Short- and Long-Term Adaptation to Ethanol Stress and Its Cross-Protective Consequences in Lactobacillus plantarum

Abstract

This paper describes the molecular responses of Lactobacillus plantarum WCFS1 toward ethanol exposure. Global transcriptome profiling using DNA microarrays demonstrated adaptation of the microorganism to the presence of 8% ethanol over short (10-min and 30-min) and long (24-h) time intervals. A total of 57 genes were differentially expressed at all time points. Expression levels of an additional 859 and 873 genes were modulated after 30 min and 24 h of exposure to the solvent, respectively. Ethanol exposure led to induced expression of genes involved in citrate metabolism and cell envelope architecture, as well as canonical stress response pathways controlled by the central stress regulators HrcA and CtsR. Correspondingly, cells grown for 24 h in medium containing 8% ethanol exhibited higher levels of citrate consumption and modified cell membrane fatty acid composition and showed invaginating septa compared with cells grown in liquid medium without ethanol. In addition, these physiological changes resulted in cross-protection against high temperatures but not against several other stresses tested. To evaluate the role of HrcA and CtsR in ethanol tolerance, ctsR and hrcA gene deletion mutants were constructed. The growth rate of the L. plantarum ΔctsR::cat strain was impaired in de Man-Rogosa-Sharpe (MRS) medium containing 8% ethanol, whereas growth of the L. plantarum ΔhrcA::cat and ΔctsR ΔhrcA::cat mutants was indistinguishable from that of wild-type cells. Overall, these results suggest that the induction of CtsR class III stress responses provides cross-protection against heat stress.

INTRODUCTION

Lactic acid bacteria (LAB) are essential for the fermentation of numerous foods and beverages, including yoghurt, sausages, olives, and wine (22, 35, 36, 50). During the application of LAB in food and beverage fermentations, these bacteria are typically required to survive and remain metabolically active under diverse environmental conditions, including specific stresses. For example, wine LAB are exposed to several stresses, such as an acidic pH, a high alcoholic content, suboptimal growth at room temperature, and growth-inhibitory compounds originating from both yeast and bacterial metabolism (50).

In order to understand the mechanisms of stress tolerance of lactobacilli, numerous studies have examined the physiological and genetic adaptations of these organisms during growth and survival in diverse environmental stresses (12, 50, 59). Recently, the availability of complete genome sequences (http://www.ncbi.nlm.nih.gov/genomes/lproks.cgi) and postgenomic approaches have accelerated our understanding of the global (genome-wide) stress responses in lactobacilli to acid, lactate, oxidative, bile, and heat stresses (7, 8, 13, 39, 44, 53). These studies have shown that lactobacilli respond rapidly to their environment by modulating expression levels of genes involved in different cellular processes, including stress response pathways, cell division, transport, and cell envelope composition. Adaptation to the harsh environmental conditions is at least partially under the control of HrcA and CtsR, canonical class I and III stress response regulators present in many Gram-positive bacteria (59).

The stress responses of the model LAB Lactobacillus plantarum WCFS1 have also been the subject of numerous reports employing transcription profiling and targeted mutation analysis of individual genes encoding either stress response proteins or their regulators (8, 44, 52). Interpretation of the results obtained in these studies has been accelerated by the availability of the L. plantarum WCFS1 genome sequence (32), its advanced gene function annotation (56), and a stoichiometry-based genome-scale metabolic model (55), as well as effective mutagenesis tools (34). Thus far, the detrimental effects of ethanol on L. plantarum are poorly understood, and ethanol toxicity is generally attributed to the interaction of ethanol with the cell membrane resulting in a loss of membrane integrity and secondary effects on metabolism and stress response pathways (60). Ethanol stress is encountered by L. plantarum in a variety of beverage fermentations, most notably beer and wine, and strains of this species have been reported to display high levels of tolerance to this solvent (21, 58).

This study aimed to identify the global adaptive and cross-protective responses of L. plantarum WCFS1 during growth in the presence of ethanol. The molecular responses of L. plantarum WCFS1 to short- and long-term exposure to 8% ethanol were investigated by whole-genome transcription profiling. Determination of specific metabolic and morphological adaptations in L. plantarum and the cross-protective effects of ethanol exposure toward other environmental stresses complemented the transcriptome-based results. In addition, mutagenesis approaches revealed that the molecular adaptations are at least partly controlled by CtsR, as previous studies revealed the direct interaction between CtsR and the promoter regions of the ctsR-clpC operon and hsp1 gene (16).

MATERIALS AND METHODS

Strains and growth conditions.

Strains used in this study are described in Table S1 in the supplemental material. Lactobacillus plantarum WCFS1 (32) was grown at 20°C in MRS (de Man-Rogosa-Sharpe) broth (Difco, West Molesey, United Kingdom) with either 8% (vol/vol) additional water or 8% (vol/vol) ethanol. Growth and cell density were determined by measurement of the optical density at 600 nm (OD₆₀₀) of the culture using a spectrophotometer (Ultraspec 2000; Pharmacia Biotech, Cambridge, United Kingdom). Citrate, lactate, formate, pyruvate, 2,3-butadiol, acetoin, succinate, acetate, propionate, and ethanol concentrations were measured in culture supernatants by high-performance liquid chromatography (HPLC) as described previously (51). Cells were harvested at an OD₆₀₀ of 1.0 for transcript profiling, cross-protection experiments, microscopy, and lipid extraction.

RNA isolation and transcriptome analysis.

Transcriptome analysis was performed in duplicate immediately before (t = 0) and after exposure to 8% (vol/vol) ethanol in MRS for 10 min, 30 min, and 24 h. RNA extraction, reverse transcription, labeling, hybridization, and data analysis were performed as described previously (41). In short, following quenching, RNA was phenol-chloroform extracted and purified using the High Pure RNA isolation kit (Roche Diagnostics, Mannheim, Germany). The quality of the RNA obtained was measured with the 2100 Bioanalyzer (Agilent Technologies, Waldbronn, Germany) using the Agilent RNA 6000 Nano kit (Agilent Technologies, Santa Clara, CA), and samples with a 23S/16S RNA ratio equal to or higher than 1.6 were taken for cDNA synthesis. cDNA was synthesized using the Superscript TMIII reverse transcriptase (RT) enzyme (Invitrogen, Carlsbad, CA), purified with the CyScribe GFX purification kit (GE Healthcare, Buckinghamshire, United Kingdom), and labeled differentially using cyanine 3 or cyanine 5 labels (Amersham; CyDye postlabeling reactive dye pack; GE Healthcare, Buckinghamshire, United Kingdom). After a second purification with the CyScribe GFX purification kit (GE Healthcare, Buckinghamshire, United Kingdom), L. plantarum WCFS1 cDNA was hybridized to oligonucleotide DNA microarrays for this strain (Agilent Technologies, Santa Clara, CA). L. plantarum WCFS1 DNA microarrays were hybridized according to a modified loop design which included comparisons of all conditions within three steps (see Fig. S1 in the supplemental material). The transcript data were normalized by local fitting of an M-A plot applying the Loess algorithm (63), using the Limma package (48) in R (http://www.R-project.org) as previously described (41), and genes with false discovery rate (FDR)-adjusted P values less than 0.05 were considered to be significantly differently expressed. To analyze the results, heat maps of gene expression levels were constructed for the transcript profiles using the Genesis platform (54). Blastn analysis was performed using http://blast.ncbi.nlm.nih.gov/Blast.cgi.

Lipid and fatty acid (FA) extraction.

Approximately 1 × 10¹¹ L. plantarum cells grown in MRS (with or without 8% [vol/vol] ethanol) until reaching an OD₆₀₀ of 1.0 at 20°C were collected by centrifugation (15,300 × g for 10 min at 23°C) and washed with phosphate-buffered saline (PBS), pH 7.4. Cell walls were degraded using 0.05 g·ml⁻¹ lysozyme (Merck, Darmstadt, Germany) and 250 units·ml⁻¹ mutanolysin (Sigma-Aldrich, St. Louis, MO) in 100 mM K₂HPO_4⁻ buffer (pH 6.2) under agitation at 10 rpm for 3 h at 44°C (RPN2511E hybridization oven/shaker; Amersham Pharmacia Biotech, Little Chalfont, United Kingdom). The cells were collected by centrifugation at 4,000 × g for 10 min at 23°C, and the cell membranes were harvested by dissolving the pellets thoroughly in 3 ml diethyl ether-heptane (1:1) acidified with 2.5 M sulfuric acid. Following centrifugation at 500 × g for 5 min at 23°C, the upper organic phase was collected for total fatty acid methyl ester (FAME) analysis. FAMEs were generated and analyzed according to the method of Badings and Dejong (3). A gas chromatograph (GC) (Mega 8060; Carlo Erba, Milan, Italy) with flame ionization detection (FID) and on-column injector was used to separate the FAMEs. The GC column (WCOT fused silica with stationary-phase CP-Wax 52 CB; Varian, The Netherlands) contained hydrogen as a carrier gas and was 15 m in length, with an inside diameter of 0.32 mm and a film thickness of 0.50 μm. Data were analyzed with EZChrom Elite, version 3.1.4 (Agilent Technologies, Santa Clara, CA).

Microscopy.

For scanning electron microscopy (SEM), round (8-mm-diameter) coverslips were coated with poly-l-lysine (0.01% [wt/vol] in water) and incubated for 30 min in L. plantarum cultures (OD₆₀₀ = 1.0). Cells adhering to the coverslips were then fixed with 4% (vol/vol) glutaraldehyde for 30 min, rinsed with water, and subsequently dehydrated by serial incubation in an acetone solution, starting from 10% acetone and going up to 30%, 50%, 70%, and 100% acetone. After critical point drying with carbon dioxide (CPD 030; BalTec, Balzers, Liechtenstein), the coverslips were affixed to a sample holder by carbon adhesive tabs (Electron Microscopy Sciences) and sputter coated with 5-nm platinum in a dedicated preparation chamber (CT 1500 HF; Oxford Instruments, Cambridge, United Kingdom). The bacteria were analyzed with a field emission scanning electron microscope (JEOL 6300 F; Tokyo, Japan) at room temperature at a working distance between 8 and 15 mm, with secondary electron detection at 3.5 kV. Images were digitally recorded (Orion 6 PCI; E.L.I. sprl., Brussels, Belgium), and contrast and brightness were optimized using Adobe Photoshop CS (Adobe, San Jose, CA).

For phase-contrast microscopy, L. plantarum cultures were examined directly by phase contrast at a magnification of 1,250-fold with a Dialux 20 microscope (Leitz, Wetzlar, Germany). Fluorescence microscopy was performed as described previously (29) with several modifications. In short, control, 30-min ethanol-exposed and 24-h ethanol-exposed cultures (OD₆₀₀ = 1.0) were 10 times diluted, incubated for 20 min on low-melting-point agarose-coated microscope slides containing 20 μg·ml⁻¹ FM4-64 (Molecular Probes, Eugene, OR) and 0.5 μl·ml⁻¹ Syto9 (Molecular Probes, Eugene, OR), and imaged by oil immersion fluorescence microscopy (BX51TRF fluorescence microscope; Olympus Corporation, Tokyo, Japan) at a 500-fold magnification.

Mutant construction.

Gene deletion mutants were constructed by using the mutagenesis vector pNZ5319 according to the method of Lambert et al. (34). The L. plantarum WCFS1 ctsR and hrcA genes were replaced with a lox66-P₃₂-cat-lox71 cassette resulting in strains NZ3410CM (ΔctsR::cat) and NZ3425CM (ΔhrcA::cat), respectively. Primers used to construct the L. plantarum WCFS1 mutants are described in Table S2 in the supplemental material. In short, upstream and downstream flanking regions of hrcA and ctsR were amplified with primers A, B, C, and D for hrcA and E, F, G, and H for ctsR. Primers B and F and primers C and G contained an overhang region homologous to the ultimate 5′ and 3′ regions of the lox66-P₃₂-cat-lox71 cassette (amplified with primers I and J), respectively, to enable the joining of the three PCR products in a splicing-by-overlap-extension (SOEing) PCR (26) with primers E and H for ctsR and A and D for hrcA (see Table S2). The obtained amplicons were blunt end ligated into Ecl136II-SwaI-digested pNZ5319 (34) and resulted in plasmids pNZ3410, pNZ3423, and pNZ3425. After introduction of the mutagenesis plasmids into competent L. plantarum WCFS1, cells were plated on MRS containing 10 μg·ml⁻¹ chloramphenicol. After 48 h, double-crossover deletion mutants were initially selected by colony PCR using primer pairs M plus O and N plus P (named 87 [34]) for ctsR and K plus O and L plus P for hrcA (see Table S2). For each mutant, a colony that generated both flanking-PCR products was selected and plated on MRS with and without 30 μg·ml⁻¹ erythromycin. A single colony for each mutant displaying the anticipated erythromycin-sensitive phenotype was selected and designated NZ3410CM (ΔctsR::cat) and NZ3425CM (ΔhrcA::cat), the latter resulting from the use of plasmid pNZ3425. The L. plantarum WCFS1 ctsR-hrcA mutant was constructed in the NZ3410CM (ΔctsR::cat) background in two steps. First, strain NZ3410 (ΔctsR) was constructed by excision of the lox66-P₃₂-cat-lox71 cassette by expression of the Cre resolvase enzyme from pNZ5348 according to methods described by Lambert et al. (34). Introduction of pNZ3423 and colony confirmation by PCR resulted in strain NZ3423CM (ΔctsR ΔhrcA::cat) (see Table S1 in the supplemental material).

To evaluate relative growth efficiency, the wild-type (WCFS1) and mutant strains NZ3410CM (ΔctsR::cat), NZ3425CM (ΔhrcA::cat), and NZ3423CM (ΔctsR ΔhrcA::cat) were inoculated at an OD₆₀₀ of 0.1 in 96-well plates and incubated in MRS with or without 8% (vol/vol) ethanol at 20°C. The OD₆₀₀ of the cultures was monitored spectophotometrically (Safire2; Tecan Austria GmbH, Grödig, Austria) in a robotic setup (Genesis Workstation 150/8; Tecan Austria GmbH, Grödig, Austria). The significance of differences in growth rates of wild type and mutants was evaluated by analysis of variance (ANOVA) using R (http://www.R-project.org). Differences were considered significant if the P value was <0.05.

Cross-protection studies.

Wild-type L. plantarum WCFS1 was grown in MRS in the absence or presence of ethanol 8% (vol/vol) until the OD₆₀₀ was 1.0 at 20°C. Cells were washed in PBS before exposure to various stresses. For all stress tolerance assays, serial dilutions of the samples were prepared immediately after stress exposure and these serial dilutions were immediately plated on MRS agar. Plates were incubated at 30°C for 2 days for CFU enumeration according to the technique described by Sieuwerts et al. (46). Oxidative stress tolerance was determined upon suspending the L. plantarum cells in PBS containing 40 mM hydrogen peroxide, a concentration which is lethal to L. plantarum WCFS1 (52). Cells were collected every 5 min for 60 min for CFU enumeration. To quantify L. plantarum survival at low pH, cells grown in MRS or MRS with 8% (vol/vol) ethanol were suspended in PBS with an adjusted pH of 2.4 (acidified by 5 M HCl) and subsequently sampled at 5-min intervals, followed by assessment of the amounts of viable cells as described above. The heat resistance of wild-type and mutant L. plantarum cultures grown in MRS in the presence or absence of 8% (vol/vol) ethanol until the OD₆₀₀ reached 1.0 was assessed after suspending the cells in PBS or PBS containing 8% (vol/vol) ethanol, followed by incubation in a thermocycler (Biometra Thermocycler; Westburg, The Netherlands) at the following temperatures: 37.0, 37.5, 39.1, 41.7, 44.4, 47.1, 49.9, and 52.6°C. Cell survival was determined every 10 min for 60 min by CFU enumeration. To analyze heat tolerance levels of L. plantarum, the log₁₀ values of the time and temperature when 1% of the original population was able to form a colony were plotted.

To determine the impact of ethanol stress adaptation on salt tolerance, L. plantarum WCFS1 was cultured in MRS with or without the addition of 8% (vol/vol) ethanol until the OD₆₀₀ reached 1.0. These cultures were inoculated into MRS broth containing 0.6, 0.7, or 0.85 M NaCl, and culture density was monitored at 20°C for 72 h with a spectrophotometer (SPECTRAmax Plus384; Molecular Devices, United Kingdom). To determine UV radiation tolerance, serial dilutions of wild-type L. plantarum broth cultures were plated on MRS agar and exposed for 0 to 180 s to UV radiation at 254 nm (E-series hand-held UV lamp; Spectroline, Westbury, NY), with a lamp height of 9 cm. After exposure, the MRS agar plates were incubated at 30°C for 2 days prior to CFU determination.

Microarray data accession numbers.

The DNA microarray design and gene expression data are available at the Gene Expression Omnibus (http://www.ncbi.nlm.nih.gov/geo/) under accession numbers GPL4318 and GSE17847, respectively.

RESULTS

L. plantarum WCFS1 growth and metabolism in the presence of 8% (vol/vol) ethanol.

Cell growth and fermentation profiles of L. plantarum WCFS1 in MRS containing either 8% (vol/vol) additional water or 8% (vol/vol) ethanol were monitored over 24 h at 20°C (see Fig. S2 and S3 in the supplemental material). The growth temperature and alcohol concentration were selected because these conditions mimic wine fermentations and L. plantarum WCFS1 was able to reach a final OD₆₀₀ close to the control condition within a few days of growth. L. plantarum WCFS1 was able to grow in the presence of 8% (vol/vol) ethanol, albeit with an approximately 5-fold-lower growth rate (0.06 ± 0.003 h⁻¹) compared with MRS cultures (0.32 ± 0.03 h⁻¹). The final optical density also was approximately 1.4-fold reduced in MRS containing ethanol, and this amount coincided with a more-than-2.3-fold-lower cell yield (see Fig. S2). Culture medium pH values when L. plantarum reached an OD₆₀₀ of 1.0 were slightly lower for the MRS cultures (pH 5.09 ± 0.02) than for cultures in ethanol-containing MRS (pH 5.15 ± 0.02). This result might have been due to the 10-fold-larger amounts of citrate consumed per 100 μmol lactate produced during L. plantarum growth in the presence of ethanol (0.59 μmol citrate consumed) compared with control cultures (0.06 μmol citrate consumed). Conversely, lactate was the primary fermentation end product of the actively dividing cultures, but also small amounts of formate and acetate were detected (see Fig. S3).

Global transcript profiles of L. plantarum WCFS1 during growth in ethanol.

The transcriptomes of L. plantarum WCFS1 after short (10-min and 30-min) and extended (24-h) incubation in ethanol-containing MRS medium were identified using DNA microarrays specific for this strain. The 24-h time point was selected because at that time L. plantarum was in mid-exponential phase of growth (OD₆₀₀ = 1.0), enabling comparisons to transcript profiles of reference MRS cultures (t = 0) harvested at the same cell density and growth phase. Genes differentially expressed by L. plantarum during exposure to ethanol were identified by comparisons to transcriptomes of L. plantarum WCFS1 cells grown on MRS. After 10 min of exposure to 8% (vol/vol) ethanol in MRS, 57 genes were significantly differentially expressed compared with MRS cultures (t = 0) (Fig. 1A). These genes constitute a core transcriptional response by L. plantarum to ethanol since their expression levels remained similarly upregulated and downregulated after 30 min and 24 h of exposure to this compound (Fig. 1A). The core ethanol response included 1.3- to 5.4-fold activation of established stress-associated genes, including groEL, groES, hsp3, grpE, and lp_0752 (putative stress-responsive transcription regulator), lp_0726 (membrane-bound protease of the CAAX family), and lp_3128 (stress-induced DNA binding protein) (Fig. 1B). In L. plantarum WCFS1, lp_3128 was upregulated after exposure to hydrogen peroxide stress (44). The gene lp_3128 shares 98% identity with the DNA starvation/stationary-phase protection protein Dps of Lactobacillus delbrueckii subsp. bulgaricus ND02. The Dps protein of Escherichia coli was previously shown to protect against DNA damage (2, 30, 57). Genes required for citrate metabolism, specifically citCDEF and fum, were also induced 1.7- to 7.0-fold at all time points (Fig. 1B). In contrast, genes coding for fatty acid biosynthesis, including fabZ1, fabH2, acpA2, and fabD, were downregulated between 1.5- and 4.0-fold (Fig. 1B).

Fig. 1.

(A) Venn diagram of the number of L. plantarum WCFS1 genes differentially expressed during 10 min, 30 min, and 24 h of incubation in MRS in the presence of 8% (vol/vol) ethanol compared with MRS incubation (0 min). Numbers before and after the slash represent up- and downregulated genes, respectively, compared with cells incubated in MRS. (B) The heat map shows expression levels of the 57 core response genes differentially expressed at all time points (10 min, 30 min, and 24 h) in MRS containing ethanol compared with MRS cultures. The lp_ number indicates gene number on L. plantarum WCFS1 chromosome (32). Genes with FDR-adjusted P values less than 0.05 were considered to be significantly differentially expressed.

Approximately 30% of the protein-encoding genes annotated in the L. plantarum genome were differentially expressed at 30 min (916 genes) and 24 h (930 genes) after inoculation into ethanol-containing MRS (Fig. 1A). At both time points, stress response pathways were induced and cell division as well as lipid and amino acid metabolism was downregulated. In the sections below, these and additional modifications in L. plantarum gene expression patterns and their associated phenotypes in response to ethanol are described.

Effects of ethanol on cell envelope composition, cell division, and morphology.

According to transcriptome analysis, L. plantarum cell membrane and cell wall components were influenced by ethanol. Expression of the dlt operon required for d-alanylation of teichoic acids was induced 1.3- to 1.7-fold by the presence of ethanol. Two tagE genes, tagE5 and tagE6, possibly involved in wall teichoic acid biosynthesis, were induced 1.4-fold after ethanol exposure for 24 h (Fig. 2A). Several lipoprotein precursor-encoding genes were induced 1.2- to 3.5-fold in the presence of ethanol after 30 min or 24 h or at both time points. In addition, three out of four L. plantarum capsular polysaccharide biosynthesis loci (cps1, cps3, and cps4) were downregulated 1.4- to 3.1-fold in the ethanol-containing MRS for 30 min or 24 h or at both time points (Fig. 2A).

Fig. 2.

Heat map of L. plantarum WCFS1 genes differentially expressed in the presence of 8% (vol/vol) ethanol for 10 min, 30 min, and 24 h. Genes are grouped based on duration of the response, gene annotation, and functional category. Gene expression levels of the cultures grown in MRS containing 8% (vol/vol) ethanol compared with control MRS cultures are shown according to annotation for cell envelope-associated functions (A), cell division (B), and genes involved in stress response pathways (C). The lp_ number indicates gene number on L. plantarum WCFS1 chromosome (32). Genes with FDR-adjusted P values less than 0.05 were considered to be significantly differentially expressed.

Ethanol stress also significantly affected the expression of L. plantarum genes associated with the fatty acid biosynthesis pathways. In general, the majority of genes required for membrane lipid biosynthesis were downregulated, including genes coding for fatty acid elongation proteins (fab) and an acyl carrier protein (ACP). The fab locus constitutes 12 genes which were repressed at least 1.5-fold starting 10 min after exposure to ethanol in MRS and remained downregulated after 30 min and 24 h in that culture medium (Fig. 2A). In contrast, expression of the two L. plantarum WCFS1 acetyl coenzyme A carboxylase (ACC) operons involved in the initiation phase of fatty acid (FA) biosynthesis differed such that acc1 was induced and acc2 was repressed. Finally, increased expression levels were observed for the gene encoding an acyl carrier protein synthase which maintains the ACP pool in its active form (acpS; 1.2-fold at 24 h) (31), and cyclopropane-fatty-acyl-phospholipid synthase (cfa2; 1.5- and 1.3-fold at 30 min and 24 h, respectively) (Fig. 2A).

Fatty acid methyl ester (FAME) analyses showed increases in the amounts of saturated fatty acids (SFA) palmitric acid (C_16:0, 1.9-fold) and stearic acid (C_18:0, 3.8-fold) in cells after 24 h of growth in ethanol-containing MRS. A 1.6-fold decrease of the amounts of the monounsaturated fatty acid C_18:1 was detected, whereas polyunsaturated acid C_18:3 increased 1.5-fold (Fig. 3). Collectively, the L. plantarum membranes from cultures grown in MRS with 8% ethanol contained an approximately 2.7-fold-lower ratio of unsaturated fatty acids (USFA) relative to saturated fatty acids (USFA/SFA = 2.85 ± 0.29) compared with control MRS cultures (USFA/SFA = 7.80 ± 1.20) (Fig. 3).

Fig. 3.

Fatty acid composition of L. plantarum WCFS1 grown in presence or absence of 8% (vol/vol) ethanol. Proportions of total membrane fatty acids were determined in mid-logarithmic cultures (OD₆₀₀ = 1.0) grown at 20°C in MRS (white bars) or MRS containing 8% ethanol (black bars). All fatty acids detected for the cells are shown. iso, isomer; conj, conjugated; USFA, unsaturated fatty acids; SFA, saturated fatty acids. The average (±standard deviation) out of four independent cultures is shown.

Because the transcript profiles indicated significant changes to the cell surface of L. plantarum in the presence of ethanol (Fig. 2A), global cell morphology and appearance were also determined for the L. plantarum cells using SEM. Mid-exponential-phase cells grown for 24 h in the presence of ethanol exhibited a rougher appearance than, and counterclockwise, spiral-shaped invaginating septa which were absent in, L. plantarum cells harvested from MRS (compare Fig. 4A and C with Fig. 4B and D). The unusual chain angles conferred by the spiral-shaped cells were also observed by phase-contrast microscopy (data not shown). Control and ethanol-exposed L. plantarum cells stained with the lipophilic cationic styryl FM4-64 dye did not show membrane lipid spirals, as was detected in Bacillus subtilis (4), nor was a difference observed between the two cultures in membrane lipid distribution (see Fig. S4 in the supplemental material). Although the physiological changes which resulted in these aberrantly shaped cells are unclear, it is likely that cell division is disturbed during ethanol exposure. This is supported by the finding that L. plantarum genes coding for septum site determination proteins MinC and MinD and the tubulin-like FtsZ protein required for establishing the site of cell division were downregulated (1.4-, 1.2-, and 1.4-fold, respectively) during growth in the presence of ethanol (Fig. 2B). Simultaneously, the gene coding for EzrA, a protein which inhibits Z-ring formation (6), was expressed at higher levels (Fig. 2B). Gene expression levels of other cell division and shape determination proteins were reduced (MreC, 1.3- and 1.5-fold for 30 min and 24 h, respectively), while several cell division-associated genes, including mreB, mreD, and rodA, were not differentially expressed in L. plantarum exposed to 8% ethanol in MRS (Fig. 2B).

Fig. 4.

SEM analysis of L. plantarum WCFS1 cultures grown in the presence or absence of ethanol. L. plantarum WCFS1 was grown at 20°C and harvested during exponential phase (OD₆₀₀ = 1.0) from MRS (A and C) or 8% (vol/vol) ethanol-containing MRS (B and D).

Induction of stress response pathways in L. plantarum during growth in ethanol.

Genes coding for the class I and class III stress response transcriptional regulators HrcA and CtsR, as well as the genes under their control, were differentially expressed in the presence of 8% ethanol. The predicted regulons of both regulators are shown in Table S3 in the supplemental material. Transcription of hrcA and two genes which are predicted to be regulated by HrcA (62), dnaK (encoding a heat shock protein) and dnaJ (a chaperone protein), was significantly upregulated in ethanol-containing MRS at 30 min (Fig. 2C). Other genes at least partially controlled by HrcA coding for chaperones GroES, GroEL, and GrpE and the putative membrane-bound protease lp_0726 were upregulated at all time points (Fig. 1B and 2C). Transcription of ctsR was significantly reduced in cells exposed to ethanol for 24 h. Genes shown to be repressed by CtsR, including clpP, clpE (encoding proteases), and hsp1 (small heat shock protein) (16, 18), were upregulated after 30 min and 24 h of ethanol exposure (Fig. 2C).

Other genes associated with tolerance to one or more environmental stresses were also differentially regulated during growth in ethanol. Stress response genes primarily known for roles in heat resistance were upregulated at all time points and include genes for a small heat shock protein (hsp3, HSP 19.3) and for a transcriptional regulator (lp_3128; stress-induced DNA binding protein) (Fig. 2C). Other genes associated with heat tolerance were intermittently upregulated in L. plantarum and include hsp2 (HSP 18.55), clpL and clpX (proteases), and tig (trigger factor) (Fig. 2C). Finally, a cell surface-localized protease encoded by htrA was also expressed at an elevated level (1.4-fold) after ethanol incubation for 30 min and 24 h. This gene was induced in Lactococcus lactis and Lactobacillus helveticus upon exposure to ethanol, NaCl, and heat (19, 47).

Genes coding for adaptation to oxidative stresses, including a glutathione peroxidase (gpo), thioredoxin (trxA1), stress-induced DNA binding protein (lp_3128 gene), catalase (kat), and a ferric uptake regulator (fur) (45), were induced in ethanol-exposed cultures after 24 h of growth. In comparison, genes in the SOS regulon important for survival under conditions which induce DNA damage were either downregulated or not differentially expressed during extended ethanol exposure (24 h). Similarly, the expression of three cell surface complexes (lp_2173 to lp_2175, lp_2975 to lp_2978, and lp_3676 to lp_3679) that were previously shown to be strongly induced during lactate stress was unaffected by ethanol stress (39).

Cross-protection of ethanol-exposed L. plantarum cells against high temperatures.

Because known stress response pathways were activated in L. plantarum WCFS1 during growth in the presence of ethanol, we examined whether this strain could withstand higher levels of other chemical or environmental stresses after exposure to ethanol compared to normally grown cells. The cross-protective stress tolerance levels of L. plantarum cultures grown for 24 h in the presence of ethanol were determined by exposing the cells to lethal levels of hydrogen peroxide (40 mM), UV radiation (254 nm, ranging from 0 to 180 s), acid pH (pH 2.4), and elevated temperatures (37°C to 53°C), as well as growth in high NaCl concentrations (0.6, 0.7, and 0.85 M).

Among the stress conditions tested, the only difference between the ethanol-exposed and control L. plantarum cultures was the increased capacity of the ethanol-exposed cells to survive at elevated temperatures. Although all cultures exhibited an exponential decay in viability in the presence of heat, L. plantarum cells grown until exponential phase in MRS containing 8% ethanol for 24 h (OD₆₀₀ = 1.0) survived longer and at higher temperatures between 37°C and 53°C over a range of 0- to 60-min exposure times than did cells harvested at the same optical density in normal MRS. This was observed by plotting the log₁₀ values of temperature and time when 1% of the starting population was still able to form a colony after heat exposure (Fig. 5). L. plantarum grown in MRS with 8% ethanol was able to survive at temperatures approximately 4°C higher than those of control MRS cultures. This cross-protective effect was observed when ethanol-exposed L. plantarum cultures were subjected to heat both in the presence and in the absence of 8% ethanol, although heat resistance was higher when ethanol was absent (Fig. 5). The viability of cells suspended in 8% (vol/vol) ethanol at the time of heat exposure declined at higher rates (between −18 log₁₀ min/°C and −19 log₁₀ min/°C) than did that of cells exposed to heat alone (−13 log₁₀ min/°C), independently of whether the cultures were grown in the presence of ethanol.

Fig. 5.

Heat resistance of L. plantarum WCFS1 grown in presence or absence of 8% (vol/vol) ethanol subjected to heat stress for 60 min. Shown are the time and temperature when 1% of the original population was able to form a colony. Cultures grown in MRS (open symbols) and in MRS containing 8% (vol/vol) ethanol (filled symbols) were subjected to heat stress with (circles) and without (diamonds) the presence of 8% (vol/vol) ethanol. Representative values of three independent cultures are shown. ○, R² of linear trend line = 0.96; •, R² = 0.95; ⋇, R² = 0.96; ♦, R² = 0.98.

ctsR and not hrcA influences growth of L. plantarum in ethanol.

To identify the roles of L. plantarum CtsR and HrcA stress response pathways under ethanol stress conditions, ctsR and hrcA deletion mutants were constructed (ΔctsR::cat, ΔhrcA::cat, and ΔctsR ΔhrcA::cat). Growth of the ΔhrcA::cat mutant was similar to that of wild-type L plantarum at 20°C. The growth rates of L. plantarum ΔctsR::cat and ΔctsR ΔhrcA::cat mutants grown in MRS at 20°C were slightly, but significantly, lower than that of the wild-type strain (1.1- and 1.2-fold, respectively) (Fig. 6A).

Fig. 6.

Growth rates of wild-type and mutant L. plantarum WCFS1 in MRS and MRS containing 8% (vol/vol) ethanol. L. plantarum WCFS1 and ΔctsR::cat, ΔhrcA::cat, and ΔctsR ΔhrcA::cat deletion mutants were grown in MRS (A) or MRS containing 8% ethanol (B) at 20°C. Significant differences in the observed growth rates of the mutants in comparison to the parental (wild-type) strain are marked by asterisks (P < 0.05). The growth rates are given as the average (±95% confidence interval) out of three independent experiments.

When grown at 20°C in MRS containing 8% ethanol, the ΔctsR::cat strain exhibited a 1.2-fold (P = 0.01)-higher growth rate than did the parental strain, whereas the ΔhrcA::cat and ΔctsR ΔhrcA::cat mutants grew similarly to wild-type cells (Fig. 6B). This indicates that CtsR negatively influences the growth rate in MRS containing ethanol at 20°C and that the growth advantage of the CtsR-deficient strain in ethanol is abolished when HrcA is absent. This result indicates an overlap in the CtsR and HrcA regulatory networks as was previously predicted (50).

DISCUSSION

Lactobacillus species are able to grow and survive under suboptimal conditions during food and beverage fermentations. Here, we unraveled the adaptations expressed by L. plantarum WCFS1 which enabled growth in medium containing 8% ethanol, a level found in some alcoholic beverages. L. plantarum WCFS1 was shown to adapt by modulating basic metabolic pathways and cell envelope composition and by inducing stress response pathways. Transcriptional responses were elicited within 10 min of exposure to 8% ethanol and expanded during extended incubation (30 min and 24 h). These adaptations resulted in cross-protection against thermal stress but not other stresses.

Ethanol is known to interfere with bacterial cell membrane integrity by interacting at the lipid-water interface. Ethanol influences membrane lipid ordering and bilayer stability and affects membrane characteristics such as permeability, fluidity, and the functioning of membrane-embedded enzymes (60). Genome-wide analyses of L. plantarum gene expression in the presence of 8% ethanol revealed that this organism responds immediately upon exposure to this solvent. This response is sustained under continuous ethanol stress and can be seen as a core response to ethanol. In addition, extended incubation in ethanol resulted in the expansion of the L. plantarum transcriptional changes beyond this core response.

The core response to ethanol stress included activation of citrate metabolism (citCDEF operon), which was accompanied by increased utilization of citrate from the medium. In L. plantarum, citrate is converted to acetate and oxaloacetate by citrate lyase and oxaloacetate is subsequently decarboxylated to form pyruvate (27). Activation of citrate metabolism in response to ethanol stress was also observed in Oenococcus oeni (38) and is probably explained by its membrane potential and pH gradient-generating effects, which can support cellular energy supplies (28, 40).

Modification of cellular FA (CFA) metabolism was another core response of L. plantarum to ethanol. Overall, the transcript profiles suggest that a reduction in FA biosynthesis led to changes in the composition of the cell membrane. Exponential-phase L. plantarum cells collected after growth in ethanol-containing MRS harbored reduced levels of C_18:1; increased levels of palmitic acid, stearic acid, and C_18:3; and an overall decrease in USFA/SFA ratios compared to cells grown in MRS. These membrane modifications resemble those observed in O. oeni ATCC BAA-1163 grown under similar conditions (24). The observed alterations in L. plantarum FA composition probably resulted from changes in de novo FA biosynthesis. Although it is possible that desaturases could modify existing phospholipid acyl chains in the membrane bilayer (64, 65), evidence that this occurred in L. plantarum is lacking. Phospholipid acyl desaturase, phospholipid cis-trans isomerase, and CFA synthase (64) are the known bacterial enzymes which catalyze FA desaturation; however, the L. plantarum genome appears to encode only a CFA synthase. CFA appears to be absent from L. plantarum membranes, and hence, CFA synthase likely does not confer a major role in the observed changes in FA composition under ethanol stress.

Transcriptome analyses also identified differential expression of several genes involved in cell wall-associated functions under ethanol stress. These adaptations included induction of the dlt operon, a locus which is involved in d-alanylation of teichoic acids. Induction of these genes was observed previously when L. plantarum was exposed to bile, another surface-active component (8). In addition, changes in expression of tagE (a gene possibly involved in wall teichoic acid biosynthesis) and certain genes coding for cell surface lipoproteins and capsular polysaccharides suggest that there were significant modifications to the cell envelope structure of L. plantarum upon ethanol exposure. The cell wall acts as a binding scaffold for enzymes and thereby has an important role in control of cell division and morphology (33). Remarkably, growth of L. plantarum in the presence of 8% ethanol resulted in invaginating spirals at the septum site of dividing cells. This phenotype resembles that of a conditional ftsZ mutant of Escherichia coli when it was grown at nonpermissive growth temperatures. The division defect of the E. coli ftsZ mutant was explained by a failure in FtsZ-ring assembly and closure (1, 5). The morphology of ethanol-exposed L. plantarum cells might have resulted from changes in FtsZ-ring assembly or other cell division-associated functions due to alterations in cell envelope composition, as was shown previously for E. coli (37).

Although there was some overlap between gene expression of L. plantarum during ethanol stress and the transcriptional responses of this organism to other environmental insults, growth of L. plantarum in the presence of ethanol cross-protected this organism exclusively against thermal stress. Similarly, exposure of Bacillus cereus to sublethal concentrations of ethanol induced cross-protection against thermal but not oxidative or high-salt stress (9). In L. plantarum, transcriptional modifications in response to ethanol included the induction of known heat shock response genes (10, 13), including hsp2 (Hsp 18.55) and hsp3 (Hsp 19.3), two genes which were previously shown to support growth of L. plantarum at elevated temperatures and in 12% ethanol (17). Heat shock responses of LAB are classified into six classes depending on their mode of transcriptional regulation in B. subtilis (43). HrcA is commonly regarded as a class I transcriptional repressor, and its regulon was predicted in L. plantarum on the basis of a cognate cis-acting element, designated CIRCE, in the promoter regions of groEL-groES, hrcA-grpE-dnaK-dnaJ, and lp_0726 (61). Transcriptional regulation by HrcA is dependent on availability of the GroELS complex such that HrcA is inactive when GroELS is unavailable during periods of cellular stress (43). Transcription of groELS, grpE, and lp_0726 was elevated in L. plantarum after 10 min, 30 min, and 24 h of incubation in MRS containing ethanol, indicating a rapid and continuous unfolding of proteins due to the presence of the alcohol. In contrast, induction of the heat shock genes dnaK-dnaJ was observed only after 30 min of exposure to ethanol. This result might be due to the differential processing of the polycistronic hrcA-grpE-dnaK-dnaJ transcript, as has been proposed as the mechanism of differential transcription of this operon in B. subtilis and Lactobacillus sakei (25, 42).

The class III heat shock regulon is controlled by CtsR, a transcriptional repressor which binds to a heptanucleotide direct repeat referred as the CtsR box (15). CtsR negatively autoregulates its own synthesis by the same mechanism (14). The CtsR regulon was previously shown to be involved in ethanol and heat stress responses in B. subtilis (23) and L. plantarum (49). Analogously, our results show that the CtsR regulon was partially induced after 30 min and 24 h of ethanol exposure and included elevated expression of ClpP-, ClpE-, and Hsp1-encoding genes. This finding suggests that the chaperonin function of GroELS was not sufficient to sustain the correct folding of proteins during ethanol stress, and the accumulation of denatured and aggregated proteins resulted in the activation of Clp-mediated proteolysis (20). The temporal activation of class I and III stress regulon members refines our knowledge of the sequential involvement of these stress regulons in the maintenance of appropriate protein functioning under ethanol stress conditions.

To further investigate the role of ctsR and hrcA in ethanol adaptation, mutants of L. plantarum WCFS1 that lack one or both of these genes were constructed. The role of the transcriptional repressor CtsR in adaptation of L. plantarum to ethanol and heat stress was observed previously (11). The slightly higher growth rate of L. plantarum ΔctsR::cat than of wild-type cells in the presence of 8% ethanol confirms the contribution of the ctsR regulon members to counteracting ethanol-induced stress. The growth rate of this mutant under normal growth conditions in MRS was slightly reduced relative to that of wild-type cells. While inactivation of hrcA did not affect the growth rate of L. plantarum in MRS culture medium with or without ethanol present, notably, in the presence of 8% ethanol the L. plantarum hrcA-ctsR mutant grew with a rate equal to that of the wild type, suggesting an interaction between the ctsR and hrcA stress response regulons in L. plantarum.

This study advances knowledge of the stress tolerance mechanisms of L. plantarum, which are important to control this organism in industrial processes that may include exposure to ethanol or similar stress conditions. Improved understanding of the adaptive behavior of bacteria under stress conditions could pave the way toward rational design of methods to maximize cell survival and targeted improvement of stress robustness in LAB.