Characterization of the biofilm phenotype of a Listeria monocytogenes mutant deficient in agr peptide sensing

Abstract Listeria monocytogenes is a food‐borne human pathogen and a serious concern in food production and preservation. Previous studies have shown that biofilm formation of L. monocytogenes and presence of extracellular DNA (eDNA) in the biofilm matrix varies with environmental conditions and may involve agr peptide sensing. Experiments in normal and diluted (hypoosmotic) complex media at different temperatures revealed reduced biofilm formation of L. monocytogenes EGD‐e ΔagrD, a mutant deficient in agr peptide sensing, specifically in diluted Brain Heart Infusion at 25°C. This defect was not related to reduced sensitivity to DNase treatment suggesting sufficient levels of eDNA. Re‐analysis of a previously published transcriptional profiling indicated that a total of 132 stress‐related genes, that is 78.6% of the SigB‐dependent stress regulon, are differentially expressed in the ΔagrD mutant. Additionally, a number of genes involved in flagellar motility and a large number of other surface proteins including internalins, peptidoglycan binding and cell wall modifying proteins showed agr‐dependent gene expression. However, survival of the ΔagrD mutant in hypoosmotic conditions or following exposure to high hydrostatic pressure was comparable to the wild type. Also, flagellar motility and surface hydrophobicity were not affected. However, the ΔagrD mutant displayed a significantly reduced viability upon challenge with lysozyme. These results suggest that the biofilm phenotype of the ΔagrD mutant is not a consequence of reduced resistance to hypoosmotic or high pressure stress, motility or surface hydrophobicity. Instead, agr peptide sensing seems to be required for proper regulation of biosynthesis, structure and function of the cell envelope, adhesion to the substratum, and/or interaction of bacteria within a biofilm.


| INTRODUC TI ON
Listeria monocytogenes is a saprophytic soil organism that is widespread in nature (Vivant, Garmyn, & Piveteau, 2013) and frequently found in food processing environments posing a threat to the food chain Muhterem-Uyar et al., 2015;NicAogáin & O'Byrne, 2016). In healthy individuals, food-borne infections with L. monocytogenes result in mild gastroenteritis or remain completely asymptomatic. However, in at-risk groups such as immunocompromised persons, elderly people and pregnant women, L. monocytogenes may cause life-threatening disease (Allerberger & Wagner, 2010;Vázquez-Boland et al., 2001).
Two characteristics that make L. monocytogenes a major concern in food processing and sanitation of the respective production lines are the ability to form surface-attached communities (also referred to as biofilm formation) and an extremely high tolerance to a wide range of environmental conditions and stresses (Ferreira, Wiedmann, Teixeira, & Stasiewicz, 2014;NicAogáin & O'Byrne, 2016).
Following initial adhesion, L. monocytogenes is able to form surface-attached communities (Carpentier & Cerf, 2011;Renier, Hébraud, & Desvaux, 2011;da Silva & De Martinis, 2013). The population density in these communities is 1-2 orders of magnitude lower than that observed for surface-attached communities of other bacteria (da Silva & De Martinis, 2013). Compared to other bacteria, biofilm formation of L. monocytogenes is not as pronounced, but may be enhanced by precolonization of surfaces by other bacteria such as Pseudomonas putida and Flavobacterium sp., probably involving the extracellular polymeric substances (EPS) produced by these bacteria (Giaouris et al., 2015). By contrast, precolonization of surfaces with, for example, Pseudomonas fragi and Serratia ssp. reduced biofilm formation of L. monocytogenes. There are conflicting results regarding the production of EPS by L. monocytogenes. Some studies conclude that L. monocytogenes biofilms generally lack EPS (Renier et al., 2011). By contrast, a recent study could show that EPS production by L. monocytogenes can be induced by elevated levels of the second messenger cyclic di-GMP and the genetic locus for EPS production was identified (Chen et al., 2014). This leaves room for interpretation as to whether or not these communities are biofilms according to the strict definition, which requires the communities to be embedded into a self-produced matrix of extracellular polymeric substances (Flemming & Wingender, 2010). Nevertheless, several studies have provided evidence for three-dimensional structures described as honey-comb or knitted chains and the presence of extracellular DNA (eDNA) and exopolysaccharides (Borucki, Peppin, White, Loge, & Call, 2003;Guilbaud, Piveteau, Desvaux, Brisse, & Briandet, 2015;Harmsen, Lappann, Knøchel, & Molin, 2010;Rieu et al., 2008;Zetzmann et al., 2015). Thus, it seems reasonable to consider surface attached communities of L. monocytogenes as biofilms.
The aim of this study was to investigate the biofilm phenotype of a L. monocytogenes mutant deficient in agr peptide sensing.

| Bacterial strains and growth conditions
In this study, L. monocytogenes strains EGD-e, its isogenic mutant EGD-e ΔagrD, and the genetically complemented strain EGD-e ΔagrD::pIMK2agrD were used. All strains have been described previously (Riedel et al., 2009). Bacteria were cultivated routinely in brain heart infusion broth (BHI, Oxoid, Altrincham, Cheshire, England) or 10-fold diluted BHI (0.1BHI) at 25 or 37°C. Precultures for functional assays were prepared by inoculation of a single colony from a fresh agar plate into 10 ml BHI and incubated aerobically on a rotary shaker (200 rpm) at 25°C overnight (o/N, i.e., approx. 16 hr).

| Quantification of surface-attached biomass
To quantify surface-attached biomass, classical crystal violet assays were performed in 96-well microtiter plates as described previously (Zetzmann et al., 2015). Where indicated, 1 unit (U) of DNase I (Thermo Scientific, Waltham, MA) or 1 mg/ml pronase (Sigma-Aldrich, Darmstadt, Germany) was added to the wells directly after inoculation. Plates were incubated at 25°C or 37°C for 24 hr. For analysis, biofilms were washed gently twice with phosphate-buffered saline (PBS) followed by staining with 0.1% (v/v) crystal violet solution (Merck, Darmstadt, Germany) for 30 min. After three further washings with PBS, crystal violet was released from biofilms by addition of 100 µl 96% (v/v) ethanol and incubated for 10 min.
Biofilm biomass was quantified by measuring absorbance at 562 nm (Abs 562 nm ) with background correction, that is, crystal violet staining in wells incubated with sterile media under the same conditions.

| Membrane and cell wall stress assays
To assess the effects of reduced osmolarity in 0.1BHI on viability of bacteria, aliquots of the preculture used for biofilm assays were diluted 1:100 in either 0.1BHI or demineralized H 2 O (dH 2 O) and viable cell counts were determined as colony-forming units per ml (CFU/ ml) by spot-plating. For this purpose, 10 µl aliquots of 10-fold serial dilutions were plated in triplicate onto BHI agar and the colonies of an appropriate dilution were counted to calculate CFU/ml. The effect of lysozyme treatment was analyzed in a similar assay except that bacteria were inoculated from a preculture into 0.1BHI, grown at 25°C to exponential growth phase (OD 600nm = 0.15-0.2), harvested by centrifugation and resuspended in 0.1BHI containing incubated in the presence of lysozyme at 25°C for the indicated time and log-reduction was calculated relative to CFU/ml at t = 0 min of an untreated control, that is, an aliquot resuspended in 0.1BHI without lysozyme.

| High hydrostatic pressure treatments
For high hydrostatic pressure (HPP) experiments, a single colony from a fresh BHI agar plate was inoculated into BHI broth and grown for 12 hr at 37°C. This preculture was diluted to an OD 600nm of 0.05 in 0.1BHI and grown for 1.5-2 hr to exponential growth phase (i.e., OD 600nm of 0.15 ± 0.02). At this stage, samples of 2 ml were loaded in Eppendorf tubes and sealed by carefully avoiding any air bubbles inside. Pressure treatments were conducted in a multivessel (four vessels of 100 ml) high-pressure equipment (Resato, Roden, the Netherlands) at 20 ± 0.5°C. As a pressure transmitting fluid a mixture of water and propylene glycol fluid (TR15, Resato) was used.
Pressure treatments were performed at 200, 300, and 400 MPa with a compression rate of 250 MPa/min and 60 s after the comeup time were considered the equilibration time necessary for each treatment. Samples were maintained for an additional 60 s at the established pressure followed by decompression of the vessels in less than 5 s. Treated samples were removed from the high-pressure vessels, and immediately afterwards, viable cell counts (CFU/ml) were quantified by spot-plating as described above.

| Motility assays
To assess motility of bacteria, precultures were prepared as described above in 0.1BHI at 25°C o/N. Of these precultures, soft agar of the same medium (0.1BHI, 0.2% agar) were inoculated by dipping an inoculation needle in the preculture and briefly stabbing onto the surface of the soft agar plate. After incubation for 24 h at 25°C, plates were imaged using a standard digital camera and the size of the zone of growth around the spot of inoculation was measured.

| Microbial adhesion to hydrocarbons
Surface hydrophobicity of all strains was evaluated using a standard assay to quantify microbial adhesion to hydrocarbons (MATH assay) (Rosenberg, 2006). Briefly, bacteria were grown in 0.1BHI at 25°C o/N, washed once in PBS and adjusted to an OD 600nm of 0.1 in PBS (OD1). Two milliliters of this suspension were mixed with 0.4 ml xylene and vortexed for 2 min. After separation of the phases, OD 600nm was again measured in the aqueous phase (OD2). Hydrophobicity (H) was then calculated as % = (OD1−OD2) OD1 × 100 .

| Statistical analysis
Statistical analysis was performed by Student's t test or analysis of variance (ANOVA) with Dunnett's posttest to adjust P-values for multiple comparisons using GraphPad Prism (version 6). Differences were considered significant at p < 0.05.

| RE SULTS AND D ISCUSS I ON
Recently, we were able to show that biomass and presence of eDNA in biofilms of L. monocytogenes EGD-e vary with growth conditions (Zetzmann et al., 2015). Additionally, a L. monocytogenes EGD-e ΔagrD deletion mutant showed reduced levels of surface-attached biomass in 0.1BHI at room temperature (Riedel et al., 2009) F I G U R E 1 Biofilm formation (a) and DNAseI sensitivity of biofilms (b) of L. monocytogenes EGD-e WT (W), EGD-e ΔagrD (Δ), and EGD-e ΔagrD::pIMK2agrD (C). Biofilms were grown in BHI or 0.1BHI at 25 or 37°C in the absence (a) or presence of DNaseI (b; 0.1BHI at 25°C only). Biofilm biomass was quantified by crystal violet staining and measuring absorbance at 562 nm (Abs 562nm ) after 24 hr of growth in polystyrene microtiter plates. All values are mean ± standard deviation of three independent experiments. Statistical analysis was performed by ANOVA with Dunnett's multiple comparisons test with L. monocytogenes EGD-e WT set as control condition (a) or Student's t test comparing biofilm of each strain in the presence and absence of DNase I (b; *p < 0.05; **p < 0.01; ***p < 0.001) followed by 0.1BHI at 25°C and lowest biofilm biomass was formed in 0.1BHI at 37°C. Interestingly, the ΔagrD mutant showed reduced biofilm formation only in 0.1BHI at 25°C. For all other conditions, no difference was observed between the three strains. Thus, agr peptide sensing is required for proper regulation of biofilm formation under specific conditions, that is, in 0.1BHI at 25°C.
Interestingly, these are the conditions under which biofilms of the WT strain showed increased abundance of eDNA and DNase I sensitivity (Zetzmann et al., 2015). This prompted us to test whether loss of agr peptide signaling is associated with altered sensitivity toward DNase I treatment (Figure 1b). However, biofilm formation of the ΔagrD mutant was reduced by DNase I to a similar extent as observed for the WT (and complemented strain) at 25°C in 0.1BHI indicating that eDNA is present in these communities and lack of eDNA is not responsible for the observed phenotype of L. monocytogenes EGD-e ΔagrD.
Since the conditions that produce the phenotype of ΔagrD mutant may cause osmotic stress due to the low nutrient and ion concentration in dH 2 O-diluted BHI (0.1BHI). In a previous study, a deletion mutant in the AgrC sensor histidine kinase of the agr system displayed increased sensitivity to high concentrations of salt (Pöntinen, Lindström, Skurnik, & Korkeala, 2017). Thus, we hypothesized that a reduced resistance to osmotic stress may lead to increased lysis of bacteria and, consequently, reduced surface-attached biomass.
In order to get a first indication whether deletion of agrD results in reduced stress resistance, we re-analyzed a previously published transcriptomic data set comparing L. monocytogenes EGD-e ΔagrD with its parental WT strain (Riedel et al., 2009).
The conditions of biofilm formation (0.1BHI, 25°C) and the transcriptomic analysis (BHI, 37°C) are different. Nevertheless, we reasoned that the transcriptional data would provide first indications as to whether or not stress related genes are affected by the lack in agr peptide signaling and any stress-related phenotype would be even more evident under for example, hypoosmotic stress (i.e., 0.1BHI). We therefore compared the differentially expressed genes to the regulon of the alternative sigma factor σ B , that is, the major regulator of the general stress response in many gram-positive bacteria including L. monocytogenes (Chaturongakul, Raengpradub, Wiedmann, & Boor, 2008;Kazmierczak, Mithoe, Boor, & Wiedmann, 2003;van Schaik & Abee, 2005). In L. monocytogenes, the σ B regulon comprises 168 genes that are positively regulated by σ B . Comparison with the 715 genes differentially expressed in L. monocytogenes EGD-e ΔagrD revealed an overlap of 132 genes, which is 78.6% of the σ B regulon and 18.5% of the agr regulated genes (Table 1 and  The four genes of the dltABCD operon are required for D-alanine esterification of teichoic acids in the cell wall of L. monocytogenes, which is involved in adhesion and virulence (Abachin et al., 2002), and a ΔdltABCD mutant showed impaired biofilm formation (Alonso et al., 2014). The entire dlt operon was differentially expressed in L. monocytogenes EGD-e ΔagrD. However, since its expression was increased in the ΔagrD mutant compared to the WT and it was thus ruled out as being responsible for the biofilm phenotype of the mutant.
Besides fliQ and motA, three other genes (flaA, fliD, and fliI) involved in flagellar motility and its regulation were shown to impact on biofilm formation by the transposon mutant screen (Alonso et al., 2014). Flagellar motility has previously been shown to play a role in adhesion and biofilm formation of L. monocytogenes (Di Bonaventura et al., 2008;Lemon, Higgins, & Kolter, 2007;Todhanakasem & Young, 2008). Interestingly, 16 of the 44 genes lmo_0675-lmo_0718 of L. monocytogenes EGD-e that encode for the flagellar apparatus were differentially regulated in the ΔagrD mutant (Supplementary File S1). Although these genes show divergent expression (i.e., some are up-and others down-regulated) in the mutant, we performed motility assays to test if this strain shows altered expression or functionality of flagella. However, no difference in swimming motility was observed between the ΔagrD mutant and the WT or complemented strain at 25°C on 0.1BHI plates containing 0.2% (w/v) agar (Figure 3).
In the absence of other indications about the possible reason for the phenotype of L. monocytogenes EGD-e ΔagrD, we further analyzed the data set of genes differentially expressed in this strain. We reasoned that impaired attachment to the substratum of the mutant and interaction with other bacteria might be involved in the observed phenotype. These processes are mediated by proteins that are either secreted into the environment (exoproteins) or attached to the bacterial cell envelope. In fact, presence of pronase completely abolished biofilm formation of all three tested strains (Appendix Figure A1).
F I G U R E 2 Resistance of L. monocytogenes EGD-e WT (W), EGD-e ΔagrD (Δ), and EGD-e ΔagrD::pIMK2agrD (C) exposed to hypoosmotic conditions (a) or high hydrostatic pressure (b). (a) Bacteria were transferred to 0.1BHI or demineralized H 2 O (dH 2 O) and viability was assessed after 60 min by determining CFU/ml. (b) Bacteria from exponential growth phase were resuspended in 0.1BHI and subjected to HPP at the indicated pressure. Changes in viability are reported as Δlog 10 (CFU/ml) compared to bacterial counts before treatment. Values are mean ± standard deviation of three independent experiments. Statistical analysis was performed by ANOVA with Dunnett's multiple comparisons test with L. monocytogenes EGD-e WT set as control condition F I G U R E 3 Motility of L. monocytogenes EGD-e WT (W), EGD-e ΔagrD (Δ), and EGD-e ΔagrD::pIMK2agrD (C). Representative images and quantification of the diameter of the zone of growth around the inoculation spot of the three strains grown on 0.1BHI soft agar (0.2%). Values are mean ± standard deviation of three experiments with independent precultures. For each preculture and strain at least three growth zones were measured. Statistical analysis was performed by ANOVA with Dunnett's multiple comparisons test with L. monocytogenes EGD-e WT set as control condition Thus, we retrieved the cellular localization of all agr-regulated proteins as annotated on the Listeriomics web page (https://listeriomics. pasteur.fr/Listeriomics/#bacnet.Listeria), which is based on an extensive in silico analysis (Renier, Micheau, Talon, Hébraud, & Desvaux, 2012). A total of 995 genes (34.8%) in the genome and 293 genes (i.e., 41.0%) of the agr-regulated genes of L. monocytogenes EGD-e encode for (predicted) extracytoplasmatic proteins (Table 2). Amongst the 715 agr-dependent genes, 19 (2.7%) encode for exoproteins (i.e., proteins secreted and released into the extracellular environment), 25 (3.5%) for lipoproteins, 27 (3.8%) for cell wall proteins, 187 (26.2%) for integral membrane proteins, and 35 (4.9%) for cytoproteins (i.e., proteins predicted to be secreted via non-classical pathways). None of the groups seems to be markedly overrepresented in the agr-regulated genes. Nevertheless, the percentages of the agr-regulated genes within these groups (except for exoproteins) were comparable or higher compared to the percentage of the respective group on the genome level suggesting that the agr system is involved in the regulation of biosynthesis, structure, and function of the cell envelope. Of note, the agr-regulated genes included 10 genes for internalins or internalin-like proteins, 15 genes for peptidoglycan-associated proteins, and a number of genes for penicillin binding proteins and proteins with (know or presumable) cell wall-hydrolyzing activity (Supplementary File 1). This indicates that the ΔagrD system is involved in regulation of cell envelope proteins that may be relevant for attachment to and interaction with abiotic surfaces as well as amongst bacterial cells.
Altered surface protein profiles may result in changes in the physicochemical properties of the bacterial surface such as charge and hydrophobicity, which were shown to play a role in adhesion and biofilm formation of L. monocytogenes (Di Bonaventura et al., 2008;Takahashi, Suda, Tanaka, & Kimura, 2010). MATH assays performed in xylene revealed that L. monocytogenes EGD-e ΔagrD did not differ in surface hydrophobicity compared to the WT or complemented strain when bacteria were grown in 0.1BHI at 25°C (Figure 4a). Similar results were obtained, when octadecene was used as solvent (data not shown).
Another functional consequence of an altered cell wall composition could be changes in the resistance to cell wall damage. To test this possibility, the resistance of L. monocytogenes EGD-e ΔagrD to treatment with 5 µg/ml lysozyme was tested in 0.1BHI (Figure 4b). Under these conditions, viability of the WT and complemented strain decreased by about 0.5 logs during the first 120 min of lysozyme challenge. More importantly, the sensitivity of the ΔagrD mutant was significantly increased at any time point measured and viable counts were reduced by about 2 logs after 120 min.
Collectively, the obtained results suggest that the biofilm phenotype of L. monocytogenes EGD-e ΔagrD is not a general feature of this mutant but is only relevant under specific conditions. The experimental conditions under which the mutant displays reduced biofilm formation include nutrient limitation and reduced osmolarity. These are the conditions similar to those encountered in difficult to access reservoirs in food processing plants (Carpentier & Cerf, 2011;Ferreira et al., 2014;da Silva & De Martinis, 2013). Thus, the agr system may be important for adaptation and survival of L. monocytogenes at such sites.
The observed phenotype of the ΔagrD mutant is not associated with differences in eDNA abundance, increased lysis in hypoosmotic conditions, flagellar motility, or surface hydrophobicity. It is more likely, that reduced biofilm formation of L. monocytogenes EGD-e ΔagrD is the result of an altered cell envelope proteome, TA B L E 2 Number and percentage of different groups of genes encoding extracytoplasmatic proteins amongst the agr-regulated genes of L. monocytogenes  F I G U R E 4 (a) Surface hydrophobicity and (b) resistance of L. monocytogenes EGD-e WT (W), EGD-e ΔagrD (Δ), and EGD-e ΔagrD::pIMK2agrD (C) exposed to lysozyme (b). (a) Surface hydrophobicity (H [%]) was evaluated using MATH assay. (b) Bacteria from exponential growth phase were resuspended in 0.1BHI containing 5 µg/ml lysozyme and incubated for the indicated time. Changes in viability are reported as Δlog 10 (CFU/ ml) compared to bacterial counts before treatment. Values are mean ± standard deviation of three independent experiments. Statistical analysis was performed by ANOVA with Dunnett's multiple comparisons test with L. monocytogenes EGD-e WT set as control condition (***p < 0.001) which manifests in reduced adhesion to the abiotic surface and/or to neighboring bacteria or the biofilm matrix and, in consequence, increased dispersal. The previously published transcriptional data (Riedel et al., 2009) provided first indications for genes and their products possibly involved in these phenotypes. In further studies, the contribution of these factors to the observed phenotype and their expression levels need to be investigated for example, by qPCR and experiments using knock-out mutants of the respective genes.

ACK N OWLED G EM ENTS
This study was partially funded within the ERA-IB2 consortium

CO N FLI C T O F I NTE R E S T S
The authors declare no conflict of interests.

AUTH O R S CO NTR I B UTI O N
CU conceived the study. MZ, FIB, PC, DB, and LGG carried out experiments. PC, DB, AIN, GMS, and CUR analyzed data. DB, AIN, and CUR drafted the manuscript and all authors contributed to preparing the final version of the manuscript. All authors read and approved the final manuscript.

E TH I C S S TATEM ENT
Not required.

DATA ACCE SS I B I LIT Y
All relevant data are presented in figures, tables, or in Supplementary File 1. Raw data used for preparation of figures will be made avail-

APPENDIX
F I G U R E A 1 Sensitivity of biofilms of L. monocytogenes EGD-e WT (W), EGD-e ΔagrD (Δ), and EGD-e ΔagrD::pIMK2agrD (C) to pronase. Biofilms were grown in BHI or 0.1BHI at 25 or 37°C in the absence (black bars) or presence (white bars) of 1 mg/ml pronase. Biofilm biomass was quantified by crystal violet staining and measuring absorbance at 562 nm (Abs 562nm ) after 24 hr of growth in polystyrene microtiter plates. All values are mean ± standard deviation of three independent experiments