AEM Accepted Manuscript Posted Online 24 April 2015 Appl. Environ. Microbiol. doi:10.1128/AEM.00648-15 Copyright © 2015, American Society for Microbiology. All Rights Reserved.

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VirR-mediated resistance of Listeria monocytogenes against food antimicrobials and cross-

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protection induced by exposure to organic acid salts

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Jihun Kang1, Martin Wiedmann1, Kathryn J. Boor1, Teresa M. Bergholz1,2*

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Department of Food Science, Cornell University, Ithaca, NY 14853

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Department of Veterinary and Microbiological Sciences, North Dakota State University, Fargo,

ND 58108

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Running title: Antimicrobial resistance mechanism in L. monocytogenes

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*

Corresponding author

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Mailing address:

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PO 6050 Dept 7690

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North Dakota State University

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Fargo, ND 58108

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Phone: 701-231-7692

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Email: [email protected]

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ABSTRACT

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Formulation of ready-to-eat (RTE) foods with antimicrobial compounds constitutes an important

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safety measure against foodborne pathogens such as Listeria monocytogenes. While the efficacy

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of many commercially-available antimicrobial compounds has been demonstrated in a variety of

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foods, current understanding of the resistance mechanisms employed by L. monocytogenes to

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counteract these stresses is limited. In this study, we screened in-frame deletion mutants of two-

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component system response regulators associated with cell envelope stress response for

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increased sensitivity to commercially-available antimicrobial compounds (nisin, lauric arginate,

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ε-polylysine, and chitosan). A virR deletion mutant showed increased sensitivity to all

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antimicrobials and significantly greater loss of membrane integrity when exposed to nisin, lauric

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arginate, and ε-polylysine (p < 0.05). The VirR-regulated operon, dltABCD, was shown to be the

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key contributor to resistance against these antimicrobial compounds, whereas another VirR-

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regulated gene mprF displayed antimicrobial-specific contribution to resistance. GUS reporter

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fusion with the dlt promoter indicated that nisin does not specifically induce VirR-dependent

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upregulation of dltABCD. Lastly, prior exposure of L. monocytogenes parent strain H7858 and

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ΔvirR to 2% potassium lactate enhanced subsequent resistance against nisin and ε-polylysine (p

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< 0.05). These data demonstrate that VirRS-mediated regulation of dltABCD is the major

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resistance mechanism used by L. monocytogenes against cell envelope-damaging food

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antimicrobials. Further, the potential for cross-protection induced by other food-related stresses

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(e.g., organic acids) needs to be considered when applying these novel food antimicrobials as

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hurdle strategy in RTE foods.

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INTRODUCTION

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Control of Listeria monocytogenes in ready-to-eat (RTE) foods is an important food safety goal

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due to the high mortality rate associated with listeriosis, particularly in susceptible populations

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such as pregnant women, the elderly, and those with a compromised immune system (1). L.

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monocytogenes is of particular concern for those RTE foods that support growth of this pathogen

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to high levels during refrigerated storage, which can potentially cause a life-threatening disease.

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L. monocytogenes harbors a variety of stress-coping mechanisms that allow it to survive in

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suboptimal environmental conditions associated with foods (e.g., acidic, osmotic, temperature

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stress) (2). The ability of L. monocytogenes to tolerate and grow under such a wide range of

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adverse conditions elevates the likelihood of foodborne transmission to a human host. Thus, a

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multi-pronged approach (e.g., prevention of post-processing contamination and reformulation of

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RTE foods with antimicrobials) to limit the number of L. monocytogenes in foods along the

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farm-to-fork continuum is critical to reduce the potential for foodborne illnesses (3).

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Natural antimicrobials are commonly applied to RTE foods to control foodborne pathogens such

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as L. monocytogenes (4). Nisin (NIS) is one of the most widely used antimicrobials, which is a

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bacteriocin naturally produced by Lactococcus lactis. Other antimicrobials that have been used

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to control L. monocytogenes growth more recently include lauric arginate (LAE; derived from

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lauric acid, L-arginine, and ethanol), ε-polylysine (EPL; produced by Streptomyces albulus), and

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chitosan (CHI; derived from crustacean exoskeletons). The primary mode of action for NIS is

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due to the binding to Lipid II (membrane-anchored cell-wall precursor) as a ‘docking molecule’

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and subsequent aggregation of nisin molecules to induce pore formation in the bacterial

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membrane, leading to dissipation of the proton motive force (5, 6). Likewise, the proposed

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mechanism of action for LAE (7, 8), EPL (9-12), and CHI (13, 14) employ the common theme of 3

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cell envelope disruption followed by concomitant disturbances in membrane-related cellular

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functions, albeit exact molecular mechanism of action remains to be further elucidated. From a

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bacterial perspective, the cell envelope not only provides structure to the cell, but also functions

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as the primary barrier to exogenous aggressions as well as virulence modulator at the pathogen-

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host interface (15). Thus, maintaining the integrity and function of the cell envelope under

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fluctuating environmental conditions is critical for ensuring bacterial survival and transmission.

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Sensing and managing of a cell envelope stress is typically facilitated through alternative sigma

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factors and/or two-component systems (TCSs) (16). In the case of TCSs, sensing of a specific

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input signal initiates autophosphorylation of a conserved histidine residue on a sensor histidine

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kinase. The phosphoryl group is then transferred to an aspartic acid residue of the cognate

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response regulator, which causes conformational change in the structure of the response

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regulator, allowing it to function as a transcriptional activator (17). L. monocytogenes harbors 15

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two-component systems and an orphan response regulator (RR) (18). Of these, 4 TCSs (liaRS,

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lisRK, cesRK, and, virRS) have been reported to play a central role in modulating cell envelope

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stress response (19). A number of genes previously implicated in L. monocytogenes nisin

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resistance are also TCSs such as liaRS (20-22), lisRK (23), virRS (24, 25), as well as genes that

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are part of TCS regulons including, telA (26), mprF (27), anrAB (24), dltABCD (28), and

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lmo2229 (21, 22). As an abundance of evidence indicate that TCSs play a central role in the cell

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envelope stress response and are induced by cationic antimicrobial peptides (CAMPs) and other

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cell wall-acting antibiotics (16, 19, 29), we sought to determine whether these TCSs also have a

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role in resistance against antimicrobials that may be used in foods to control L. monocytogenes.

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Better understanding of resistance mechanisms used by L. monocytogenes against these

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membrane-damaging agents may provide further insight on effective hurdle strategy as well as

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development of inhibitors targeting these TCSs to improve the pathogen control in foods.

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MATERIALS AND METHODS Bacterial strains, mutant construction, antimicrobials, and growth conditions. All L.

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monocytogenes strains used in this study are listed in Table 1. Non-polar deletion mutants were

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constructed from the parent strain H7858 using the splicing-by-overlap-extension (SOE) method

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(30). All deletions mutants were confirmed by PCR and subsequent sequencing of the

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chromosomal copy of the deletion allele. The stock antimicrobial concentrations were 500 µg/ml

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(NIS), 1000 µg/ml (LAE), 50000 µg/ml (EPL), and 50000 µg/ml (CHI) and were prepared in

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sterile H2O except CHI which was prepared in 1% (v/v) acetic acid as previously described (31).

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L. monocytogenes strains were maintained in Brain Heart Infusion (BHI) broth at -80oC with 15%

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glycerol. Prior to each experiment, strains were streaked onto BHI agar and incubated at 37oC for

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24 h. A single colony was used to inoculate 5 ml BHI, followed by incubation at 37oC for 16 h

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with shaking (230 rpm). Overnight cultures were transferred into 10 ml BHI (1:100) and

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incubated until log phase (OD600 = 0.2-0.3) at 7oC.

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Minimum Inhibitory Concentration (MIC) determination. The MIC experiment was

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conducted to determine if the mutants had increased sensitivity to the antimicrobial effects (i.e.,

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bactericidal and/or bacteriostatic) of selected compounds. On the day of the experiment,

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antimicrobial stock solutions were diluted in sterile H2O and 100 µl was loaded into a sterile 96-

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well flat-bottom polystyrene microtiter plate (Corning Inc., Corning, NY). Log phase cultures of

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L. monocytogenes H7858 and its isogenic response regulator mutants grown at 7oC were diluted 5

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1:10 in double strength (2X) BHI and 100 µl was loaded into corresponding wells using a multi-

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channel pipette. The microtiter plate was covered with an adhesive film (Breathe-Easy®,

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Diversified Biotech, Dedham, MA) to prevent contamination and the optical density (OD600) was

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measured immediately with the Synergy H1 microplate reader (BioTek, Winooski, VT). The

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microplate was stored at 7oC for 7 d and the OD600 was measured again. The MICs were defined

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as the lowest concentration that completely inhibited growth (OD600 increase ≤ 0.05) after 7 d

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incubation at 7oC (32, 33). MIC determination was based on 3 biological replicates for all strains.

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Membrane integrity assay. The assay utilizes the nucleic acid stains SYTO 9 (Green

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fluorescent) and propidium iodide (Red fluorescent). SYTO 9 can penetrate cells with intact as

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well as compromised membranes, whereas propidium iodide can only penetrate those with

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compromised membranes. The ratio of the green to red fluorescence thus can be used as an

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indicator of the relative membrane integrity under antimicrobial stress. Prior to each experiment,

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log phase cultures of L. monocytogenes H7858 and ΔvirR grown at 7oC were aliquoted into 1.5

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ml tubes and stock solutions of NIS, LAE, EPL, and CHI were transferred 1:1 (500 µl

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antimicrobial into 500 µl cells) into tubes at the previously determined MIC concentration for

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ΔvirR except for NIS which was tested at 0.5 µg/ml. Higher concentrations of NIS resulted in the

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saturation of the propidium iodide penetration, possibly due to the strong pore-forming activity

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of NIS. Following incubation at 7oC for 24 h, cells (1 ml) were harvested by centrifugation

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(6,800 X g for 5 min), resuspended in 1 ml 0.85% NaCl, and the assay was carried out using

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these cells according to the manufacturer’s protocol (Live/Dead BacLight Bacterial Viability kit,

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Molecular Probes, Inc., Eugene, OR). Washed cells (100 µl) were loaded into a 96-well flat-

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bottom polystyrene microtiter plate (Corning Inc.), mixed with 100 µl 2X staining solution 6

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(mixture of SYTO 9 and propidium iodide), and incubated at room temperature for 15 min in the

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dark. The fluorescence intensity of each well was measured with a Synergy H1 microplate reader

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(BioTek); SYTO 9 intensity was measured with the excitation wavelength of 485 nm and

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emission wavelength of 530 nm while propidium iodide intensity was measured with the

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excitation wavelength of 485 nm and emission wavelength of 630 nm. The raw fluorescence

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readings were used to quantify the relative dye ratios for each treatment. The relative dye ratios

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were calculated as the ratio of SYTO 9 to propidium iodide and divided by the fluorescence ratio

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of the untreated parent strain (for the parent strain relative dye ratio) or by the fluorescence ratio

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of untreated ΔvirR (for ΔvirR relative dye ratio). The membrane integrity assay was replicated

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three times for all strain and treatment combinations.

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Survival of H7858, ∆virR, ∆dltA, ∆mprF, and ∆anrAB exposed to NIS, LAE, EPL,

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and CHI. The relative survival experiment was performed to determine the contributions of each

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VirR regulon member to resistance to each antimicrobial compound. Cultures of L.

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monocytogenes parent strain H7858 and in-frame deletion mutants of virR and VirR-regulated

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genes dltA, mprF, and anrAB were grown to log phase at 7oC as described above. These cultures

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were diluted in phosphate-buffered saline (PBS) and enumerated on BHI agar plates using a

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spiral plater (Autoplate 4000, Spiral Biotech, Inc., Norwood, MA) to obtain the initial cell counts

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(T0). The antimicrobial solutions (100 µl) were directly added to cultures to achieve the final

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concentration of 0.5 µg/ml NIS, 20 µg/ml LAE, 500 µg/ml EPL, and 50 µg/ml CHI. These

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antimicrobial concentrations were selected in order to specifically measure bacterial inactivation

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rather than growth inhibition (i.e., MIC), and antimicrobial concentrations different than those

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used in the MIC assay were needed to observe differences between the parent strain and the 7

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mutants. Cells treated with antimicrobials were incubated at 7oC for 24 h and viable cells were

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enumerated on BHI agar plates. The BHI agar plates were incubated at 37oC overnight and

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colonies were counted using the Q-count colony counter (Spiral Biotech, Inc.). Viable cell counts

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were transformed to log10CFU/ml and the log reduction in cell counts was determined as the

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difference in log10CFU/ml between T24 and T0 for each strain. The survival experiment was

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replicated three times for all strain and treatment combinations.

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Cytochrome c binding assay. Cytochrome c binding assay was performed as previously

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described (34-36) to assess whether deletion of virR and VirR-regulated genes (i.e., dltA and

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mprF) leads to altered cell envelope charge. Briefly, cells grown to log phase at 7oC were

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collected by centrifugation at 4,000 rpm and washed twice with 20 mM MOPS (3-(N-

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morpholino)propanesulfonic acid) buffer (pH 7). The washed cells were adjusted to a

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concentration of 108 CFU/ml in the same buffer and cytochrome c (Sigma-Aldrich, St. Louis,

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MO) was added to a final concentration of 50 µg/ml. After 10 min incubation at room

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temperature, the cell suspension was centrifuged at 13,000 rpm for 5 min and the supernatant

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absorbance was measured at 410 nm. The absorbance of cytochrome c in the absence of cells

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was compared to the absorbance of cytochrome c with cells (i.e., maximum unbound cytochrome

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c) as a measure of relative cytochrome c binding using the following equation:

% cytochrome bound = 100 −

OD OD

,

× 100

,

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GUS (β-glucuronidase) activity assay. The Pdlt-GUS transcriptional fusion was

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constructed to assay for the activation of VirR upon exposure to NIS (Table 1). The quantitative

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determination of GUS activity directed from VirR-dependent dlt promoter was conducted as

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previously described (37). Cells were grown to log phase at 7oC in BHI and were then exposed to

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a sublethal concentration (100 ng/ml) of NIS or sterile H2O for 12 h at 7oC. Cells (1 ml) were

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harvested prior to NIS exposure and following 12 h incubation by centrifugation and washed

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with 1 ml ABlight buffer (60 mM K2HPO4, 40 mM KH2PO4, 0.1 M NaCl), followed by

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resuspension in 1 ml ABlight buffer. Aliquotes (100 µl) were taken from each sample and

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enumerated on BHI agar plates supplemented with 10 µg/ml chloramphenicol. The remaining

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cells were frozen at -80oC. Prior to GUS measurements, frozen cells were thawed and 135 µl of

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CellLytic B reagent (Sigma-Aldrich) was added to lyse cells for 10 min at room temperature.

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Lysed cells were loaded into 96-well flat-bottomed black polystyrene plates (Corning Inc.) in

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duplicate and 20 µl of MUG (Sigma-Aldrich) was added for a final concentration of 0.4 mg/ml.

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For a parallel set of cells, the same volume of H2O was added instead of MUG to measure the

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background fluorescence from the cells in the absence of the substrate. The enzymatic

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fluorescent byproduct 4-methylumbelliferone (MU; Sigma-Aldrich) was used to generate a

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standard curve from which respective GUS activity was inferred. The enzymatic reaction was

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stopped after 30 min by the addition of the Stop solution (1 M Na2CO3). The fluorescence was

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measured in the Synergy H1 plate reader (BioTek) with the excitation wavelength of 365 nm and

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the emission wavelength of 460 nm. The background fluorescence from the cells without MUG

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was subtracted from the corresponding wells and the MU standard curve was used to calculate

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GUS activity, reported as nM MU/log CFU/min.

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Organic acid-induced cross-protection against NIS, LAE, EPL, and CHI. L.

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monocytogenes strains H7858 and ΔvirR grown to log phase at 7oC were transferred (1 ml) to 9 9

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ml BHI, BHI + potassium lactate (PL), and sodium diacetate (SD) for the final concentration of 2%

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PL and 0.14% SD. BHI containing no acid (CTRL) and acid-supplemented BHI were all

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adjusted to pH 6.0 to separate the effects of pH from that of the acid salts. The initial pH of the

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BHI treatment media were 7.35 ± 0.01, 7.35 ± 0.01, 6.65 ± 0.06 for plain BHI, BHI + PL, and

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BHI + SD, respectively. Since the salts of organic acids were used, the direct impact on the pH

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of BHI media was not as profound as using lactic acid or acetic acid. L. monocytogenes strains

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were exposed to acid conditions for 8 h at 7oC prior to antimicrobial addition at the final

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concentration of 1 µg/ml NIS, 20 µg/ml LAE, 1000 µg/ml EPL, 100 µg/ml CHI. Acid-exposed

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cells were enumerated prior to antimicrobial challenge and following 24 h incubation at 7oC in

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the presence of indicated antimicrobial concentrations as described above. To examine organic

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acid-induced cross protection, antimicrobial concentrations sufficient to cause at least 1 log

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reduction of the parent strain were tested for each antimicrobial.

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Statistical analysis. The MIC of the mutants was compared to the MIC of the parent

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strain using the ‘Interval’ package in R-Studio v 0.98.1103, which performs logrank tests for

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interval-censored data based on non-parametric maximum likelihood estimation of the survival

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distributions (38). For statistical analysis of the membrane integrity assay, VirR regulon mutant

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survival assay, and cytochrome c binding assay, one-way analysis of variance (ANOVA) was

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carried out for each antimicrobial using the relative dye ratio, decrease in log10CFU/ml, and

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relative binding, respectively, as the response variable. The strain genotype and replicate were

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modeled as fixed effects. GUS activity assay was analyzed with a three-way ANOVA model

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using GUS activity as the response variable and the strain genotype, NIS concentration, assay

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time, and their interaction as fixed effects. Similar two-way ANOVA model with the decrease in 10

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log10CFU/ml as the response variable and the strain genotype, acid treatment, and their

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interaction as fixed effects was used to assess the cross-protection induced by organic acids. The

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Tukey multiple comparison procedure was applied to all ANOVA results. Adjusted p values of


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0.05) whereas both PL and SD had significant protective effect against subsequent LAE

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exposure for ΔvirR with a mean decrease in cell density of 3.2 ± 0.0 and 4.3 ± 0.2 log10CFU/ml,

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respectively, compared to 4.7 ± 0.1 log10CFU/ml without acid exposure (p < 0.05). Organic acid-

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induced cross-protection against EPL was similar to NIS; prior exposure to PL increased EPL

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resistance (p < 0.05) whereas SD had no significant effect on subsequent EPL resistance (p >

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0.05) for both the parent strain and ΔvirR. The potential cross-protective effect of organic acids

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against CHI was less pronounced. Prior exposure to PL had no effect on the parent strain and

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significant (p < 0.05) but numerically limited (0.3 log10CFU/ml) cross-protective effect on ΔvirR.

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Prior exposure to SD slightly increased sensitivity of the parent strain and ΔvirR (by 0.41 ± 0.23

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and 0.22 ± 0.03 log10CFU/ml, respectively) to CHI but this increased sensitivity was also

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numerically limited.

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DISCUSSION

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The TCS response regulator VirR plays a key role in resistance to a range of food

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antimicrobials. In this study, we have identified VirR as a critical transcriptional regulator for

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resistance against antimicrobials used in foods. Reduced membrane integrity for ΔvirR under

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NIS, LAE, and EPL stress suggests that the absence of VirR renders cells more susceptible to

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membrane-perturbing effects of these compounds, likely due to increased binding. While ΔvirR

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exhibited increased sensitivity to CHI stress, the loss of membrane integrity for ΔvirR was

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similar to the parent strain under CHI stress. This observation is consistent with a previous report

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on CHI mechanism of action in which teichoic acids were proposed to be the main molecular

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target of CHI, followed by subsequent disruption of membrane-related functions (13).

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In L. monocytogenes EGD, VirR has been reported to regulate 12 genes, and along with VirS, its

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cognate sensor kinase, this signal transduction system has been implicated as a critical virulence

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determinant in L. monocytogenes through modulating the interaction between L. monocytogenes

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cells and components of the innate immune system (25). Further, the VirRS regulon is composed 16

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of genes with previously ascribed roles in resistance against animal- and plant-derived cationic

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antimicrobial peptides (CAMP). While VirRS has been studied extensively in regard to

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virulence-related functions in the host and resistance against therapeutic antibiotics, relatively

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little is known about its role in resistance against antimicrobials used in foods. Our findings of

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the universal contribution of VirR toward antimicrobial resistance relevant to food preservation

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further highlight the important functional role of VirR and its regulon in pathogen survival and

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transmission in different environments (e.g., foods vs human host). Since the contribution of

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VirR in resistance to antimicrobials used in foods was assessed in a laboratory medium, further

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examination in appropriate food systems is warranted. Moreover, the evaluation of potential

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growth phase-dependent effects on VirR-mediated antimicrobial resistance may be of importance

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as there may be overlapping mechanisms that could contribute to resistance due to the activation

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of the stress response in stationary phase (46).

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VirR-regulated dltABCD is the main contributor to resistance against bactericidal

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effects of food antimicrobials. Members of the VirR regulon (25) such as dltABCD (28, 35, 47),

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mprF (27, 48, 49), and anrAB (24) are conserved in many Gram positive bacteria and are

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important for conferring resistant to a variety of antimicrobial compounds including NIS and

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other cationic peptides of animal, plant, and microbial origin (39). The relative contribution of

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VirR to resistance to each antimicrobial is summarized in Figure 6. The side-by-side comparison

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of these mutants against the parent strain and ΔvirR indicated that DltABCD is the major

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contributor of VirR-mediated resistance, as ΔdltA was as sensitive as ΔvirR in the presence of all

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4 antimicrobials tested here. A possible explanation for this observation is that the increased

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electropositive charge facilitated by DltABCD results in electrostatic repulsion of cationic

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antimicrobials, thus preventing antimicrobials from binding to their targets (35, 39, 50). Recently, 17

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Saar-Dover et al. has further refined this notion with a newly proposed mechanism of Dlt-

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mediated increase in cell wall density (51). This resistance mechanism based on cell-wall

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thickening has also been suggested from transcriptomic analysis of a NIS-resistant Lactococcus

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lactis strain (50).

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MprF shares a functional similarity with DltABCD in that the protein alters cell envelope charge

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by modifying the membrane lipid phosphatidylglycerol with L-lysine (27, 39). The relative

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resistance contribution by MprF was highly varied among antimicrobials. The relative

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contribution of MprF to NIS resistance was less, compared to VirR, consistent with a previous

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report, which showed that the MIC of ΔmprF under NIS stress was 2-fold higher compared to

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ΔvirR (24). Our results also demonstrated a minor effect of mprF deletion on LAE and CHI

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resistance. In Gram positive bacteria, the cell envelope is composed of the phospholipid bilayer

391

shrouded by multiple layers of cross-linked murein which are further decorated with cell wall

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glycopolymers such as teichoic acids (52, 53). Since MprF is responsible for reduced anionicity

393

of the membrane (as opposed to the cell wall), we hypothesize that the presence of DltABCD in

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ΔmprF imparts a sufficient ‘shielding effect’ which can reduce local concentration of

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antimicrobials in the vicinity of cell envelopes. Interestingly, ΔmprF was shown to be more

396

susceptible to EPL compared to ΔvirR or ΔdltA. The greater sensitivity of ΔmprF compared to

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ΔvirR suggests a possible co-regulation of mprF by other regulators, under EPL stress, despite its

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initial classification into the VirR regulon (25).

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VirRS signal transduction pathway is not induced by NIS. We also sought to

400

determine whether the presence of NIS specifically induces VirRS-dependent transcriptional

401

regulation of dltABCD. In the absence of VirR or VirS, dltABCD promoter activity was minimal

402

or negligible, consistent with its reported dependence on VirRS (25). While there was an obvious 18

403

increase in GUS activity during 12 h incubation at 7oC, the increase in GUS activity due to NIS

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was not significant, indicating that NIS only weakly induces or does not specifically induce

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VirR-dependent regulation of dltABCD. Although NIS has been shown to induce dlt operon

406

expression in B. subtilis (54) and Clostridium difficile (55), it was shown to be weakly induced in

407

these studies. Further, no significant up-regulation of dltA was observed in spontaneous leucocin-

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or pediocin-resistant mutants of L. monocytogenes, even though a higher D-alanine content in

409

teichoic acids was observed in these strains (36). Thus, the VirR-dependent increase in dlt

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transcription observed here appears to be the result of a growth-dependent effect rather than a

411

specific induction by NIS. Growth phase-dependent regulation of dltABCD and mprF has also

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been shown in Staphylococcus aureus in which the expression of these genes is upregulated

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during the exponential phase whereas the opposite occurs during the stationary phase (34).

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Pre-exposure to potassium lactate increases subsequent resistance to nisin and ε-

415

polylysine. L. monocytogenes typically encounters multiple stresses in a food environment and

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adaptation to a stress condition may lead to cross-protection against a subsequent stress. For

417

example, osmotic stress and acid stress have been reported to induce cross protection against NIS

418

in L. monocytogenes (20, 42). Nevertheless, this potential cross-protective effect has not been

419

assessed for other relevant food antimicrobial compounds. Further, the evidence of VirR-regulon

420

induction under acid stress prompted us to hypothesize that a potential cross-protective effect

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would be VirR-mediated (43-45). Our results indicated that exposure to 2% PL increased

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subsequent NIS and EPL resistance of both the parent strain and ΔvirR. NIS resistance has

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previously been associated with alterations in cytoplasmic membrane fatty acid compositions,

424

reflecting more rigid membrane fluidity (42, 56, 57). Further, alterations in membrane fatty acid

425

profiles (e.g., higher branched to straight chain fatty acid ratio) during growth at 10oC were 19

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associated with more fluid membrane and resulted in increased NIS sensitivity in L.

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monocytogenes (58). Acid adaptation in L. monocytogenes also alters membrane fatty acid

428

composition that is suggestive of more rigid membrane fluidity, which may act as a defense

429

mechanism by restricting the diffusion of acids across the membrane (59, 60). Thus, the

430

increased rigidity of the cytoplasmic membrane during adaptation to potassium lactate could at

431

least partially increase subsequent resistance against NIS.

432

Transcriptional profiling of L. monocytogenes during adaptation to lactic, acetic, hydrochloric

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acid has also previously indicated a consistent pattern of alterations in cell membrane including

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down-regulation of genes involved in branched chain fatty acid synthesis (43). Compared to SD,

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PL was shown to have a stronger antilisterial activity as well as more pronounced down-

436

regulation of branched-chain fatty acid-related genes (43) even when the concentration of

437

undissociated form of SD was higher (3.7 mmol/L for SD compared to 1.4 mmol/L for PL),

438

which determines the primary inhibitory effects (60). In our study, the estimated concentration of

439

undisscoiated form of PL and SD were 1.1 mmol/L and 0.54 mmol/L, respectively using the

440

Henderson-Hasselbalch equation. Thus, relatively stronger effect of PL can be expected

441

compared to SD, which may explain profound PL-induced cross-protective effect. As the mode

442

of action for EPL also involves cell membrane disruption (9, 11, 12), it is reasonable to speculate

443

that the membrane fatty acid modification is linked with PL-induced cross-protection against

444

EPL.

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The pre-exposure of L. monocytogenes to PL induced cross-protection against LAE only for

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ΔvirR but not the parent strain. Consistent with the observation that dltABCD is the major

447

resistance determinant for LAE, it may be the case that the regulation of cell wall glycopolymers

448

in the parent strain provides protection against LAE whereas in the absence of VirR-dependent 20

449

regulation of cell wall structures, acid-induced changes in cytoplasmic membrane affords

450

additional protection. In the case of CHI, relatively minor effects of either acid on subsequent

451

CHI resistance is consistent with a previous report that the primary molecular target of CHI is

452

likely teichoic acids with subsequent extraction of membrane lipids (13). It is worth noting that

453

the increased sensitivity of ΔvirR under CHI stress (100 µg/ml) was not as evident compared to

454

the survival experiment with 50 µg/ml CHI (Fig. 2). This could be due to the concentration-

455

dependent saturation of the bactericidal effect of CHI, upon which there is no additional lethality

456

even with increasing concentrations (61).

457

Conclusions. This study shows a critical role of the L. monocytogenes two-component

458

system response regulator VirR in resistance against antimicrobials intended for food

459

preservation. Improved understanding of resistance mechanisms may facilitate the development

460

of effective hurdle strategies utilizing different molecular targets (33). Additionally, the potential

461

for cross-protection induced by a food-relevant stress (e.g., organic acids) should be taken into

462

consideration to avoid unintended over-estimation of preservative strength in a food system (20,

463

42). A mechanistic understanding of the interaction between antimicrobials and the target

464

pathogen also facilitates better delineation of how other food constituents may enhance (57) or

465

decrease (62) the antimicrobial efficacy. Lastly, identification and characterization of TCSs with

466

respect to stress tolerance and virulence have potential implication in development of better

467

control strategies for the pathogen as novel antimicrobial agents may be developed to target these

468

molecular targets (63, 64). In light of recent advances in high-throughput sequencing

469

technologies and decreasing cost, future efforts in understanding global changes in the

470

transcriptome of L. monocytogenes under antimicrobial challenge can be expected to provide

471

more comprehensive view of bacterial stress response and resistance mechanisms (65). 21

472

ACKNOWLEDGEMENTS

473

This work was supported by New York Sea Grant R/SHH-16, funded under award

474

NA07OAR4170010 from the National Sea Grant College Program of the U.S. Department of

475

Commerce’s National Oceanic and Atmospheric Administration, to the Research Foundation of

476

State University of New York, and by Agriculture and Food Research Initiative grant 2010-

477

65201-20575 from the U.S. Department of Agriculture, National Institute of Food and

478

Agriculture, Food Safety Program.

479

We would like to thank Barbara Bowen and Rebecca Schmidt for constructing the

480

deletion mutants used in this study, and Matthew Stasiewicz for assistance with the analysis of

481

MIC data.

482

22

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688 689 690

32

691

Table 1. Strains and plasmids used in this study. Strain / Plasmid Strains FSL F6-0366 FSL B2-0315 FSL B2-0377 FSL B2-0379 FSL B2-0380 FSL K5-0022 FSL K5-0024 FSL K5-0025 FSL K5-0026 FSL K5-0027 FSL K5-0028 FSL K5-0029 Plasmids pJK1 pPL2

Strain Alias / Relevant Genotype

Reference

H7858; parent strain; serotype 4b H7858 ΔliaR H7858 ΔlisR H7858 ΔvirR H7858 ΔcesR H7858 ΔvirS Pdlt-GUS FSL F6-0366 Pdlt-GUS FSL B2-0379 (ΔvirR) Pdlt-GUS FSL K5-0022 (ΔvirS) H7858 ΔdltA H7858 ΔmprF H7858 ΔanrAB

(66) (67) This work This work This work This work This work This work This work This work This work This work

Pdlt-uidA(GUS) on pPL2 Integrative shuttle vector (Cmr)

This work (68)

692 693

33

694 695

Table 2. MIC determinations for the parent strain (H7858) and its isogenic TCS response

696

regulator mutantsa,b

Antimicrobial (µg/ml) NIS LAE EPL CHI

H7858 12.5 12.5 100 >200

ΔliaR 6.25 12.5 50 >200

MIC for ΔlisR 12.5 6.25 100 >200

ΔvirR 1.56 6.25 25 100

ΔcesR 12.5 12.5 50 >200

697 698 699 700

a

MICs are determined from three biological replicates and represented as the mode.

b

MICs in bold are statistically different compared to the parent strain with p value = 0.1 as

determined by the logrank test.

701 702

34

703

FIGURE LEGENDS

704

Figure 1. Membrane integrity of the parent strain H7858 and ΔvirR strain under NIS, LAE, EPL,

705

CHI exposure for 24 h at 7oC. Black and gray bars represent the relative dye ratio of the parent

706

strain and ΔvirR, respectively. The relative dye ratios are calculated as the fluorescence ratio

707

(SYTO9 to propidium iodide) for antimicrobial-treated parent strain and ΔvirR divided by the

708

fluorescence ratio of untreated parent strain and ΔvirR. Asterisks indicate the relative dye ratios

709

for ΔvirR under each antimicrobial stress that are significantly (p < 0.05) different compared to

710

the parent strain. The data represent the mean and the standard deviation of three biological

711

replicates.

712

Figure 2. Log decrease in cell density for the parent strain H7858, ΔvirR, ΔdltA, ΔmprF, and

713

ΔanrAB exposed to NIS (0.5 µg/ml), LAE (20 µg/ml), EPL (500 µg/ml), and CHI (50 µg/ml) for

714

24 h at 7oC. Within each antimicrobial, strains with the same letter are not significantly different

715

from each other (p < 0.05). The data represent the mean and the standard deviation of three

716

biological replicates.

717

Figure 3. Relative binding of cytochrome c for the parent strain H7858, ΔvirR, ΔdltA, and

718

ΔmprF grown to log phase at 7oC. Strains with the same letter are not significantly different from

719

each other (p < 0.05). The data represent the mean and the standard deviation of three biological

720

replicates.

721

Figure 4. GUS activities for dlt promoter-GUS reporter fusions expressed in the parent strain,

722

ΔvirR, ΔvirS backgrounds in the presence of 100ng/ml NIS at 7oC. Asterisks indicate

723

significantly (p < 0.05) different GUS activity levels in each genetic background. The data

724

represent the mean and the standard deviation of three biological replicates. 35

725

Figure 5. Log decrease in cell density for the parent strain H7858 and ΔvirR strains pre-exposed

726

to no acid (CTRL), 2% potassium lactate (PL), and 0.14% sodium diacetate (SD) for 8 h at 7oC

727

prior to 24 h challenge with NIS, LAE, EPL, and CHI at 7oC. Closed circles and open circles

728

represent the log decrease in cell density for the parent strain and ΔvirR strains, respectively.

729

Asterisks above and below the dots indicate the log decrease in cell numbers for acid-exposed

730

parent strain and ΔvirR, respectively, that are significantly (p < 0.05) different compared to no

731

prior acid exposure (CTRL). The data represent the mean and the standard deviation of three

732

biological replicates.

733

Figure 6. Model of VirR-mediated resistance to food antimicrobials based on our data and

734

previously proposed modes of action for these antimicrobials. The figure was adapted from a

735

previous publication (53). The L. monocytogenes cell envelope consisting of the cell wall (gray)

736

and the cytoplasmic membrane (below gray area) is shown. Teichoic acids that are anchored to

737

the cell wall and the cytoplasmic membrane represent wall teichoic acids and lipoteichoic acids,

738

respectively. D-alanylation of teichoic acids (mediated by DltABCD) and L-lysinylation of

739

membrane phospholipids (mediated by MprF) are indicated by the letter A or L in a circle,

740

respectively. Letters below each strain designate the phenotype when exposed to each

741

antimicrobial as inferred from the survival assays: R, resistant; S, sensitive; MS, most sensitive.

742

Potential electrostatic repulsion or exclusion of antimicrobials facilitated by teichoic acid as well

743

as phospholipid modifications are indicated by deflected arrows with a horizontal line whereas a

744

plain arrow indicates antimicrobial penetration to the target site. The combination of two arrows

745

indicates partial resistance provided by the specific modification.

746

36