Applied Microbiology and Biotechnology

, Volume 99, Issue 8, pp 3547–3558 | Cite as

Tyramine biosynthesis is transcriptionally induced at low pH and improves the fitness of Enterococcus faecalis in acidic environments

  • Marta Perez
  • Marina Calles-Enríquez
  • Ingolf Nes
  • Maria Cruz Martin
  • Maria Fernandez
  • Victor Ladero
  • Miguel A. Alvarez
Applied microbial and cell physiology

Abstract

Enterococcus faecalis is a commensal bacterium of the human gut that requires the ability to pass through the stomach and therefore cope with low pH. E. faecalis has also been identified as one of the major tyramine producers in fermented food products, where they also encounter acidic environments. In the present work, we have constructed a non-tyramine-producing mutant to study the role of the tyramine biosynthetic pathway, which converts tyrosine to tyramine via amino acid decarboxylation. Wild-type strain showed higher survival in a system that mimics gastrointestinal stress, indicating that the tyramine biosynthetic pathway has a role in acid resistance. Transcriptional analyses of the E. faecalis V583 tyrosine decarboxylase cluster showed that an acidic pH, together with substrate availability, induces its expression and therefore the production of tyramine. The protective role of the tyramine pathway under acidic conditions appears to be exerted through the maintenance of the cytosolic pH. Tyramine production should be considered important in the adaptability of E. faecalis to acidic environments, such as fermented dairy foods, and to survive passage through the human gastrointestinal tract.

Keywords

Enterococcus faecalis Tyramine tdc cluster expression Gastrointestinal stress Internal pH 

Introduction

The ability of Enterococcus faecalis to tolerate wide ranges of pH, temperature, and osmotic conditions allows it to colonize environments as different as water, soil, and foodstuffs especially fermented food products where it can be present in raw materials or contaminate them (Agudelo Higuita and Huycke 2014; Giraffa 2003; Lebreton et al. 2014). It is also a commensal of both human and animal gastrointestinal tracts (GITs). Some enterococcal strains, however, can also act as opportunistic pathogens, causing nosocomial infections such as endocarditis and bacteremia, usually following the colonization of the GIT (Agudelo Higuita and Huycke 2014; Paulsen et al. 2003; Ubeda et al. 2010). In fact, hospital-adapted, multiantibiotic-resistant enterococci have spread dramatically in recent decades; vancomycin-resistant (VRE) E. faecalis strains in particular can colonize healthy people and farm animals (Bonten et al. 2001), who along with certain foodstuffs (dairy and meat products) may act as VRE reservoirs (Giraffa 2003; Mathur and Singh 2005).

Little is known about the mechanisms used by VRE enterococci to colonize the human gut (Lebreton et al. 2014; Ubeda et al. 2010), although the intrinsic robustness of E. faecalis to different stresses may contribute towards its adaptability (Solheim et al. 2014). In lactic acid bacteria (LAB) and pathogens such as Listeria monocytogenes and Escherichia coli, amino acid decarboxylation is thought to provide an acid resistance system that helps them face the challenges of colonizing GIT environments (Castanie-Cornet and Foster 2001; Gahan and Hill 2014; Pessione 2012). Strains of enterococci of clinical, human, and food origin can all decarboxylate the amino acid tyrosine to produce tyramine; indeed, the biosynthesis of tyramine is a general species trait of E. faecalis (Ladero et al. 2012).

Tyramine is a biogenic amine (BA) that can accumulate in foodstuffs via the action of microbial decarboxylases (Linares et al. 2011). The consumption of large amounts can cause toxicological effects including migraines and hypertension and sometimes problems as serious as cerebral haemorrhages (EFSA 2011; Ladero et al. 2010a; Pessione 2012). These symptoms are together known as the ‘cheese effect’ (Ladero et al. 2010a) since tyramine is one of the most commonly found and abundant BA in dairy products (Fernandez et al. 2007a; Linares et al. 2011, 2012a). Enterococci are among the microorganisms responsible for tyramine biosynthesis in cheeses, constituting a serious food safety concern (Ladero et al. 2010b; Linares et al. 2012a). Tyramine is formed from tyrosine by the action of the enzyme tyrosine decarboxylase (TdcA). Tyramine is further secreted from the cell in exchange for tyrosine by the antiporter TyrP. The proteins involved in the tyramine pathway are encoded in the tdc cluster, which has been described in E. faecalis JH2-2 (Connil et al. 2002), Enterococcus durans IPLA655 (Ladero et al. 2013) and Enterococcus faecium RM58 (Marcobal et al. 2006a) among others. The tdc cluster has also been annotated in the genome sequence of other LAB (Linares et al. 2011), as well as in that of the clinically important VRE strain E. faecalis V583 (Paulsen et al. 2003). All the sequences share the same genetic organization, which comprises four genes (Fig. 1a): tyrS, an aminoacyl transfer RNA (tRNA) synthetase-like gene; tdcA, which encodes the tyrosine decarboxylase; tyrP, which codes for the tyrosine/tyramine exchanger; and nhaC-2, which encodes an Na+/H+ antiporter, the involvement of which in the biosynthesis of tyramine remains unknown (Linares et al. 2011; Lucas et al. 2003).
Fig. 1

Genetic organization and transcriptional analysis of the tdc cluster of E. faecalis V583. a Diagram showing the genetic organization of the tdc cluster and flanking regions. Putative promoters are indicated by broken arrows, and secondary structures and transcription termination regions by lollipops. RT-PCR-targeted intergenic regions and expected mRNA are indicated. b RT-PCR amplification of the intergenic regions: fragment A (ef0632-tyrS), fragment B (tyrS-tdcA), fragment C (tdcA-tyrP), fragment D (tyrP-nhaC-2) and fragment E (nhaC-2-ef0636). Negative controls (minus sign) were performed without reverse transcriptase and positive controls (plus sign) with chromosomal DNA. M molecular weight markers (GeneRuler DNA ladder mix, Fermentas)

Tyramine production in food-borne E. durans and E. faecium strains has been related to tolerance to low pH. The coupled reactions of decarboxylation and tyrosine/tyramine exchange have been proposed as a mechanism for adapting to acidic environments, as well as an indirect way of obtaining metabolic energy via proton motive force generation (Fernandez et al. 2007b; Marcobal et al. 2006b; Pereira et al. 2009). The possible roles of tyramine production in GIT resistance, immunomodulation and the adhesion of pathogens to enterocytes have all been examined (Fernandez de Palencia et al. 2011; Lyte 2004; Pereira et al. 2009). However, little is known about the regulation and physiological role of the tyramine production pathway in E. faecalis.

In this work, a tdc knockout mutant was constructed in order to characterize the tdc cluster of the tyramine-producing strain E. faecalis V583. A transcriptional study under different environmental conditions was performed, and the physiological role of tyramine production under stress conditions, including those encountered in GIT passage, was examined. Tyramine production via tyrosine decarboxylation is here suggested to provide a cytosolic pH maintenance mechanism that helps cope with acid stress.

Materials and methods

Strains, media and growth conditions

E. coli Gene-Hogs (Invitrogen, Paisley, UK) was used as an intermediate host for the pAS222 cloning vector (Jonsson et al. 2009) and derived plasmid (pAS222 TDC, this work). The strain was cultured at 37 °C with aeration in Luria-Bertani medium (Green and Sambrook 2012) supplemented with 100 mg mL−1 of ampicillin (USB Corporation, Cleveland, OH, USA) when necessary.

The wild-type E. faecalis V583 strain (hereafter referred to as ‘wt’) was used as a model strain since its genomic sequence was the first to become available for an E. faecalis strain, and it is deposited in the American Type Culture Collection under the accession number ATCC 700802. The wt and the derived mutant E. faecalis V583 Δtdc (hereafter referred to as ‘Δtdc’) were grown routinely in M17 medium (Oxoid, Hampshire, UK) supplemented with 5 g L−1 glucose (Merck, Darmstadt, Germany) (GM17) at 37 °C under aerobic conditions with an initial inoculum of 0.1 %.

When indicated, 10 mM tyrosine (Sigma-Aldrich, St. Louis, MO, USA) was added (GM17 + T). The latter medium was used to study the factors that affect the growth of the wt and Δtdc strains by reducing the sugar concentration to 1 g L−1 glucose and/or the pH to 5 (initially pH 6.8) as indicated. Tyrosine consumption and tyramine production were checked after 12 h of growth

To test the effect of tyrosine concentration on gene expression, wt cells were grown in 50 mL of chemically defined medium (CDM) (Poolman and Konings 1988) supplemented with different tyrosine concentrations at 37 °C for 4 h. To measure gene expression and tyramine production under controlled pH conditions, the wt strain was cultivated in a Six-Fors bioreactor (Infors AG, Bottmingen, Switzerland) in GM17 supplemented with tyrosine at a non-limiting concentration (15 mM, GM17 + T15) for 6 h. The reactor was maintained at 37 °C, 50-rpm stirring and with zero air input. The pH was maintained by automatically adding 2 N NaOH or 5 N HCl as needed. All data are the means for at least three cultures independently grown under each condition.

DNA isolation

Total DNA was extracted from 2 mL of an overnight culture using the GenElute™ Bacterial Genomic DNA Kit (Sigma-Aldrich), following the manufacturer’s instructions. Plasmid extraction was performed following standard procedures (Green and Sambrook 2012).

PCR amplification and sequencing

PCR amplifications were performed in 25-μL reaction volumes with 1 μL of DNA as a template (typically 200 ng), 400 nM of each primer, 200 μM of dNTP (GE Healthcare, Little Chalfont, UK), the reaction buffer and 1 U of Taq polymerase (Phusion High-Fidelity DNA Polymerase, Thermo Scientific, Madrid, Spain). All primers (Table 1) were designed based on the E. faecalis V583 genome sequence (GenBank accession number: AE016830) and synthesized by Macrogen (Seoul, Korea). Amplifications were performed using a MyCycler apparatus (Bio-Rad, Hercules, CA, USA) under the following conditions: 94 °C for 5 min, 35 cycles of 94 °C for 30 s, 55 °C for 45 s, 72 °C for 1 min and a final extension step at 72 °C for 5 min. The amplifications were analyzed by agarose gel electrophoresis; purification, when needed, was performed using the GenElute PCR Clean-Up Kit (Sigma-Aldrich). Sequencing of the PCR fragments was performed at Macrogen.
Table 1

Primers used in this study

Primers

Function

Sequence (5′ to 3′)

Reference

cardiolRT F

ef0631-tyrS RT-PCR

CTCCAGAAGTTGTTCGCGACAT

This work

tyrSRT R

ef0631-tyrS RT-PCR

CTGTAAGTTCTCTTAGTCCTTC

This work

tyrS3 F

tyrS-tdcA RT-PCR

TGCAGTCGATCCAACACAACATT

This work

tyrS4 R

tyrS-tdcA RT-PCR

TTGTAGCTCATTAAGTGAGCAAATTCATG

This work

tdcART F

tdcA-tyrP RT-PCR

GAATGGAACCGTGCAGGTAAAG

This work

tyrPRT R

tdcA-tyrP RT-PCR

GTTGAGGGCCACCTTCTTGAGGAAG

This work

tyrPRT 2 F

tyrP-nhaC-2 RT-PCR

GTGACTGATGCAGTCTTAGTTGC

This work

nhaC2RT R

tyrP-nhaC-2 RT-PCR

CTGTCATCGCATTGTCGAATCC

This work

nhaC2RT F

nhaC-2-ef0637 RT-PCR

CCCATTGCTTTGTCCCATTATCACCG

This work

ef0637 R

nhaC-2-ef0637 RT-PCR and tdc deletion check

GATCCGCTTGTGAAGTTGTCGCTGCAG

Ladero et al. (2012)

tdcV583q F

tdcA expression analysis

CTGCTGATATTATCGGTATCGGTT

This work

tdcV583q R

tdcA expression analysis

GTAGTTATGGTCAACTGGTACTGGG

This work

tyrSq F

tyrS expression analysis

AAACGTGAAGCACAAAGACGCT

This work

tyrSq R

tyrS expression analysis

TTTTGCGCTTCTTCTAATGCTG

This work

recA F

recA internal control

CAAGGCTTAGAGATTGCCGATG

This work

recA R

recA internal control

ACGAGGAACTAACGCAGCAAC

This work

EFV583-tuf F

tuf internal control

CAGGACATGCGGACTACGTTAA

This work

EFV583-tuf R

tuf internal control

TAGGACCATCAGCAGCAGAAAC

This work

T1 F

Δtdc mutant construction

TCGATCCAACTGGAGATAGCATGCATA

This work

T2 R

Δtdc mutant construction

AGTATTTGATGACATCACGATCAT

This work

T3 F

Δtdc mutant construction

AACAATGTAATCGGTGAAATCCAGAATCCTAGGATTCTGGATTTCACCGATTACATTGTT

This work

T4 R

Δtdc mutant construction

TGACGGTGATAATGGGACAAAGCAAT

This work

Card F

tdc deletion check

GATGATAGTGTCTTGGCTGCTTTAAAGG

Ladero et al. (2012)

F forward, R reverse

Construction of the E. faecalis tdc knockout mutant

An E. faecalis V583 non-tyramine-producing mutant, i.e. with a tdc cluster deletion from tyrS (793 nt from its start codon) to nhaC-2 (691 nt from its start codon), was achieved by double-crossover homologous recombination with the cloning vector pAS222 following a previously described protocol (Jonsson et al. 2009). Briefly, the flanking fragments of the tdc cluster were amplified by splicing by overlap extension PCR (Horton et al. 1989) and two PCR reactions performed with primers T1 F, T2 R and T3 F, T4 R (Table 1). The amplicons were purified, and a mix used as a template for PCR amplification with the outer primers T1 F and T4 R. The inner primer carrying regions of homology for the fusion step was T3 F (Table 1). The PCR product was cloned into the SnaBI (Fermentas, Vilnius, Lithuania) site of pAS222 to generate pAS222 TDC, which was propagated in E. coli Gene-Hogs cells. pAS222 TDC was transformed into electrocompetent E. faecalis V583 cells obtained following a previously described protocol (Holo and Nes 1989) using 4 % glycine in the growth medium. E. faecalis V583 cells harbouring pAS222 TDC were grown in GM17 under previously described conditions (Biswas et al. 1993) in order to select bacteria showing evidence of double-crossover events. The deletion of tdc was checked by PCR amplification and further sequencing at Macrogen, using card F and ef0637 R primers (Table 1). The absence of tyramine biosynthesis was checked in the supernatant of overnight cultures in GM17 + T as described below. A positive deletion mutant (E. faecalis V583 Δtdc) was confirmed by both methods and selected for further analysis.

RNA isolation

E. faecalis cells were grown in the required medium for each experiment, as previously indicated. Adequate culture volumes (adjusted to a cell density of approximately optical density (OD)600 = 2) were harvested by centrifugation in a refrigerated benchtop microcentrifuge (Eppendorf, Hamburg, Germany) running at maximum speed. Total RNA was extracted using TRI reagent (Sigma-Aldrich) as previously described (Linares et al. 2009). To eliminate any DNA contamination, 2 μg of total RNA samples were treated with 2 U of DNAse I (Fermentas) for 2 h. Control PCR to ensure that no contaminant DNA remains was performed using specific primers to amplify recombinase A (recA). The total RNA concentration was determined in an Epoch Microplate Spectrophotometer (BioTek, Winooski, VT, USA).

Reverse transcription PCR (RT-PCR)

Total complementary DNA (cDNA) was synthesized from 0.5 μg of RNA using the reverse transcription (RT) iScript™ cDNA Synthesis kit (Bio-Rad) and 1 μL used as a template for PCR reactions involving 400 nM of each primer (Table 1), 200 μM of dNTP, the reaction buffer and 1 U of Taq polymerase (DreamTaq, Fermentas). Five pairs of primers (Table 1) were used to amplify regions spanning the gene junctions.

Gene expression quantification by RT-qPCR

Gene expression analysis was performed by reverse transcription–quantitative real-time PCR (RT-qPCR) in a 7500 Fast Real-Time PCR System (Applied Biosystems, Carlsbad, CA, USA) using SYBR Green PCR Master Mix (Applied Biosystems). Fourfold dilutions of the cDNA samples were used as a template (4 μL) with 700 nM of each primer and SYBR Green PCR Master Mix in a 20-μL final volume. Amplifications were performed with specific primers (Table 1) based on internal sequences of the tyrS and tdcA genes designed using Primer Express software (Applied Biosystems). Specific primers for recA and elongation factor thermo-unstable (tufA) genes were used as internal controls to normalize the RNA concentration. The linearity and amplification efficiency of the reactions were tested for each primer pair using six 10-fold serial dilutions of total E. faecalis V583 DNA. A positive control with total E. faecalis V583 DNA was included for each run, and the resulting melting curves for the samples were compared with that of this positive control. A negative control with all the reaction components except cDNA was included. Amplifications were performed using the default cycling settings suggested by Applied Biosystems. The abundance of messenger RNA (mRNA) species was calculated following the 2-ΔΔCT method described by Livak and Schmittgen (2001). The condition with the lowest level of expression was selected as the calibrator for all experiments. RT-qPCR analysis was performed on RNA purified from at least three independent cultures for each condition.

Determination of tyramine biosynthesis

Medium supernatants were recovered from centrifuged cultures from which RNA was obtained and filtered through 0.45-μm polytetrafluoroethylene (PTFE) filters (VWR, Barcelona, Spain) for tyrosine and tyramine quantification by ultra-high-performance liquid chromatography (UHPLC). The filtered supernatants were derivatized with diethyl ethoxymethylenemalonate (Sigma-Aldrich) and further separated in a UPLC® system (Waters, Milford, MA, USA) using previously described column, solvent and gradient conditions (Redruello et al. 2013). Data were acquired and analyzed using Empower 2 software (Waters). The tyrosine and tyramine concentrations provided are the average of at least three independent cultures.

Gastrointestinal transit tolerance assay

Simulation of the digestion conditions influencing the survival of the microorganisms during their transit through the human GIT was performed as previously described (Fernández de Palencia et al. 2008) with the following modifications. Cells of the wt and Δtdc strains from late exponential phase cultures in GM17 + T (approximately 1010 colony-forming units (cfu) mL−1) were harvested and resuspended in the electrolyte solution supplemented with 10 mM tyrosine. After cell exposure to lysozyme, gastric (G) stress conditions were mimicked by treating cells with pepsin and a successively decreasing pH. Gastrointestinal (GI) stress analysis was simulated by exposure of the samples incubated at pH 5, 4.1 and 3 to bile salts and pancreatin at pH 8. Finally, to mimic colonic stress (Van den Abbeele et al. 2010), the GI pH 3 sample was adjusted to pH 7 and incubated overnight. Cell viability under each set of conditions was determined using the LIVE/DEAD® BacLight™ fluorescent stain (Molecular Probes, Leiden, the Netherlands) adhering to previously described conditions (Fernández de Palencia et al. 2008). The correlation between the green (live)/red (dead) bacteria fluorescent ratio (G/R) and viable cell numbers was previously established by plate counting. The values presented are the mean of three replicates from independent cultures, expressed as a percentage of the untreated control. Tyramine accumulation was also quantified by UHPLC as described above.

Measurement of intracellular pH

Cytosolic pH measurements were performed using carboxyfluorescein succinimidyl ester (cFSE, Sigma-Aldrich) (an internally conjugated fluorescence pH probe) following a previously described protocol (Sanchez et al. 2006) with slight modifications. The wt and Δtdc strains were grown in GM17 + T for 6 h. After collecting cells from 1 mL of culture and washing in creatine phosphokinase (CPK) buffer (sodium citrate 50 mM, disodium phosphate 50 mM, potassium chloride 50 mM) at pH 7.0, they were resuspended in 1 mL of CPK buffer adjusted to different pH values and incubated at 30 °C for 30 min in the presence of the cFSE probe. They were then washed again in CPK buffer and resuspended in 1 mL of the same plus 15 mM glucose at the pH required and maintained for 15 min at 30 °C. The cells were then washed once again in CPK buffer at the pH required and resuspended in 100 μL of the same, also at the required pH. Finally, 100 μL of CPK buffer supplemented with 5 mM tyrosine (final concentration 2.5 mM) was added to the treated cells, and 100 μL of CPK buffer without tyrosine to the control cells. Fluorescence intensities were measured for 10 min (intervals of 0.25 s) in a Cary Eclipse fluorescence spectrophotometer (Varian Inc., Palo Alto, CA, USA) with the excitation and emission values indicated by Breeuwer et al. (1996). Background fluorescence levels were assessed by measuring non-fluorescent control cells; these values were subtracted from the fluorescence results. The cytosolic pH values were determined from the ratio of the fluorescence signal at 440/490 nm taken from a calibration curve constructed using buffers at pH 4.5–8.0, after equilibrating the internal (pHin) and external (pHout) pH with 0.1 % Triton (Molenaar et al. 1991). The value given for each condition is the average of three independent replicates (each the mean of values obtained over 8 min of monitoring).

Statistical analysis

Means ± standard deviations were calculated from at least three independent replicates as indicated. Means were compared by the Student t test or ANOVA and the Tukey post hoc test when indicated. Significance was set at p < 0.05.

Results

Physiological role of the tdc cluster in E. faecalis

To study the physiological role of tyramine production in E. faecalis, a deletion mutant of the tdc cluster was obtained as indicated above. One clone—termed E. faecalis Δtdc—was selected after checking for the deletion of the cluster by PCR using primers card F and ef0637 R (Table 1). Analysis by UHPLC of the supernatants from overnight cultures of Δtdc in GM17 + T showed it to be unable to produce tyramine (data not shown).

To determine whether tyramine biosynthesis offers some advantage in terms of the growth of E. faecalis V583, the OD600 of wt and Δtdc cultures in GM17 + T was monitored (Fig. 2a). No differences were seen between the growth of both strains in these conditions. The influence of tyramine production was also examined under the stress condition of limited carbon source, monitoring the growth of the wt and Δtdc strains in M17 + T with 1 g L−1 glucose. Although both strains showed a reduction in the maximum OD600 reached (Fig. 2b) when grown with 5 g L−1 glucose (Fig. 2a), no difference was detected between the wt and Δtdc strains. The significance of tyrosine decarboxylation under acid stress conditions was studied by comparing the growth of the wt and Δtdc strains, adjusting the initial pH of the GM17 + T medium to pH 5.0. Figure 2c shows that the tyramine-producing wt strain achieved a higher OD600 than the Δtdc strain (1.5 vs 0.9) and showed a steeper exponential phase slope. Finally, an experiment combining a reduced carbon source and an acidic initial pH was performed. Cells were cultured in M17 + T with 1 g L−1 glucose, adjusted to an initial pH of 5.0. The OD600 values recorded (Fig. 2d) were slightly lower than those obtained under acidic pH conditions (Fig. 2c). The OD600 returned by wt was twice that of Δtdc (1.2 vs 0.6), showing that tyrosine decarboxylation enabled the wt strain to grow more quickly.
Fig. 2

Influence of different factors on the growth of E. faecalis V583 (continuous line) and E. faecalis V583 Δtdc (discontinuous line), in the presence of 10 mM tyrosine. a Effect of tyramine biosynthesis on cells grown in GM17 + T. b Influence of carbon source depletion in cultures propagated in M17 + T supplemented with glucose 1 g L−1. c Effect of acidic pH on cells cultured in GM17 + T adjusted to an initial pH of 5.0. d Influence of carbon source depletion and acidic pH on cultures grown in M17 + T with glucose 1 g L−1 and an initial pH adjusted to 5.0. The growth curves were monitored over 12 h by measurement of the OD600

These results suggest that tyramine biosynthesis might play an important role in E. faecalis acid resistance by improving cell growth under acidic conditions, such as those encountered in GIT environments.

The tyrosine decarboxylation pathway improves survival under highly acidic gastric conditions

A gastric and gastrointestinal tolerance assay was performed for the wt and Δtdc strains in the presence of tyrosine. Analysis of tyramine production in the wt strain (Fig. 3) showed that it was able to produce tyramine under all the conditions assayed, with stronger production under the more acidic gastric conditions (pH 3.0, 2.1 and 1.8).
Fig. 3

Response of E. faecalis V583 and E. faecalis V583 Δtdc in the gastrointestinal tolerance assay. Survival (%) of the wt (grey bars) and Δtdc (white bars) strains after gastric (G), gastrointestinal (GI) and colonic stresses in the presence of 10 mM tyrosine. C untreated cells (control). Survival was measured using the LIVE/DEAD® BacLight™ fluorescent stain. Values are expressed as a percentage of the control value (the 100 % control values of the G/R ratio of untreated wt and Δtdc strains were, respectively, 6.9 and 6.8, corresponding to 7.6 × 1010 and 7.5 × 1010 cfu mL−1). Cells from cultures propagated with 10 mM tyrosine for 6 h were used. The asterisk indicates statistically significant difference (p < 0.05; Student t test). The tyramine produced by the wt strain under each set of conditions is indicated

The viability of wt and Δtdc cells under gastrointestinal stress was assessed using the LIVE/DEAD® BacLight™ fluorescent stain. Under G stress, the wt strain showed reduced viability (of around 10 %) at pH 3.0, 2.1 and 1.8 compared to the untreated controls; at these pH values, greater tyramine production was detected (Fig. 3). The Δtdc cells showed reduced viability under all the conditions assayed, significantly so at pH 2.1 and 1.8 (p < 0.05), at which approximately only 65 % of the cells survived. The conditions under which tyramine production by strain wt was highest were those under which the survival of the Δtdc mutant strain was poorest. Under GI and colonic stress conditions (exposure to proteolytic enzymes and bile salts), the survival of both populations was reduced to around 15 %, with no difference observed between the strains, even though wt was still able to produce tyramine.

These results show that E. faecalis is probably able to survive GIT passage and that tyramine biosynthesis, which has been shown to take place under these conditions, enhances cell survival (especially under G stress). Therefore, tyramine production may improve the fitness of E. faecalis under acidic conditions, potentially contributing towards in situ tyramine production and accumulation in the GIT. The influence of pH and tyrosine concentration on the regulation of tdc cluster transcription was therefore examined.

The catabolic genes tdcA, tyrP and nhaC-2 are co-transcribed as a polycistronic mRNA

Before starting the transcriptional analysis of factors affecting tdc cluster expression, its transcriptional organization in E. faecalis V583 was examined. To determine whether the tdc cluster genes are co-transcribed, cDNA from total RNA of cultures grown in GM17 + T was used in RT-PCR amplifications with five sets of primers (Table 1) designed to amplify the intergenic and flanking regions of the tdc cluster (Fig. 1a). As expected, PCR products were obtained in RT-PCR amplifications neither of tyrS and ef0632 nor of the nhaC-2 and ef0636 intergenic regions (Fig. 1b) since these do not belong to the tdc cluster. Two amplification products were obtained (Fig. 1b), showing that tdcA, tyrP and nhaC-2 are co-transcribed. No amplification was obtained for the tyrS and tdcA intergenic region, indicating that although tyrS belongs to the tdc cluster, it is not included in the catabolic operon. mRNA covering tdcA, tyrP and nhaC-2 seemed to run from the putative tdcA promoter to the putative rho-independent terminator hairpin downstream of nhaC-2 (ΔG = −11.5 kcal) (Fig. S1). As indicated by the RT-PCR results, tyrS mRNA is individually transcribed in a monocistronic mRNA covering its own promoter to its putative rho-independent terminator hairpin (ΔG = −21.3 kcal) (Fig. S1).

tyrS expression is repressed by high tyrosine concentrations

Initially, the influence of the amino acid substrate on tyrS expression was evaluated in CDM at different tyrosine concentrations (Fig. 4a). The highest concentration of tyrosine assayed was 5 mM; higher concentrations resulted in its precipitation. tyrS expression was quantified by RT-qPCR after 4 h of incubation. As shown in Fig. 4a, tyrS was maximally transcribed in the absence of tyrosine (an inverse correlation was seen with tyrosine concentration). The expression of tyrS diminished progressively with the tyrosine concentration; at 5 mM tyrosine, minimum induction was observed. Analysis of the sequence upstream of tyrS showed strong homology with the structural features described for the tyrS leader region in E. durans (Linares et al. 2012b) (Fig. S1). These results suggest that, in E. faecalis, a similar transcription antitermination mechanism mediated by tyrosine regulates tyrS transcriptional repression in the presence of tyrosine.
Fig. 4

Quantification of gene expression measured by RT-qPCR, and tyramine production quantified by UHPLC. a Effect of different tyrosine concentrations on the expression of tyrS (white bars) and tdcA (grey bars), and on tyramine production, in E. faecalis V583 grown in CDM supplemented with 0, 0.1, 0.25, 0.5, 1, 1.5, 2.5 and 5 mM tyrosine. The lowest expression level for each gene was normalized to 1 and used as the reference condition. Bars with the same letter indicate statistically significant differences in relative expression with respect to the no-tyrosine condition (ANOVA and the Tukey post hoc test). b Influence of pH (5.0 vs 7.0) on tdcA transcription (grey bar) in E. faecalis V583 grown in GM17 + T15 for 6 h. The expression at pH 7.0 was normalized to 1 and used as the reference condition. The asterisk indicates statistically significant difference in relative induction (p < 0.01). a and b indicate statistically significant differences (p < 0.05) in tyramine production and OD600, respectively (Student t test)

The expression of tdcA is enhanced by tyrosine

Since the tdc catabolic genes of E. faecalis V583 are co-transcribed in a polycistronic mRNA, only the expression of tdcA was studied. The same cDNA samples obtained for the aforementioned tyrS expression assay following 4 h of incubation with different tyrosine concentrations were used to quantify tdcA expression. In contrast to that seen for tyrS, tdcA expression correlated positively with the tyrosine concentration until 0.5 mM tyrosine (Fig. 4a), after which no further induction was observed. At the same time point (after 4 h of incubation), tyramine production measured by UHPLC showed an increase as the tyrosine concentration increased (Fig. 4a). The concentrations of tyramine produced indicate that E. faecalis decarboxylates tyrosine efficiently, even at low concentrations of the substrate. This, plus the aforementioned result indicating tyrS to be maximally transcribed in the absence of tyrosine, meant that only the expression of tdcA under tyramine production conditions (substrate availability) was further studied.

Acidic pH increases tdcA expression and tyramine production

Results obtained by RT-qPCR analysis of tdcA expression (Fig. 4b) showed an approximate 10-fold upregulation in the culture at pH 5.0 compared to that at pH 7.0 (p < 0.01). Accordingly, tyramine production also reached its maximum under the acidic condition: 8.37 versus 4.38 mM at pH 7.0 (p < 0.05). It is noteworthy that while the OD600 achieved at pH 7.0 was 2.04, the culture grown at pH 5.0 only reached an OD600 of 0.55 (p < 0.05). These results highlight how an acidic pH can induce tdcA expression and tyramine biosynthesis in E. faecalis.

Tyramine biosynthesis counteracts acidification of the cytosol in acidic environments

Although the mechanism underlying the resistance to acid conferred by the production of tyramine remains unclear, several authors have indicated connections between decarboxylation reactions and the maintenance of pH homeostasis in acidic environments (Pereira et al. 2009; Romano et al. 2014). To confirm the function of tyrosine decarboxylation as a mechanism for neutralizing acidic conditions, cytosolic pH changes were monitored in the wt and Δtdc strains at different pH (from 7.0 to 4.5) in the absence/presence of tyrosine (2.5 mM). Figure 5a indicates that the wt strain was able to maintain a neutral intracellular pH even at the lowest pH tested (pHout 4.5) when in the presence of tyrosine. Compared to the control (tyrosine absence), the difference between the internal pH of the cells in the presence of tyrosine increased as the extracellular pH fell. This difference was significant even at pHout 7.0. In contrast, the cytosolic pH of the Δtdc strain fell as the extracellular pH decreased, both in the presence and absence of tyrosine (Fig. 5b), until eventually dropping below neutral at the lowest pH tested. These results indicate that the effect of the E. faecalis tdc cluster on pH homeostasis is greater at lower extracellular pH values and that the production of tyramine counteracts the intracellular acidification produced by acidic pH challenge.
Fig. 5

Variation in the intracellular pH (pHin) at different extracellular pH (pHout) (7, 6.5, 6, 5.5, 5 and 4.5) measured using a cFSE probe in resting cells of aE. faecalis V583 (continuous line) and bE. faecalis V583 Δtdc (discontinuous line), in the presence (black circles; 2.5 mM tyrosine) and absence (white circles; control condition) of tyrosine. Cells from cultures propagated with 10 mM tyrosine for 6 h were used. Asterisks indicate statistically significant differences (*p < 0.05; **p < 0.01; Student t test)

Discussion

Enterococci are LAB highly adapted to the GIT of human and animals, and it is also an important member of fermented food microbiota. Although usually commensals, they have emerged as a cause of multidrug-resistant, nosocomial infections. Indeed, those caused by VRE can be severe (Lebreton et al. 2014). Colonization of the GIT by VRE has been indicated to significantly increase the risk of suffering a systemic enterococal infection (Ubeda et al. 2010). Understanding colonization of both commensal and opportunistic pathogen enterococci requires a better knowledge of the mechanisms by which these bacteria cope with the acidic environment of the stomach. The decarboxylation of amino acids has been indicated as a mechanism by which LAB and human pathogenic bacteria can resist acidic conditions (Lund et al. 2014; Romano et al. 2014). Enterococci, such as E. faecalis, E. faecium and E. durans, have been shown to decarboxylate tyrosine to form tyramine, a toxic BA that can accumulate in food (Ladero et al. 2012)—specially in cheese where enterococci are one of the bacteria mainly responsible for tyramine accumulation (Ladero et al. 2010b). In fact, the capability to decarboxylate tyramine could be an advantage for the microorganism against acidification during the fermentation process. Therefore, the present work examined the role of tyramine production by the strain E. faecalis V583 as a means of resisting the effects of acid during GIT passage. The influence of environmental factors in the transcriptional regulation of tyramine production was tested, and evidence is provided that the tyramine biosynthetic pathway confers acid resistance by maintaining the intracellular pH stable.

The physiological significance of tyramine production—which remains under discussion—was studied by constructing a knockout deletion mutant of the tdc cluster of E. faecalis V583. This mutant was unable to produce tyramine, confirming the involvement of the tdc cluster in tyramine biosynthesis. The comparison of the growth fitness of wt and the non-tyramine-producing Δtdc in the presence of tyrosine and under different stress conditions (carbon source limitation and/or acidic pH) showed that tyramine production improved cell growth under acidic conditions. This indicates that tyramine biosynthesis may help counteract acid stress (Fig. 2c, d). No significant advantage was observed for either strain under conditions of sugar restriction (Fig. 2b). Previous comparative proteomic studies of E. faecalis suggest that tyrosine decarboxylation does not compete with other energy-supplying routes (Pessione et al. 2009). The present results are therefore consistent with studies that suggest that amino acid decarboxylation affords a means of counteracting acid stress (Pereira et al. 2009; Trip et al. 2012) rather than it being a mechanism for obtaining energy.

Tyramine biosynthesis in E. faecalis might then be considered an acid resistance mechanism that improves cell growth under acidic conditions. Microbes face the challenge of harsh acidic conditions in the GIT environment, and amino acid decarboxylation might play a role in their survival. The analysis of E. faecalis survival in an in vitro gastrointestinal model, and the production of tyramine under such conditions, was therefore tested. The results (Fig. 3) reveal that E. faecalis V583, like E. durans and Lactobacillus brevis strains (Fernandez de Palencia et al. 2011; Russo et al. 2012), is able to produce tyramine when exposed to GI stress. Whereas some 50 % of E. durans populations survive under G stress at pH 3.0 (Fernandez de Palencia et al. 2011), 85 % of the present E. faecalis population survived. Similarly, when faced with highly acidic gastric conditions (pH 2.1 and pH 1.8), the survival of the wt and Δtdc strains showed E. faecalis resistance to be enhanced by the presence of a functional tyramine biosynthetic pathway. This agrees with the finding that the tyramine producer E. faecium E17 conserves 91 % of its viability in a medium buffered at pH 2.5 in the presence of tyrosine (Pereira et al. 2009). The resistance to acidic conditions improved by the tyramine pathway might explain why E. faecalis, followed by E. faecium, is likely the dominant enterococcus in the human GIT (Nes et al. 2014). Altogether, these findings indicate that tyramine production should be considered an important characteristic that contributes to the colonization of the human GIT by opportunistic enterococci.

Since tyrosine decarboxylation improved E. faecalis fitness under acidic conditions, the effect of medium pH and substrate availability on the regulation of the tdc cluster transcription was examined. Different transcriptional organizations of the tdc cluster have been found in different strains. In E. durans IPLA655, tdcA and tyrP are elements of a single operon, while tyrS is transcribed independently (Linares et al. 2009). However, in E. faecalis JH2-2, the existence of a polycistronic mRNA covering tyrStdcAtyrP has been described (Connil et al. 2002). Similarly, in L. brevis IOEB 9890, a polycistronic mRNA covering tyrS, tdcA, tyrP and nhaC-2 has been indicated (Lucas et al. 2003). The present findings in E. faecalis V583 reveal a monocistronic mRNA covering tyrS and a polycistronic mRNA covering the operon formed by tdcAtyrPnhaC-2 (Fig. 1a). The relative high abundance of the transcript tdcAtyrP compared to the transcript tyrPnhaC-2 indicated that tdcA and tyrP genes could be expressed from both a short (tdcAtyrP) and a long mRNA (tdcAtyrPnhaC-2), as the transcriptional analysis of L. brevis IOEB 9890 tdc cluster has been suggested (Lucas et al. 2003). A potential weakest transcriptional terminator was found in the corresponding intergenic region (Figs. 1a and S1) supporting this possibility. Thus, the expression analysis of each transcript—tyrS and tdcAtyrPnhaC-2—was performed separately by RT-qPCR. The expression of tyrS, which encodes a tyrosyl-tRNA synthetase-like enzyme, under different tyrosine concentrations revealed an inverse correlation between tyrS transcription level and tyrosine concentration, with the maximum expression seen in the absence of tyrosine (Fig. 4a). This agrees with other results published by our group (Linares et al. 2012b) that indicate E. durans tyrS to be repressed by tyrosine concentration. tRNA synthetase genes are strictly regulated—via a termination–antitermination system—by the corresponding amino acid. If its concentration is low, it does not become bound to the tRNA, thus ensuring amino acid availability to protein synthesis and growth. In the present work, the tyrS upstream region of E. faecalis showed the typical structural motifs (Fig. S1) of a transcription antitermination system involving tyrosine (Grundy et al. 2002; Linares et al. 2012b), suggesting that a similar mechanism may be involved in the regulation of tyrS expression in this species. Tyrosine is a substrate amino acid for protein biosynthesis, and tyrS could be a sensor of the intracellular tyrosine pool for use in the regulation of tyrosine decarboxylation (Fernandez et al. 2004; Linares et al. 2012b); the role of tyrS in the regulation of the tyramine operon, however, is unclear.

Several authors have shown that decarboxylation reactions depend on amino acid substrates being available (Calles-Enriquez et al. 2010; Coton et al. 2011; Linares et al. 2009). The effect of increasing the concentration of tyrosine on the tdcA expression profile was analyzed in E. faecalis V583 and showed it to be transcriptionally upregulated in response. An increase in tyramine production was therefore observed (Fig. 4a). This regulation by tyrosine has also been seen in the tyramine-producing E. durans IPLA655 and Sporolactobacillus sp. 3PJ strains (Coton et al. 2011; Linares et al. 2012b). However, the relative induction levels observed were very low, and saturation in the expression was observed at tyrosine concentrations above 0.5 mM. Nevertheless, tdcA upregulation was very sensitive since even very low tyrosine levels (0.1 mM) were enough to increase it. Thus, the cells are able to decarboxylate tyrosine not only when it is in excess, as proposed in order to ensure protein biosynthesis (Linares et al. 2012b), but also when it is at low substrate concentrations. The fact that E. faecalis is not auxotrophic for tyrosine (it grew in CDM in the absence of tyrosine) might explain the functionality of the tdc operon even at low tyrosine concentrations.

The crucial induction factor in tyramine biosynthesis seems to be an acidic pH (Fernandez et al. 2007b; Linares et al. 2009; Marcobal et al. 2006b). The present results show a critical effect of low pH on the induction of tdcA and tyramine production in E. faecalis (Fig. 4b), confirming it to be a key factor in tyramine biosynthesis. The mechanism by which tyrosine decarboxylation exerts its role in acidic resistance remains unclear. Consistent with previous results in E. faecium (Pereira et al. 2009), the present data reveal tyramine production able to neutralize any acidification of the intracellular pH, the extent of tyrosine decarboxylation depending on external pH (Fig. 5a, b). It is noteworthy that, in the absence of tyrosine, the wt and Δtdc strains were able to maintain their internal pH above 6.5, suggesting that other mechanisms are also active, such as F0F1 ATPase activity (Pereira et al. 2009). These results are consistent with those obtained by other authors (Romano et al. 2014; Trip et al. 2012; Wolken et al. 2006) who indicate that amino acid decarboxylation pathways may be involved in cytoplasmic pH homeostasis through the alkalinizing effect of the decarboxylation reaction.

The present work provides evidence of a physiological role for tyramine biosynthesis in E. faecalis. It appears to be involved in resistance to acidic pH, since (i) the tdc cluster improves this bacterium’s growth in acidic media, (ii) it enhances its survival under GIT conditions, especially at low pH, and (iii) the expression of tdcA is induced by acidic pH. The protective effect seems to be mediated via the maintenance of intracellular pH. The present results highlight the importance of the tyramine pathway of E. faecalis in survival under acidic conditions, such as those encountered in passage through the GIT, against which it showed resistance and the continued ability to produce tyramine. Thus, tyramine production might be considered an important characteristic that contributes towards adaptability and that aids in the colonization of the human digestive tract by commensal and opportunistic pathogen enterococci. The increase in tyramine production under acidic conditions might also have food safety implications since enterococci are the major tyramine producers in many cheeses, where acid pH conditions are found due to the fermentation process.

Notes

Acknowledgments

This work was funded by the Ministry of Economy and Competitiveness, Spain (AGL2013-45431-R) and the Spanish National Research Council (CSIC201270E144). M.P. is beneficiary of an FPU fellowship from the Spanish Ministry of Education. We thank Pilar Fernández de Palencia and Paloma López for their help in the GIT survival experiments. The authors also thank Adrian Burton for language and editing assistance.

Supplementary material

253_2014_6301_MOESM1_ESM.pdf (237 kb)
ESM 1(PDF 236 kb)

References

  1. Agudelo Higuita N, Huycke M (2014) Enterococcal disease, epidemiology, and implications for treatment. In: Gilmore MS, Clewell DB, Ike Y, Shankar N (eds) Enterococci: from commensals to leading causes of drug resistant infection. Massachusetts Eye and Ear Infirmary, BostonGoogle Scholar
  2. Biswas I, Gruss A, Ehrlich SD, Maguin E (1993) High-efficiency gene inactivation and replacement system for gram-positive bacteria. J Bacteriol 175(11):3628–3635PubMedCentralPubMedGoogle Scholar
  3. Bonten MJ, Willems R, Weinstein RA (2001) Vancomycin-resistant enterococci: why are they here, and where do they come from? Lancet Infect Dis 1(5):314–325CrossRefPubMedGoogle Scholar
  4. Breeuwer P, Drocourt J, Rombouts FM, Abee T (1996) A novel method for continuous determination of the intracellular ph in bacteria with the internally conjugated fluorescent probe 5 (and 6-)-carboxyfluorescein succinimidyl ester. Appl Environ Microbiol 62(1):178–183PubMedCentralPubMedGoogle Scholar
  5. Calles-Enriquez M, Eriksen BH, Andersen PS, Rattray FP, Johansen AH, Fernandez M, Ladero V, Alvarez MA (2010) Sequencing and transcriptional analysis of the Streptococcus thermophilus histamine biosynthesis gene cluster: factors that affect differential hdcA expression. Appl Environ Microbiol 76(18):6231–6238. doi:10.1128/AEM. 00827-10 CrossRefPubMedCentralPubMedGoogle Scholar
  6. Castanie-Cornet MP, Foster JW (2001) Escherichia coli acid resistance: cAMP receptor protein and a 20 bp cis-acting sequence control pH and stationary phase expression of the gadA and gadBC glutamate decarboxylase genes. Microbiology 147:709–715PubMedGoogle Scholar
  7. Connil N, Le Breton Y, Dousset X, Auffray Y, Rince A, Prevost H (2002) Identification of the Enterococcus faecalis tyrosine decarboxylase operon involved in tyramine production. Appl Environ Microbiol 68(7):3537–3544CrossRefPubMedCentralPubMedGoogle Scholar
  8. Coton M, Fernandez M, Trip H, Ladero V, Mulder NL, Lolkema JS, Alvarez MA, Coton E (2011) Characterization of the tyramine-producing pathway in Sporolactobacillus sp. P3J. Microbiol 157:1841–1849. doi:10.1099/mic. 0.046367-0 CrossRefGoogle Scholar
  9. EFSA (2011) Scientific Opinion on risk based control of biogenic amine formation in fermented foods. EFSA Panel on Biological Hazards (BIOHAZ). EFSA J 9(10):2393–2486Google Scholar
  10. Fernandez de Palencia P, Fernandez M, Mohedano ML, Ladero V, Quevedo C, Alvarez MA, Lopez P (2011) Role of tyramine synthesis by food-borne Enterococcus durans in adaptation to the gastrointestinal tract environment. Appl Environ Microbiol 77(2):699–702. doi:10.1128/AEM. 01411-10 CrossRefPubMedCentralPubMedGoogle Scholar
  11. Fernández de Palencia P, López P, Corbí A, Peláez C, Requena T (2008) Probiotic strains: survival under simulated gastrointestinal conditions, in vitro adhesion to Caco-2 cells and effect on cytokine secretion. Eur Food Res Technol 227(5):1475–1484. doi:10.1007/s00217-008-0870-6 CrossRefGoogle Scholar
  12. Fernandez M, Linares DM, Alvarez MA (2004) Sequencing of the tyrosine decarboxylase cluster of Lactococcus lactis IPLA 655 and the development of a PCR method for detecting tyrosine decarboxylating lactic acid bacteria. J Food Prot 67(11):2521–2529PubMedGoogle Scholar
  13. Fernandez M, Linares DM, Del Rio B, Ladero V, Alvarez MA (2007a) HPLC quantification of biogenic amines in cheeses: correlation with PCR-detection of tyramine-producing microorganisms. J Dairy Res 74(3):276–282. doi:10.1017/S0022029907002488 CrossRefPubMedGoogle Scholar
  14. Fernandez M, Linares DM, Rodriguez A, Alvarez MA (2007b) Factors affecting tyramine production in Enterococcus durans IPLA 655. Appl Microbiol Biotechnol 73(6):1400–1406. doi:10.1007/s00253-006-0596-y CrossRefPubMedGoogle Scholar
  15. Gahan CG, Hill C (2014) Listeria monocytogenes: survival and adaptation in the gastrointestinal tract. Front Cell Infect Microbiol 4:9. doi:10.3389/fcimb.2014.00009 CrossRefPubMedCentralPubMedGoogle Scholar
  16. Giraffa G (2003) Functionality of enterococci in dairy products. Int J Food Microbiol 88(2–3):215–222CrossRefPubMedGoogle Scholar
  17. Green MR, Sambrook J (2012) Molecular cloning: a laboratory manual, 4th edn. Cold Spring Harbor Laboratory Press, Cold Spring HarborGoogle Scholar
  18. Grundy FJ, Moir TR, Haldeman MT, Henkin TM (2002) Sequence requirements for terminators and antiterminators in the T box transcription antitermination system: disparity between conservation and functional requirements. Nucleic Acids Res 30(7):1646–1655CrossRefPubMedCentralPubMedGoogle Scholar
  19. Holo H, Nes IF (1989) High-frequency transformation, by electroporation, of Lactococcus lactis subsp. cremoris grown with glycine in osmotically stabilized media. Appl Environ Microbiol 55(12):3119–3123PubMedCentralPubMedGoogle Scholar
  20. Horton RM, Hunt HD, Ho SN, Pullen JK, Pease LR (1989) Engineering hybrid genes without the use of restriction enzymes: gene splicing by overlap extension. Gene 77(1):61–68CrossRefPubMedGoogle Scholar
  21. Jonsson M, Saleihan Z, Nes IF, Holo H (2009) Construction and characterization of three lactate dehydrogenase-negative Enterococcus faecalis V583 mutants. Appl Environ Microbiol 75(14):4901–4903CrossRefPubMedCentralPubMedGoogle Scholar
  22. Ladero V, Calles-Enríquez M, Fernández M, Alvarez MA (2010a) Toxicological effects of dietary biogenic amines. Curr Nutr Food Sci 6:145–156CrossRefGoogle Scholar
  23. Ladero V, Fernandez M, Calles-Enriquez M, Sanchez-Llana E, Canedo E, Martin MC, Alvarez MA (2012) Is the production of the biogenic amines tyramine and putrescine a species-level trait in enterococci? Food Microbiol 30(1):132–138. doi:10.1016/j.fm.2011.12.016 CrossRefPubMedGoogle Scholar
  24. Ladero V, Fernandez M, Cuesta I, Alvarez MA (2010b) Quantitative detection and identification of tyramine-producing enterococci and lactobacilli in cheese by multiplex qPCR. Food Microbiol 27(7):933–939. doi:10.1016/j.fm.2010.05.026 CrossRefPubMedGoogle Scholar
  25. Ladero V, Linares DM, Del Rio B, Fernandez M, Martin MC, Alvarez MA (2013) Draft genome sequence of the tyramine producer Enterococcus durans strain IPLA 655. Genome Announc 1:e00265–13. doi:10.1128/genomeA. 00265-13 PubMedCentralPubMedGoogle Scholar
  26. Lebreton F, Willems R, Gilmore M (2014) Enterococcus diversity, origins in nature, and gut colonization. In: Gilmore MS, Clewell DB, Ike Y, Shankar N (eds) Enterococci: from commensals to leading causes of drug resistant infection. Massachusetts Eye and Ear Infirmary, BostonGoogle Scholar
  27. Linares DM, Del Rio B, Ladero V, Martinez N, Fernandez M, Martin MC, Alvarez MA (2012a) Factors influencing biogenic amines accumulation in dairy products. Front Microbiol 3:180. doi:10.3389/fmicb.2012.00180 CrossRefPubMedCentralPubMedGoogle Scholar
  28. Linares DM, Fernandez M, Del-Rio B, Ladero V, Martin MC, Alvarez MA (2012b) The tyrosyl-tRNA synthetase like gene located in the tyramine biosynthesis cluster of Enterococcus durans is transcriptionally regulated by tyrosine concentration and extracellular pH. BMC Microbiol 12:23. doi:10.1186/1471-2180-12-23 CrossRefPubMedCentralPubMedGoogle Scholar
  29. Linares DM, Fernandez M, Martin MC, Alvarez MA (2009) Tyramine biosynthesis in Enterococcus durans is transcriptionally regulated by the extracellular pH and tyrosine concentration. Microb Biotechnol 2(6):625–633. doi:10.1111/j.1751-7915.2009.00117.x CrossRefPubMedCentralPubMedGoogle Scholar
  30. Linares DM, Martin MC, Ladero V, Alvarez MA, Fernandez M (2011) Biogenic amines in dairy products. Crit Rev Food Sci Nutr 51(7):691–703. doi:10.1080/10408398.2011.582813 CrossRefPubMedGoogle Scholar
  31. Livak KJ, Schmittgen TD (2001) Analysis of relative gene expression data using real-time quantitative PCR and the 2(−delta delta C(T)) method. Methods 25(4):402–408. doi:10.1006/meth.2001.1262 CrossRefPubMedGoogle Scholar
  32. Lucas P, Landete J, Coton M, Coton E, Lonvaud-Funel A (2003) The tyrosine decarboxylase operon of Lactobacillus brevis IOEB 9809: characterization and conservation in tyramine-producing bacteria. FEMS Microbiol Lett 229(1):65–71CrossRefPubMedGoogle Scholar
  33. Lund P, Tramonti A, De Biase D (2014) Coping with low pH: molecular strategies in neutralophilic bacteria. FEMS Microbiol Rev 4(10):1574–6976. doi:10.1111/1574-6976.12076 Google Scholar
  34. Lyte M (2004) The Biogenic amine tyramine modulates the adherence of Escherichia coli O157: H7 to intestinal mucosa. J Food Prot 67(5):878–883PubMedGoogle Scholar
  35. Marcobal A, de las Rivas B, Munoz R (2006a) First genetic characterization of a bacterial beta-phenylethylamine biosynthetic enzyme in Enterococcus faecium RM58. FEMS Microbiol Lett 258(1):144–149. doi:10.1111/j.1574-6968.2006.00206.x CrossRefPubMedGoogle Scholar
  36. Marcobal A, Martin-Alvarez PJ, Moreno-Arribas MV, Munoz R (2006b) A multifactorial design for studying factors influencing growth and tyramine production of the lactic acid bacteria Lactobacillus brevis CECT 4669 and Enterococcus faecium BIFI-58. Res Microbiol 157(5):417–424. doi:10.1016/j.resmic.2005.11.006 CrossRefPubMedGoogle Scholar
  37. Mathur S, Singh R (2005) Antibiotic resistance in food lactic acid bacteria-a review. Int J Food Microbiol 105(3):281–295. doi:10.1016/j.ijfoodmicro.2005.03.008 CrossRefPubMedGoogle Scholar
  38. Molenaar D, Abee T, Konings WN (1991) Continuous measurement of the cytoplasmic pH in Lactococcus lactis with a fluorescent pH indicator. Biochim Biophys Acta 1115(1):75–83CrossRefPubMedGoogle Scholar
  39. Nes IF, Diep DB, Ike Y (2014) Enterococcal bacteriocins and antimicrobial proteins that contribute to niche control. In: Gilmore MS, Clewell DB, Ike Y, Shankar N (eds) Enterococci: from commensals to leading causes of drug resistant infection. Massachusetts Eye and Ear Infirmary, BostonGoogle Scholar
  40. Paulsen IT, Banerjei L, Myers GS, Nelson KE, Seshadri R, Read TD, Fouts DE, Eisen JA, Gill SR, Heidelberg JF, Tettelin H, Dodson RJ, Umayam L, Brinkac L, Beanan M, Daugherty S, DeBoy RT, Durkin S, Kolonay J, Madupu R, Nelson W, Vamathevan J, Tran B, Upton J, Hansen T, Shetty J, Khouri H, Utterback T, Radune D, Ketchum KA, Dougherty BA, Fraser CM (2003) Role of mobile DNA in the evolution of vancomycin-resistant Enterococcus faecalis. Science 299(5615):2071–2074CrossRefPubMedGoogle Scholar
  41. Pereira CI, Matos D, Romao MVS, Crespo MTB (2009) Dual role for the tyrosine decarboxylation pathway in Enterococcus faecium E17: response to an acid challenge and generation of a proton motive force. Appl Environ Microbiol 75(2):345–352. doi:10.1128/AEM. 01958-08 CrossRefPubMedCentralPubMedGoogle Scholar
  42. Pessione E (2012) Lactic acid bacteria contribution to gut microbiota complexity: lights and shadows. Front Cell Infect Microbiol 2:86. doi:10.3389/fcimb.2012.00086 CrossRefPubMedCentralPubMedGoogle Scholar
  43. Pessione E, Pessione A, Lamberti C, Coisson DJ, Riedel K, Mazzoli R, Bonetta S, Eberl L, Giunta C (2009) First evidence of a membrane-bound, tyramine and beta-phenylethylamine producing, tyrosine decarboxylase in Enterococcus faecalis: a two-dimensional electrophoresis proteomic study. Proteomics 9(10):2695–2710. doi:10.1002/pmic.200800780 CrossRefPubMedGoogle Scholar
  44. Poolman B, Konings WN (1988) Relation of growth of Streptococcus lactis and Streptococcus cremoris to amino acid transport. J Bacteriol 170(2):700–707PubMedCentralPubMedGoogle Scholar
  45. Redruello B, Ladero V, Cuesta I, Alvarez-Buylla JR, Martin MC, Fernandez M, Alvarez MA (2013) A fast, reliable, ultra high performance liquid chromatography method for the simultaneous determination of amino acids, biogenic amines and ammonium ions in cheese, using diethyl ethoxymethylenemalonate as a derivatising agent. Food Chem 139(1–4):1029–1035. doi:10.1016/j.foodchem.2013.01.071 CrossRefPubMedGoogle Scholar
  46. Romano A, Ladero V, Alvarez MA, Lucas PM (2014) Putrescine production via the ornithine decarboxylation pathway improves the acid stress survival of Lactobacillus brevis and is part of a horizontally transferred acid resistance locus. Int J Food Microbiol 175:14–19. doi:10.1016/j.ijfoodmicro.2014.01.009 CrossRefPubMedGoogle Scholar
  47. Russo P, Fernandez de Palencia P, Romano A, Fernandez M, Lucas P, Spano G, Lopez P (2012) Biogenic amine production by the wine Lactobacillus brevis IOEB 9809 in systems that partially mimic the gastrointestinal tract stress. BMC Microbiol 12:247. doi:10.1186/1471-2180-12-247 CrossRefPubMedCentralPubMedGoogle Scholar
  48. Sanchez B, de los Reyes-Gavilan CG, Margolles A (2006) The F1F0-ATPase of Bifidobacterium animalis is involved in bile tolerance. Environ Microbiol 8(10):1825–1833CrossRefPubMedGoogle Scholar
  49. Solheim M, La Rosa SL, Mathisen T, Snipen LG, Nes IF, Brede DA (2014) Transcriptomic and functional analysis of NaCl-induced stress in Enterococcus faecalis. PLoS One 9:e94571. doi:10.1371/journal.pone.0094571 CrossRefPubMedCentralPubMedGoogle Scholar
  50. Trip H, Mulder NL, Lolkema JS (2012) Improved acid stress survival of Lactococcus lactis expressing the histidine decarboxylation pathway of Streptococcus thermophilus CHCC1524. J Biol Chem 287(14):11195–11204. doi:10.1074/jbc.M111.330704 CrossRefPubMedCentralPubMedGoogle Scholar
  51. Ubeda C, Taur Y, Jenq RR, Equinda MJ, Son T, Samstein M, Viale A, Socci ND, van den Brink MR, Kamboj M, Pamer EG (2010) Vancomycin-resistant Enterococcus domination of intestinal microbiota is enabled by antibiotic treatment in mice and precedes bloodstream invasion in humans. J Clin Invest 120(12):4332–4341. doi:10.1172/JCI43918 CrossRefPubMedCentralPubMedGoogle Scholar
  52. Van den Abbeele P, Grootaert C, Marzorati M, Possemiers S, Verstraete W, Gerard P, Rabot S, Bruneau A, El Aidy S, Derrien M, Zoetendal E, Kleerebezem M, Smidt H, Van de Wiele T (2010) Microbial community development in a dynamic gut model is reproducible, colon region specific, and selective for Bacteroidetes and Clostridium cluster IX. Appl Environ Microbiol 76(15):5237–5246. doi:10.1128/AEM. 00759-10 CrossRefPubMedCentralPubMedGoogle Scholar
  53. Wolken WA, Lucas PM, Lonvaud-Funel A, Lolkema JS (2006) The mechanism of the tyrosine transporter TyrP supports a proton motive tyrosine decarboxylation pathway in Lactobacillus brevis. J Bacteriol 188(6):2198–2206. doi:10.1128/JB.188.6.2198-2206.2006 CrossRefPubMedCentralPubMedGoogle Scholar

Copyright information

© Springer-Verlag Berlin Heidelberg 2014

Authors and Affiliations

  • Marta Perez
    • 1
  • Marina Calles-Enríquez
    • 1
  • Ingolf Nes
    • 2
  • Maria Cruz Martin
    • 1
  • Maria Fernandez
    • 1
  • Victor Ladero
    • 1
  • Miguel A. Alvarez
    • 1
  1. 1.Instituto de Productos Lácteos de Asturias (IPLA-CSIC)VillaviciosaSpain
  2. 2.Norwegian University of Life Sciences UMBÅsNorway

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