The glutamate decarboxylase system (GAD2) is activated by osmotic stress in a GadR-dependent manner in Listeria monocytogenes

Abstract

Under acidic conditions, the glutamate decarboxylase system GAD2 protects Listeria monocytogenes by consuming intracellular protons. GAD2 is encoded by the gadT2D2 operon and regulated by the transcriptional regulator GadR. In this study we investigated whether osmotic stress, which is commonly used as a hurdle in food preservation, influences GadR activity. Exposure to 0.5 M or 1 M NaCl rapidly increased gadT2 transcription in a GadR-dependent manner. This induction was independent of Na⁺ or Cl^- ions as other osmotically active molecules (e.g. KCl, Na₂SO₄, sucrose) produced similar effects, suggesting that this regulation is responsive to changes in osmolarity. Two putative GadR binding sites in PgadT2 were indispensable for this osmotic induction. Furthermore, GadR-dependent GAD2 expression is elevated in NaCl-treated cultures and it confers protection against lethal acid stress. Taken together these data showed that osmotic stress induces GadR-dependent GAD2 expression, which confers a significant survival advantage when cells are subsequently exposed to acid stress, likely due to an improved capacity for intracellular pH homeostasis. Thus, GadR is activated by two conditions commonly used in food preservation, mild acid pH and elevated osmolarity, which may increase bacterial resistance to gastric acid, posing a potential risk to food safety.

Listeria monocytogenes activates the glutamate decarboxylase system GAD2 in response to osmotic stress in a GadR-dependent manner, promoting survival at low pH.

Introduction

Listeria monocytogenes (L. monocytogenes) is a significant food-borne pathogen that can present a major health risk in ready-to-eat (RTE) food products. This bacterium can withstand important food preservative stresses [e.g. low water activity (a_W), low pH, low temperature] (van der Veen et al. 2008). Upon the transit from contaminated food products to consumers, L. monocytogenes faces a significant host defense system provided by acidic stomach fluid. Survival under these conditions is essential for establishing an infection in the host, termed listeriosis (Lecuit 2020). This disease, despite its low occurrence, has an estimated mortality rate of 20%–30% in outbreaks and is among food-borne illnesses that pose the most severe disease burden worldwide (The European Union One Health 2023).

RTE food industries often implement a combination of food preservation measures to achieve bacteriostatic/bactericidal effect, which is termed “hurdle technology” (Leistner 2000). In the RTE food categories (e.g. dairy, deli meat, fish) in which L. monocytogenes contaminations are found to be prevalent (The European Union One Health 2023), low pH and low a_W are often among the intrinsic factors that help to prevent the growth of pathogens. However, mild stress stimuli often induce tolerance/resistance in food-borne pathogens that enhances survival in otherwise lethal conditions (Bergholz et al. 2012, Jordan et al. 1999, Yeak et al. 2023). In L. monocytogenes, it has been well-established that mild acid stress stimuli (pH 5) induces strong protection against lethal acidic challenge (pH < 3) (Davis et al. 1996, Ferreira et al. 2003, Guerreiro et al. 2022, Wu et al. 2024). Interestingly, salt stress is also shown to simulate protection against lethal stress conditions (Bergholz et al. 2012). This poses a challenge to the “hurdle technology” as this pathogen, if not successfully eliminated by these stresses, is more likely to survive exposure to gastric acid and therefore more likely to cause infection (Guerreiro et al. 2022). Understanding the regulatory mechanisms involved in cross protection by mild stress can inform the design of improved food preservation strategies.

Listeria monocytogenes can grow at pH values as low as 4.5 and can survive transiently at pH 2.3–2.5 (Davis et al. 1996, van der Veen et al. 2008, Wu et al. 2022). This is largely due to the various proton consuming metabolic reactions that serve to buffer the intracellular pH (pH_i) in this bacterium (Wu et al. 2024). A recently discovered transcriptional regulator, GadR, governs the expression of the GAD2 glutamate decarboxylase system (one of three GAD systems in L. monocytogenes) and controls the acid survival of this pathogen (Wu et al. 2023a). Furthermore, this transcriptional regulator is responsible for the mild acid stress-induced protection against lethal acidic challenge. The glutamate decarboxylase system GAD2 consists of a glutamate decarboxylase (GadD2) and a glutamate/gamma-aminobutyric acid (GABA) antiporter (GadT2). GadD2 catalyses the proton-consuming decarboxylation reaction and the main decarboxylation product GABA is exported by GadT2 in exchange for extracellular glutamate. Intracellular GABA can also be metabolized through a pathway known as the GABA-shunt, which bypasses the α-ketoglutarate to succinate steps in TCA cycle that are missing in L. monocytogenes (Feehily et al. 2013). Glutamate is not only the most abundant amino acid in cytoplasm but also a critical metabolite in osmotic homeostasis. Glutamate and potassium are taken up as the primary response to osmotic up-shift stress (Sleator, Gahan et al. 2003, Bremer and Krämer 2019). In L. monocytogenes, glutamate can be accumulated up to 280 mM intracellularly when growing with 7.5% NaCl (Patchett et al. 1992). In a ΔpdeA mutant that has higher c-di-AMP level and perturbated osmotic homeostasis, gadT2D2 is upregulated and this likely explains the hyper-acid resistance under lethal acid challenge (Witte et al. 2013). These observations suggest that osmotic stress likely influences the ability of surviving lethal acidic challenge by influencing gadT2D2 expression.

In this study, we aimed to investigate how osmotic stress influences the expression of GAD2. We showed that osmotic stress induces the GadR-dependent GAD2 expression, which provides protection against lethal acidic challenge. The study has implications for understanding how L. monocytogenes responds to food preservations stresses and how this response can potentially influence survival of the pathogen in gastric fluid.

Material and methods

Bacterial strains and growth conditions

Listeria monocytogenes and Escherichia coli TOP10 strains and plasmids used in this study are listed in Table 1. Listeria monocytogenes strains (Wu et al. 2023a) were cultured in sterile 50 ml falcon tubes containing 5 ml Brain Heart Infusion (BHI) broth (LAB M LAB048) at 37°C with agitation at 150 rpm. To prepare exponential phase cultures, 1.5 ml overnight cultures of L. monocytogenes were centrifuged and resuspended in fresh BHI broth and then inoculated to 5 ml BHI in polypropylene containers (100 ml) to achieve an initial OD600nm of 0.05. The cultures were incubated for approximately 3 h until reaching mid-exponential phase (OD600nm ≈ 0.4). Where indicated, chloramphenicol (cml) was added to the medium at a concentration of 10 µg/ml to arrest protein synthesis. 50 µg/ml kanamycin (kan) was added to growth media where necessary.

Table 1.

Strains and plasmids used in this study.

Strains or plasmids	Reference or source
Strains
Escherichia coli One Shot TOP10	Invitrogen
Listeria monocytogenes EGD-e (gadR-)	Dr Pascal Piveteau
Listeria monocytogenes EGD-e gadR+	(Wu et al. 2023a)
Listeria monocytogenes EGD-e gadR+ PgadT2_V4	(Wu et al. 2023a)
Listeria monocytogenes EGD-e gadR+ PgadT2_V5	(Wu et al. 2023a)
Listeria monocytogenes EGD-e::pIMK-PgadT2::egfp	This study
Listeria monocytogenes EGD-e gadR+::pIMK-PgadT2::egfp	This study
Plasmids
pIMK2; Kanr	(Monk et al. 2008)
pIMK-PgadT2::egfp; Kanr	This study

Molecular techniques

Plasmids and primers used in this study are listed in Tables 1 and 2.To construct the reporter system, the promoter region (−1 to −383 bp relative to gadT2 start codon) of gadT2 was amplified (Phusion polymerase, ThermoFisher) from strain EGD-e. The egfp gene was amplified from the plasmid pKSV7-P_lmo2230::egfp (Utratna et al. 2012). The two PCR products were then fused using splicing by overlap extension PCR. The resulting fusion product and the vector pIMK2 were digested (FastDigest, ThermoFisher) and ligated (T4 DNA ligase Roche).

Table 2.

Primers used in this study.

Primer name	Primer sequence (5’-3’)
Cloning
Plmo2362-F	TCGAGAGCTCCCTCCCTCTTCCCATTATTACCT
Plmo2362-R	TGTGAATAATTCCTCTCCTTTACTAACCATTGTATGTTACCTCCCAATTTTTTG
EGFP-F	CGCAAACAAAAAATTGGGAGGTAACATACAATGGTTAGTAAAGGAGAGGAATTAT
EGFP-R	CGATGGATCCTTATTTGTATAATTCATCCATTCCT
T3-F	AATTAACCCTCACTAAAGGGAAC
pIMK3-insert-check-F	CCAGGAATTGGGGATCGGAA
PL95-R	ACATAATCAGTCCAAAGTAGATGC
PL102	TATCAGACCTAACCCAAACCTTCC
qPCR primer
16S_RT_F	TGGGGAGCAAACAGGATTAG
16S_RT_R	TAAGGTTCTTCGCGTTGCTT
lmo2362_RT_F	ATCCAACATTTGCCACTTCC
lmo2362_RT_R	AAGAAGATTGCGGCAAAACC

Five microliters of the ligation mixture were used to transform E. coli TOP10 chemically competent cells via heat shock. Subsequently cells were plated on LB-kanamycin (LB^kan) plates and incubated overnight at 37°C. A positive transformant carrying the correct insert was selected and grown overnight in LB^kan medium to amplify the plasmid for purification. The purified plasmid (pIMK-P_lmo2362::egfp) was then electroporated into electrocompetent L. monocytogenes strains. Transformants were selected on BHI^kan plates as previously described. Integration of the reporter system into the genome was achieved by one-step integration, as described in earlier studies (Monk et al. 2008). The genomic integration of the reporter system was confirmed by colony PCR (Table 2).

Osmotic stress treatment

For osmotic stress treatment, prewarmed 5 ml BHI medium with a double concentration of the desired osmolyte was added to an equal volume of exponential phase culture (OD600nm ≈ 0.4) and incubated at 37°C for the durations specified (15 min, 30 min, 45 min, 1 h, 24 h). When using different osmotically active solutes, 0.5 M KCl, 0.33 M Na₂SO₄, or 2 M sucrose was applied to achieve an equivalent osmolality to 0.5 M NaCl. As negative controls, 5 ml of prewarmed BHI media without added osmolyte were added to equal volume of exponential phase cultures.

Transcriptional analysis

Transcript levels were determined using RT-qPCR following previously described methods with minor modifications (Guerreiro et al. 2022). For RNA extraction, 1 ml of culture was mixed with 5 ml RNA Later (Sigma) and incubated at ambient temperature (∼18°C) for RNA stabilization. RNA extraction and cDNA synthesis were performed following previously described protocols (Wu et al. 2022). Briefly, cells were recovered by centrifugation (8000 rpm 5 min, 4°C) and resuspended in 700 μl RNA lysis buffer RLT (Qiagen RNeasy). The mixture was transferred to lysis matrix B tubes (Fisher Scientific) for mechanical lysis (40 s at 6 m·s^-1, twice). RNA extraction was completed following the manufacturer’s instructions (Qiagen RNeasy). DNA was depleted (DNase Turbo, Invitrogen) before reverse transcription was performed (SuperScript III, Invitrogen). Quantitative PCR (LightCycler 480 SYBR Green I Master Roche) was performed with 500 × diluted cDNA samples using Roche LightCycler 480 system. Three independent experiments were performed, each with two technical replicates. Relative gene transcription levels were calculated using Q-gene software (Simon 2003), using 16S rRNA as a reference gene.

GABA assay

The extracellular GABA was measured using a previously established method with minor modifications (O’Byrne et al. 2011). For stationary phase bacteria, exponential phase cultures were grown for 24 h with or without exposure to osmotic stress and 2 ml was resuspended (10 000 g, 5 min) in equal volume of BHI^cml 0.5 M NaCl (pH 3.25) and incubated at 37°C for 30 min. For exponential phase bacteria, the cultures were exposed to either 1 h osmotic stress (0.5 M NaCl) or pH 5.0 acid stress. For the pH 5.0 treatment, 5 ml exponential phase cultures were mixed with 5 ml BHI, then acidified to pH 5.0 using 5 M HCl (62 μl) and incubated at 37°C for 1 h. Then 2 ml of culture that had been exposed to 1 h osmotic stress or pH 5.0 acid stress was resuspended (10 000 g, 5 min) in 200 µl of BHI^cml (pH 3.25), and incubated at 37°C for 30 min.

Before and following incubation, samples were taken, serial diluted (10-fold, in phosphate-buffered saline), and 10 µl from each dilution was spotted onto BHI agar to determine viable cell counts. Subsequently samples were centrifuged at 10 000 g for 5 min, and the supernatant was collected for GABA quantification. To measure GABA levels, 5 µl of the supernatant was added to 95 µl of a freshly prepared reaction master mix containing 80 mM Tris buffer, 750 mM sodium sulfate, 10 mM dithiothreitol, 1.4 mM NADP⁺, 2 mM α-ketoglutarate, and 0.1 g/l GABase (Sigma–Aldrich). The mixture was incubated at 37°C for 1 h in a temperature-controlled plate reader (Labtech, LT-5000MS ELISA Reader) before OD340nm value was measured. Standard curves were established using 0–10 mM GABA solutions prepared in BHI (pH 3.25). Three independent experiments were performed, each with two technical replicates Extracellular GABA levels were expressed as GABA pmol/10⁶ CFU for both strains.

Reporter fluorescence assay

To detect EGFP fluorescence signals in stationary phase, bacterial cultures were grown in the presence of osmotic stress for 24 h. Following treatment, 2 ml of culture was washed (10 000 rpm, 5 min) once and resuspended with 2 ml PBS^cml or PBS^cml 0.5 M NaCl respectively.

For exponential-phase bacteria, cultures were exposed to either osmotic stress (0.5 M NaCl) or acid stress (pH 5.0) for 1 h. For acid stress treatment, 5 ml of exponential-phase culture was mixed with 5 ml of BHI medium, acidified to pH 5.0 using 62 μl of 5 M HCl, and then incubated at 37°C for 1 h. After treatment, 2 ml of each stressed culture was centrifuged (10 000 rpm, 5 min) and washed once with 2 ml of PBS^cml or PBS^cml 0.5 M NaCl. The pellet was then resuspended in 0.5 ml of the corresponding buffer as well.

To measure fluorescence, 200 μl of each resuspension was transferred into wells of a 96-well microplate. OD600nm and EGFP fluorescence (excitation at 485 nm, emission at 528 nm) were measured using a plate reader (BioTek Synergy H1). Three independent experiments were conducted, each with technical duplicates. Fluorescence levels were normalized and expressed as relative fluorescence units (RFU) normalized by OD600nm.

Acid survival experiments

The ability of osmotic stress exposed L. monocytogenes strains to survive in lethal acidic conditions was tested as previously described (Wu et al. 2023b) with minor adjustments. To examine the acid resistance, 100 µl of culture was mixed with 900 µl pH 2.3 BHI with or without 0.5 M NaCl in 1.5 ml microcentrifuge tubes and incubated statically at 37°C. Samples were taken at the specified time points, serial diluted (10-fold, in phosphate-buffered saline), and 10 µl of cell suspensions from each dilution was then spotted onto BHI agar and incubated at 37°C to quantify the number of viable cells. Acid resistance was presented as percentage survival at each time point relative to the starting cell count (prior to acid challenge). Three independent experiments were carried out, each with technical triplicates.

Statistical analysis

All statistical analysis, were performed using GraphPad Prism 8. The statistical tests used are described in the figure legends.

Results

Osmotic stress induces gadT2 transcription in a GadR-dependent manner

Previously we established that the RofA-like transcriptional regulator GadR plays a critical role in regulating the gadT2D2 operon in response to mild acid stress. Here we investigated whether osmotic stress influenced the transcription of this operon and whether GadR is required for this regulation. Wild type strain EGD-e (gadR^-) and its gadR⁺ derivative (Wu et al. 2023a) were grown in BHI at 37°C to mid-exponential phase when 0.5 M or 1.0 M NaCl was added. After 1 h incubation, the transcript levels of gadT2 from NaCl treated cultures were compared with untreated cultures by RT-qPCR. In the gadR⁺ strain there was a substantial and significant increase in the gadT2 transcript level following exposure to 0.5 M and 1.0 M NaCl (< 0.0001 and < 0.0001, respectively; Fig. 1A). The level of gadT2 mRNA was also moderately increased in the gadR⁺strain compared to the wild type strain in the absence of NaCl (P = 0.0047; Fig. 1A). Next, the transcription profile of gadT2 was examined over time following exposure to 0.5 M NaCl. gadT2 transcription was strongly and rapidly induced in the gadR⁺strain but not in wild type EGD-e (Fig 1B). To test whether this observation is dependent on Na⁺ or Cl^-, the transcriptional response of gadT2 to three other osmolytes (KCl, Na₂SO₄, and sucrose) were tested. When they were each supplied at an osmotically equivalent concentrations to 0.5 M NaCl, gadT2 transcription was induced to similar levels (Fig. 1C).

Figure 1.

Osmotic stress activates gadT2 transcription in a GadR-dependent manner. (A) The transcript levels of gadT2 in EGD-e WT or gadR⁺ strains with or without 1 h NaCl stress (0.5 M or 1 M) treatment during exponential phase are shown, expressed relative to EGD-e WT without NaCl. (B) The transcript levels of gadT2 in EGD-e WT or gadR⁺ strains at 15, 30, 45, 60 min following 0.5 M NaCl treatment during exponential phase, with each strain expressed relative to its respective untreated control. (C) The transcript levels of gadT2 were measured in EGD-e WT and gadR⁺ exponential phase cultures after 30 min treatment using different osmolytes (at osmotically equivalent concentrations 0.5 M NaCl, 0.5 M KCl, 0.33 M Na₂SO₄, or 1 M sucrose). Exponential phase cultures without treatment or those treated with medium were included as controls. Transcripts levels are shown relative to the EGD-e WT without treatment control. (D) The transcript levels of gadT2 in exponential phase culture were measured with or without 0.5 M NaCl treatment for 1 h in EGD-e strains (WT, gadR⁺, gadR⁺ ΔBox1, and gadR⁺ ΔBox2) expressed relative to EGD-e WT without NaCl stress condition. Three independent experiments were performed with technical duplicates using 16S as reference gene. Statistically significant differences were determined by two-way ANOVA with Sidak’s multiple comparisons test (ns, not significant; *, P < 0.05; **, P < 0.01; ***, P < 0.001; and ****, P < 0.0001).

Next, we investigated whether the putative GadR binding boxes in the gadT2 promoter region (Wu et al. 2023a) are required for the activation of transcription by osmotic stress. Deletion of either of the two GadR boxes (Wu et al. 2023a) (BOX1 −209 to −226 bp TCTCAAAAATTTTTCCTG; BOX2 −178 to −195 bp CTGGAATAATTTTCGAAA relative to gadT2 start codon) abolished the osmotic induction of gadT2 (Fig. 1D). Thus, both of these 18 bp predicted binding sites of GadR, are needed for the osmotic induction of the operon. Taken together these data show that the induction of gadT2 transcription in the presence of osmotic stress requires GadR and its putative binding boxes.

Osmotic stress activates the GAD2 system

We sought to test whether osmotic stress induced gadT2D2 transcription leads to functional expression of GAD2 system. For this, we fused PgadT2 (including ribosome binding site) with a codon-optimised version of egfp gene (Utratna et al. 2012) to generate PgadT2::egfp reporter. This construct was integrated into the genomes of EGD-e WT (gadR^-) and the gadR⁺ derivative of EGD-e WT and fluorescence was measured following the addition of 0.5 M NaCl. One hour after NaCl addition there was no detectable increase in the expression from the PgadT2, whereas the pH 5.0 acidification resulted in a significant increase in fluorescence after 1 h (P < 0.0001; Fig. 2A). However, after extended incubation for 24 h the NaCl-treated culture showed a significant increase in EGFP expression compared to the untreated control (P < 0.0001; Fig. 2B). The EGD-e WT strain showed only background fluorescence in all conditions and this was not affected by the inclusion of NaCl in the medium. Thus, GadR-dependent expression of gadT2D2 operon was positively affected by osmotic stress in stationary phase cultures.

Figure 2.

The GAD2 system is activated by osmotic stress. EGD-e and EGD-e gadR⁺ strains carrying the PgadT2::egfp reporter integrated into the genome were exposed to BHI with or without 0.5 M NaCl for 1 h (A) or 24 h (B), or to acidified BHI (pH 5) for 1 h (A). Subsequently, OD600nm and RFU were measured and are expressed as RFU/OD for both strains. EGD-e WT and EGD-e gadR⁺ strains were exposed to BHI with or without 0.5 M NaCl for 1 h (C) or 24 h (D), or to acidified BHI (pH 5) for 1 h (C). Subsequently, strains were exposed to BHI^cml pH 3.25 for 30 min, and extracellular GABA levels were measured and presented as GABA pmol/10⁶ CFU for both strains. Three independent experiments were performed with technical duplicates. Statistically significant differences were determined by two-way ANOVA with Sidak’s multiple comparisons test (ns, not significant; *, P < 0.05; **, P < 0.01; and ****, P < 0.0001).

Next, we sought to test the activity of the GAD2 system under osmotic stress. The secretion of extracellular GABA under acidic conditions (where GAD2 is active) was assayed as a measure of GAD2 activity (Karatzas et al. 2010). When EGD-e WT strain was exposed to either low pH or osmotic stress (0.5 M NaCl) for 1 h during mid-exponential phase and subsequently incubated at pH 3.25 for 30 min, no increase in GAD2 activity was detected (Fig. 2C). The EGD-e gadR⁺ strain showed small but not significant increase in GAD2 activity following NaCl exposure. In contrast, exposure to pH 5.0 acid stress resulted in a marked increase in GAD2 activity (P = 0.0015; Fig. 2C). In line with the observation that GadR-dependent expression is activated by osmotic stress in stationary phase, prolonged NaCl exposure (24 h) significantly induced GAD2 activity in EGD-e gadR⁺ strain (P < 0.0001; Fig. 2D). The EGD-e WT strain did not produce GABA under these conditions, confirming that an active GadR is required for activation of GAD2 (Fig. 2D). Next, we investigated whether activation of GAD2 by osmotic stress conferred a survival advantage under lethal acidic conditions. We challenged stationary phase cultures of both strains at pH 2.3 and measured survival over time by performing viable cell counts. The EGD-e gadR⁺ cultures survived much better under these conditions than the parental EGD-e WT strain, consistent with previous observations (Wu et al. 2023a). The presence of NaCl during growth had no impact on the survival of EGD-e WT strain but significantly enhanced the survival of the EGD-e gadR⁺strain under these conditions (Fig. 3). Together these data suggest that osmotic stress activates the GAD2 system and this can produce a significant effect on acid survival.

Figure 3.

Osmotic stress induces GadR-dependent acid resistance. EGD-e WT (blue symbols) and EGD-e gadR⁺ (green symbols) strains were cultivated in BHI (circles) or BHI with 0.5 M NaCl (squares) for 24 h, and subsequently challenged in BHI at pH 2.3 (acidified with HCl). The survival rates of both strains were monitored over time. The survival rates of EGD-e WT cultivated in BHI or BHI with 0.5 M NaCl fell below the detection limit after 50 min and 40 min of acid challenge, respectively; Therefore, no additional time points were included thereafter. Three independent experiments were performed with technical triplicates. Statistically significance was calculated for EGD-e gadR⁺ with or without osmotic stress treatment using two-way ANOVA with Sidak’s multiple comparisons test. (*, P < 0.05; and ****, P < 0.0001).

Discussion

This study establishes that osmotic stress induces the expression of the GAD2 system, comprising the glutamate decarboxylase GadD2 and the Glu-GABA antiporter GadT2, in manner that depends on the GadR transcriptional regulator. Since the GAD2 system is known to function as a system for pH_i homeostasis in response to acid stress (Cotter et al. 2001) this raises the question of whether this system is also needed to cope with osmotic stress. During osmotic stress bacterial cells lose water with a consequent drop in turgor pressure, both of which impact growth negatively (O’Byrne and Booth 2002). Listeria monocytogenes has several systems that allow it to cope with osmotic stress and indeed this pathogen has a very high tolerance for environments with low a_W (Cole et al. 1990). For example, both potassium and glutamate rapidly accumulate to high concentrations in the cytoplasm in response to osmotic shock (Patchett et al. 1992, Amezaga et al. 1995, Feehily and Karatzas 2013). Subsequently, compatible solutes such as carnitine and glycine betaine are transported into the cytoplasm where they can reach high concentrations and serve to reverse the water loss and restore turgor pressure by raising the cytoplasmic osmolarity (Sleator et al. 2003, O’Byrne and Karatzas 2008). It is possible that during the osmotic balancing that takes place there is an impact on pH_i that also needs to be counteracted. Indeed there is some evidence that pH_i is reduced during osmotic stress in L. monocytogenes, although the effect is quite small (∼0.1 pH units) at 0.5 M NaCl (Fang et al. 2004). Osmotic stress is also known to cause cytoplasmic acidification in Salmonella Typhimurium and Escherichia coli (Chakraborty et al. 2017), and it induces GAD system expression in E. coli (De Biase et al. 1999). Thus, it is possible that the GadR induces gadT2D2 expression as a response to reduced pH_i under osmotic stress. pH_i measurements will be required to fully investigate this hypothesis.

The mechanism that leads to activation of GadR has not yet been established. The C-terminal domain of GadR has 2 PTS Regulatory Domains (PRD) and an enzyme IIB-like domain (Wu et al. 2023a, 2024) suggesting that the protein might be regulated by interacting with one or more of the PTS sugar transport systems of the cell, as has been suggested for other members of this family of regulators (Galinier and Deutscher 2017, Rom et al. 2021). Listeria monocytogenes has 29 complete PTS sugar transport systems (Stoll and Goebel 2010, Sauer et al. 2019) giving this saprophytic bacterium the capacity to utilize a wide range of carbon and energy sources from its environment. It is known that PTS gene expression is significantly altered during conditions of osmotic stress in L. monocytogenes (Bae et al. 2012). These changes in expression could potentially provide a signal that alters GadR activity leading to upregulated gadT2D2 transcription. Alternatively, changes in pH_i (in response to acid or osmotic stress) might act as a stimulus to induce GadR activity prompting pH_i homeostasis via the GAD2 system.

Water activity (a_w) and pH are two of the main parameters that food processors consider when developing foods, both to ensure resistance to spoilage and to limit the growth of pathogens. Since GadR can activate the GAD2 system in response to either low pH (Wu et al. 2023a) or reduced a_w (this study) it could be an important determinant of how L. monocytogenes behaves in RTE foods and subsequently in the stomach after ingestion. The gadR gene is known to have a high rate of premature stop codons (PMSC) across the phylogenetic tree of L. monocytogenes based on an analysis of approximately 40 K whole genome sequences. In lineage II strains approximately 8% of strains carry a PMSC, whereas in lineage I strains the rate is around 1% (Wu et al. 2023a). The occurrence of a PMSC in the well-studied strain EGD-e is known to produce an acid sensitive phenotype. Here we show that a strain carrying a gadR PMSC also fails to activate gadT2D2 transcription in response to NaCl. Since GAD2 plays a significant role in allowing L. monocytogenes to survive the gastric acid conditions (Cotter et al. 2001), knowledge of the gadR genotype might help food producers assess the risk presented by individual strains. Further research is needed to test whether GadR influences the behavior of L. monocytogenes in food matrices with reduced pH and/or low a_W.

Conflicts of interest

None declared.

Funding

This project was supported by the Research Ireland Frontiers for the Future Programme (21/FFP-P/10 078).