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. Author manuscript; available in PMC: 2017 Sep 1.
Published in final edited form as: Mol Microbiol. 2016 Jul 4;101(6):942–953. doi: 10.1111/mmi.13433

The LysR-type transcriptional regulator, CidR, regulates stationary phase cell death in Staphylococcus aureus

Sujata S Chaudhari 1,§, Vinai Chittezham Thomas 1,§, Marat R Sadykov 1, Jeffrey L Bose 3, Daniel Jehong Ahn 1, Matthew C Zimmerman 2, Kenneth W Bayles 1,*
PMCID: PMC5014633  NIHMSID: NIHMS809565  PMID: 27253847

Summary

The Staphylococcus aureus LysR-type transcriptional regulator, CidR, activates the expression of two operons including cidABC and alsSD that display pro- and anti-death functions, respectively. Although several investigations have focused on the functions of different genes associated with these operons, the collective role of the CidR regulon in staphylococcal physiology is not clearly understood. Here we reveal that the primary role of this regulon is to limit acetate-dependent potentiation of cell death in staphylococcal populations. Although both CidB and CidC promote acetate generation and cell death, the CidR-dependent co-activation of CidA and AlsSD counters the effects of CidBC by redirecting intracellular carbon flux towards acetoin formation. From a mechanistic standpoint, we demonstrate that CidB is necessary for full activation of CidC, whereas CidA limits the abundance of CidC in the cell.

Keywords: Staphylococcus aureus, cell death, biofilm, pyruvate: menaquinone oxidoreductase, acetolactate synthase, CidA, CidB

Graphical Abstract

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Introduction

Infections caused by Staphylococcus aureus remain a major cause of morbidity and mortality worldwide (Archer, 1998). The ability of this pathogen to develop into matrix-encased communities (biofilms) aids its recalcitrance towards host immunity and antimicrobials (Otto, 2008). Although biofilms may co-opt specific host components as part of their matrix, it also consists of a variety of secreted and recycled bacterial components (proteins, nucleic acids and carbohydrates) that help stabilize the maturing biofilm architecture (Otto, 2008). Intriguingly, much of the recycled components that make up the biofilm matrix are derived from an active suicidal process that is tightly regulated at the population level (Bayles, 2007, Rice & Bayles, 2008).

We recently observed major roles for two overflow metabolic pathways in regulating cell death within a biofilm microenvironment (Thomas et al., 2014). Both pathways, CidC (pyruvate:menaquinone oxidoreductase) and AlsSD (α-acetolactate synthetase/decarboxylase) utilized the same substrate, pyruvate and yet affected cell death, antithetically (Thomas et al., 2014, Patton et al., 2005, Tsang et al., 2008). Acetate resulting from CidC activity potentiated cell death by inducing cytoplasmic acidification of target cells (Thomas et al., 2014). Cell death was characterized by multiple hallmarks of eukaryotic apoptosis including reduced respiration, increased generation of oxygen radicals and DNA fragmentation (Thomas et al., 2014). In contrast, AlsSD activity favored cell survival by diverting carbon flux away from CidC and by maintaining intracellular pH homeostasis due to the inherent proton scavenging activity of this pathway (Thomas et al., 2014). At the transcriptional level, both cidC and alsSD are positively regulated by the LysR-type transcriptional regulator (LTTR), CidR (Yang et al., 2006). This raised the possibility that CidR regulated both pro- and anti-cell death mechanisms simultaneously.

In the present study, we test the above hypothesis and arrive at the surprising conclusion that CidR primarily has an anti-death function despite it transcriptionally activating the pro-death factor, CidC. We also advance new insights on the role of two additional members of the CidR regulon, CidA and CidB in modulating cell death by altering cellular metabolic function.

Results

CidR activity is necessary for optimal survival in stationary phase

Following growth in the presence of excess glucose (35 mM), S. aureus culture supernatants are acidic due to a buildup of acetic acid (Patton et al., 2005). This eventually leads to the generation of reactive oxygen species (ROS) and the potentiation of stationary phase cell death (Thomas et al., 2014, Patton et al., 2005). Consistent with this interpretation, cell death can be arrested by inactivating the acetogenic CidC pathway (Thomas et al., 2014, Patton et al., 2005). Given that CidR positively regulates both alsSD and cidC expression, it might be expected that cidR inactivation would generate a mutant strain that would phenocopy the ΔcidC mutant, primarily because AlsSD function in maintaining intracellular pH homeostasis and promoting cell survival would be unnecessary in the absence of CidC-dependent weak acid stress. However, previous results indicate this is not the case as the cidR mutant was shown to exhibit increased cell death during stationary phase compared to the wild-type strain (Yang et al., 2005). Consistent with these results, the ΔcidR mutant not only exhibited an increased rate of cell death in stationary phase (Fig 1A), but also generated excess ROS compared to the levels observed in the ΔcidC mutant (Fig 1B).

Figure 1. Stationary phase cell death associated with a cidR mutation does not result from intracellular pyruvate toxicity.

Figure 1

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A. S. aureus survival at stationary phase was monitored at 24 h intervals over a period of five days in TSB-35mM glucose. B. Whole cell EPR analysis of S. aureus after 72 h of growth. ROS generation was determined by utilizing the membrane permeable and ROS sensitive spin probe 1-hydroxy-3-methoxycarbonyl-2,2,5,5-tetramethylpyrrolidine (CMH). C. Acetoin concentrations were determined from culture supernatants of various S. aureus strains after 6 h of growth in TSB-35mM glucose. D. The effect of pyruvate on various strains under acidic conditions (TSB-35mM glucose, pH 4.5) were determined by monitoring growth (OD600) for a 24h period according to a previously published method (Thomas et al., 2013). ns, P>0.05; **P<0.01; ***P<0.001; ****P<0.0001; One-way ANOVA with Tukey’s multiple comparisons test.

Interestingly, the ΔcidR mutant phenotypically resembled the ΔalsSD mutant rather than the ΔcidC mutant, with an increased rate of cell death relative to the wild-type strain (Fig 1A) and decreased production of acetoin (Fig 1C). Although it is reasonable to expect depleted acetoin levels in supernatants of the ΔcidR mutant (as CidR also transcriptionally activates the AlsSD pathway), the effects on cell death following cidR mutation were counterintuitive, since this mutant exhibited decreased cidC expression.

Increased rate of cell death in the ΔcidR mutant does not result from intracellular pyruvate toxicity

One potential cause for the cell death observed in the ΔcidR mutant could result from toxicity induced by increased intracellular concentrations of pyruvate. Given that CidR regulates both CidC and AlsSD pathways that consume pyruvate as a substrate, its inactivation could plausibly result in increased levels of intracellular pyruvate. Consistent with this notion we observed an increase in intracellular concentrations of pyruvate of the CidR mutant relative to the wildtype strain at late stationary phase (72h) (Fig S1A). The cellular toxicity attributed to pyruvate under acid stress is controversial. While some studies point to a beneficial role for intracellular pyruvate following acidic stress, others have alluded to a more toxic nature for this metabolite (Wu et al., 2014, Repizo et al., 2011, Tsau et al., 1992, Harvey & Collins, 1963). Hence, to test whether intracellular pyruvate affects S. aureus under acidic stress, we challenged the wild-type strain with increasing concentrations of sodium pyruvate in TSB that was acidified to an initial pH of 4.5 and monitored growth at a wavelength of 600 nm (OD600) for 24 h. Because of the weak acid properties of pyruvic acid (pKa= 2.49), this approach not only allows a small percentage (<1%; estimate based on the Henderson-Hasselbalch equation) of extracellular pyruvic acid to diffuse freely into cells and ensure cytoplasmic acidification, but also simultaneously increases the levels of intracellular pyruvate (upon disassociation of pyruvic acid within the cytoplasm) in the acid stressed cells. The relative amount of growth (fractional area) of the wild-type strain was calculated from the ratio of the area under the growth curve of test samples (samples supplemented with various concentrations of pyruvate) to that of the corresponding control (sample without pyruvate supplementation) and displayed as a function of pyruvate concentration (Fig 1D). Under these conditions, it is important to note that the wildtype strain does not significantly grow due to a low extracellular pH (pH, 4.5). If pyruvate is toxic, its supplementation would only add to the toxicity of existing acidic stress and growth inhibition should be maintained. However, contrary to this argument, our results clearly demonstrate that pyruvate supplementation promoted growth of the wild-type strain in a concentration dependent manner due to detoxification of the prevailing acidic conditions (Fig 1D). This strongly argues against a role for pyruvate in promoting cellular toxicity. Notably, our results suggest that CidR dependent alsSD expression is necessary for the wildtype strain to sustain pyruvate dependent acid detoxification processes as neither the ΔalsSD mutant that promotes intracellular pH homeostasis (Thomas et al., 2014), nor the ΔcidR mutant were able to grow in the presence of pyruvate under the same conditions (Fig 1D). Collectively, these results suggest that although CidR may transcriptionally activate the pro-death factor, CidC, its physiological function is to limit cell death by ensuring intracellular pH homeostasis through AlsSD activity.

CidR-dependent expression of CidC does not result in significant acetate production

In addition to CidR, the alternate sigma factor, sigma B (σB) controls cidC transcription during growth of S. aureus (Rice et al., 2004). Together, their activity results in two distinct transcripts (Fig 2A; (Rice et al., 2005, Rice et al., 2004)). During the early exponential growth phase, the cidBC transcript is abundant due to σB-dependent expression (Rice et al., 2004). However, during the post-exponential phase a switch to cidABC transcription (Fig 2A) is favored through CidR (Rice et al., 2004). This led us to investigate the relative contribution of both transcripts to CidC-mediated acetate production. In S. aureus the Pta-AckA pathway is the major route for acetate synthesis during aerobic growth (Sadykov et al., 2013). Hence, to clearly distinguish the level of acetate contributed by CidC activity alone, we carried out experiments in a Δpta mutant background in which the Pta-AckA pathway has been inactivated. As expected the majority of acetate generated in the Δpta mutant was due to CidC activity (Fig 2B). We next compared the rate of acetate generated by CidC in a Δpta mutant and its isogenic ΔptaΔcidR double mutant. Surprisingly, both the Δpta mutant and ΔptaΔcidR double mutant excreted acetic acid at similar rates (Fig 2B). Furthermore, qRT-PCR analysis revealed that although the cidABC transcript levels were significantly diminished in the ΔptaΔcidR double mutant relative to Δpta mutant, the cidBC transcript was expressed at similar levels in both strains (Fig 2C). These results suggest that CidR-mediated transcriptional activation of CidC does not result in the generation of significant amounts of acetic acid, and the majority of CidC-generated acetate produced is a consequence of σB-dependent cidBC transcription. Consistent with this conclusion, we were unable to detect a significant difference in either the extracellular concentrations of acetate (Fig S1B) or the pH (Fig S1C) of the ΔcidR mutant relative to the wildtype strain until late stationary phase. Furthermore, in agreement with the proposed role for SigB dependent cidBC expression in promoting cell death, the ΔsigB mutant exhibited increased stationary phase survival relative to the wildtype strain (Fig S2).

Figure 2. CidR minimally influences acetate generation.

Figure 2

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A. Although CidR is the canonical transcriptional regulator of the cidABC operon, a second transcript cidBC is under the indirect control of the alternate sigma factor B (Rice et al., 2004). B. The contribution of CidR towards acetate generation was determined in the Δpta background to minimize noise in signal. Rates of acetate generation were calculated at exponential growth of various strains from harvested culture supernatants using a commercially available kit (R-biopharm). C. qRT-PCR analysis to differentiate between cidABC (cidA specific primers) and cidBC transcripts (cidB or cidC specific primers) were performed using cells in the exponential growth phase. ns, P>0.05; *P<0.05; One-way ANOVA with Tukey’s multiple comparisons test.

CidA and CidB are modulators of stationary phase cell death

Although a role for AlsSD in limiting CidC-mediated cell death was earlier proposed (Thomas et al., 2014), the functional significance of neither CidA nor CidB was examined in this context. To investigate the effects of CidA and CidB on acetate-dependent cell death, we first monitored the viability of in-frame isogenic and markerless ΔcidA, ΔcidB and ΔcidAB double deletion mutants following extended growth in TSB supplemented with 35 mM glucose. Over a period of 5 days we observed that the ΔcidA mutant exhibited an increased rate of cell death relative to the wild-type strain (Fig 3A). In contrast, the ΔcidB mutant survived markedly better than the wild-type strain over the same period, indicating antithetical functions for both proteins with respect to the modulation of cell death. Given that the ΔcidAB double mutant partially phenocopied the ΔcidA mutant, these results suggest that despite opposing functions, CidA activity might exhibit some degree of functional dominance over CidB (Fig 3A).

Figure 3. CidA and CidB are antithetical modulators of cell death.

Figure 3

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A. The colony forming units (cfu/ ml) of various S. aureus strains (WT, mutant and complemented strains) at 120 h were determined by plating on TSB. Values in the graph represent fold changes in cfu/ml of the mutants and complemented strains (denoted by ‘c’) relative to the WT strain. Acetoin concentrations (B) and acetate excretion rates (C) were determined from culture supernatants of various S. aureus strains. D. Pyruvic acid sensitivity of the WT and various cid mutants were carried out in TSB-35mM glucose supplemented with 30mM pyruvic acid and growth (OD600) was monitored at maximum aeration in an automated plate reader (TECAN M200) at 37°C. E. Western blot analysis of S. aureus membrane associated proteins (1μg) fractionated from various strains were performed using CidC specific polyclonal antibodies. Protein band intensities were quantified using ImageJ software and are depicted in the graph. ns, P>0.05; *P<0.05; **P<0.01; ***P<0.001; ****P<0.0001; One-way ANOVA with Tukey’s multiple comparisons test.

Although similar in trends, the phenotypic differences in stationary phase viabilities of the ΔcidA and ΔcidB mutants (Fig 3A) were not as pronounced as those observed for the ΔalsSD and ΔcidC mutants, which exhibited nearly two and five order of magnitude differences, respectively, in stationary phase viabilities compared to the wild-type strain (Thomas et al., 2014). This suggested that both CidA and CidB might have a role in regulating cell viability by modulating carbon flux through CidC and AlsSD pathways. To test this hypothesis, we measured the levels of acetate and acetoin excreted by ΔcidA and ΔcidB mutants. While acetoin levels in cell supernatants could be measured directly, acetate levels were measured only after inactivating the Pta-AckA pathway in the ΔcidA and ΔcidB mutants. We reasoned that the latter approach would eliminate any interference from the Pta-AckA pathway and would clarify the contribution of CidC-dependent acetate generation in the ΔcidA and ΔcidB mutant backgrounds. Consistent with the increased cell death observed in the ΔcidA mutant, we observed increased rates of acetate (Fig 3B) generation and decreased levels of acetoin (Fig 3C) in culture supernatants of this strain. Additionally, the increased survival of the ΔcidB mutant in the stationary phase (Fig 3A) was associated with decreased acetate (Fig 3B) and increased acetoin levels (Fig 3C). The observed differences in cell viability and excreted metabolite levels of the ΔcidA, ΔcidB and ΔcidAΔcidB mutants could be complemented to wild-type levels (Figs 3A and C), ruling out secondary site mutations as the cause of these phenotypes. Furthermore, the antithetical trends in the acetate and acetoin excretion profiles of the ΔcidA and ΔcidB mutants were also reflected in their susceptibilities to 30 mM pyruvic acid (Fig 3D). We have previously shown that the growth of S. aureus UAMS-1 in TSB supplemented with pyruvic acid is dependent on the activities of CidC and AlsSD (Thomas et al., 2014). Following acidification of the culture media with 30 mM pyruvic acid (pH ~4.5), CidC-dependent excreted acetic acid was shown to enhance cellular toxicity by cytoplasmic acidification resulting in growth inhibition. Under these very same conditions, AlsSD activity can counter CidC-dependent cytoplasmic acidification and limit growth inhibition (Thomas et al., 2014). In agreement with a role for CidA in increasing the acetate to acetoin ratio in culture supernatants, we observed increased susceptibility of the ΔcidA mutant to the weak acid effects of pyruvic acid relative to the wild-type strain (Fig 3D). In contrast, the ΔcidB mutant displayed a modest increase in resistance to pyruvic acid, consistent with the decreased acetate to acetoin ratio generated by this strain (Fig 3D). Further, in accordance with earlier phenotypes (Fig 3A–C), the ΔcidAB mutant appeared to be more susceptible to pyruvic acid relative to the wild-type strain, although the degree of susceptibility was not as profound as that observed for the ΔcidA mutant.

Clarified role of CidA in the cell lysis pathway

The increased cell death of the ΔcidA mutant is in direct contrast to a previous study from our laboratory (Rice et al., 2007) where CidA was implicated as an effector of murein hydrolase activity and the induced cell lysis pathway. The UAMS-1 cidA::ermR mutant KB1050 was found to display reduced lysis in stationary phase and as a result, exhibited decreased biofilm adherence due to decreased extracellular DNA (Rice et al., 2007). Based on these and other findings in cidA mutants of laboratory strains, it was concluded that CidA is a pro-lytic protein responsible for cell lysis and biofilm formation. Interestingly, analysis of KB1050 demonstrated the presence of atl mRNA but an absence of Atl protein by northern blot and western blot, respectively (data not shown). Based on these findings, we predicted that there was a previously unidentified atl mutation in strain KB1050. The atl open reading frame of both KB1050 and UAMS-1 was amplified by PCR and sequenced. Four independent sequencing trials confirmed that KB1050 had a single thymine deletion within a cluster of four thymines at position 582, when compared to the UAMS-1 sequence, leading to a frameshift mutation that disrupted the C-terminal region of the Atl propeptide that precedes the catalytic domains. As a result of this mutation, it would be expected that KB1050 would be lysis deficient as a result of its inability to produce functional Atl, thus questioning the previously identified role of CidA as a positive effector of murein hydrolase activity. To address this, we utilized the new ΔcidA mutant created in the current study, which possesses an internal in-frame deletion spanning amino acid residues 2 through 52 and leaves the σB-dependent promoter for the downstream genes cidB and cidC intact. In addition, we restored the wild-type cidA allele in KB1050, generating KB1050RC. These strains were compared to UAMS-1 and a Δatl mutant KB5000 to examine the impact of CidA on murein hydrolase activity and biofilm formation. As seen in Figure 4A, colonies of the ΔcidA mutant, KB1065, on TSA containing 2 mg ml−1 Micrococcus cells showed murein hydrolase activity similar to UAMS-1 while KB1050 and KB1050RC were indistinguishable from the Δatl mutant. In addition, static biofilm formation was shown to be dependent on the presence or absence of Atl and independent of CidA (Fig. 4A–C). Together, these data demonstrate that the previous model for the positive effect of cidA on cell death and lysis was based on a strain that contained a secondary site mutation, thus, leading to an inaccurate conclusion on its function.

Figure 4. Characterization of ΔcidA and Δatl mutants.

Figure 4

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A) Qualitative murein hydrolase analysis of UAMS-1, KB1050 (cidA::ermR), KB1050 RC (KB1050 with cidA restored), Δatl, and ΔcidA grown on TSA containing 2 mg ml−1 Micrococcus lysodeikticus at 37°C. B) Representative image and C) quantification of static biofilm assay of UAMS-1 and mutants grown under static conditions for 24 h at 37°C. ns, P>0.05; ***P<0.001; ****P<0.0001; One-way ANOVA with Tukey’s multiple comparisons test.

Control of CidC activity

Overall, the data presented in the current study provide a new perspective on CidA and CidB function and suggest that these proteins play antithetical roles in modulating stationary phase cell death by controlling the levels of excreted acetate and acetoin. However, since both CidA and CidB are membrane proteins, it is not readily apparent how they regulate central metabolism. It is unlikely that the differences in acetate and acetoin in culture supernatants of the cidA and cidB mutants resulted from altered transcription of cidC or alsSD as qRT-PCR analysis did not reveal any significant changes in the expression of these genes relative to the wild-type control (Fig S3A). Given that the homolog of CidC in E. coli, PoxB is a peripheral membrane protein whose pyruvate:quinone oxidoreductase activity is realized only after interaction with membrane lipids (Russell et al., 1977), we tested whether CidA or CidB influenced the association of CidC to the membrane. To test this hypothesis, cell membranes were isolated from UAMS-1 and its isogenic ΔcidA, ΔcidB and ΔcidAB double mutants and the presence of CidC was detected using polyclonal antibodies specific to this protein. Inactivation of cidA either by itself or in combination with cidB resulted in a significant increase in the levels of CidC associated with the membrane (Fig 3E). However, a similar increase in the total CidC protein concentration within cells (Fig S3B) was also observed in the ΔcidA and ΔcidAB mutants so it is unclear if the cidA mutation has a direct effect on the membrane localization of CidC, or if it effects CidC production and/or stability. Surprisingly, CidC-dependent acetate generation in the ΔcidAB double mutant was significantly lower than the ΔcidA mutant (Fig 3B) despite high levels of CidC associated with the membrane (Fig 3E), suggesting that CidB may play a role in influencing the activity of CidC, independent of its localization to the membrane.

Discussion

It is well accepted that eukaryotic organisms are uniquely adapted to take advantage of cellular death mechanisms for developmental purposes (Cooper GM, 2009). Similarly, multiple studies have recently revealed that single-celled bacteria can also manipulate pathways that promote cell death within clonal multicellular populations such as a biofilm (Sadykov & Bayles, 2012, Bayles, 2014, Rice & Bayles, 2008). Employing S. aureus as a model organism, our group has previously established that a subpopulation of cells within a developing staphylococcal biofilm undergoes cell death to promote biofilm maturation (Rice et al., 2007). In this case, death and subsequent lysis of a few cells was reported to release sufficient DNA into the environment to act as a scaffold for cell attachment in the maturing biofilm (Rice et al., 2007). Only recently did we find evidence to suggest why death affected only a subpopulation of the biofilm (Thomas et al., 2014). According to our model, acetic acid, a byproduct of glucose metabolism, accumulates within biofilm microcolonies resulting in localized acidic zones within these structures. Since acetate is a weak acid, an environmental pH that trends towards the pKa of acetate (~4.8) would result in the generation of protonated acetate species (CH3COOH) capable of penetrating a subpopulation of cells in microcolonies. Given that intracellular pH is close to neutral, acetic acid translocated from the external environment would dissociate in the cytoplasm resulting in acidification and potentiation of cell death. Additionally, cells also possess mechanisms that maintain intracellular pH homeostasis and limit the potentiation of cell death. In S. aureus the genetic loci that controls cell death is primarily associated with the cidABC and alsSD operons. While CidC is capable of potentiating cell death via acetate generation and ensuing intracellular acidification, AlsSD counters cytoplasmic acidification by consuming protons and producing the neutral metabolite, acetoin (Thomas et al., 2014, Tsang et al., 2008). A LysR-type transcriptional regulator, CidR, transcriptionally activates both cidABC and alsSD operons, leading us to speculate that both pro- and anti-death functions are co-regulated. However, in the present study, we demonstrate that CidR primarily has an anti-death function.

The effects of CidABC and AlsSD on cell physiology are readily observed in the stationary phase following aerobic growth in TSB containing excess glucose (35 mM). We have previously established that the dynamics of cidABC expression observed in biofilm microcolonies that develop under flow cell conditions (with media containing 7 mM glucose) can be sufficiently mimicked during planktonic growth following addition of 35 mM glucose (Thomas et al., 2014). Under these conditions, two alternate transcripts arise from the cidABC locus. During the early exponential growth phase, the cidBC transcript predominates. It was earlier shown that expression of this transcript is positively regulated by the alternate sigma factor, σB (Rice et al., 2004). In the present study, we provide additional evidence to suggest that σB-regulated cidBC expression produces significant concentrations of acetic acid and is responsible for the cell death phenotype observed in the stationary phase of growth. While inactivation of cidC resulted in an expected decrease in acetate production, in-frame deletion of cidB also caused a partial reduction of acetate accumulation in culture supernatants suggesting a role for CidB in promoting CidC function. Similarly, the levels of acetoin were elevated following inactivation of both cidC and cidB presumably due to carbon redirection through the AlsSD pathway. Consistent with the reduced acetate and increased acetoin excretion observed, the ΔcidB mutant also exhibited a reduced rate of cell death in the stationary phase similar to the ΔcidC mutant. It is unlikely that these phenotypes attributed to the ΔcidB mutant would have resulted from a polar effect on cidC as the phenotype was not only complementable, but also similar levels of CidC protein was observed following cidB mutation relative to the wild-type strain (Figs. 3E and S1). Although there is a clear effect of CidB on CidC function, the molecular mechanism underlying this effect remains unknown.

Once acetate levels in the culture supernatant peak during the early post-exponential phase, CidR-dependent cidABC transcription is activated (Rice et al., 2005, Rice et al., 2004). Disruption of the cidR gene unexpectedly resulted in an increased rate of cell death relative to the wild-type strain (Yang et al., 2005). This phenotype is consistent with a pro-survival role for the CidR regulon. How does CidR promote survival despite transcriptional activation of pro-death proteins (CidB and CidC)? It does so by co-expressing cidA and alsSD to counteract the activities of the CidB and CidC proteins. While we have previously shown a role for AlsSD in countering CidC-dependent acetate generation (Thomas et al., 2014), the role of CidA in this process had not been tested. We have now discovered that inactivation of cidA results in increased acetate and decreased acetoin excretion. One possible reason for this phenotypic outcome may be due to the increased abundance of CidC in the cytoplasm and its ensuing localization to the membrane, in the absence of CidA. It is conceivable that in the wild-type strain, CidA may promote instability and increased turnover of CidC by interacting with its presumed binding partner, CidB. Although this may explain why cellular CidC levels increase in ΔcidA mutants, this hypothesis remains to be tested. Regardless of the mechanism by which CidA modulates CidC function, the ΔcidA mutant exhibited an increased rate of cell death relative to the wild-type strain consistent with a pro-survival function for CidA. The observed function for CidA in this study is in contrast to the previously hypothesized role for this protein as a pro-death regulator (Rice et al., 2007). We recently discovered that an earlier ΔcidA mutant (KB1050) had developed a secondary mutation in the gene encoding the major autolysin of S. aureus resulting in incorrect interpretation of its function. Thus, while it is true that CidA exhibits some holin-like properties including similarities in size, membrane localization and oligomerization (Ranjit et al., 2011), as well as pore formation in cidA-overexpressing cells and in reconstituted membranes containing purified CidA (manuscript submitted), its activity appears to promote cell survival by counteracting the pro-death activity of CidB and limiting acetate generation rather than promote cell death and lysis akin to a bacteriophage-encoded holin as previously envisioned (Rice & Bayles, 2008, Wang & Bayles, 2013).

Recent studies in our laboratory have also pointed to CidB as the primary effector of cell death in this system. In these studies, we observed that the increased stationary phase cell death associated with a srrAB mutation was dependent on de-repression of cidABC in this mutant (Windham et al., 2016). Interestingly, increased cidABC expression in the srrAB mutant did not result in increased production of acetate, consistent with a role for CidA in countering CidC activity. However, cell death in the srrAB mutant appeared to result from CidB-dependent ROS generation suggesting that an increased abundance of CidB may be toxic to cells. Collectively, the available evidence indicates that CidB may promote cell death by increasing CidC activity, and by promoting the generation of ROS. Additional studies are underway to clarify the mechanism by which CidB mediates cell death.

Being integral membrane proteins, the exact mechanism by which CidA and CidB modulate metabolic output is unclear. It is also not known whether their effects on CidC are mediated through direct interaction or through alternate indirect means. Although initially studied in Arabidopsis thaliana, functional homologs of CidA-like proteins have been identified in several plant species (Wang & Bayles, 2013). In A. thaliana, mutation of the AtLrgB homolog resulted in intraveinal chlorosis and necrosis in leaves suggesting their involvement in regulating leaf senescence (Yang et al., 2012). Interestingly, a link to carbohydrate metabolism for AtlrgB was also established in these studies (Yang et al., 2012). However, subsequent studies revealed AtlrgB to be a chloroplast protein whose effects on cell death and carbon partitioning were presumably due to accumulation of glycerate or glycolate to toxic concentrations (Pick et al., 2013). Based on these observations and their ability to form higher order oligomers in cell membranes, we speculate that CidA and/or CidB may also have transport related functions that may influence central metabolism and consequently, the activity of CidC on the membrane.

A role for the alternate sigma factor, σB, in biofilm development has previously been demonstrated (Lauderdale et al., 2009). While inactivation of σB resulted in decreased biofilm development, its overexpression was shown to increase the frequency of microcolonies in the biofilm (Bateman et al., 2001, Rachid et al., 2000). It is tempting to speculate that cell death due to σB-mediated cidBC expression may at least partially account for the observed phenotypes. Indeed, consistent with this hypothesis we previously observed decreased biofilm microcolony development in the ΔcidC mutant (Thomas et al., 2014).

In conclusion, the present study provides clear evidence that the stationary phase cell death phenotype observed following growth in the presence of excess glucose is dependent on the activities of CidB and CidC proteins (Fig 5). During the exponential phase, expression of cidBC results in the conversion of excess pyruvate into acetate. While this may be beneficial at this stage as functional pyruvate:menaquinone oxidoreductase has been shown to be coupled to aerobic respiration due to its ability to shuttle electrons to quinone intermediates (Koland et al., 1984), the excess acetate generated results in cytoplasmic acidification during the post-exponential phase and eventually potentiates cell death (Rice et al., 2005, Thomas et al., 2014). To modulate the rate of cell death in the population, S. aureus activates the expression of cidA and alsSD, both members of the CidR regulon. While CidA may destabilize CidC and promote its degradation, AlsSD counters CidC mediated cytoplasmic acidification by redirecting excess pyruvate to the neutral metabolite, acetoin and by consuming cytoplasmic protons during this process (Thomas et al., 2014). Our model confirms that both CidA and CidB antithetically affect the extracellular concentrations of acetate and acetoin, consistent with their ability to modulate cell death by altering glucose catabolism. These observations reveal a previously unacknowledged and critical role for membrane-anchored members of the CidR regulon in controlling cell death.

Figure 5. Physiological role of CidR.

Figure 5

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Under conditions of high glycolytic flux, a σB dependent upregulation of cidBC expression promotes the generation of acetate (overflow metabolism). Although CidC by itself is capable of producing acetate once it is localized to the membrane, its full activation requires the presence of CidB. Due to its weak acid nature, acetate can in turn potentiate staphylococcal cell death in the stationary phase by cytoplasmic acidification. To limit the rate of cell death, cells upregulate the expression of cidABC and alsSD via. the LysR type transcriptional activator, CidR. While AlsSD consumes protons and competes with CidC for its substrate, pyruvate (not shown in figure), the activity of CidA suppresses cytoplasmic abundance of CidC through an as yet unknown mechanism.

Experimental Procedures

Bacterial strains, plasmids and growth conditions

The bacterial strains used in this study are listed in Table S1. S. aureus strains were cultured in Tryptic Soy broth (TSB) supplemented with either 0.25% or 0.63% glucose. For long-term growth, bacterial cultures were aerobically grown at 37°C in Erlenmeyer flasks fitted with bug stoppers to minimize evaporation. E. coli strains were cultured in Luria Bertani (LB) broth. When appropriate, antibiotics were added to cultures as follows: ampicillin (100 μg/ml); erythromycin (5 μg/ml); tetracycline (10 μg/ml); and chloramphenicol (10 μg/ml).

Chromosomal deletion mutations were generated as previously described (Bose et al., 2012). Generation of the Δpta and ΔackA mutants are described in a previous study (Sadykov et al., 2013). Double chromosomal mutations in the Δpta and ΔackA genetic backgrounds were generated by phi11-mediated phage transduction. Previously, we reported the construction of ΔcidA and ΔcidB mutants (Windham et al., 2016) The cidC gene was sequenced from both ΔcidA and ΔcidB mutants to confirm the absence of any polar effects that may have occurred during the gene deletion process. The cidAB mutant was constructed by amplifying the cidB deletion from KB1060 and ligating into pJB60, which contains the upstream of cidA (Windham et al., 2016). This plasmid, pJB88, was used for allelic exchange to generate KB1068, with deletion of cidB and ~38% of cidA, effectively deleting these genes while leaving both the PcidABC and PcidBC promoters intact. Complementation of ΔcidA, ΔcidB and ΔcidAB mutants were carried out by markerless allelic replacement of the mutated sites within the cid locus using pVCT11. The plasmid, pVCT11 was created by PCR amplifying cidAB from S. aureus UAMS-1 chromosome using primers (Compl. CidA/B- F 5′-AGTCGAATTCGCATGTCGGCAGTCATGAAT-3′ and (Compl. CidA/B- R 5′-CATGGTCGACGCCATTAATCGTTCTGGACG-3′) engineered with EcoR1 and Sal1 restriction sites. Following restriction with EcoR1 and Sal1, the PCR product was ligated into EcoR1/ Sal1 restricted pJB38 (Bose et al., 2013) and resulting plasmid was maintained in E. coli Electro 10 Blue (Stratagene). All plasmids used for mutagenesis were temperature-sensitive and allelic exchange was performed as previously described (Bose et al., 2013, Bose, 2014).

Electron paramagnetic resonance (EPR) spectroscopy

EPR analysis was carried out as previously described (Thomas et al., 2014). Briefly, three-day old stationary phase bacterial cells were resuspended to an OD600 of 10 units in 1 ml KDD buffer (Krebs-HEPES buffer, pH 7.4; 99 mM NaCl, 4.69 mM KCl, 2.5 mM CaCl2, 1.2 mM MgSO4, 25 mM NaHCO3, 1.03 mM KH2PO4, 5.6 mM D-glucose, 20 mM HEPES, 5 μM DETC and 25 μM deferoxamine). The bacterial samples were then mixed with 200 μM cell-permeable ROS sensitive spin probe 1-hydroxy-3-methoxycarbonyl-2,2,5,5-tetramethylpyrrolidine (CMH; Noxygen Science Transfer and Diagnostics, Elzach, Germany) and incubated for 15 minutes at ambient temperature. EPR analysis was carried out using a Bruker e-scan EPR spectrometer with the following settings: field sweep width, 60.0 gauss; microwave frequency, 9.75 kHz; microwave power, 21.90 mW; modulation amplitude, 2.37 gauss; conversion time, 10.24 ms; time constant, 40.96 ms.

Growth and viability assays

For long-term cell viability assays, sample aliquots were taken every 24 h up to a maximum of 5 days and plated on TSB. Following incubation at 37°C, bacterial colonies were counted and colony-forming units determined. To monitor growth, S. aureus cultures were resuspended to an OD600 of 0.06 in TSB (35 mM glucose) and dispensed into 96-well microtiter plates. Bacterial cultures were grown for 24 h at 37°C in a Tecan infinite 200 spectrophotometer under maximum aeration. The optical density (OD600) was recorded every 30 minutes for the entire period of growth. In these experiments bacteria were either challenged with 30mM pyruvic acid or the pH of the media adjusted to pH 4.5 with concentrated hydrochloric acid followed by supplementation of sodium pyruvate.

Metabolite assays

Aliquots of bacterial cultures (1 ml) were centrifuged for 3 min at 14,000 rpm at 4°C. The supernatants were removed and stored at −20°C until use. Acetate was measured from culture supernatants using a commercial kit (R-Biopharm, Marshall, MI), according to the manufacturer’s instructions. Acetoin was measured at 560nm as previously described (Thomas et al., 2014).

qPCR analysis

Quantitative Reverse-transcriptase PCR was performed as described previously (Sadykov et al., 2013). Briefly, cDNA was synthesized from 500 ng of total RNA using the Quantitect Reverse Transcription Kit (Qiagen). The cDNA samples were then diluted 1:20 and subsequently used as template for PCR reactions. PCR amplification was carried out using the LightCycler DNA Master SYBR green I kit (Roche Applied Science) following the manufacturer’s protocol. The relative transcript levels were calculated using the comparative threshold cycle (CT) method and normalized to the amount of housekeeping Sigma factor A (sigA) transcripts present in the RNA samples.

Cell fractionation and CidC detection

S. aureus membrane preparations were prepared as previously described with the following modifications (Ranjit et al., 2011). Bacterial strains were inoculated into fresh TSB to an optical density at 600nm of 0.06 and allowed to grow at 37°C and 250 rpm for 6 hours. Cells were harvested by centrifugation (4°C) at 7000 x g for 15 minutes and resuspended in 1ml phosphate-buffered saline (PBS) supplemented with 100 U/ml of lysostaphin, 10 U/ml DNase I (Sigma) and 1X protease inhibitor mixture (Complete minikit, EDTA free; Roche). The cultures were incubated at 37°C for 40 minutes and then lysed with two passes through a high-pressure homogenizer (EmulsiFlex-C3; Avestin Inc., Germany) at 15,000 lb/in2. Cell debris was removed by centrifugation at 10000 × g at 4°C for 12 minutes. Crude membranes were pelleted by centrifugation at 100000 × g for 1 h at 4°C using a Beckman L8 60M centrifuge and 45Ti rotor. The pellet, composed of the crude membrane fraction, was resuspended in membrane buffer (100 mM Tris-HCl [pH 7.5], 100 mM NaCl, 10 mM MgCl2, 10% glycerol, and 1% dodecyl maltoside [DDM]) to extract and solubilize the membrane proteins. This was again centrifuged at 100000 × g as described above to remove the detergent-insoluble material. The concentration of proteins in the resultant supernatant, containing the detergent-soluble membrane proteins, was measured using a Bio-Rad protein assay kit. For CidC detection, total membrane associated protein concentration of 1 μg was separated on a 8% Tris-glycine SDS-polyacrylamide gel and western blotting was performed using a 1:1000 dilution of rabbit primary polyclonal antibodies raised against staphylococcal CidC protein.

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Acknowledgments

This work was funded by NIH grant no. R01-A1038901, P01-AI083211 to K.W.B. and American Heart Association postdoctoral fellowship to V.C.T. The EPR spectroscopy core is supported, in part, by a NIH Center of Biomedical Research Excellence (COBRE) grant (1P30GM103335) awarded to the University of Nebraska’s Redox Biology Center. The funders had no role in study-design, data collection, interpretation and decision to submit this work for publication.

Footnotes

The authors declare they have no conflict of interest.

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