VirR-Mediated Resistance of Listeria monocytogenes against FoodAntimicrobials and Cross-Protection Induced by Exposure to OrganicAcid Salts

Jihun Kang,a Martin Wiedmann,a Kathryn J. Boor,a Teresa M. Bergholza,b

Department of Food Science, Cornell University, Ithaca, New York, USAa; Department of Veterinary and Microbiological Sciences, North Dakota State University, Fargo,North Dakota, USAb

Formulations of ready-to-eat (RTE) foods with antimicrobial compounds constitute an important safety measure against food-borne pathogens such as Listeria monocytogenes. While the efficacy of many commercially available antimicrobial compoundshas been demonstrated in a variety of foods, the current understanding of the resistance mechanisms employed by L. monocyto-genes to counteract these stresses is limited. In this study, we screened in-frame deletion mutants of two-component system re-sponse regulators associated with the cell envelope stress response for increased sensitivity to commercially available antimicro-bial compounds (nisin, lauric arginate, �-polylysine, and chitosan). A virR deletion mutant showed increased sensitivity to allantimicrobials and significantly greater loss of membrane integrity when exposed to nisin, lauric arginate, or �-polylysine (P <0.05). The VirR-regulated operon, dltABCD, was shown to be the key contributor to resistance against these antimicrobial com-pounds, whereas another VirR-regulated gene, mprF, displayed an antimicrobial-specific contribution to resistance. An experi-ment with a �-glucuronidase (GUS) reporter fusion with the dlt promoter indicated that nisin does not specifically induce VirR-dependent upregulation of dltABCD. Lastly, prior exposure of L. monocytogenes parent strain H7858 and the �virR mutant to2% potassium lactate enhanced subsequent resistance against nisin and �-polylysine (P < 0.05). These data demonstrate thatVirRS-mediated regulation of dltABCD is the major resistance mechanism used by L. monocytogenes against cell envelope-dam-aging food antimicrobials. Further, the potential for cross-protection induced by other food-related stresses (e.g., organic acids)needs to be considered when applying these novel food antimicrobials as a hurdle strategy for RTE foods.

Control of Listeria monocytogenes in ready-to-eat (RTE) foodsis an important food safety goal due to the high mortality rate

associated with listeriosis, particularly in susceptible populations,such as pregnant women, the elderly, and those with a compro-mised immune system (1). L. monocytogenes is of particular con-cern for those RTE foods that support growth of this pathogen tohigh levels during refrigerated storage, which can potentially causea life-threatening disease. L. monocytogenes harbors a variety ofstress coping mechanisms that allow it to survive under subopti-mal environmental conditions associated with foods (e.g., acidic,osmotic, and/or temperature stress) (2). The ability of L. monocy-togenes to tolerate and grow under such a wide range of adverseconditions elevates the likelihood of foodborne transmission to ahuman host. Thus, a multipronged approach (e.g., prevention ofpostprocessing contamination and reformulation of RTE foodswith antimicrobials) to limit L. monocytogenes in foods along thefarm-to-fork continuum is critical to reduce the potential forfoodborne illnesses involving this organism (3).

Natural antimicrobials are commonly applied to RTE foods tocontrol foodborne pathogens such as L. monocytogenes (4). Nisin(NIS) is one of the most widely used antimicrobials; it is a bacte-riocin naturally produced by Lactococcus lactis. Other antimicro-bials that have been used to control L. monocytogenes growth morerecently include lauric arginate (LAE; derived from lauric acid,L-arginine, and ethanol), ε-polylysine (EPL; produced by Strepto-myces albulus), and chitosan (CHI; derived from crustacean exo-skeletons). The primary mode of action for NIS is due to the bind-ing to lipid II (a membrane-anchored cell wall precursor) as adocking molecule and subsequent aggregation of nisin moleculesto induce pore formation in the bacterial membrane, leading to

dissipation of the proton motive force (5, 6). Likewise, the pro-posed mechanisms of action for LAE (7, 8), EPL (9–12), and CHI(13, 14) employ the common theme of cell envelope disruptionfollowed by concomitant disturbances in membrane-related cel-lular functions, though the exact molecular mechanisms of actionremain to be further elucidated. From a bacterial perspective, thecell envelope not only provides structure to the cell but also func-tions as the primary barrier to exogenous aggressions as well as avirulence modulator at the pathogen-host interface (15). Thus,maintaining the integrity and function of the cell envelope underfluctuating environmental conditions is critical for ensuring bac-terial survival and transmission.

Sensing and managing of cell envelope stress is typically facil-itated through alternative sigma factors and/or two-componentsystems (TCSs) (16). In the case of TCSs, sensing of a specificinput signal initiates autophosphorylation of a conserved histi-dine residue on a sensor histidine kinase. The phosphoryl group is

Received 27 February 2015 Accepted 21 April 2015

Accepted manuscript posted online 29 April 2015

Citation Kang J, Wiedmann M, Boor KJ, Bergholz TM. 2015. VirR-mediatedresistance of Listeria monocytogenes against food antimicrobials and cross-protection induced by exposure to organic acid salts. Appl Environ Microbiol81:4553–4562. doi:10.1128/AEM.00648-15.

Editor: M. W. Griffiths

Address correspondence to Teresa M. Bergholz, Teresa.bergholz@ndsu.edu.

Copyright © 2015, American Society for Microbiology. All Rights Reserved.

doi:10.1128/AEM.00648-15

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then transferred to an aspartic acid residue of the cognate responseregulator, which causes conformational change in the structure ofthe response regulator, allowing it to function as a transcriptionalactivator (17). L. monocytogenes harbors 15 two-component sys-tems and an orphan response regulator (RR) (18). Of these, 4TCSs (liaRS, lisRK, cesRK, and virRS) have been reported to play acentral role in modulating the cell envelope stress response (19). Anumber of genes previously implicated in L. monocytogenes nisinresistance are also TCSs, such as liaRS (20–22), lisRK (23), andvirRS (24, 25), as well as genes that are part of TCS regulons,including telA (26), mprF (27), anrAB (24), dltABCD (28), andlmo2229 (21, 22). As an abundance of evidence indicates thatTCSs play a central role in the cell envelope stress response and areinduced by cationic antimicrobial peptides (CAMPs) and othercell wall-acting antibiotics (16, 19, 29), we sought to determinewhether these TCSs also have a role in resistance against antimi-crobials that may be used in foods to control L. monocytogenes.Better understanding of resistance mechanisms used by L. mono-cytogenes against these membrane-damaging agents may providefurther insight into an effective hurdle strategy as well as develop-ment of inhibitors targeting these TCSs to improve pathogen con-trol in foods.

MATERIALS AND METHODSBacterial strains, mutant construction, antimicrobials, and growthconditions. All L. monocytogenes strains used in this study are listed inTable 1. Nonpolar deletion mutants were constructed from the parentstrain H7858 by using the splicing by overlap extension (SOE) method(30). All deletions mutants were confirmed by PCR and subsequent se-quencing of the chromosomal copy of the deletion allele. The stock anti-microbial concentrations were 500 �g/ml (NIS), 1,000 �g/ml (LAE),50,000 �g/ml (EPL), and 50,000 �g/ml (CHI), and they were prepared insterile H2O except for CHI, which was prepared in 1% (vol/vol) acetic acidas previously described (31). L. monocytogenes strains were maintained inbrain heart infusion (BHI) broth at �80°C with 15% glycerol. Prior toeach experiment, strains were streaked onto BHI agar and incubated at37°C for 24 h. A single colony was used to inoculate 5 ml BHI, followed byincubation at 37°C for 16 h with shaking (230 rpm). Overnight cultureswere transferred into 10 ml BHI (1:100) and incubated until log phase(optical density at 600 nm [OD600], 0.2 to 0.3) at 7°C.

MIC determinations. The MIC experiment was conducted to deter-mine if the mutants had increased sensitivities to the antimicrobial effects(i.e., bactericidal and/or bacteriostatic) of selected compounds. On theday of the experiment, antimicrobial stock solutions were diluted in sterileH2O, and 100 �l was loaded into a sterile 96-well flat-bottom polystyrenemicrotiter plate (Corning Inc., Corning, NY). Log-phase cultures of L.monocytogenes H7858 and its isogenic response regulator mutants grownat 7°C were diluted 1:10 in double -strength (2�) BHI, and 100 �l wasloaded into corresponding wells by using a multichannel pipette. Themicrotiter plate was covered with an adhesive film (Breathe-Easy; Diver-sified Biotech, Dedham, MA) to prevent contamination, and the OD600

was measured immediately with the Synergy H1 microplate reader(BioTek, Winooski, VT). The microplate was stored at 7°C for 7 days, andthe OD600 was measured again. The MICs were defined as the lowestconcentration that completely inhibited growth (OD600 increase of�0.05) after 7 days of incubation at 7°C (32, 33). MIC determinationswere based on results with 3 biological replicates for all strains.

Membrane integrity assay. The membrane integrity assay utilizes thenucleic acid stains SYTO 9 (green fluorescence) and propidium iodide(red fluorescence). SYTO 9 can penetrate cells with intact or with com-promised membranes, whereas propidium iodide can only penetratethose with compromised membranes. The ratio of green to red fluores-cence thus can be used as an indicator of the relative membrane integrityunder antimicrobial stress. Prior to each experiment, log-phase culturesof L. monocytogenes H7858 and the �virR mutant grown at 7°C werealiquoted into 1.5-ml tubes, and stock solutions of NIS, LAE, EPL, andCHI were transferred 1:1 (500 �l of antimicrobial into 500 �l of cells) intotubes at the previously determined MIC for H7858 �virR except for NIS,which was tested at 0.5 �g/ml. Higher concentrations of NIS resulted inthe saturation of the propidium iodide penetration, possibly due to thestrong pore-forming activity of NIS. Following incubation at 7°C for 24 h,cells (1 ml) were harvested by centrifugation (6,800 � g for 5 min) andresuspended in 1 ml 0.85% NaCl, and the assay was carried out using thesecells according to the manufacturer’s protocol (Live/Dead BacLight bac-terial viability kit; Molecular Probes, Inc., Eugene, OR). Washed cells (100�l) were loaded into a 96-well flat-bottom polystyrene microtiter plate(Corning Inc.), mixed with 100 �l 2� staining solution (mixture of SYTO9 and propidium iodide), and incubated at room temperature for 15 minin the dark. The fluorescence intensity of each well was measured with aSynergy H1 microplate reader (BioTek); SYTO 9 intensity was measuredwith an excitation wavelength of 485 nm and emission wavelength of 530nm, while propidium iodide intensity was measured with an excitationwavelength of 485 nm and emission wavelength of 630 nm. The raw flu-orescence readings were used to quantify the relative dye ratios for eachtreatment. The relative dye ratios were calculated as the ratio of SYTO 9 topropidium iodide and divided by the fluorescence ratio of the untreatedparent strain (for the parent strain relative dye ratio) or by the fluores-cence ratio of untreated H7858 �virR (for the H7858 �virR relative dyeratio). The membrane integrity assay was replicated three times for allstrain and treatment combinations.

Survival of H7858 and the �virR, �dltA, �mprF, and �anrAB mu-tant strains exposed to NIS, LAE, EPL, and CHI. The relative survivalexperiment was performed to determine the contributions of each VirRregulon member to resistance to each antimicrobial compound. Culturesof L. monocytogenes parent strain H7858 and derivatives thereof with in-frame deletions of virR and VirR-regulated genes dltA, mprF, and anrABwere grown to log phase at 7°C as described above. These cultures werediluted in phosphate-buffered saline (PBS) and enumerated on BHI agarplates by using a spiral plater (Autoplate 4000; Spiral Biotech, Inc., Nor-wood, MA) to obtain the initial cell counts (T0). The antimicrobial solu-tions (100 �l) were directly added to cultures to achieve final concentra-tions of 0.5 �g/ml NIS, 20 �g/ml LAE, 500 �g/ml EPL, and 50 �g/ml CHI.These antimicrobial concentrations were selected in order to specificallymeasure bacterial inactivation rather than growth inhibition (i.e., MIC),and antimicrobial concentrations different than those used in the MIC

TABLE 1 Strains and plasmids used in this study

Strain orplasmid Strain alias/relevant genotype Reference

StrainsFSL F6-0366 H7858; parent strain; serotype 4b 66FSL B2-0315 H7858 �liaR 67FSL B2-0377 H7858 �lisR This workFSL B2-0379 H7858 �virR This workFSL B2-0380 H7858 �cesR This workFSL K5-0022 H7858 �virS This workFSL K5-0024 Pdlt-GUS FSL F6-0366 This workFSL K5-0025 Pdlt-GUS FSL B2-0379 (�virR) This workFSL K5-0026 Pdlt-GUS FSL K5-0022 (�virS) This workFSL K5-0027 H7858 �dltA This workFSL K5-0028 H7858 �mprF This workFSL K5-0029 H7858 �anrAB This work

PlasmidspJK1 Pdlt-uidA(GUS) on pPL2 This workpPL2 Integrative shuttle vector (Cmr) 68

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assay were needed to observe differences between the parent strain and themutants. Cells treated with antimicrobials were incubated at 7°C for 24 h,and viable cells were enumerated on BHI agar plates. The BHI agar plateswere incubated at 37°C overnight and colonies were counted using theQ-count colony counter (Spiral Biotech, Inc.). Viable cell counts weretransformed to log10 CFU per milliliter values, and the log reduction incell counts was determined as the difference in the log10 CFU per millilitervalue between T24 and T0 for each strain. The survival experiment wasreplicated three times for all strain and treatment combinations.

Cytochrome c binding assay. A cytochrome c binding assay wasperformed as previously described (34–36) to assess whether deletionof virR and VirR-regulated genes (i.e., dltA and mprF) led to an alteredcell envelope charge. Briefly, cells grown to log phase at 7°C werecollected by centrifugation at 4,000 rpm and washed twice with 20 mMMOPS [3-(N-morpholino)propanesulfonic acid] buffer (pH 7). Thewashed cells were adjusted to a concentration of 108 CFU/ml in thesame buffer, and cytochrome c (Sigma-Aldrich, St. Louis, MO) wasadded to a final concentration of 50 �g/ml. After a 10-min incubationat room temperature, the cell suspension was centrifuged at 13,000rpm for 5 min and then the supernatant absorbance was measured at410 nm. The absorbance of cytochrome c in the absence of cells wascompared to the absorbance of cytochrome c with cells (i.e., maximumunbound cytochrome c) as a measure of relative cytochrome c binding,using the following equation: % cytochrome c bound � 100 � [(OD410

with cells)/(OD410 without cells)] � 100.GUS activity assay. The Pdlt-GUS transcriptional fusion was con-

structed to assay for the activation of VirR upon exposure to NIS (Table1). The quantitative determination of GUS activity directed from theVirR-dependent dlt promoter was conducted as previously described(37). Cells were grown to log phase at 7°C in BHI and then exposed to asublethal concentration (100 ng/ml) of NIS or sterile H2O for 12 h at 7°C.Cells (1 ml) were harvested prior to NIS exposure and following 12 h ofincubation by centrifugation and washed with 1 ml ABlight buffer (60mM K2HPO4, 40 mM KH2PO4, 0.1 M NaCl), followed by resuspension in1 ml ABlight buffer. Aliquots (100 �l) were taken from each sample andenumerated on BHI agar plates supplemented with 10 �g/ml chloram-phenicol. The remaining cells were frozen at �80°C. Prior to GUS mea-surements, frozen cells were thawed and 135 �l of CellLytic B reagent(Sigma-Aldrich) was added to lyse cells for 10 min at room temperature.Lysed cells were loaded into 96-well flat-bottom black polystyrene plates(Corning Inc.) in duplicate, and 20 �l of 4-methylumbelliferyl-�-D-glucuronide hydrate (MUG; Sigma-Aldrich) was added to a final concen-tration of 0.4 mg/ml. For a parallel set of cells, the same volume of H2Owas added instead of MUG, to measure the background fluorescence fromthe cells in the absence of the substrate. The enzymatic fluorescent by-product 4-methylumbelliferone (MU; Sigma-Aldrich) was used to gener-ate a standard curve from which respective GUS activity was inferred. Theenzymatic reaction was stopped after 30 min by the addition of the Stopsolution (1 M Na2CO3). The fluorescence was measured in the SynergyH1 plate reader (BioTek) with an excitation wavelength of 365 nm and anemission wavelength of 460 nm. The background fluorescence for the cellswithout MUG was subtracted from the corresponding treated wells, andthe MU standard curve was used to calculate GUS activity, reported as thenanomolar concentration of MU per log CFU per minute.

Organic acid-induced cross-protection against NIS, LAE, EPL, andCHI. L. monocytogenes H7858 and the �virR mutant grown to log phase at7°C were transferred (1 ml) to 9 ml BHI, BHI plus potassium lactate (PL),or BHI plus sodium diacetate (SD) for final concentrations of 2% PL and0.14% SD. BHI containing no acid (CTRL) and acid-supplemented BHIwere adjusted to pH 6.0 to separate the effects of pH from those of the acidsalts. The initial pH of the BHI treatment media were 7.35 � 0.01, 7.35 �0.01, and 6.65 � 0.06 (means � standard deviations) for plain BHI, BHIplus PL, and BHI plus SD, respectively. Since salts of organic acids wereused, the direct impact on the pH of BHI medium was not as profound aswhen lactic acid or acetic acid are used. L. monocytogenes strains were

exposed to acid conditions for 8 h at 7°C prior to antimicrobial addition atfinal concentrations of 1 �g/ml NIS, 20 �g/ml LAE, 1,000 �g/ml EPL, and100 �g/ml CHI. Acid-exposed cells were enumerated prior to antimicro-bial challenge and following 24 h of incubation at 7°C in the presence ofthe indicated antimicrobial concentrations described above. To examineorganic acid-induced cross-protection, antimicrobial concentrations suf-ficient to cause at least a 1-log reduction of the parent strain were tested foreach antimicrobial.

Statistical analysis. The MICs for the mutants were compared to theMIC of the parent strain by using the Interval package in R-Studio v0.98.1103, which performs log rank tests for interval-censored data basedon nonparametric maximum likelihood estimation of the survival distri-bution (38). For statistical analysis of the results of the membrane integ-rity assay, VirR regulon mutant survival assay, and cytochrome c bindingassay, one-way analysis of variance (ANOVA) was carried out for eachantimicrobial using the relative dye ratio, decrease in the log10 CFU permilliliter, and relative binding, respectively, as the response variable.The strain genotype and replicate were modeled as fixed effects. TheGUS activity assay results were analyzed with a three-way ANOVAmodel using GUS activity as the response variable and the strain geno-type, NIS concentration, assay time, and their interaction as fixed ef-fects. A similar two-way ANOVA model with the decrease in the log10

CFU per milliliter as the response variable and the strain genotype,acid treatment, and their interaction as fixed effects was used to assessthe cross-protection induced by organic acids. The Tukey multiplecomparison procedure was applied to all ANOVA results. Adjusted Pvalues of 0.05 were considered significant. ANOVA models and mul-tiple comparisons were performed using JMP statistical software(JMP10; SAS Institute, Inc., Cary, NC).

RESULTSDeletion of virR increases sensitivity to the food antimicrobialsNIS, LAE, EPL, and CHI at 7°C. To determine whether TCS re-sponse regulators were important for resistance against selectedantimicrobials, we screened in-frame deletion mutants of selectedTCS response regulators (i.e., �liaR, �lisR, �virR, and �cesRstrains) for increased sensitivity using a MIC assay. We specificallyfocused on the role of response regulators rather than their cog-nate histidine kinases to eliminate potential cross talk with otherhistidine kinases or alternative activation mechanisms (25). Incomparison to the parent strain, deletion of virR decreased theMIC by 2- to 8-fold for all antimicrobials tested (P � 0.1), indi-cating that VirR potentially plays a role in resistance against testedantimicrobials (Table 2). In addition to VirR-dependent resis-tance, antimicrobial-specific sensitivity was identified for otherresponse regulators. The MIC of H7858 �liaR was 2-fold lowerthan that of the parent strain under NIS (P � 0.1), whereas theMIC of H7858 �lisR was 2-fold lower than that of the parent strain

TABLE 2 MIC determinations for the parent strain (H7858) and itsisogenic TCS response regulator mutantsa

Antimicrobial

MIC (�g/ml) for indicated strain

H7858�liaRmutant

�lisRmutant

�virRmutant

�cesRmutant

NIS 12.5 6.25 12.5 1.56 12.5LAE 12.5 12.5 6.25 6.25 12.5EPL 100 50 100 25 50CHI 200 200 200 100 200a MICs were determined from three biological replicates and the data shown are themodes. MIC values shown in boldface were statistically different from the MIC for theparent strain (P � 0.1) as determined by the log rank test.

Antimicrobial Resistance of L. monocytogenes

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when grown in the presence LAE (P � 0.1). Although these resultsindicate a potential contribution of LiaR and LisR to NIS and LAEresistance, respectively, further experiments are necessary to as-certain these findings due to minor (i.e., 2-fold) differences in theMICs.

Since H7858 �virR was most sensitive to all tested antimicro-bials, the membrane integrity assay was used to assess whether theit was specifically more sensitive to the membrane-perturbing ef-fect of the antimicrobials. Our results indicated that the mem-brane integrity was compromised upon addition of the antimicro-bial compounds compared to the untreated control membranes(Fig. 1). Deletion of virR resulted in greater loss of membraneintegrity under NIS, LAE, and EPL stress than in the parent strain,as indicated by significantly lower dye ratios in the �virR strain(P 0.05). Interestingly, the loss of membrane integrity underCHI stress was not significantly different for the parent strain andH7858 �virR (Fig. 1). In summary, the data from the MIC deter-minations and membrane integrity assay demonstrate a centralrole of VirR in resistance against food antimicrobials as well as anantimicrobial-specific contribution of other response regulatorsto antimicrobial resistance. Further, the increased sensitivity ofthe �virR mutant is likely associated with the increased loss ofmembrane integrity during exposure to NIS, LAE, and EPL, butnot CHI.

VirR-regulated genes contribute to protection against thebactericidal actions of food antimicrobials. To determine thespecific resistance contribution of VirR-regulated genes againstthe effects of antimicrobials, we compared survival of the parentstrain, the �virR mutant, and deletion mutants of VirR-regulatedgenes (i.e., �dltA, �mprF, and �anrAB strains) under antimicro-bial stress. The functional role of DltABCD and MprF is to reducethe net negative charge of the cell envelope by the modification ofteichoic acids with D-alanine residues or by the modification of

membrane phospholipids with L-lysine, respectively (39). Thesemolecular modifications of cell envelope constituents are essentialfor cationic antimicrobial peptide (CAMP) resistance and viru-lence in L. monocytogenes (27, 28). AnrAB is an ABC transporterwhich also facilitates CAMP resistance, presumably via effluxpumping of toxic compounds (24). As expected, H7858 �virRshowed increased sensitivity to all antimicrobials compared to theparent strain after 24 h exposure at 7°C (Fig. 2). Relative to theparent strain, the mean decrease in cell density of the �virR mu-tant was greater by 5.2 � 0.4, 1.0 � 0.1, 1.5 � 0.1, and 1.9 � 0.1log10 CFU/ml for NIS, LAE, EPL, and CHI, respectively (P 0.05). Similarly, H7858 �dltA was significantly more sensitive toall antimicrobials (P 0.05) than the parent strain. In all cases, thesensitivity of the �dltA mutant was not significantly different fromthat of the �virR strain, indicating that DltABCD is the key com-ponent of VirR-mediated resistance to antimicrobials. While de-letion of mprF also influenced antimicrobial sensitivity, the rela-tive contribution to resistance varied between antimicrobials. Theincreased sensitivity of H7858 �mprF was highest under EPLstress, as that strain had the greatest log reduction in cell numbercompared to results for all other mutants, including H7858 �virR.The mean decrease in cell density of H7858 �mprF under NISstress was greater than that for the parent strain (by 3.0 � 0.4 log10

CFU/ml; P 0.05) but was considerably less than that of the �virRor �dltA mutant (Fig. 2), suggesting that MprF only contributespartially to VirR-mediated resistance against NIS. MprF wasshown to play no significant role in resistance against LAE and arelatively minor role in resistance against CHI stress, as the meandecrease in cell density of H7858 �mprF under CHI stress wasgreater than that for the parent strain by 0.4 � 0.1 log10 CFU/ml(P 0.05). No significant effect of anrAB deletion was observedwith regard to sensitivity against all 4 of the antimicrobials. Takentogether, D-alanylation of teichoic acids facilitated by DltABCDappears to be the most critical resistance determinant againsttested food antimicrobials, while L-lysinylation of membranephospholipids confers various degrees of resistance against differ-

FIG 1 Membrane integrity of the parent strain H7858 and the �virR mutantstrain under NIS, LAE, EPL, or CHI exposure for 24 h at 7°C. The relative dyeratios were calculated as the fluorescence ratio (SYTO9 to propidium iodide)for the antimicrobial-treated parent strain and the �virR mutant divided bythe fluorescence ratio of the untreated parent strain and the �virR mutant.Asterisks indicate the relative dye ratios for H7858 �virR under each antimi-crobial stress that were significantly (P 0.05) different from the ratios for theparent strain. The data represent the means and the standard deviations ofthree biological replicates.

FIG 2 Log decrease in cell density for the parent strain H7858 and the �virR,�dltA, �mprF, and �anrAB mutants exposed to NIS (0.5 �g/ml), LAE (20�g/ml), EPL (500 �g/ml), or CHI (50 �g/ml) for 24 h at 7°C. For each anti-microbial, strains with the same letter were not significantly different fromeach other (P 0.05). The data represent the means and the standard devia-tions of three biological replicates.

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ent antimicrobials, with the most significant contribution againstEPL stress.

Deletion of virR, dltA, and mprF alters the cell envelopecharge of L. monocytogenes. As DltABCD and MprF have pre-viously reported roles in modification of electrostatic charge ofthe cell envelope, relative changes in cell envelope charge in theparent strain and the mutants were measured by quantifyingthe amount of cytochrome c binding (34, 36). Compared to theparent strain, the �virR, �dltA, and �mprF mutants boundsignificantly more cytochrome c, with relative binding levels of18.0% � 2.4%, 16.1% � 5.1%, 15.1% � 1.2%, respectively,whereas the binding affinity of the parent strain was signifi-cantly lower with a relative binding of 3.2% � 3.0% (P 0.05)(Fig. 3). This indicates that deletion of genes with a role in cellsurface charge modification has a profound effect on cell enve-lope charge and plausibly results in altered electrostatic inter-actions between cationic antimicrobial compounds and the an-ionic cell surface.

The presence of NIS does not significantly induce VirR-de-pendent upregulation of dltABCD. To determine whether resis-tance to antimicrobials (e.g., NIS) is the consequence of transcrip-tional regulation of VirR-regulated genes (i.e., dltABCD), wecreated a transcriptional fusion of the promoter of dltABCDwith a GUS gene as a reporter (37). This promoter was selectedto specifically determine VirRS-dependent transcription ofdltABCD, as this operon has been reported to be dependent onVirR (25). As expected, GUS activity was not detected in theabsence of VirR or VirS, confirming that dltABCD is dependenton VirRS for its transcriptional activation. Compared to theinitial level of GUS activity, there was significantly higher GUSactivity (P 0.05) after 12 h at 7°C, even in the absence of NIS(Fig. 4). Though a slightly higher level of GUS activity wasobserved in the presence of 100 ng/ml NIS after 12 h at 7°C, thechange in GUS activity level was not significantly differentfrom the control without NIS. Taken together, the GUS activitydata suggest that NIS does not specifically induce VirR-medi-ated upregulation of dltABCD.

Preexposure to organic acids induces cross-protection in anantimicrobial-dependent manner. Organic acids such as lactateand diacetate may be incorporated into a food formulation aseffective L. monocytogenes growth inhibitors (31, 40, 41). Ideally,the combination of hurdles should exhibit synergistic effects oninactivating or inhibiting growth of undesirable microorganisms.In contrast, there have been reports of a cross-protective phenom-enon, where prior exposure to a stress may increase resistanceagainst a subsequent stress (20, 42). Additionally, acid-inducedtranscriptome profiling in L. monocytogenes has suggested up-regulation of stress- and virulence-related genes, including theVirR regulon, as part of acid stress adaptation (43–45). To test thehypothesis that prior exposure to organic acids induces cross-protection against the selected antimicrobials and is mediated byVirR, we exposed the L. monocytogenes parent strain and H7858�virR to BHI (no acid control), 2% PL, and 0.14% SD, followed bysubsequent challenge with antimicrobial compounds. Our resultsindicated that preexposure to PL increased NIS resistance of boththe parent strain and the �virR mutant; preexposure of the parentstrain cells to PL resulted in no change in cell density when subse-quently challenged with NIS, whereas no prior PL exposure led toa mean decrease in cell density of 0.9 � 0.3 log10 CFU/ml (P 0.05) (Fig. 5). Similarly, PL-exposed �virR cells had a mean de-crease in cell density of 5.7 � 0.2 log10 CFU/ml, compared to cellswith no prior PL exposure, which had a mean decrease in celldensity of 6.6 � 0.2 log10 CFU/ml (P 0.05), indicating that theobserved cross-protection is likely independent of VirR. Prior ex-posure to SD did not have a significant effect on NIS resistance foreither the parent strain or the �virR mutant (Fig. 5). For LAEstress, prior acid exposure had no effect on LAE resistance of theparent strain (P 0.05), whereas both PL and SD had significantprotective effects against subsequent LAE exposure for H7858�virR, with a mean decrease in cell density of 3.2 � 0.0 and 4.3 �0.2 log10 CFU/ml, respectively, compared to 4.7 � 0.1 log10

CFU/ml without acid exposure (P 0.05). Organic acid-inducedcross-protection against EPL was similar to that with NIS; prior

FIG 3 Relative binding of cytochrome c for the parent strain H7858 and the�virR, �dltA, and �mprF mutants grown to log phase at 7°C. Strains with thesame letter were not significantly different from each other (P 0.05). Thedata represent the means and the standard deviations of three biological rep-licates.

FIG 4 GUS activities for dlt promoter-GUS reporter fusions expressed in theparent strain or in the �virR or �virS background in the presence of 100 ng/mlNIS at 7°C. Asterisks indicate significantly (P 0.05) different GUS activitylevels for each genetic background. The data represent the means and thestandard deviations of three biological replicates.

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exposure to PL increased EPL resistance (P 0.05), whereas SDhad no significant effect on subsequent EPL resistance (P 0.05)for either the parent strain or the �virR mutant. The potentialcross-protective effect of organic acids against CHI was less pro-nounced. Prior exposure to PL had no effect on the parent strainand a significant (P 0.05) but numerically limited (0.3 log10

CFU/ml) cross-protective effect on H7858 �virR. Prior exposureto SD slightly increased sensitivity of the parent strain and the�virR mutant (by 0.41 � 0.23 and 0.22 � 0.03 log10 CFU/ml,respectively) to CHI. but this increased sensitivity was also numer-ically limited.

DISCUSSIONThe TCS response regulator VirR plays a key role in resistance toa range of food antimicrobials. In this study, we have identifiedVirR as a critical transcriptional regulator for resistance againstantimicrobials used in foods. Reduced membrane integrity forH7858 �virR under NIS, LAE, and EPL stress suggests that theabsence of VirR renders cells more susceptible to the membrane-perturbing effects of these compounds, likely due to increasedbinding. While H7858 �virR exhibited increased sensitivity toCHI stress, the loss of membrane integrity for that strain was sim-ilar to that for the parent strain under CHI stress. This observationis consistent with a previous report on the CHI mechanism ofaction, in which teichoic acids are proposed to be the main mo-lecular target of CHI, followed by subsequent disruption of mem-brane-related functions (13).

In L. monocytogenes EGD, VirR has been reported to regulate12 genes, and along with VirS, its cognate sensor kinase, this signaltransduction system has been implicated as a critical virulencedeterminant in L. monocytogenes through modulating the interac-tion between L. monocytogenes cells and components of the innateimmune system (25). Further, the VirRS regulon is composed ofgenes with previously ascribed roles in resistance against animal-and plant-derived CAMPs. While VirRS has been studied exten-sively with regard to virulence-related functions in the host andresistance against therapeutic antibiotics, relatively little is knownabout its role in resistance against antimicrobials used in foods.Our findings of the universal contribution of VirR toward antimi-crobial resistance relevant to food preservation further highlightthe important functional role of VirR and its regulon in pathogensurvival and transmission in different environments (e.g., foodsversus human hosts). Since the contribution of VirR in resistanceto antimicrobials used in foods was assessed in a laboratory me-dium, further examination in appropriate food systems is war-ranted. Moreover, the evaluation of potential growth phase-de-pendent effects on VirR-mediated antimicrobial resistance may beof importance, as there may be overlapping mechanisms thatcould contribute to resistance due to the activation of the stressresponse in stationary phase (46).

FIG 5 Log decreases in cell density for the parent strain H7858 and �virRstrains preexposed to no acid (CTRL), 2% potassium lactate (PL), and 0.14%sodium diacetate (SD) for 8 h at 7°C prior to 24 h challenge with NIS, LAE,EPL, and CHI at 7°C. Closed circles and open circles represent the log decreasein cell density for the parent strain and �virR strains, respectively. Asterisksabove and below the dots indicate the log decrease in cell numbers for theacid-exposed parent strain and the �virR mutant, respectively, which weresignificantly (P 0.05) different from results with the group with no prior acidexposure (CTRL). The data represent the means and the standard deviations ofthree biological replicates.

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VirR-regulated dltABCD is the main contributor to resis-tance against bactericidal effects of food antimicrobials. Mem-bers of the VirR regulon (25), such as dltABCD (28, 35, 47), mprF(27, 48, 49), and anrAB (24), are conserved in many Gram-posi-tive bacteria and are important for conferring resistant to a varietyof antimicrobial compounds, including NIS and other cationicpeptides of animal, plant, and microbial origins (39). The relativecontribution of VirR to resistance to each antimicrobial is sum-marized in Fig. 6. The side-by-side comparison of these mutantsagainst the parent strain and the �virR mutant indicated that Dlt-ABCD is the major contributor of VirR-mediated resistance, asH7858 �dltA was as sensitive as H7858 �virR in the presence of all4 antimicrobials tested here. A possible explanation for this obser-vation is that the increased electropositive charge facilitated byDltABCD results in electrostatic repulsion of cationic antimicro-bials, thus preventing antimicrobials from binding to their targets(35, 39, 50). Recently, Saar-Dover et al. further refined this notionwith a newly proposed mechanism of Dlt-mediated increase in cellwall density (51). This resistance mechanism based on cell wallthickening has also been suggested from a transcriptomic analysisof a NIS-resistant Lactococcus lactis strain (50).

MprF shares a functional similarity with DltABCD in that theprotein alters cell envelope charge by modifying the membranelipid phosphatidylglycerol with L-lysine (27, 39). The relative re-sistance contribution by MprF was highly varied among antimi-crobials. The relative contribution of MprF to NIS resistance wasless, compared to VirR, consistent with a previous report, whichshowed that the MIC of H7858 �mprF under NIS stress was 2-foldhigher than that for the �virR strain (24). Our results also dem-onstrated a minor effect of mprF deletion on LAE and CHI resis-

tance. In Gram-positive bacteria, the cell envelope is composed ofthe phospholipid bilayer shrouded by multiple layers of cross-linked murein, which are further decorated with cell wall glyco-polymers, such as teichoic acids (52, 53). Since MprF is responsi-ble for reduced anionicity of the membrane (as opposed to the cellwall), we hypothesize that the presence of DltABCD in the �mprFmutant imparts a sufficient “shielding effect” which can reducelocal concentration of antimicrobials in the vicinity of cell enve-lopes. Interestingly, H7858 �mprF was shown to be more suscep-tible to EPL than was the �virR or �dltA strain. The greater sen-sitivity of H7858 �mprF compared to the �virR mutant suggests apossible coregulation of mprF by other regulators, under EPLstress, despite its initial classification into the VirR regulon (25).

The VirRS signal transduction pathway is not induced byNIS. We also sought to determine whether the presence of NISspecifically induces VirRS-dependent transcriptional regulationof dltABCD. In the absence of VirR or VirS, dltABCD promoteractivity was minimal or negligible, consistent with its reporteddependence on VirRS (25). While there was an obvious increase inGUS activity during 12 h of incubation at 7°C, the increase in GUSactivity due to NIS was not significant, indicating that NIS onlyweakly induces or does not specifically induce VirR-dependentregulation of dltABCD. Although NIS has been shown to inducedlt operon expression in B. subtilis (54) and Clostridium difficile(55), it was shown to be weakly induced in these studies. Further,no significant upregulation of dltA was observed in spontaneousleucocin- or pediocin-resistant mutants of L. monocytogenes, eventhough a higher D-alanine content in teichoic acids was observedin these strains (36). Thus, the VirR-dependent increase in dlttranscription observed here appears to be the result of a growth-

FIG 6 Model of VirR-mediated resistance to food antimicrobials, based on our data and previously proposed modes of action for these antimicrobials. The figurewas adapted from a previous publication (53). The L. monocytogenes cell envelope consisting of the cell wall (gray) and the cytoplasmic membrane (below grayarea) is shown. Teichoic acids that are anchored to the cell wall and the cytoplasmic membrane represent wall teichoic acids and lipoteichoic acids, respectively.D-Alanylation of teichoic acids (mediated by DltABCD) and L-lysinylation of membrane phospholipids (mediated by MprF) are indicated by the letters A or Lin circles, respectively. Letters below each strain designate the phenotype when exposed to each antimicrobial as inferred from the survival assays: R, resistant; S,sensitive; MS, most sensitive. Potential electrostatic repulsion or exclusion of antimicrobials facilitated by teichoic acid as well as phospholipid modifications areindicated by deflected arrows with a horizontal line, whereas a plain arrow indicates antimicrobial penetration to the target site. The combination of two arrowsindicates partial resistance provided by the specific modification.

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dependent effect rather than a specific induction by NIS. Growthphase-dependent regulation of dltABCD and mprF has also beenshown in Staphylococcus aureus, in which the expression of thesegenes is upregulated during the exponential phase, whereas theopposite occurs during the stationary phase (34).

Preexposure to potassium lactate increases subsequent resis-tance to nisin and �-polylysine. L. monocytogenes typically en-counters multiple stresses in a food environment, and adaptationto a stress condition may lead to cross-protection against a subse-quent stress. For example, osmotic stress and acid stress have beenreported to induce cross-protection against NIS in L. monocyto-genes (20, 42). Nevertheless, this potential cross-protective effecthas not been assessed for other relevant food antimicrobial com-pounds. Further, the evidence of VirR regulon induction underacid stress prompted us to hypothesize that a potential cross-pro-tective effect would be VirR mediated (43–45). Our results indi-cated that exposure to 2% PL increased subsequent NIS and EPLresistance of both the parent strain and the �virR mutant. NISresistance has previously been associated with alterations in cyto-plasmic membrane fatty acid compositions, reflecting more rigidmembrane fluidity (42, 56, 57). Further, alterations in membranefatty acid profiles (e.g., higher-branched versus straight-chainfatty acid ratio) during growth at 10°C were associated with morefluid membranes and resulted in increased NIS sensitivity in L.monocytogenes (58). Acid adaptation in L. monocytogenes also al-ters membrane fatty acid composition that is suggestive of morerigid membrane fluidity, which may act as a defense mechanismby restricting the diffusion of acids across the membrane (59, 60).Thus, the increased rigidity of the cytoplasmic membrane duringadaptation to potassium lactate could at least partially increasesubsequent resistance against NIS.

Transcriptional profiling of L. monocytogenes during adapta-tion to lactic, acetic, or hydrochloric acid has also previously in-dicated a consistent pattern of alterations in cell membrane, in-cluding downregulation of genes involved in branched-chain fattyacid synthesis (43). Compared to SD, PL was shown to have astronger antilisterial activity as well as a more pronounceddownregulation of branched-chain fatty acid-related genes(43), even when the concentration of the undissociated form ofSD was higher (3.7 mmol/liter for SD, compared to 1.4 mmol/liter for PL), which determines the primary inhibitory effects(60). In our study, the estimated concentration of the undiss-coiated form of PL and SD were 1.1 mmol/liter and 0.54 mmol/liter, respectively, obtained using the Henderson-Hasselbalchequation. Thus, a relatively stronger effect of PL can be ex-pected versus SD, which may explain the profound PL-inducedcross-protective effect. As the mode of action for EPL also in-volves cell membrane disruption (9, 11, 12), it is reasonable tospeculate that the membrane fatty acid modification is linkedwith PL-induced cross-protection against EPL.

The preexposure of L. monocytogenes to PL induced cross-pro-tection against LAE only for H7858 �virR but not the parentstrain. Consistent with the observation that dltABCD is the majorresistance determinant for LAE, it may be the case that the regu-lation of cell wall glycopolymers in the parent strain provides pro-tection against LAE, whereas in the absence of VirR-dependentregulation of cell wall structures, acid-induced changes in the cy-toplasmic membrane afford additional protection. In the case ofCHI, relatively minor effects of either acid on subsequent CHIresistance are consistent with a previous report that the primary

molecular target of CHI is likely teichoic acids with subsequentextraction of membrane lipids (13). It is worth noting that theincreased sensitivity of the �virR strain under CHI stress (100�g/ml) was not as evident compared to the survival experimentwith 50 �g/ml CHI (Fig. 2). This could be due to the concentra-tion-dependent saturation of the bactericidal effect of CHI, uponwhich there is no additional lethality, even with increasing con-centrations (61).

Conclusions. This study shows a critical role of the L. monocy-togenes two-component system response regulator VirR in resis-tance against antimicrobials intended for food preservation. Im-proved understanding of resistance mechanisms may facilitate thedevelopment of effective hurdle strategies utilizing different mo-lecular targets (33). Additionally, the potential for cross-protec-tion induced by a food-relevant stress (e.g., organic acids) shouldbe taken into consideration to avoid unintended overestimationof preservative strength in a food system (20, 42). A mechanisticunderstanding of the interaction between antimicrobials and thetarget pathogen also facilitates better delineation of how otherfood constituents may enhance (57) or decrease (62) the antimi-crobial efficacy. Lastly, identification and characterization of TCSswith respect to stress tolerance and virulence have potential im-plications in development of better control strategies for thispathogen as novel antimicrobial agents may be developed to targetthese molecular targets (63, 64). In light of recent advances inhigh-throughput sequencing technologies and decreasing costs,future efforts in understanding global changes in the transcrip-tome of L. monocytogenes under antimicrobial challenge can beexpected to provide a more comprehensive view of bacterial stressresponse and resistance mechanisms (65).

ACKNOWLEDGMENTS

This work was supported by New York Sea Grant R/SHH-16, fundedunder award NA07OAR4170010 from the National Sea Grant CollegeProgram of the U.S. Department of Commerce’s National Oceanic andAtmospheric Administration, to the Research Foundation of the StateUniversity of New York and by Agriculture and Food Research Initiativegrant 2010-65201-20575 from the U.S. Department of Agriculture, Na-tional Institute of Food and Agriculture, Food Safety Program.

We thank Barbara Bowen and Rebecca Schmidt for constructing thedeletion mutants used in this study and Matthew Stasiewicz for assistancewith the analysis of MIC data.

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