- Open Access
Identification of a highly active tannase enzyme from the oral pathogen Fusobacterium nucleatum subsp. polymorphum
- Julen Tomás-Cortázar†1,
- Laura Plaza-Vinuesa†3,
- Blanca de las Rivas3,
- José Luis Lavín2,
- Diego Barriales1,
- Leticia Abecia1,
- José Miguel Mancheño4,
- Ana M. Aransay2, 5,
- Rosario Muñoz3,
- Juan Anguita1, 6Email authorView ORCID ID profile and
- Héctor Rodríguez1Email author
© The Author(s) 2018
- Received: 2 October 2017
- Accepted: 22 February 2018
- Published: 26 February 2018
Tannases are tannin-degrading enzymes that have been described in fungi and bacteria as an adaptative mechanism to overcome the stress conditions associated with the presence of these phenolic compounds.
We have identified and expressed in E. coli a tannase from the oral microbiota member Fusobacterium nucleatum subs. polymorphum (TanBFnp). TanBFnp is the first tannase identified in an oral pathogen. Sequence analyses revealed that it is closely related to other bacterial tannases. The enzyme exhibits biochemical properties that make it an interesting target for industrial use. TanBFnp has one of the highest specific activities of all bacterial tannases described to date and shows optimal biochemical properties such as a high thermal stability: the enzyme keeps 100% of its activity after prolonged incubations at different temperatures up to 45 °C. TanBFnp also shows a wide temperature range of activity, maintaining above 80% of its maximum activity between 22 and 55 °C. The use of a panel of 27 esters of phenolic acids demonstrated activity of TanBFnp only against esters of gallic and protocatechuic acid, including tannic acid, gallocatechin gallate and epigallocatechin gallate. Overall, TanBFnp possesses biochemical properties that make the enzyme potentially useful in biotechnological applications.
We have identified and characterized a metabolic enzyme from the oral pathogen Fusobacterium nucleatum subsp. polymorphum. The biochemical properties of TanBFnp suggest that it has a major role in the breakdown of complex food tannins during oral processing. Our results also provide some clues regarding its possible participation on bacterial survival in the oral cavity. Furthermore, the characteristics of this enzyme make it of potential interest for industrial use.
- Phenolics biotransformation
- Oral pathogen
Tannins are high molecular weight secondary phenolic metabolites of plant origin that have been traditionally considered antinutrients due to their capacity to bind and precipitate protein [1, 2]. This group of chemicals can be found in tea, wine, berries, fruits and chocolate among other dietary components. They are toxic compounds for a variety of microorganisms because of their protein and iron binding capacity, and may interfere with many biological processes that are essential for their growth . In turn, some microorganisms have developed strategies to overcome tannin toxicity, including the presence of genes in their genomes encoding degrading enzymes such as tannase enzymes . Initially, tannases, also known as tannin acyl hydrolases (EC 22.214.171.124), were described in fungi and studied primarily because of their industrial use in processes related to food detannification and the removal of pollutants, in the leather industry or for the production of gallic acid, which is an important intermediate in the synthesis of the antibiotic trimethoprim [3, 4]. Recently, tannases have also been isolated from bacteria that populate environments rich in vegetable content, although just a few of the genes encoding bacterial tannases have been described and even fewer have been characterized biochemically. The most studied tannases are present in bacteria isolated from the rumen, gut microbiota or soils with abundant vegetation [5–11]. Two different types of tannases have been described so far that are encoded by two different genes. Extracellular tannases (encoded by the tanA gene) contain a signal peptide and a molecular size of around 66 kDa. These include TanASl from Staphylococcus lugdunensis [5, 9, 12, 13], TanALp from Lactobacillus plantarum , and TanASg from Streptococcus gallolyticus . In addition, 50 kDa intracellular bacterial tannases (encoded by tanB genes) have also been described. These proteins are found in L. plantarum (TanBLp) [6, 15] or S. gallolyticus (TanBSg) . Recently, an extracellular tannase that is nevertheless encoded by a tanB gene has been described in Streptomyces sviceus (TanBSs) .
The gut and the oral cavity harbor distinctive populations of bacteria permanently exposed to a vast array of chemical compounds present in food, including tannins. The human microbiota is a massive and largely unexplored source of enzymes with new and/or improved activities ; however, limited research has been performed to identify microbes capable of degrading tannins within the inhabitants of the human body. Although bacteria that contain enzymes with tannase activity, such as L. plantarum, S. gallolyticus, or S. lugdunensis, have been described in the human gastrointestinal tract [9, 17–19], many questions remain about the oral metabolism of food tannins. As the oral cavity is continuously exposed to tannins, we hypothesized that oral bacteria might harbor tannin-degrading genes in their genomes. Previously, a tannase (TanAAp) from Atopobium parvulum, a species abundant in the oral cavity, was characterized . This protein exhibited the lowest specific activity among bacterial tannases and was unable to hydrolyze complex tannins. Therefore, the biochemical properties of TanAAp discarded its role in the breakdown of complex food tannins. Herein, we have identified and characterized the first tannase enzyme described within the genus Fusobacterium and overall, from a pathogenic bacterium. The study of its biochemical properties and the substrates among different relevant tannin derivatives present in food demonstrate that it is among the most active tannases described so far possessing a wide range of substrate specificity that includes several derivatives of gallic acid, protocatechuic acid and complex tannins.
Presence of a putative tannase in members of the genus Fusobacterium
TanBFnp is among the most active bacterial tannases identified to date
Biochemical characterization of TanBFnp
We then determined the biochemical properties of TanBFnp using the purified protein. Its optimum activity pH was measured at 30 °C in 50 mM phosphate buffer. Figure 4a shows an optimum pH for TanBFnp of 7 and suboptimal activity of the enzyme at pH values ranging from 6 to 8, although it still retained around 80% of the maximal activity. The optimal activity temperature was determined incubating the protein in 50 mM phosphate buffer pH 6.5. TanBFnp showed the highest activity at 55 °C, albeit it retained around 80% of the maximal activity at all temperatures tested between 22 and 55 °C (Fig. 4b). The thermal stability of the enzyme was also determined by pre-incubation at different temperatures ranging from 22 to 65 °C for increasing lengths of time up to 20 h, followed by activity determination. Surprisingly, pre-incubation of the enzyme between 22 and 45 °C did not substantially affect the activity of the enzyme, maintaining methyl gallate hydrolysis rates above 60% of the maximum activity and in some cases, increasing the enzymatic activity. On the contrary, incubation at temperatures higher than 45 °C induced a dramatic decrease in TanBFnp activity after 30 min of incubation (Fig. 4c).
Relative activity of TanBFnp in the presence of different additives
Additive (1 mM)
Relative activity (%)
Substrate specificity of TanBFnp
Activity of TanBFnp against different substrates
The search for new tannase enzymes with improved activity and new substrate specificities constitutes a permanent focus for industrial microbiology research due to their extensive use in the food and pharmaceutical industries. Indeed, the search for enzymes with tannase activity has recently been expanded to the human microbial population. Herein, we describe a new tannase enzyme from the oral pathogen F. nucleatum susbp. polymorphum. Oral bacteria need to have extensive metabolic resources to face the presence of food components with antimicrobial properties that they may encounter. We previously described a tannase from A. parvulum, an inhabitant of the human oral cavity . However, this tannase (TanAAp) possessed low specific activity and was unable to hydrolyze complex tannins. Therefore, it is unlikely that this tannase contributes to tannin breakdown. Because the presence of enzymes with tannase activity is one of the preferred mechanisms to overcome phenolic-related stress, we sought and identified a homologous protein of the tannase from L. plantarum, TanBLp, within the Fusobacterium genus. Phylogenetically, TanBFnp shows a closer relationship with TanBLp than with all other bacterial tannases. TanBLp has been previously associated with a cluster of tannases unrelated to those of fungal origin . Therefore, TanBFnp might be included within the same cluster of bacterial tannin-degrading enzymes.
The biochemical characterization of TanBFnp reveals that it is an enzyme with potential industrial applications. Its activity against methyl gallate is among the highest described in bacteria so far and considerably higher than other tannases previously reported [7, 8, 10, 11, 15, 20]. Other biotechnological features that are important for its potential industrial application include its temperature range of activity. The optimal temperature of most tannases varies between 30 and 40 °C . Strikingly, TanBFnp is highly active in a much wider range of temperatures, from 22 to 60 °C, with a maximum activity peak at 60 °C, similar to S. gallolyticus (TanBSg) and S. sviceus tannases (TanBSs) [7, 11]. Accordingly, TanBFnp thermal stability is much higher than that of all previously described tannases. Similar to TanBSg, TanBFnp is able to keep high activity rates after it has been exposed for long time intervals to temperatures ranging from 22 to 45 °C while most bacterial tannases suffer a deep decrease in their activity after long exposures to temperatures equal to or higher than 37 °C. As previously suggested , it is possible that for extracellular tannases, such as TanBFnp, a better adaptability to the harsh extracellular environment has induced the development of more resistant enzymes. Both the high performance temperature and its thermal resistance to unfolding or denaturalization in the absence of substrate are key features that support TanBFnp biotechnological use. These characteristics may likely improve its interaction with substrates, favor high mass transfer rates, and lower the risk of contamination .
Like all bacterial tannases characterized to date, TanBFnp showed the highest activity rates at pH between 6 and 8, which overlaps the pH of the oral cavity (between 6.7 and 7.3). The neutral pH range of activity seems to be a common characteristic of bacterial tannases in contrast with those of fungal origin that present activity peaks under acidic conditions .
The search for enzymes with new activities is also an important target in tannase research due to the high variety of tannin substrates. We performed a substrate characterization using 27 different compounds from different tannin families. The results presented are identical to the substrate specificity previously described for TanBLp, the most similar tannase to TanBFnp. While other tannases such as TanBSg and TanALp only degrade esters of phenolic acid with short chain aliphatic alcohols [7, 8], TanBFnp is capable of degrading esters of phenolic acids with long chain alcohols such as lauryl gallate. The hydrolyzing activity of TanBFnp against longer esters points to a markedly different substrate-binding site in this enzyme that would permit the access of these bulkier compounds.
Fusobacterium nucleatum subsp. polymorphum is an inhabitant of the human oral cavity frequently isolated from dental plaque biofilms . As a pathogen, it has been repeatedly associated with peridontitis and extraoral infections including preterm births and colorectal cancer . However, little is known about the metabolic arsenal that allows them to thrive in these environments. The extracellular tannase identified in F. nucleatum susps. polymorphum, TanBFnp, might provide an ecological advantage to bacteria thriving in a niche permanently exposed to phenolic stress from food sources. The enzyme is still highly active (80% of its optimal activity) at body temperature and at pH usually found within the human body. Vegetable food residues are permanently deposited in the oral cavity as a consequence of food intake. An extracellular tannase could be an important survival factor for F. nucleatum with a dual role during bacterial colonization of teeth: the detoxification of toxic compounds and the provision of a source of sugar moieties resulting from the hydrolyzation of complex tannins. The wide distribution of tannase enzymes among oral inhabitants in comparison to microbial species in other niches suggests that its presence may constitute an adaptative advantage to compete in an environment that is permanently receiving tannins from diet sources. Therefore, TanBFnp may constitute a putative virulence factor for F. nucleatum and a potential target of therapeutic intervention for this pathogen.
The description of TanBFnp increases our knowledge about tannin breakdown in the human gastrointestinal tract. Oral and intestinal tannases could contribute to tannin digestion. At least five different species of oral or intestinal bacteria have been described to possess active tannases, including A. parvulum, L. plantarum, S. gallolyticus, S. lugdunensis and, in this work, Fusobacterium nucleatum subsp. polymorphum. Further testing would be required to define the metabolism of these phenolic compounds comprehensively. In particular, the activity of the complex communities of microorganisms present in all parts of the human digestive tract would need to be examined. Moreover, as tannase-producing bacteria have been identified in several human cancer microbiomes , the study of the association between dietary tannin intake and tumor recurrence or regression, may be critical in understanding the role of gut bacteria on the anti-cancer effects of dietary polyphenols.
In this work, we describe an enzyme with tannase activity in the oral pathogen Fusobacterium nucleatum subsp. polymorphum. TanBFNP is one of the most active tannases described to date. This fact combined with an extraordinary thermal stability and a wide range of temperature activity makes TanBFNP a suitable candidate for industrial applications. Moreover, since F. nucleatum subsp. polymorphum is a pathogen associated with oral and extra-oral diseases our research increases the knowledge about a putative niche adaptation determinant that might be involved in virulence associated with the transformation of diet antimicrobial compounds.
Bacterial strains and growth conditions
Escherichia coli strains DH5α and BL21(DE3) were used as transformation and expression hosts, respectively, for the pHISp-fusotan vector. The bacteria were grown in LB medium containing ampicillin (100 µg/mL) at 37 °C with agitation.
Genomic DNA from F. nucleatum subsp. polymorphum Strain F0401 was obtained from BEI Resources Repository. Standard molecular biology procedures  were followed to clone the tannase gene avoiding the signal peptide of the protein. The gene was PCR-amplified using phusion hot start II polymerase (Thermo Fisher Scientific, Waltham, MA) with primers Fusotannase-f (5′-CGC CAT GGG CGT AAA AAA TGA GTA TGA TT-3′) and Fusotannase-r (5′-CGG TCG ACT TAT TTT TTT ACA ACA CCA TC-3′). The PCR product was purified using a PCR purification kit (Thermo Fisher Scientific). The 1.5 kb PCR product and the vector pHis parallel 2 (Addgene) were digested with NcoI and SalI and then ligated for 16 h using T4 DNA ligase (Promega, Madison, WI). The ligation mixture was transformed in DH5α cells and positive colonies were verified by colony PCR and sequencing. Positive plasmids (pHISp-fusotan) were finally transformed into Escherichia coli BL21(DE3) cells for protein production.
Protein production and purification
Sequence-confirmed TanB Fnp clones in E. coli BL21(DE3) were grown in Luria–Bertani medium supplemented with ampicillin and induced with 1 mM IPTG for 16 h at 20 °C. The His-tag fusion protein was then purified by nickel affinity chromatography (GE Healthcare, Uppsala, Sweden) and eluted in 20 mM Tris, pH 7.5 with 150 mM NaCl and 250 mM Imidazole. For a second purification step using gel filtration chromatography, fractions containing TanBFnp identified by SDS-PAGE were pooled, concentrated and loaded onto a HiLoad 10/300 GL Superdex 75 column (GE Healthcare) pre-equilibrated in 20 mM Tris pH 7.5; 150 mM NaCl, using an AKTA chromatography system (GE Healthcare). Fractions with the protein of interest were pooled and the protein was concentrated and stored at − 80 °C until its use. Protein concentration was determined using the BCA protein assay kit (Thermo Fisher Scientific).
Determination of tannase activity
The generation of gallic acid in hydrolysis reactions was determined in triplicate with the following assay: TanBFnp (10 μg) in 700 μL of 50 mM phosphate buffer, pH 6.5, was incubated with 40 μL of 25 mM methyl gallate (1 mM final concentration) for 5 min at 37 °C. After incubation, 150 μL of a methanolic rhodanine solution (0.667% rhodanine in 100% methanol) was added to the reaction mixture. After 5 min of incubation at 30 °C, 100 μL of 0.5 M KOH was added and the absorbance at 520 nm was measured on a spectrophotometer. A standard curve using gallic acid concentrations ranging from 0.125 to 1 mM was prepared. One unit of tannase activity was defined as the amount of enzyme required to release 1 μmol of gallic acid per minute under standard reaction conditions.
Activities of TanFnp from F. nucleatum subsp. polymorphum F401 were measured at 4, 20, 30, 37, 45, 55, and 65 °C to determine the optimal temperature for enzymatic activity. The optimum pH value for tannase activity was determined by studying its pH dependence within the pH range between 3 and 10. The buffers used were: acetic acid-sodium acetate buffer for pH 3–5, citric acid-sodium citrate for pH 6, sodium phosphate for pH 7, Tris–HCl for pH 8, glycine–NaOH for pH 9, and sodium carbonate-bicarbonate for pH 10. A 100 mM concentration was used in all the buffers. The rhodanine assay was used for the optimal pH characterization of TanBFnp . Since the rhodanine-gallic acid complex forms only under basic conditions, after the enzymatic degradation of methyl gallate, 100 μL of 0.5 M KOH was added to the reaction mixture to ensure that the same pH value (pH 11) was achieved in all samples assayed.
For temperature stability measurements, TanBFnp was incubated in 50 mM phosphate buffer, pH 6.5, at 22, 30, 37, 45, 55, and 65 °C for 15 min, 30 min, and 1, 2, 5, and 18 h. After incubation, the residual activity was measured as described above.
To test the effects of metals and ions on the activity of TanBFnp, the enzymatic activity was measured in the presence of different additives at a final concentration of 1 mM. The additives analyzed were MgCl2, KCl, CaCl2, HgCl2, ZnCl2, Triton X-100, urea, Tween 80, EDTA, dimethyl sulfoxide (DMSO) and β-mercaptoethanol. All the determinations were done in triplicate.
TanBFnp substrate specificity
The activity of TanBFnp against 27 potential substrates was analyzed. The substrates assayed were gallic esters (methyl gallate, ethyl gallate, propyl gallate, and lauryl gallate), benzoic esters (methyl benzoate and ethyl benzoate), hydroxybenzoic esters (methyl 4-hydroxybenzoate, ethyl 4-hydroxybenzoate, propyl 4-hydroxybenzoate and butyl 4-hydroxybenzoate), a vanillic ester (methyl vanillate), dyhydroxybenzoic esters (methyl 2,4-dihydroxybenzoate, ethyl 3,4-dihydroxybenzoate or protocatechuic acid ethyl ester and ethyl 3,5-dihydroxybenzoate), a gentisic ester (methyl gentisate), a salicylic ester (methyl salicylate) and ferulic esters (ferulic methyl ester and ferulic ethyl ester). Tannic acid and epigallocatechin gallate were also assayed as potential substrates. Recombinant tannase (10 μg/mL) was incubated at 37 °C in ammonium acetate 50 mM, pH 5 in the presence of the substrate (1 mM). As controls, acetate buffers containing the reagents but not the enzyme were incubated under the same conditions. After incubation, the samples were analyzed by high-performance liquid chromatography with diode array detection (HPLC–DAD). A Thermo chromatograph (Thermo Fisher Scientific) equipped with a P4000 SpectraSystem pump, an AS3000 autosampler, and a UV6000LP photodiode array detector was used. A gradient of solvent A (water-acetic acid, 98:2 [vol/vol]) and solvent B (water-acetonitrile-acetic acid, 78:20:2 [vol/vol/vol]) was applied to a reversed-phase Nova-pack C18 cartridge (25 cm by 4.0-mm inside diameter [i.d.]; 4.6-μm particle size) at room temperature as follows: 0–55 min, 80% B linear, 1.1 mL/min; 55–57 min, 90% B linear, 1.2 mL/min; 57–70 min, 90% B isocratic, 1.2 mL/min; 70–80 min, 95% B linear, 1.2 mL/min; 80–90 min, 100% linear, 1.2 mL/min; 100–120 min, washing, 1.0 mL/min; and re-equilibration of the column under the initial gradient conditions. Samples were injected onto the cartridge after being filtered through a 0.45-μm PVDF filter. Detection of the substrates and the degradation compounds was performed spectrophotometrically by scanning from 220 to 380 nm. The identification of degradation compounds was carried out by comparing the retention times and spectral data of each peak with those of standards from commercial suppliers.
The sequences of tanases analyzed in this work were inspected for conserved functional domains with the CDD web tool . Then, the region harboring the functional domain related to the tanase activity was extracted. These sequences were the input for a Multiple Sequence Analysis using CLUSTAL omega , in order to identify conserved amino acid patterns among them, and use it as input for the phylogenetic analysis carried out by the Phylogeny.fr web-service  including the following steps: the CLUSTAL omega alignment was used as input, followed by the removal of poorly aligned/gapped regions using Gblocks v0.91b  with default settings for both tools. Phylogenetic trees were reconstructed using the maximum likelihood method with the PhyML program v3.0  using the WAG substitution model and 100 bootstrap replicates for inner branch accuracy. The graphical representation and edition of the phylogenetic tree was performed with TreeDyn v198.3 . Sequence logos were obtained using Python’s WebLogo package  using the CLUSTAL omega’s alignment file as input. The scale of the logo was measured in bits, which are units of measure with a precise thermodynamic relationship to energy. Display error bars indicate an approximate Bayesian 95% confidence interval.
In order to study the distribution of tannase ortholog genes among Fusobacterium spp. (in dental plaque and gut microflora) and other oral bacteria, we performed a BLASTp  search against the non-redundant database limited to bacteria taxa. Results were filtered by e-value (< 1e−20). Those with a query coverage > 50% and sequence identity > 20% were retained as putative orthologs (Additional file 1: Table S1).
JA and HR designed the study. JT, LPV, BDR and DB performed the experimental work. JLL and AMA performed bioinformatic analysis. HR, JA, LA, JLL, JMM and RM analyzed the results and wrote the manuscript. All authors read and approved the final manuscript.
The following reagent was obtained through BEI Resources, NIAID, NIH: Genomic DNA from Fusobacterium nucleatum subsp. polymorphum, Strain F0401.
The authors declare that they have no competing interests.
Availability of data and materials
All data generated or analysed during this study are included in this published article. The plasmids and strains obtained during the current study are available from the corresponding author on reasonable request.
Consent for publication
Ethics approval and consent to participate
Supported by a Grant from the Spanish Ministry of Economy and Competitiveness/FEDER—MINECO—to H.R. (SAF2015-73549-JIN) and R.M. (AGL2014-52911-R).
We thank the MINECO for the Severo Ochoa Excellence accreditation (SEV2016-0644), the Basque Department of Industry, Tourism and Trade (Etortek and Elkartek programs), the Innovation Technology Department of the Bizkaia Province and the CIBERehd network.
D.B. is the recipient of a fellowship for Formación de Personal Investigador from MINECO.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.
- Chung KT, Wong TY, Wei CI, Huang YW, Lin Y. Tannins and human health: a review. Crit Rev Food Sci Nutr. 1998;38:421–64.View ArticleGoogle Scholar
- Smeriglio A, Barreca D, Bellocco E, Trombetta D. Proanthocyanidins and hydrolysable tannins: occurrence, dietary intake and pharmacological effects. Br J Pharmacol. 2017;174:1244–62.View ArticleGoogle Scholar
- Aguilar CN, Rodriguez R, Gutierrez-Sanchez G, Augur C, Favela-Torres E, Prado-Barragan LA, Ramirez-Coronel A, Contreras-Esquivel JC. Microbial tannases: advances and perspectives. Appl Microbiol Biotechnol. 2007;76:47–59.View ArticleGoogle Scholar
- Chávez-González M, Rodríguez-Duránb LV, Balagurusamy N, Prado-Barragán A, Rodríguez R, Contreras JC, Aguilar CN. Biotechnological advances and challenges of tannase: an overview. Food Bioprocess Technol. 2012;5:445–9.View ArticleGoogle Scholar
- Chaitanyakumar A, Anbalagan M. Expression, purification and immobilization of tannase from Staphylococcus lugdunensis MTCC 3614. AMB Express. 2016;6:89.View ArticleGoogle Scholar
- Iwamoto K, Tsuruta H, Nishitaini Y, Osawa R. Identification and cloning of a gene encoding tannase (tannin acylhydrolase) from Lactobacillus plantarum ATCC 14917(T). Syst Appl Microbiol. 2008;31:269–77.View ArticleGoogle Scholar
- Jimenez N, Barcenilla JM, de Felipe FL, de las Rivas B, Munoz R. Characterization of a bacterial tannase from Streptococcus gallolyticus UCN34 suitable for tannin biodegradation. Appl Microbiol Biotechnol. 2014;98:6329–37.View ArticleGoogle Scholar
- Jimenez N, Esteban-Torres M, Mancheno JM, de las Rivas B, Munoz R. Tannin degradation by a novel tannase enzyme present in some Lactobacillus plantarum strains. Appl Environ Microbiol. 2014;80:2991–7.View ArticleGoogle Scholar
- Noguchi N, Ohashi T, Shiratori T, Narui K, Hagiwara T, Ko M, Watanabe K, Miyahara T, Taira S, Moriyasu F, Sasatsu M. Association of tannase-producing Staphylococcus lugdunensis with colon cancer and characterization of a novel tannase gene. J Gastroenterol. 2007;42:346–51.View ArticleGoogle Scholar
- Sharma KP, John PJ. Purification and characterization of tannase and tannase gene from Enterobacter sp. Process Biochem. 2011;46:240–4.View ArticleGoogle Scholar
- Wu M, Wang Q, McKinstry WJ, Ren B. Characterization of a tannin acyl hydrolase from Streptomyces sviceus with substrate preference for digalloyl ester bonds. Appl Microbiol Biotechnol. 2015;99:2663–72.View ArticleGoogle Scholar
- Noguchi N, Fukuzawa M, Wajima T, Yokose K, Suzuki M, Nakaminami H, Kawai T, Moriyasu F, Sasatsu M. Specific clones of Staphylococcus lugdunensis may be associated with colon carcinoma. J Infect Public Health. 2017;11(1):39–42.View ArticleGoogle Scholar
- Noguchi N, Goto K, Ro T, Narui K, Ko M, Nasu Y, Utsumi K, Takazawa K, Moriyasu F, Sasatsu M. Using the tannase gene to rapidly and simply identify Staphylococcus lugdunensis. Diagn Microbiol Infect Dis. 2010;66:120–3.View ArticleGoogle Scholar
- Jimenez N, Reveron I, Esteban-Torres M, de Felipe FL, de las Rivas B, Munoz R. Genetic and biochemical approaches towards unravelling the degradation of gallotannins by Streptococcus gallolyticus. Microb Cell Fact. 2014;13:154.View ArticleGoogle Scholar
- Curiel JA, Rodriguez H, Acebron I, Mancheno JM, de las Rivas B, Munoz R. Production and physicochemical properties of recombinant Lactobacillus plantarum tannase. J Agric Food Chem. 2009;57:6224–30.View ArticleGoogle Scholar
- Koppel N, Rekdal VM, Balskus EP. Chemical transformation of xenobiotics by the human gut microbiota. Science. 2017;356(6344):eaag2770.View ArticleGoogle Scholar
- Abdulamir AS, Hafidh RR, Abu Bakar F. The association of Streptococcus bovis/gallolyticus with colorectal tumors: the nature and the underlying mechanisms of its etiological role. J Exp Clin Cancer Res. 2011;30:11.View ArticleGoogle Scholar
- Abdulamir AS, Hafidh RR, Mahdi LK, Al-jeboori T, Abubaker F. Investigation into the controversial association of Streptococcus gallolyticus with colorectal cancer and adenoma. BMC Cancer. 2009;9:403.View ArticleGoogle Scholar
- Rusniok C, Couve E, Da Cunha V, El Gana R, Zidane N, Bouchier C, Poyart C, Leclercq R, Trieu-Cuot P, Glaser P. Genome sequence of Streptococcus gallolyticus: insights into its adaptation to the bovine rumen and its ability to cause endocarditis. J Bacteriol. 2010;192:2266–76.View ArticleGoogle Scholar
- Jimenez N, Santamaría L, Esteban-Torres M, De Las Rivas B, Muñoz R. Contribution of a tannase from Atopobium parvulum DSM 20469T in the oral processing of food tannins. Food Res Int. 2014;62:397–402.View ArticleGoogle Scholar
- Sanchez-Mangas R, Garcia-Ferrrer A, de Juan A, Arroyo AM. The probability of death in road traffic accidents. How important is a quick medical response? Accid Anal Prev. 2010;42:1048–56.View ArticleGoogle Scholar
- Ren B, Wu M, Wang Q, Peng X, Wen H, McKinstry WJ, Chen Q. Crystal structure of tannase from Lactobacillus plantarum. J Mol Biol. 2013;425:2737–51.View ArticleGoogle Scholar
- Zhao H, Chu M, Huang Z, Yang X, Ran S, Hu B, Zhang C, Liang J. Variations in oral microbiota associated with oral cancer. Sci Rep. 2017;7:11773.View ArticleGoogle Scholar
- Inoue KH, Hagerman AE. Determination of gallotannin with rhodanine. Anal Biochem. 1988;169:363–9.View ArticleGoogle Scholar
- Turner P, Mamo G, Karlsson EN. Potential and utilization of thermophiles and thermostable enzymes in biorefining. Microb Cell Fact. 2007;6:9.View ArticleGoogle Scholar
- Lekha PK, Lonsane BK. Production and application of tannin acyl hydrolase: state of the art. Adv Appl Microbiol. 1997;44:215–60.View ArticleGoogle Scholar
- Karpathy SE, Qin X, Gioia J, Jiang H, Liu Y, Petrosino JF, Yerrapragada S, Fox GE, Haake SK, Weinstock GM, Highlander SK. Genome sequence of Fusobacterium nucleatum subspecies polymorphum—a genetically tractable fusobacterium. PLoS ONE. 2007;2:e659.View ArticleGoogle Scholar
- Han YW, Wang X. Mobile microbiome: oral bacteria in extra-oral infections and inflammation. J Dent Res. 2013;92:485–91.View ArticleGoogle Scholar
- Lopez de Felipe F, de las Rivas B, Munoz R. Bioactive compounds produced by gut microbial tannase: implications for colorectal cancer development. Front Microbiol. 2014;5:684.Google Scholar
- Maniatis T, Fritsch EF, Sambrook J, editors. Molecular cloning: a laboratory manual. Cold Spring Harbor: Cold spring harbor laboratory press; 1989.Google Scholar
- Marchler-Bauer A, Derbyshire MK, Gonzales NR, Lu S, Chitsaz F, Geer LY, Geer RC, He J, Gwadz M, Hurwitz DI, et al. CDD: NCBI’s conserved domain database. Nucleic Acids Res. 2015;43:D222–6.View ArticleGoogle Scholar
- Sievers F, Wilm A, Dineen D, Gibson TJ, Karplus K, Li W, Lopez R, McWilliam H, Remmert M, Soding J, et al. Fast, scalable generation of high-quality protein multiple sequence alignments using Clustal Omega. Mol Syst Biol. 2011;7:539.View ArticleGoogle Scholar
- Dereeper A, Guignon V, Blanc G, Audic S, Buffet S, Chevenet F, Dufayard JF, Guindon S, Lefort V, Lescot M, et al. Phylogeny.fr: robust phylogenetic analysis for the non-specialist. Nucleic Acids Res. 2008;36:W465–9.View ArticleGoogle Scholar
- Castresana J. Selection of conserved blocks from multiple alignments for their use in phylogenetic analysis. Mol Biol Evol. 2000;17:540–52.View ArticleGoogle Scholar
- Anisimova M, Gascuel O. Approximate likelihood-ratio test for branches: a fast, accurate, and powerful alternative. Syst Biol. 2006;55:539–52.View ArticleGoogle Scholar
- Chevenet F, Brun C, Banuls AL, Jacq B, Christen R. TreeDyn: towards dynamic graphics and annotations for analyses of trees. BMC Bioinform. 2006;7:439.View ArticleGoogle Scholar
- Crooks GE, Hon G, Chandonia JM, Brenner SE. WebLogo: a sequence logo generator. Genome Res. 2004;14:1188–90.View ArticleGoogle Scholar
- Camacho C, Coulouris G, Avagyan V, Ma N, Papadopoulos J, Bealer K, Madden TL. BLAST+: architecture and applications. BMC Bioinform. 2009;10:421.View ArticleGoogle Scholar