Modulation of Lactobacillus plantarum Gastrointestinal Robustness by Fermentation Conditions Enables Identification of Bacterial Robustness Markers

Background Lactic acid bacteria (LAB) are applied worldwide in the production of a variety of fermented food products. Additionally, specific Lactobacillus species are nowadays recognized for their health-promoting effects on the consumer. To optimally exert such beneficial effects, it is considered of great importance that these probiotic bacteria reach their target sites in the gut alive. Methodology/Principal Findings In the accompanying manuscript by Bron et al. the probiotic model organism Lactobacillus plantarum WCFS1 was cultured under different fermentation conditions, which was complemented by the determination of the corresponding molecular responses by full-genome transcriptome analyses. Here, the gastrointestinal (GI) survival of the cultures produced was assessed in an in vitro assay. Variations in fermentation conditions led to dramatic differences in GI-tract survival (up to 7-log) and high robustness could be associated with low salt and low pH during the fermentations. Moreover, random forest correlation analyses allowed the identification of specific transcripts associated with robustness. Subsequently, the corresponding genes were targeted by genetic engineering, aiming to enhance robustness, which could be achieved for 3 of the genes that negatively correlated with robustness and where deletion derivatives displayed enhanced survival compared to the parental strain. Specifically, a role in GI-tract survival could be confirmed for the lp_1669-encoded AraC-family transcription regulator, involved in capsular polysaccharide remodeling, the penicillin-binding protein Pbp2A involved in peptidoglycan biosynthesis, and the Na+/H+ antiporter NapA3. Moreover, additional physiological analysis established a role for Pbp2A and NapA3 in bile salt and salt tolerance, respectively. Conclusion Transcriptome trait matching enabled the identification of biomarkers for bacterial (gut-)robustness, which is important for our molecular understanding of GI-tract survival and could facilitate the design of culture conditions aimed to enhance probiotic culture robustness.


Introduction
According to the world health organization (WHO) probiotics are defined as live microorganisms which, when administered in adequate amounts, confer a health benefit on the host [1]. The most widely applied probiotic strains belong to the genera Lactobacillus and Bifidobacterium [2,3]. Probiotics are most commonly provided as freshly fermented food products, non-fermented food products to which probiotics are added, or as dried bacterial preparations [3,4]. The viability of bacteria is considered an important trait for probiotic functionality, justifying the interest to unravel the mechanism(s) involved in gastrointestinal (GI)-tract survival at the molecular level [5,6,7,8].
During passage through the GI-tract, probiotics encounter several stresses including acidity in the stomach which may reach a pH as low as 1 during fasting [7]. This low extracellular pH affects the proton motive force of the bacterial cells, thereby disrupting the energy supply required for processes such as membrane transport [9]. In addition, lower intracellular pH values caused by acidic conditions may inhibit specific pathways by damaging acid-sensitive associated enzyme functions [9]. After stomach passage probiotic strains reach the small intestine, where bile acids act primarily as a surfactant that can disrupt bacterial membranes [10] and damage macromolecules such as RNA and DNA through the generation of free oxygen radicals [11]. Moreover, protonated bile acids can freely pass bacterial cell membranes and release protons intracellularly which might lead to lowering of the intracellular pH, analogous to acid stress [9].
Among the lactobacilli, Lactobacillus plantarum is encountered in a plethora of fermentations, ranging from vegetables to dairy, meat and sourdough [12,13]. L. plantarum is also frequently encountered as a natural inhabitant of the GI-tract of several mammals, including humans [14]. In addition, L. plantarum NCIMB8826 was demonstrated to effectively survive passage of the human stomach, reached the ileum in high numbers, and was detected in the colon [15]. A single colony isolate of this strain (designated L. plantarum strain WCFS1) was the first Lactobacillus strain of which the full genome sequence was published [16]. Subsequently, sophisticated bioinformatics tools were developed for this LAB strain, including an advanced genome annotation [17], genome-based metabolic models [18], as well as effective mutagenesis tools [19]. This enables the molecular investigation of gene-regulatory mechanisms underlying the observed GI-tract persistence of L. plantarum WCFS1.
The availability of full genome sequences has also enabled the exploration of genomic diversity among L. plantarum strains and its association to differential phenotypes [13,20,21,22,23]. To enable the identification of genes of which the relative expression level is correlated to the phenotype of interest, we recently developed a complementary transcriptome-phenotype matching strategy for L. plantarum (see accompanying paper by Bron et al.). Here, we employed this fermentation genomics platform to correlate transcriptome data to GI-tract survival. These correlations led to the identification of 13 candidate effector molecules for GI-tract persistence. A subsequent gene deletion strategy established a definite role in GI-tract persistence for the AraC-family transcription regulator encoded by lp_1669, the penicillin-binding protein Pbp2A involved in peptidoglycan biosynthesis, and the Na + /H + antiporter NapA3.

Gastric Acidity is a Critical Determinant of L. plantarum Survival
An in vitro assay was developed that allows high-throughput assessment of bacterial GI-tract survival ( fig. 1A). Two independent reference L. plantarum WCFS1 cultures that were harvested during logarithmic phase of growth (OD 600 = 1.0) displayed a 6log decrease in CFU?ml 21 in the GI-tract assay ( fig. 1B). The survival curves of these reference cultures demonstrated the major impact on survival exerted by gastric juice on L. plantarum viability and the relatively minor effect of the conditions which resembled the small intestine ( fig. 1B). This differential effect on survival during the two stages within the GI-tract assay was consistently observed for all cultures tested, irrespective of the fermentation conditions applied or the growth phases from which bacterial cells were harvested.
The strongest determinant in the loss of survival during the gastric juice treatment appeared to be pH. For screening log-phase cells of L. plantarum, a pH of 2.4 was used for cells, as lowering or increasing of the gastric juice pH by 0.1 pH unit resulted in death or survival of almost all cells, respectively (over 7-log differences, data not shown). L. plantarum cells harvested at the stationary phase of growth consistently displayed a higher tolerance to the gastric juice treatment, which is exemplified by their higher survival rate in the GI-tract assay when a reduced pH of 2.3 was used (fig. 1B) at which the cells harvested from the logarithmic phase of growth were nearly all killed within 60 minutes of incubation.

Fermentation-enhanced Digestive Tract Survival
We examined the effects of different growth conditions on L. plantarum WCFS1 GI-tract survival by applying samples derived from the fermentation-genomics platform described in the accompanying paper by Bron et al. to our in vitro GI-tract assay. The results demonstrate that fermentation conditions used to culture L. plantarum WCFS1 conferred a profound influence on the GI-tract survival. Variable fermentation conditions resulted in major differences (a reduction of 7 logs for the logarithmic population and 5 logs for stationary cells) in L. plantarum WCFS1 survival after incubation in gastric juice ( fig. 2). Notably, survival of cultures grown in different fermentation conditions strongly exceeded the levels of variation in survival observed in independent GI-tract assays ( fig. 1B).
To identify the fermentation conditions that significantly affected the survival rate in the simulated GI-tract conditions, a L. plantarum WCFS1 cultures were grown aerobically at 28uC in 26CDM containing normal acid concentration, at a pH of 5.8 and without NaCl. The cultures were harvested at mid-exponential phase (OD 600 = 1.0) and subjected to an upper GI-tract mimicking assay (A): After 60 min incubation in gastric juice containing pepsin and lipase at a pH of 2.4 (logarithmic cells) or 2.3 (stationary cells), cultures were neutralized with NaHCO 3 and pancreatic juice containing pancreatin and bile acids was added and incubation continued for 60 min (see materials and methods for details). Preceding and during incubation, samples were taken for CFU determination (aligned arrows). Panel B shows the relative survival of two independent cultures in logarithmic phase (solid lines) and stationary phase (dashed lines) during the GI-tract mimicking assay. Input (CFU determination immediately prior to the GI assay) is set at 0 Log 10 CFU ml 21 , data presented are averages of technical sextuplicates (+standard deviation). doi:10.1371/journal.pone.0039053.g001 Mann-Whitney U test-based correlation analysis was performed in FermDB on all time points measured (See accompanying manuscript by Bron et al. for correlation analyses details). The presence of 300 mM additional NaCl in the growth medium resulted in a significant (P,0.05) negative influence on L. plantarum GI-tract survival irrespective whether cells were analyzed after collection from either logarithmic or stationary phase of growth (shown for 60 min incubation in fig. 3A and B). L. plantarum grown Figure 2. Relative GI-tract survival of differently grown L. plantarum WCFS1. Log 10 CFU ml 21 determination of L. plantarum WCFS1 in logarithmic phase (A) and stationary phase (B) after 20 (light grey), 40 (dark grey), and 60 min (black) gastric juice incubation. Input is set at 0 Log 10 CFU ml 21 , # = fermentation number, cultures were grown in 26CDM with (300 mM) or without (0) NaCl; with normal amino acid concentration (2) or reduced (1.1); at 28uC or 37uC; medium buffered at a pH of 5.2, 5.8, or 6.4; and aerobically (O 2 ) or anaerobically (N 2 ). Data presented are averages of technical sextuplicates (+ standard deviation). doi:10.1371/journal.pone.0039053.g002 in more acidic conditions (pH 5.2 instead of pH 6.4) and harvested in stationary phase showed a significantly (P,0.05) enhanced the gastric juice survival rate (shown for 60 min in fig. 3C).

Transcriptome to Phenotype Association Identifies Candidate Effector Molecules for GI-tract Survival
In parallel with the GI-tract survival patterns, transcriptome profiles were obtained for logarithmic cells harvested from all fermentation conditions employed in this study (see accompanying paper by Bron et al.). To investigate whether high-and low-rate surviving cultures in the GI-tract assay could be distinguished based on the expression of specific genes, the cultures were first ranked on their GI-tract survival after gastric juice incubation (t = 60 min). For cultures that had retained undetectable survival rates after 60 min of gastric incubation, the relative survival rates after 20 min and 40 min of gastric incubation, were employed to refine their relative survival ranking ( fig. 2A).
The transcriptomes of the eight cultures with the highest survival rates and the eight cultures with the lowest survival in the GI-tract assay were clearly distinguishable according to principal component analysis (PCA) ( fig. 4). This result indicated that the transcriptomes contained information (genes) within the first two components of the PCA which might allow the discrimination between high-and low-survival rates in the GI-tract assay. To identify specific transcripts that discriminate between low and high GI-tract survival, and thus can be regarded as candidate robustness markers, the random forest algorithm was applied (see the accompanying paper by Bron et al.). This allowed the identification of transcripts that have a high contribution to accurately predict the low-and high-survival outcomes (table S1).

Validation of Target GI-tract Survival Effector Molecules by Mutagenesis
To validate the association of the expression level of specific genes in L. plantarum with GI-tract survival, the 13 genes with the highest ranking based on the criteria described in the Materials and Methods section were targeted by genetic engineering (table 1), aiming to improve GI-tract survival beyond the levels observed with the wild-type strain. Therefore, the direction of the  Genes targeted for sakacin-inducible overexpression were thrC, lp_0149, hicD3, and lp_1357 (table 1). For folB overexpression, we used a previously constructed mutant that overexpresses the entire folB-folK-folE-folC2-xtp2-folP cluster [24,25]. Sakacin P induced overexpression of the cytoplasmic hicD3 and thrC and the downstream lp_2759 gene products could readily be confirmed by SDS-PAGE analysis of cell-free extracts of induced cultures (figure S1). In contrast, overproduction of the membraneanchored (lp_1357) and transmembrane proteins (lp_0148-0150) were not distinguishable by SDS-PAGE (data not shown).
Although overexpression could only be demonstrated for two of the genetic loci, we applied all overexpression strains to our GItract assay. The constructed overexpression and gene deletion mutants were grown until the logarithmic growth phase and subjected to the GI-tract assay. The survival of the Sakacin P induced overexpression mutants was anticipated to improve when compared to a control strain harboring the empty induction plasmid ( fig. 6). Although not significant, the contrary seemed to be the case, since the slight effects that were observed in some of the experiments suggested that the expression of the cloned genes reduced the survival capacity of these cells rather than improved.
In contrast, the L. plantarum Dpbp2A::cat, Dlp_1669::cat, and DnapA3::cat mutants showed significantly improved survival in the GI-tract assay, as compared to their parental strain ( fig. 7). These strains harbored disruptions in genes associated with poor survival in gastric stress. Notably, we have combined the individual mutants described here to construct Dpbp2A-DnapA3::cat and DnapA3-Dlp_1669::cat. However, these double gene deletion derivatives displayed robustness phenotypes comparable to the single DnapA3::cat gene deletion derivative, indicating that the positive effect on GI robustness of these mutations appeared not cumulative (data not shown). Nevertheless, these results establish the involvement of certain fermentation-condition dependent gene products in GI survival.
Pbp2A is annotated as a penicillin-binding protein involved in peptidoglycan biosynthesis, Lp_1669 is predicted to be a transcription regulator, and NapA3 is homologous to Na + /H + antiporters. To gain more insight in the mechanisms by which these proteins influence robustness, growth of the parental strain and the Dpbp2A::cat, Dlp_1669::cat, and DnapA3::cat derivatives was monitored under standard-and stress-conditions. At 28uC in complex culture medium (MRS), the growth rates of the mutants did not differ from the wild-type, nor did the addition of H 2 O 2 (1 to 5 mM), lysozyme (0.025 to 3.2 g/ml), or SDS (0.9 to 30 g/l) induce differences in growth rate of the mutants compared with the wild type strain. However, the presence of bile salts (10 to 50 mM) in the culture medium reduced the maximum growth rate of Dpbp2A::cat to 20% as compared to the parental strain (data not shown). This result indicates that Pbp2A contributes to the survival capacity of L. plantarum in low-pH, stomach like conditions, but also improves bile tolerance, but not to tolerance to detergents in general.
The addition of NaCl to the growth medium reduced the growth rate of DnapA3::cat to 20% (400 mM) and 80% (1 M) of the wild type (data not shown). As NapA3 is a Na + /H + antiporter which might be affected by extracellular pH, the growth of the DnapA3::cat mutant was monitored under different starting pH conditions (pH 4.6 to 6.4) in the presence and absence of NaCl (300 mM). The growth rate of the mutant appeared unaltered during growth in the absence of salt. Only the presence of NaCl reduced the growth rate of DnapA3::cat under all measured conditions (data not shown). These results support a role of this function in salt tolerance, which in our experiments, appeared to be independent of the pH.
Contrary to DnapA3::cat and Dpbp2A::cat, a specific phenotype was not established for the transcription regulator Lp_1669. To elucidate the regulon associated with this regulator, the transcriptome profile of the NZ3417 CM (Dlp_1669::cat) strain was compared to that of the wild-type strain grown in 26CDM [17] or MRS. The results showed that the Lp_1669-deficient strain displayed enhanced expression of genes belonging to the main functional class of cell envelope associated functions, and more specifically to its subclass of surface polysaccharides, lipopolysaccharides, and antigens. This effect of the mutation was observed independent of the medium used ( fig. 8 and table S2 and S3). Analysis at the individual transcript level revealed that the capsular polysaccharide (CPS) clusters cps2, cps3, and cps4 were induced in the MRS-grown Lp_1669-deficient strain as compared to the wildtype, suggesting that the regulatory function   encoded by lp_1669 is involved, either directly or indirectly, in the regulation of CPS biosynthesis. Notably, especially the expression of the cps2 cluster was induced in 26 CDM grown Lp1669 deficient cells (table S2 and S3). Analysis of monosaccharide composition revealed minor changes in CPS sugar composition of the Lp_1669deficient strain in comparison to the wild type strain (table 2). Galactosamine was only detected in the mutant strain, whereas arabinose was found only in the wild-type strain. Rhamnose and glucosamine also tended to be slightly more abundant in the wild type L. plantarum WCFS1. Moreover, the average molar mass of Dlp_1669::cat strain-derived CPS was 1.5-fold higher compared to the wild type (Table 2). This indicates that Lp_1669 seems to be involved in subtle CPS modification, specifically in chain length determination. These observations might also (partially) explain the observed increased gastrointestinal survival of the L. plantarum Lp_l669-deficient strain.

Discussion
This study demonstrates that the production method, medium composition, and stage of growth strongly influenced the GIsurvival efficacy of L. plantarum WCFS1. Combining the fermentation and survival data pinpointed to specific fermentation conditions that may enhance robustness (low salt and low pH), whereas genome association analysis of the transcriptome and survival data revealed 13 genes potentially involved in GI-survival.
As reported previously [26], cells harvested from stationary phase generally were more robust than logarithmically growing cells, and in particular, those cells displayed enhanced survival in gastric juice which overall had a dramatically larger impact on survival compared to pancreatic juice. The influence of acidity on GI-tract survival was also emphasized by the observation that lowering the gastric juice pH by as little as 0.1 unit had a pronounced impact on survival. Differences among bacterial species in their sensitivity to gastric and intestinal secretions have been observed before [27,28,29] and a higher sensitivity for acid than bile stress was also noted for L. rhamnosus, as well as for other L. plantarum strains [30,31]. The finding that exposure to low pH during growth enhances GI-survival is in agreement with earlier observations that pre-adaptation to sublethal stress conditions enhances the subsequent robustness of bacteria to lethal stress conditions [9], supporting the suitability of the fermentation genomics platform and bioinformatics tools employed in this study and the accompanying paper by Bron et al. For salt it is known that it can protect against, but also increase susceptibility to, other stresses [32,33]. Moreover, these results clearly establish that fermentation conditions have a major impact on the GI-tract associated stress tolerance of bacterial cultures, and that specifically mild salt stress and lower pH adaptation may elicit adaptive responses that reduce and support such stress tolerance, respectively.
Overexpression and gene deletion derivatives of the parental strain were constructed, depending on the direction of the predicted correlation to GI-tract survival and aiming to enhance this trait. Three of the five constructed gene deletion derivatives displayed enhanced GI-tract survival, confirming the predicted role of the targeted genes. By contrast, none of the overexpression derivatives displayed improved robustness behavior in the GI-tract assay, and all had survival characteristics that were virtually identical to those of the parental strain. A possible explanation for these observations may be found in the potential disruption of a gene-regulatory network by the deletion of a single gene in that network, while overexpression of a single element from a complementary gene-function network may not provide the same effect as the enhanced expression of all elements in the network. The fact that 3 out of 5 gene deletion derivatives displayed enhanced survival is in line with earlier observations [21,22,23] and can be explained by the fact that the random forest algorithm also leads to the identification of non-causal relationships, reiterating the importance of follow-up mutagenesis approaches to establish a definite role for candidate biomarkers identified with this algorithm.
The 3 genes for which the importance in GI-tract survival could be confirmed by gene deletion encode a AraC family regulator (Lp_1669), a Na + /H + antiporter; NapA3, and a penicillin binding protein; Pbp2A. Notably, all three proteins are associated with cell wall modification and transport, and their mutation may lead to cell envelope modulation. This finding per se, may not be qualified as surprising, because the cell envelope is the first line of defense against stresses [34]. Moreover, the resistance to acid and adaptation to bile stress in L. plantarum WCFS1 has been associated with membrane integrity and cell envelope modifications, respectively [35,36]. The AraC family of regulators to which Lp_1669 belongs [16] is characterized by transcriptional regulators that act mostly as activators. However, in some cases these regulators serve as repressors of transcription or as both activators and repressors [37]. The observed effect of Lp_1669 on GI-tract survival is likely to be indirect, possibly via CPS remodeling, because the Lp_1669-deficient strain had CPS with a higher molar mass that might result in a thicker CPS layer around the cells. It has been demonstrated that the presence of EPS/CPS improved the in vivo GI survival of L. rhamnosus GG [38]. The Na + /H + antiporter NapA3 might affect GI survival via a role in pH homeostasis. Because disruption of napA3 improved GI-tract survival, it seems likely that NapA3 exports sodium ions associated with the influx of protons, thereby decreasing its internal pH and proton motive force due to the acid stomach conditions. This is also in line with our observation that the gene deletion derivative is only reduced during growth in the presence of sodium salts. Finally, pbp2A encodes the penicillin binding protein 2A which is annotated to be involved in peptidoglycan biosynthesis [16]. Disruption of pbp2A improved the acid stomach condition survival, while it decreased the growth rate in the presence of bile. Noteworthy in this respect is the finding that the compositions of peptidoglycan directly affects the integrity of the cells and can influence the acid-and bile-tolerance [9,39,40,41]. Moreover, transcriptome analysis of L. acidophilus NCFM and L. plantarum WCFS1 demonstrated that many genes related to cell membrane and peptidoglycan biosynthesis displayed altered expression profiles during exposure to bile [39,42]. An increased acid sensitivity by the inactivation of penicillin binding proteins is found in Lactococcus lactis and L. reuteri [9,40]. However, we found the deletion of pbp2A improves the GI-tract survival, which suggests that disruptions in peptidoglycan biosynthesis genes could either improve or decrease the survival of probiotics, reiterating the general concept of subtle inter-strain and species differences in survival mechanisms.
In conclusion, this study demonstrated that fermentation conditions have a large influence on the GI-tract survival of L. plantarum. We showed that transcriptome-trait matching enables the identification of genetic loci involved in gastrointestinal robustness and this approach can also be employed to rationally design fermentation and process conditions that aim for the production of probiotics with improved GI survival and consequently have a higher potential to achieve their desired healthbeneficial effects on the consumer.

GI-tract Assay and Correlation to Transcriptome Data
Cells were harvested from the fermentation-genomics platform at OD 600 = 1.0 for full-genome transcriptome profiling (see accompanying paper by Bron et al.), while the GI-tract survival was determined in the same cells, as well as in cells that were harvested 25 h after inoculation. One set of fermentations (F19-F24, see accompanying paper by Bron et al.) was excluded from the data analysis as the GI-tract survival data appeared unreliable, likely caused by minor deviations in the pH of the batch of GJ applied which is known to heavily influence GI survival. For GI-tract survival analysis, cultures were washed with prewarmed (37uC) PBS and resuspended in prewarmed (37uC) filter sterilized gastric juice [53 mM NaCl, 15 mM KCl, , 10 mM NaHCO 3 , 30 mg ml 21 pancreatin (Sigma; derived from porcine stomach) and bile acid mixture (latter two components were added fresh to pancreatic juice immediately prior to the treatment)] was added, followed by continued incubation for another 60 min (agitation at 10 rpm, 37uC). The bile acid mixture consisted of 3.0 mM (final concentration in assay) sodium glycocholate hydrate, 1.3 mM sodium glycodeoxycholate, 2.4 mM sodium glycochenodeoxycholate, 1.0 mM taurocholic acid sodium salt hydrate, 0.4 mM sodium taurodeoxycholate hydrate and 1.0 mM sodium taurochenodeoxycholate to mimic human bile components and concentrations [43]. Preceding and during GI-tract assay incubation (t = 0, 20, 40, 60, 90, and 120), samples were taken for colony forming unit (CFU) enumeration by spot-plating [44]. A reduction of 8 logs could be detected with this method. Relative GI-tract survival of the different cultures was expressed as the fraction of the corresponding input numbers of viable cells (t = 0 was set at 1.00). The transcriptome and GI-tract survival data are available in FermDB (www.cmbi.ru.nl/fermdb).
The initial list of genes predicted by the random forest algorithm [45] (see the accompanying paper by Bron et al. for details on correlation analyses) to be associated with GI-tract survival was further refined by application of several selection criteria that are based on transcript ranking. Firstly, only transcripts with an importance factor higher than 1 according to the random forest algorithm were selected for further analysis. Secondly, the quantitative correlation of individual transcripts with the survival rate observed in individual cultures was evaluated, selecting those transcripts (genes) that had the highest quantitative correlation with survival (expressed in R 2 in table S1, see figure 5 for two examples). Lastly, genes encoding prophage associated functions that are typically hypervariable among L. plantarum strains were discarded [13,20]. The remaining transcripts and their associated genes (table 1) were considered to have the strongest correlation with the measured gastric juice tolerance and were therefore selected for validation by gene deletion or overexpression.

Overexpression Mutant Construction and SDS-PAGE Analysis
Gene overexpression mutants were constructed using the expression vector pSIP411 [48]. For the candidate genes selected for overexpression that were part of a predicted operon [49], the whole operon was cloned in the sakacin induction vector (table 1). Primers were designed (table S5) to introduce a restriction enzyme site for cloning the target gene(s) into the expression vector pSIP411 at the NcoI site. The lp_1357 and thrC+lp_2759 overexpression mutants were designed with BspHI site, which has compatible ends with NcoI site. The target gene(s) were amplified by PCR using corresponding primers for each mutant (F1/F2, G1/G2, H1/H2 and I1/I2 for lp_1357, lp_2349, thrC+lp_2759, and lp_0148,0150 mutants, respectively). The reactions were carried out with KOD polymerase (Novagen, Darmstadt, Germany) according to the instructions of the manufacturer. The purified PCR products were digested by restriction enzymes (Invitrogen, Molecular probes, Inc, USA) for which sites were introduced in the primers (see table S4) and cloned in NcoI-SmaI digested pSIP411. Ligation mixtures were transformed to E. coli, and re-isolated from primary transformants. Correctly assembled overexpression plasmids were identified by PCR, restriction and sequence analysis. Re-isolated plasmids were propagated into L. plantarum WCFS1 and transformants were selected on MRS containing 30 mg?ml 21 erythromycin (table 3).
For protein analysis of the overexpression mutants, the induction and sample preparation procedures were modified from the description by Sørvig et al. [48]. The 19-amino-acid inducing peptide (of Met-Ala-Gly-Asn-Ser-Ser-Asn-Phe-Ile-His-Lys-Ile-Lys-Gln-Ile-Phe-Thr-His-Arg [50]) was custom-synthesized by BACHEM (Budendorf, Switzerland). The inducing peptide was dissolved in degassed water, as recommended by BACHEM to avoid oxidation of the peptides. The overnight cultures of the overexpression strains were diluted 50-fold and then incubated at 37uC. After OD 600 had reached 0.3, the inducing peptide was added to the cultures at varying concentrations of 0, 0.1, 1, 10, and 50 ng/ml. Incubation was continued at 37uC for another 4 h until the OD 600 had reached approximately 1.8. Bacterial cells were collected by centrifugation at 5,2006g for 10 min, followed by resuspension of the cell pellet in 50 mM Sodium-phosphate buffer pH 7. The cells were disrupted with 1 g zirconium beads by using a FastPrep TM (Qbiogene Inc, Cedex, France). After the disruption, the samples were centrifuged 5 min at 20,8006g to obtain cell-free extracts for analysis by SDS-PAGE.

DNA Microarray Analysis and Data Visualization
DNA microarray analysis were performed to compare global transcriptome profiles of NZ3417 CM (Dlp_1669::cat) and the wildtype. RNA isolation from L. plantarum, subsequent cDNA synthesis and indirect labeling, as well as DNA microarray hybridizations were performed as described in the accompanying paper by Bron et al. The hybridization scheme is presented in figure S3. Genes of the Lp_1669 regulon with FDR-adjusted p-values less than 0.05 together with a fold-change higher than 2.0 or lower than 0.5 were considered to be significantly differently expressed. All microarray data is MIAME compliant and is available in the GEO database under accession number GSE31254. The biomolecular interaction network of the Lp_1669 regulon in 26 CDM and MRS was visualised using the Cytoscape software (version 2.8.1) [51], and the Biological Networks Gene Ontology (BiNGO) tool [52] was employed to detect significantly overrepresented categories in the regulon of Lp_1669. See the accompanying paper by Bron et al. for details.

Phenotypic Assays of Mutant Strains
Gene deletion mutants were analyzed for their gastrointestinal survival characteristics in a procedure identical to that described for the wild-type (see above). To evaluate the relative GI-tract survival of the overexpression mutants, the mutant strain SIP411B (empty vector) and the overexpression mutants were sakacininduced (50 ng/ml) (see above). Additionally, to measure the relative GI-tract survival of the folate overexpression strain, strains NZ7021 (empty vector) and NZ7026 (folate overproducing strain) [25] were inoculated at OD 600 = 0.1 in MRS containing 80 mg/ ml chloramphenicol and 0 or 10 mg/ml p-aminobenzoic acid (pABA) according to Wegkamp et al. [24], grown at 37uC until OD 600 was 1.0, and subjected to the GI-tract survival assay. To evaluate relative growth efficiency of the deletion mutants, the parental strain (WCFS1) and mutant strains NZ3412 CM (Dpbp2A::cat), NZ3417 CM (Dlp_1669::cat), and NZ3416 CM (DnapA3::cat) were inoculated at OD 600 = 0.1 in 96-wells plates and incubated in MRS broth at 28uC. OD 600 of the cultures was monitored spectophotometrically (GENios, Tecan Austria GmbH, Grödig, Austria).

Capsular Polysaccharide Isolation and Determination
Capsular polysaccharide (CPS) was purified and chain lengths and sugar composition were determined essentially as described before [53]. Briefly, 500 ml cultures of L. plantarum WCFS1 and NZ3417 CM (Dlp_1669::cat) were grown in 26 CDM at 37uC until stationary phase (25 h). After 1 h incubation at 55uC, the cells were separated from the CPS containing growth medium by centrifugation for 15 min (60006g) and to prevent overgrowth during dialysis, erythromicine was added to the supernatant to a final concentration of 10 mg/ml. A dialyzing tube 12-1400 Da (Fisher Scientific) was prepared by boiling twice 2% NaHCO 3 / 2 mM EDTA, and once in reverse osmosis water. After overnight dialysis against running tap water followed by 4 h dialysis using reverse osmosis water, the samples were freeze-dried and stored at 220uC until further analysis.
The samples were dissolved in eluent (100 mM NaNO 3 +0.02% NaN 3 ), filtered over 0.2 mm, and placed in a thermally controlled sample holder at 10uC and 200 ml was injected (model 231 Bio, Gilson) on the columns connected in series and remained at 35uC with a temperature control module (Waters, Milford, USA) to perform size exclusion chromatography (SEC) [TSK gel PWXL guard column, 6.0 mm64.0 cm, TSK gel G6000 PWXL analytical column, 7.8 mm630 cm, 13.0 mm and TSK gel G5000 PWXL analytical column, 7.8 mm630 cm, 10 mm (TosoHaas, King of Prussio, USA)]. Light scattering was measured at 632.8 nm at 15 angles between 32u and 144u (DAWN DSP-F, Wyatt Technologies, Santa Barbara, USA). UV absorption was measured at 280 nm (CD-1595, Jasco, de Meern, The Netherlands) to detect proteins. The specific viscosity was measured with a viscosity detector (ViscoStar, Wyatt Technologies, Santa Barbara, USA) at 35uC and sample concentration was measured by refractive index detection, held at a fixed temperature of 35uC (ERC-7510, Erma Optical Works, Tokyo, Japan).
During the analysis with SEC the polysaccharide peak was collected (2 min60.5 mL/min = 1 mL). The acid hydrolyses of the collected polysaccharide was carried out for 75 min at 120uC with 2 M trifluoro acetic acid under nitrogen. Following hydrolyses, the solutions were dried overnight under vacuum and dissolved in water. High Performance Anion Exchange Chromatography with Pulsed Amperometric Detection (HPAEC-PAD) on a gold electrode was used for the quantitative analyses of the monosaccharides rhamnose, galactosamine, arabinose, glucosamine, galactose, glucose, mannose, xylose, galacturonic acid, and glucuronic acid. The analyses were performed with a 600E System controller pump (Waters, Milford, USA) with a helium degassing unit and a model 400 EC detector (EG&G, Albuquerque, USA). With a 717 autosampler (Waters, Milford, USA), 20 ml of the sample was injected on a Dionex Carbopac PA-1, 25064 mm , column thermostated at 30uC. The monosaccharides were eluted at a flow rate of 1.0 mL/min. The monosaccharides were eluted isocratic with 16 mM sodium hydroxide, followed by the elution of the acid monosaccharides starting at 20 min with a linear gradient to 200 mM sodium hydroxide +500 mM sodium acetate in 20 minutes. Data analysis was done with Dionex Chromeleon software version 6.80. Quantitative analyses were carried out using standard solutions of the monosaccharides (Sigma-Aldrich, St. Louis, USA). Figure S1 SDS-PAGE of cell-free extracts logarithmic L. plantarum strains overexpressing hicD3 (lp_2349) and overexpressing thrC (lp_2758) and lp_2759. The arrows indicate protein bands increasing with increasing amounts of Sakacin P (inducing peptide, IP). Empty vector = pSIP411B. L. plantarum harboring pNZ3431 (over-hicD3), and pNZ3432 (over-thrC+lp_2759). Marker sizes are indicated in kDalton (kDa).