching assay (CBA) in structure- 25 26 C 27 11 radical scavenging activity studies of selected phenolic compounds. J. Agric. 28 o 29 12 Food Chem. 2006 , 54, 9347-9356. 30 n 31 13 (25) Ramirez-Coronel, M. A.; Viniegra-González, G.; Darvill, A.; Augur, C. A novel 32 f 33 i 34 14 tannase from Aspergillus nige r with β-glucosidased activity. Microbiology 2003 , 35 36 15 149, 2941-2946. e 37 38 n 39 16 (26) Haslam, E.; Stangroom, J. E. The esterase and depsidase activities of tannase. 40 t 41 17 Biochem. J . 1966 , 99, 28-31. i 42 a 43 18 (27) Iibuchi, S.; Minoda, Y.; Yamada, K. Hydrolyzing pathway, substrate specificity 44 l 45 46 19 and inhibition of tannin acyl hydrolase of Asp. oryzae nº 7. Agr.- Biol. Chem . 47 48 20 1972 , 36, 1553-1562. A 49 50 21 C 51 52 S 53 22 54 55 23 This work was supported by grants AGL2005-00470, AGL2008-01052, BFU2007- 56 57 58 24 67404/BMC, Consolider INGENIO 2010 CSD2007-00063 FUN-C-FOOD and 59 60 20 ACS Paragon Plus Environment Page 21 of 27 Submitted to Journal of Agricultural and Food Chemistry

1 2 3 1 Consolider INGENIO 2007 Factoria de Cristalización (CICYT), RM2008-00002 4 5 6 2 (INIA), and S-0505/AGR/000153 (CAM). H. Rodríguez and J. A. Curiel were 7 F 8 3 recipients of predoctoral fellowships from the I3P-CSIC Program and FPI-MEC, 9 o 10 4 respectively. 11 r 12 13 5 R 14 15 6 16 e 17 v 18 19 i 20 e 21 22 w 23 24 . 25 26 C 27 28 29 o 30 31 n 32 f 33 i 34 d 35 36 e 37 38 n 39 40 t 41 i 42 a 43 44 l 45 46 - 47 48 A 49 50 C 51 52 S 53 54 55 56 57 58 59 60 21 ACS Paragon Plus Environment Submitted to Journal of Agricultural and Food Chemistry Page 22 of 27

1 2 3 Table 1 4 5 6 7 F 8 9 o 10 11 r 12 T 13 TableR 1. Effect of additives on recombinant L. plantarum CECT 748 tannase activity 14 15 16 e Additions Relative activity 17 v(1 mM) (%) 18 19 Controli 100 20 EDTA e 107 21 KCl 126 22 w HgCl 2 22 23 CaCl 181 24 2 MgCl . 88 25 2 26 ZnCl 2 C 82 27 Triton X 100 80 28 DMSO o 91 29 Tween 80 80 30 Urea 48 31 n 32 β-mercaptoethanol 15f 33 Ethyl gallate 52 i 34 d 35 36 e 37 38 n 39 40 t 41 i 42 a 43 44 l 45 46 - 47 48 A 49 50 C 51 52 S 53 54 55 56 57 58 59 60 22 ACS Paragon Plus Environment Page 23 of 27 Submitted to Journal of Agricultural and Food Chemistry

1 2 3 Figure 1 4 5 6 7 F 8 9 o 10 11 r 12 13 R 14 15 e 12345678 16 kDa 17 v 18 97 - 19 i 66 - 20 e 21 45 - 22 w 23 24 31 - . 25 26 C 27 28 29 o 30 31 n 32 Figure 1 . SDS-PAGE analysis of the expression and purification of the tannase protein 33 f T i 34 from L. plantarum CECT 748 . Analysis of solubled cell extracts of IPTG-induced E. coli 35 36 JM109 (DE3) (pURI3) (lane 1) or E. coli JM109 (DE3)e (pURI3-TanLp1) cultures (lane 2), 37 38 n 39 or fractions eluted after His-Trap-FF crude chelating affinity column (lines 3 to 8). SDS- 40 t 41 Polyacrylamide gel (12%) was stained with Coomassie blue. Thei positions of molecular 42 a 43 mass markers (Bio-Rad) are indicated on the left. 44 l 45 46 - 47 48 A 49 50 C 51 52 S 53 54 55 56 57 58 59 60 23 ACS Paragon Plus Environment Submitted to Journal of Agricultural and Food Chemistry Page 24 of 27

1 2 3 Figure 2 4 5 A 6 7 F 100 8 80 9 o 60 10 . 40 11 r 20 12 0 13 R activity (%) Enzyme 3 4 5 6 7 8 910 14 pH 15 16 e 17 Bv 18 19 i 100 20 e80 21 60 22 w40 23 20 24 0 . 25 EnzymeEnzymeactivity activity(%) . (%) . 20 30 40 50 60 70 26 CTemperature (ºC) 27 28 29 o 30 C n 31 100 32 80 33 f 60 i 34 d 35 40 36 20 e 37 0 38 (%) . activity Enzyme 0 5 10 15n 20 39 Time (h) 40 t 41 i 42 a 43 44 l 45 Figure 2 . Biochemical properties of recombinant L. plantarum CECT 748 T tannase. (A) 46 - 47 48 Relative activity of the tannase versus pH. Enzyme activity was assayed at 37A ºC. (B) 49 50 Relative activity of tannase versus temperature. Enzyme activity was assayed at pHC 6.5. (C) 51 52 Residual activities of the recombinant L. plantarum tannase after preincubation at 25 SºC 53 54 55 (♦), 30 ºC ( ■), 37 ºC ( ▲) or 45 ºC ( ●) in phosphate buffer pH 6.5 during 15 or 30 min, and 56 57 1, 2, 3, 4, and 20 h. In all the cases, the observed maximum activity was defined as 100%. 58 59 60 24 ACS Paragon Plus Environment Page 25 of 27 Submitted to Journal of Agricultural and Food Chemistry

1 2 Control Tannase 3 400400 4 A A 10001000 5 GA 6 200 mAU200 mAU MG 7 F 500500 8 9 o 00 00 0 20 40 60 80 100 120 0 20 40 60 80 100 120 10 B 400400 B 11 r 10001000 12 200200 EG mAU

mAU 13 R500500 GA 14

15 00 00

mAU e 0 20 40 60 80 100 120 16 1500 0 20 40 60 80 100 120 1500 1500 Minutes C 1500 Minutes C 17 v 18 10001000 10001000 mAU 19 mAU i PG GA 20 500500 e 500500

21 00 00 22 20 40 w60 80 100 120 0 20 40 60 80 100 120 15001500 1500 Minutes D 1500 Minutes 23 D D 24 10001000 . 10001000 mAU 25 LG mAU 500500 GA 26 C 500500 27 00 28 00 00 2020 40 60 60 80 100100 o120 00 20 40 60 80 100100 120 20 60 29 Minutes Minutes 30 200200 31 n E 10001000 E 32 EP f 33 100100 PA mAU mAU mAU 500500 i 34 d 35 36 00 00 0 20 40 60 80 e 0 20 40 60 80 0 2040 60 8080 37 Minutes 38 Time (min) nTime (min) 39 40 t 41 i 42 Figure 3 . HPLC chromatograms of recombinant L. plantarum tannasea on different gallic 43 44 l T 45 and protocatechuic esters. Tannase enzyme from L. plantarum CECT 748 was incubated 46 - 47 for 10 min in the presence of methyl gallate (A), ethyl gallate (B), propyl gallate (C), lauryl 48 A 49 gallate (D) or protocatechuic acid ethyl ester (E). Chromatograms without protein (controls) 50 C 51 52 are also showed. The chromatograms were recorded at 280 nm. MG, methyl gallate; EG,S 53 54 ethyl gallate; PG, propyl gallate; LG, lauryl gallate; EP, or 55 56 protocatechuic acid ethyl ester; GA, gallic acid; PA, protocatechuic acid. 57 58 59 60 25 ACS Paragon Plus Environment Submitted to Journal of Agricultural and Food Chemistry Page 26 of 27

1 2 3 Figure 4 4 5 6 7 F 8 9 o Control Tannase 10 15001500 11 r 10001000 A A 12 TA 10001000 13 500500 mAU R mAU GA 14 500500 15 e 16 00 00 0 20 40 60 80 17 500500 v B B 18 GCG GA 50 50 mAU

19 mAU mAU 250250 i 20 e GC 21 00 00 22 0 20 w40 60 80 0 20 40 60 80 15001500 23 C C GA 24 10001000 100100

EGCG mAU mAU . 25 EGC 500500 26 C 27 00 00 0 20 40 60 8080 0 20 40 60 80 28 0 20 40 60 0 20 40 60 80 29 Time (min)o Time (min) 30 31 n 32 f 33 Figure 4 . HPLC chromatograms of recombinant iL. plantarum tannase on tannins. Tannase 34 d 35 enzyme from L. plantarum CECT 748 T was incubated for 10 min in the presence of tannic 36 e 37 acid (A), gallocatechin gallate (B) or epigallocatechin gallate (C). Chromatograms without 38 n 39 40 protein (controls) are also showed. The chromatograms were recordedt at 280 nm. TA, 41 i 42 tannic acid; GCG, gallocatechin gallate; EGCG, epigallocatechin agallate; GA, gallic acid; 43 44 l 45 GC, gallocatechin; EGC, epigallocatechin. 46 - 47 48 A 49 50 C 51 52 S 53 54 55 56 57 58 59 60 26 ACS Paragon Plus Environment Page 27 of 27 Submitted to Journal of Agricultural and Food Chemistry

1 2 3 Figure 5 4 5 6 7 F 8 9 o 10 11 r 12 13 R 14 15 O R1 16 e 17 v 18 19 i 20 e 21 22 w 23 24 . R2 OH 25 26 C 27 OH 28 29 o 30 31 n 32 Figure 5 . Schematic representation of the potentialf substrates of L. plantarum CECT 748 T 33 i 34 d 35 tannase. The R 1 substituents could be –OCH 3, -OCH 2-CH 3, -OCH 2-CH 2-CH 3, - 36 e 37 OCH 2(CH 2)10 -CH 3, glucose, gallic acid, gallocatechin, and epigallocatechin. The R 2 38 n 39 substituents could be –H or –OH. 40 t 41 i 42 a 43 44 l 45 46 - 47 48 A 49 50 C 51 52 S 53 54 55 56 57 58 59 60 27 ACS Paragon Plus Environment