CodY controls the SaeR/S two-component system by modulating branched-chain fatty acid synthesis in Staphylococcus aureus

ABSTRACT

Staphylococcus aureus is a Gram-positive, opportunistic human pathogen that is a leading cause of skin and soft tissue infections and invasive disease worldwide. Virulence in this bacterium is tightly controlled by a network of regulatory factors. One such factor is the global regulatory protein CodY. CodY links branched-chain amino acid sufficiency to the production of surface-associated and secreted factors that facilitate immune evasion and subversion. Our previous work revealed that CodY regulates virulence factor gene expression indirectly in part by controlling the activity of the SaeRS two-component system (TCS). While this is correlated with an increase in membrane anteiso-15:0 and −17:0 branched-chain fatty acids (BCFAs) derived from isoleucine, the true mechanism of control has remained elusive. Herein, we report that CodY-dependent regulation of SaeS sensor kinase activity requires BCFA synthesis. During periods of nutrient sufficiency, BCFA synthesis and Sae TCS activity are kept relatively low by CodY-dependent repression of the ilv-leu operon and the isoleucine-specific permease gene brnQ2. In a codY null mutant, which simulates extreme nutrient limitation, de-repression of ilv-leu and brnQ2 directs the synthesis of enzymes in redundant de novo and import pathways to upregulate production of BCFA precursors. Overexpression of brnQ2, independent of CodY, is sufficient to increase membrane anteiso BCFAs, Sae-dependent promoter activity, and SaeR ~P levels. Our results further clarify the molecular mechanisms by which CodY controls virulence in S. aureus.

IMPORTANCE

Expression of bacterial virulence genes often correlates with the exhaustion of nutrients, but how the signaling of nutrient availability and the resulting physiological responses are coordinated is unclear. In S. aureus, CodY controls the activity of two major regulators of virulence—the Agr and Sae two-component systems (TCSs)—by unknown mechanisms. This work identifies a mechanism by which CodY controls the activity of the sensor kinase SaeS by modulating the levels of anteiso branched-chain amino acids that are incorporated into the membrane. Understanding the mechanism adds to our understanding of how bacterial physiology and metabolism are linked to virulence and underscores the role virulence in maintaining homeostasis. Understanding the mechanism also opens potential avenues for targeted therapeutic strategies against S. aureus infections.

INTRODUCTION

Staphylococcus aureus is a Gram-positive bacterium found commonly on human skin and in the anterior nares. Up to 30% of the human population may be stably colonized by this pathogen without experiencing any symptoms. As an opportunistic human pathogen, S. aureus is the leading cause of skin and soft tissue infections. These infections can progress to devastating invasive infections including infectious endocarditis, osteomyelitis, and sepsis (1, 2). S. aureus nosocomial infections have been observed in hospitals for decades, but S. aureus now circulates in communities (so-called community-acquired [CA] infections). Methicillin-resistant Staphylococcus aureus (MRSA) and strains resistant to other antimicrobials have compounded the problem. Over the past decade, CA-MRSA has killed tens of thousands of people (3), and CA-MRSA infections cost the U.S. healthcare system between $560 million and $2.7 billion in annual treatment-associated costs (4).

S. aureus can infect nearly every organ of the body. The success of S. aureus as a pathogen has been attributed to its vast repertoire of surface-associated and secreted virulence factors that enhance host colonization and facilitate evasion of the host immune response (5). However, from a resource and energy conservation perspective, the production of all virulence factors simultaneously is likely to be costly to the bacterium. Therefore, the expression of virulence factors is tightly regulated by a network of transcription factors, two-component systems, and small regulatory RNAs, whose activities are influenced by the environment of the infection niche (6–9). Interfering with the regulation and production of virulence factors may be our best hope for combating pathogens resistant to available antimicrobial therapies (10). For instance, in collaboration with Dufresne and colleages, we recently identified a compound that interferes with toxic shock syndrome toxin-1 production (11).

The production of virulence factors in S. aureus is controlled in large part by the Sae two-component system (TCS). Sae is known to regulate the expression of over 20 surface-associated and secreted virulence genes such as hla (ɑ- hemolysin), coa (coagulase), nuc (thermonuclease), and multiple protease genes (12–15). The sae locus consists of a weak constitutive P3 promoter that drives the expression of the response regulator gene saeR and the histidine kinase gene saeS. In addition, an inducible P1 promoter controls the transcription of two auxiliary genes saeP and saeQ along with saeRS. SaeS is an atypical histidine kinase (16–18). Unlike typical histidine kinases that contain a large extracellular domain that binds a ligand to transduce signals across the membrane, SaeS belongs to the intramembrane family of histidine kinases (19) that lack this domain. Indeed, SaeS only has a nine-amino acid residue extracellular-facing peptide that links two transmembrane helices. The linker and membrane helices comprise the N-terminal domain. This N-terminal domain constrains SaeS kinase activity in the cytosol at the C-terminus (20). In the presence of neutrophil-produced factors such as human neutrophil peptides 1, 2, and 3 (HNP1-3), SaeS is phosphorylated on a conserved histidine residue (21). Activation by HNPS occurs via an unknown mechanism. The phosphoryl group is then transferred to a specific, conserved aspartate residue in SaeR (to generate SaeR ~P). SaeP is a lipoprotein, and SaeQ is a transmembrane protein. Previous work has shown that SaeP and SaeQ form a complex with SaeS and stimulate the phosphatase activity of SaeS toward SaeR by an unknown mechanism. This lowers cellular SaeR ~P levels and returns the Sae TCS to the pre-induced state (22). SaeR ~P binds directly to dozens of targets with variable affinity (12, 23). This includes the sae P1 promoter, triggering the production of SaeP and SaeQ, creating a negative feedback loop to prevent Sae activity from becoming unlimited.

Branched-chain fatty acids (BCFAs) are the most abundant membrane fatty acids in staphylococcal membranes, and we previously reported that BCFA levels are correlated with SaeS kinase activity (24). The synthesis of BCFAs in S. aureus begins with the amino acids isoleucine, leucine, and valine (ILV). These amino acids can be imported into the cell through specific transporters such as BrnQ1, BrnQ2, and BcaP, or they can be synthesized within the cell under certain conditions (25–28). After ILV import and transamination by the enzyme IlvE, the resulting branched-chain α-keto acids undergo oxidative decarboxylation to form branched-chain carboxylic acids (BCCAs) that are subsequently activated to their acyl-CoA derivatives. Under laboratory conditions, this series of reactions is catalyzed by the branched-chain α-keto acid dehydrogenase complex (BKDH) and associated coenzymes (29). The acyl-CoAs are then primed by the FabH enzyme, which catalyzes their condensation with malonyl-ACP. The resulting β-ketoacyl-ACP is further elongated by the type II fatty acid synthase (FASII) before being incorporated into phospholipids (30). In S. aureus, the most abundant BCFAs in the cell membrane are iso (i) fatty acids and anteiso (a) fatty acids derived from isoleucine, specifically a15:0 and a17:0 (25, 31).

CodY (AKA, Controller of dpp) was first discovered in Bacillus subtilis as a repressor of dipeptide permease (dpp) and then later a repressor of a variety of permease genes, biosynthetic genes, and alternative nutrient processing enzyme genes during rapid growth (i.e., metabolic genes) (32–34). We now know that CodY is a global regulatory protein that controls the expression of dozens of genes in many low G + C Gram-positive bacterial genera, including Bacillus, Listeria, Stapylococcus, Clostridioides, Lactococcus, and Streptococcus (35–45). In pathogens like S. aureus, CodY also regulates the expression of virulence genes (42, 43, 46). As such, CodY serves as an important linkage between metabolism and pathogenic potential (47). CodY is activated as a DNA-binding protein when bound to ILV and GTP (42, 43, 48–53). CodY interacts with a site-specific DNA sequence essential for binding target genes (AATTTTCWGAAAATT) originally determined in Lactococcus lactis and validated in S. aureus (36, 42). CodY prioritizes gene expression based in part on DNA-binding activity (controlled by availability of the corepressors) and affinity of CodY for its many sites to facilitate the bacterium’s adaptation to its environment (37, 54–56). For many of the metabolic genes, CodY directly represses gene expression at target promoters. In contrast, CodY control of virulence in S. aureus is indirect via two major regulators of virulence—the Agr quorum sensing system and the Sae TCS (24, 42, 57, 58). This control mediates the production of secreted digestive enzymes and cytotoxins to replenish nutrients during infection when amino acid and energy reserves are low. In support, deleting codY results in hypervirulence in mouse models of skin and soft tissue infection (SSTI) and necrotizing pneumonia (59). At least for SSTIs, this is due to CodY-dependent regulation of hla (α-toxin) (24).

Although CodY adjusts the expression of the sae locus by direct and indirect mechanisms at the sae P1 promoter (58), we previously showed that this transcriptional control is dispensable for virulence gene upregulation in the codY mutant. Rather, CodY controls the activity of the SaeS kinase by an unknown mechanism correlated with changes to membrane fatty acid content (24). Herein, we report that CodY-deficient strains disrupted for de novo ILV synthesis and isoleucine import exhibit reduced Sae-dependent gene expression and SaeS kinase activity. This is correlated with lower levels of anteiso 15:0 and 17:0 branched-chain fatty acids derived from isoleucine. This low Sae TCS activity phenotype is complemented genetically and also chemically by supplementing mutant cells with exogenous branched-chain carboxylic acids. We show that overexpression of the isoleucine-specific permease brnQ2 is sufficient to activate Sae TCS activity in wild-type (WT) cells during laboratory growth, and this increased activity is correlated with the levels of the anteiso 15:0 and 17:0 BCFAs in the membrane. Our results further clarify the mechanism by which CodY controls a major virulence regulator to optimize virulence factor production during nutrient depletion.

RESULTS

Branched-chain fatty acid synthesis is required to upregulate Sae TCS activity when CodY activity is reduced

We previously demonstrated that BCFAs are essential for Sae TCS activity, and BCFA content is increased in ΔcodY mutant cells (24). We wondered whether a15:0 BCFAs are required to upregulate Sae TCS activity when codY is disrupted. To test this, we used a mutant strain that synthesizes i14:0 BCFAs for growth, but not a15:0 BCFAs to promote Sae activity, and performed an epistasis experiment using the Sae-dependent saeP1-gfp reporter fusion to indirectly measure Sae activity. The strain (ΔlpdA mbcS1) is devoid of BKDH activity and overexpresses a gene coding for methylbutryl-CoA synthetase (MbcS). MbcS activates exogenously supplied branched carboxylic acids (i.e., isobutyric acid; iC₄) (60, 61). As expected, when we disrupted codY in an otherwise WT background, saeP1-gfp promoter activity increased twofold during growth in rich, complex medium (i.e., tryptic soy broth [TSB]). We measured very low promoter activity in the ΔlpdA mbcS1 mutant (Fig. 1, compare lpdA mbcS1 and codY to WT). When we disrupted codY in the lpdA mbcS1 background, promoter activity was indistinguishable from that measured in the lpdA mbcS1 strain. Promoter activity was restored when we supplemented ΔlpdA mbcS1 double-mutant and ΔlpdA mbcS1 ΔcodY triple-mutant cells with 0.5 mM of the individual short, branched-chain carboxylic acids aC₅ (2-methylbutyric acid) and iC₄ (isobutyric acid), but not iC₅ (3-methylbutyric acid). We saw no additive or negative effects when we provided a mixture of all three carboxylic acid precursors (Fig. 1). These data indicate that ΔlpdA mbcS1 is epistatic to ΔcodY and that BCFAs are required for CodY-dependent upregulation of Sae TCS activity. The effect is specific for some but not all BCFAs, but we did not test whether there was a preference for anteiso BCFAs over iso BCFAs.

Fig 1.

CodY control of Sae TCS activity requires BCFAs. The indicated USA300 LAC strains harboring the saeP1–gfp reporter fusion were grown in TSB or TSB supplemented with 0.5 mM BCFA precursors aC₅ (2-methylbutyric acid), iC₅ (3-methylbutyric acid), or iC₄ (isobutyric acid) for 16 hours. Data are plotted as mean ± SEM of three biological replicates. *P  <  0.05, **P  <  0.01, ***P  <  0.001, and ****P  <  0.0001; one-way ANOVA with Tukey’s multiple comparison test within each treatment group. ns, not significant.

De novo amino acid biosynthetic or salvaging pathways are required for CodY-dependent regulation of Sae TCS activity

During laboratory growth in ILV-replete medium, BCFAs are synthesized from ILV, and the membrane fatty acid composition depends in part on the levels of exogenous amino acids and selectivity of the BKDH complex for isoleucine (25). CodY represses genes coding for the enzymes that direct de novo ILV biosynthesis (ilv-leu) and the ILV permeases brnQ1 and brnQ2 (42, 55). BrnQ1 transports all three branched-chain amino acids (BCAAs), whereas BrnQ2 transports isoleucine specifically (26). We hypothesized that CodY restricts BCFA synthesis during conditions of BCAA sufficiency and keeps Sae TCS activity low. During BCAA limitation, reduced CodY activity would result in upregulation of ILV import and synthesis pathways. This would lead to increased branched-chain ɑ-keto acid levels that would feed branched-chain fatty acid synthesis and upregulate Sae activity when incorporated into the membrane. To test this hypothesis, we disrupted de novo ILV biosynthesis or import genes in the ΔcodY mutant and monitored Sae activity indirectly using the Sae-dependent nuc–gfp reporter fusion (15, 55) during growth in chemically defined medium (CDM). We used CDM to simplify the experimental conditions as TSB contains peptides as well as free amino acids. As expected, nuc-gfp promoter activity increased by ~fourfold in the ΔcodY mutant. Deleting ilvD or disrupting brnQ1 or brnQ2 in the ΔcodY mutant background does not significantly reduce nuc-gfp activity (Fig. 2A, compare double mutants to the codY mutant). However, synthesis and import are redundant pathways for maintaining ILV levels and BCFA precursor pools. Therefore, we constructed triple mutant strains. While disrupting ilvD and brnQ1 in the ΔcodY mutant did not alter nuc-gfp promoter activity, when we mutated both ilvD and brnQ2 in the ΔcodY mutant, we measured a twofold drop in promoter activity (Fig. 2A). We did not mutate additional permease genes in this background (i.e., bcaP) as this would likely result in a confounding growth defect in CDM. We then analyzed the membrane fatty acid content of the ΔcodY ΔilvD and ΔcodY ΔilvD ΔbrnQ2 strains using gas chromatography analysis of fatty acid methyl esters (GC-FAME). Sae-dependent promoter activity was correlated with anteiso BCFA levels. Specifically, we measured a decrease in a15:0 and a17:0 BCFAs in the triple mutant strain. nuc-gfp promoter activity, BCFA content, and secreted nuclease production phenotypes were complemented chemically when we included a mixture of the carboxylic acid precursors in the medium or genetically when we expressed brnQ2⁺ from the multicopy plasmid pRMC2 (Fig. 2B through D).

Fig 2.

CodY control of Sae TCS activity requires de novo α-keto acid synthesis or isoleucine import. (A and B) The indicated LAC strains were grown for 16 hours in chemically defined medium (CDM), at which time nuc-gfp promoter activity was measured. (C and D) The indicated strains were grown to exponential phase in CDM, and then secreted nuclease activity was measured and membrane fatty acid content was analyzed using GC-FAME. For all panels, data are plotted as mean +/- SEM of at least three independent experiments. *P  <  0.05; **P  <  0.01; ***P <  0.001, and ****P <  0.0001. Significance was determined using one-way ANOVA with repeated measures and Tukey’s post-test for (A and C) or two- way ANOVA for (B and D). BCCAs, 0.5 mM each of aC₅, iC₄, and iC₅; ns, not significant. VOC, vector-only control.

If CodY controls the Sae TCS by controlling the synthesis of BCFAs, then uncoupling brnQ2 or ilv-leu expression from CodY regulation and overexpressing the gene(s) in a WT background should phenocopy the ΔcodY mutant for heightened Sae activity. To test this, we expressed brnQ2⁺ using the inducible P_tet promoter in WT cells and monitored Sae TCS activity using the saeP1-gfp reporter fusion. As expected, promoter activity increased > twofold in the ΔcodY mutant relative to WT; the fusion is Sae-dependent (Fig. S1). Compared to the vector control, we observed a dose-dependent increase in saeP1-gfp promoter activity in WT cells as we increased the concentration of anhydrotetracycline in the medium. At higher levels of aTc (e.g., 25 ng mL⁻¹), the promoter activity exceeded that measured in the ΔcodY mutant (Fig. 3A; Fig. S1). This increase in SaeS activity was not observed in WT cells containing saeP1-gfp and the pRMC2 vector-only control (Fig. 3A). Again, increased promoter activity was correlated with a significant increase in 15:0 anteiso BCFAs. We observed a trend toward increased 17:0 anteiso BCFA levels in brnQ2-expressing cells, though the difference was not statistically significant compared to the vector-only control (Fig. 3B). We note that the growth conditions used maximize CodY’s DNA-binding activity. As such, overexpression of brnQ2 does not affect the transcription of the sae and ilv genes under the conditions tested (Fig. S2).These data strongly suggest that de novo amino acid biosynthetic or salvaging pathways are required to upregulate Sae activity when CodY activity is reduced. Furthermore, overexpression of brnQ2 results in the same activation phenotype, implying that increasing BCFA abundance in the membrane, but not differential production of membrane proteins under CodY control, is required. The mechanism is consistent with those found in our previous work, revealing that Sae TCS activity is affected by isoleucine-derived phospholipids (24).

Fig 3.

Overexpressing the isoleucine permease gene brnQ2 results in higher levels of anteiso BCFAs and SaeS activity. Strains were grown to the exponential phase in CDM with and without anhydrotetracycline (aTC). Cells were collected, and (A) saeP1-gfp promoter activity, (B) membrane fatty acid composition, and (C) intracellular SaeR protein species were measured. SaeR was detected in SDS-polyacrylamide gels with and without Phos-Tag reagent using polyclonal antibodies raised against SaeR. SrtA was detected as a loading control (upper; representative gel). Densitometric quantification was performed using ImageJ (lower). Data are plotted as mean +/- SEM of at least three independent experiments. *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001; one-way ANOVA with Tukey’s multiple comparison test. VOC, vector-only control; ns, not significant; nd, not detected.

Unphosphorylated SaeR does not bind target promoters. Rather, activated SaeR (SaeR ~P) binds to stimulate gene transcription (23). Furthermore, each histidine kinase–response regulator pair functions autonomously. Crosstalk between noncognate HK-RR pairs rarely occurs (62). Therefore, SaeS kinase activity solely determines the cellular levels of SaeR ~P. To test directly whether overexpressing brnQ2 increases SaeS kinase activity, we used Phos-Tag electrophoresis (63) and Western blotting with SaeR antisera to measure the relative levels of SaeR and SaeR ~P in WT cells expressing brnQ2 from the inducible P_tet promoter. As expected, the fraction of SaeR ~P was ~10% in WT cells in the pre-induced state; this increased to ~20% in the ΔcodY mutant. Compared to vector controls, when we overexpressed brnQ2⁺ in WT cells, SaeR ~P levels were essentially identical to those measured in the ΔcodY mutant. This was correlated with a modest increase in total SaeR levels, likely due to autoregulation of sae expression via the sae P1 promoter (Fig. 3C; Fig. S3) (24). Thus, SaeS kinase activity increases when isoleucine import via BrnQ2 increases.

The mechanism of BCFA-dependent upregulation of Sae activity by CodY appears to be specific and causative

We wondered whether other conditions known (or have the potential) to affect BCFA content also affect Sae activity. For instance, Bacillus subtilis and Listeria monocytogenes can adapt to cold temperatures by increasing levels of 15:0 anteiso BCFAs (64, 65). L. monocytogenes can grow at refrigeration temperature, an ability that contributes to the bacterium’s notoriety as a foodborne pathogen. B. subtilis (but not S. aureus) can also desaturate pre-existing fatty acids via a cold-shock-induced, membrane-bound phospholipid desaturase called Des (66, 67). S. aureus can survive on fomites to aid transmission between hosts and may survive in extreme cold conditions for short periods of time (29, 68). The latter condition is likely rare. However, the bacterium does change its transcriptome and proteome when shifting between three physiologically relevant temperatures: 34°C (the nares), 37°C (internal body), and 40°C (extreme pyrexia [AKA fever]). Mining these data, we failed to find a strong association between upregulated gene expression or protein production of direct, SaeR targets during these temperature transitions (69) (data not shown). As we did not test it, we cannot exclude the possibility that temperatures < 34°C result in increased BCFA production and heightened Sae activity.

Daptomycin is a cationic, cyclic lipopeptide membrane-targeting antibiotic currently used as a last resort for drug-resistant S. aureus infections (70–72). Treatment failure is attributed, in many cases, to resistance to daptomycin. Resistance is conferred by mutations in the membrane phospholipid biosynthesis pathway or in regulatory proteins that mediate cell envelope stress response and membrane homeostasis (73, 74). It was shown recently that strains with evolved high-level daptomycin resistance express a mutant allele of pgsA (phosphatidylglycerol synthase). The strains have increased levels of branched-chain fatty acids in their membrane phospholipids, increased membrane fluidity, and thickened cell walls (73, 75). We wondered if these daptomycin-resistant strains exhibit increased Sae activity. As shown in Fig. S4; Fig. 4A, the daptomycin-resistant strain in fact showed reduced secreted nuclease activity and reduced saeP1-gfp promoter activity, suggesting SaeS kinase activity was actually impaired, not enhanced (compare D8 to N315 parent strain, solid bars). In both strains, SaeS could be induced using HNP1 (Fig. 4B). We reasoned that the defect in phospholipid synthesis may result in unincorporated free fatty acids, which have been shown to inhibit the Sae TCS (76, 77). To test this hypothesis, we used fatty acid-free bovine serum albumin (BSA) to remove fatty acids from these cells by providing an extracellular sink for these lipids (77). As expected, BSA treatment resulted in a modest but reproducible increase in Sae activity in WT cells. There was a trend toward a slight increase in the Sae activity in the daptomycin-resistant mutant, but this failed to reach significance (Fig. 4A, compare striped bars vs. solid bars). Increasing BSA had no further effect (data not shown). The defect in Sae-dependent nuclease production and secretion was complemented with a wild-type copy of pgsA in trans (Fig. S4). As we have discovered that multiple lipids are required to maintain full Sae activity (24, 78), one possibility is that additional changes to the membrane may be masking the effect of increased membrane BCFAs. We note that exposure of WT cells to sub-inhibitory concentrations of daptomycin also did not induce Sae activity, which is in agreement with the observations of previous studies (21) (data not shown).

Fig 4.

A daptomycin-resistant strain with increased membrane BCFA levels has reduced Sae activity. (A) The indicated strains were grown for 16 hours in TSB with or without fatty acid-free bovine serum albumin (BSA) supplementation. Then, saeP1-gfp promoter activity was measured. (B) Indicated strains were grown in TSB to the exponential phase, at which time cells were exposed to HNP1 for 2 hours. Subsequently, cell supernatants were collected, and secreted nuclease activity was measured. For all panels, data are plotted as mean +/- SEM of at least three biological replicates. *P < 0.05; ****P < 0.0001. (A) One-way ANOVA with Tukey’s post-test. ns, not significant, (B) Unpaired t-test, with Holm–Šídák post-test.

DISCUSSION

SaeS is a member of the intramembrane family of histidine kinases for which relatively little is known. A defining feature of these kinases is the lack of a large extracellular ligand/signal-binding domain that perceives environmental stimuli (19). Rather, a short peptide links two transmembrane helices. The prevailing view for SaeS is that the conformation of the N-terminal domain controls the activity of the C-terminal cytoplasmic kinase domain (20). In this context, it is conceivable that any signal, internal or external, can influence SaeS kinase activity to upregulate the expression of virulence genes by altering the membrane environment. Indeed, human neutrophil peptides, which are produced by immune cells and perforate the bacterial membrane, induce Sae TCS activity by an unknown mechanism. We previously discovered that CodY controls nearly all S. aureus virulence genes indirectly, in part, by controlling the activity of the Sae TCS (55, 58). However, the mechanism by which CodY upregulates Sae activity has remained unclear. Herein, we show that the upregulation of Sae TCS activity in ΔcodY mutant cells depends on anteiso BCFA synthesis using exogenous or endogenous branched-chain carboxylic acid precursors. Endogenous synthesis of the precursors is controlled by both de novo synthesis via the ILV biosynthetic pathway or by ILV import and transamination by IlvE. The result is an increase in SaeS kinase activity and cellular SaeR ~P levels to stimulate Sae-dependent promoters.

Our findings build upon those of previous work by Pendleton et al., demonstrating the essential role of BCFAs in Sae TCS activity and informing our working model (Fig. 5). The question of how exactly BCFAs affect the SaeS signaling complex remains. As described above, the prevailing view is that the conformation of the transmembrane portion of SaeS controls activation, but how HNPs alter the conformation is an open question. A growing body of research suggests that the transmembrane helices connect the IM-HKs to additional membrane proteins (79). SaeS was reported to be localized in functional membrane microdomains (FMMs), regions of low fluidity in an otherwise fluid membrane (80). One possibility is that anteiso BCFAs specifically facilitate the diffusion of an activator protein through the bilayer into the FMM or facilitate SaeS diffusion out of the FMM to interact with the activator protein. On the other hand, anteiso BCFAs may facilitate the dispersal of inhibitor proteins out of the SaeS signaling complex. Indeed, SaeS is known to interact with auxiliary membrane proteins SaeP and SaeQ, which induce SaeS phosphatase activity by an unknown mechanism (22). This returns Sae TCS activity to its pre-induced state. However, at least for CodY-dependent regulation, SaeP and SaeQ are not involved, as CodY control is retained in an SaePQ-deficient strain (24). We cannot exclude the possibility that HNP-mediated induction occurs in an SaePQ-dependent manner. Another variation on this theme is that other protein–protein interactions are affected in an anteiso BCFA-dependent manner that somehow indirectly affects the activity of SaeS. We are currently working to identify proteins that interact with the SaeS signaling complex.

Fig 5.

Working model for proposed mechanisms of SaeS activation by BCFA in Staphylococcus aureus. During ILV limitation, the inactivation of CodY leads to the de-repression of brnQ2 and the ilv-leu operon, enhancing the import and biosynthesis of isoleucine. Subsequent metabolic conversion of isoleucine to its α-keto acid results in the production of a15:0 and a17:0 BCFAs via BKDH-dependent and BKDH-independent pathways that upregulate SaeS kinase activity and increase cellular SaeR ~P levels. BCFA activation of SaeS may be direct or indirect, altering protein–protein or protein–lipid interactions at the SaeS signaling complex. Green box, potential activator protein; the solid line denotes the BKDH-dependent BCFA synthesis pathway; the dotted arrow denotes a second, BKDH-independent and MbcS-dependent route to BCFA synthesis, as described previously (61).

Lipids have also emerged as important regulatory effectors in prokaryotic and eukaryotic membrane receptors. Indeed, in addition to BCFAs, we have shown that cardiolipin is essential for full SaeS activity (78). BCFAs may bind directly to SaeS to facilitate dimerization, promote kinase activity, or act as an allosteric modulator to induce a conformational change. A BCFA binding site specific to a15:0 (or i14:0) might explain the lack of chemical complementation with iC5. Additionally, titrations with aC5 and iC4 are necessary to determine if there is a preference for fatty acids derived from these carboxylic acids (Fig. 1). Alternatively, a15:0 (or i14:0) may act indirectly either because one of its minor secondary metabolites interacts with SaeS or because it regulates a modification of SaeS. Non-native mass spectrometry of SaeS would reveal the latter, and powerful new lipidomics techniques would reveal the former (81, 82). Thus, key lipids and membrane proteins may act as the true sensors for SaeS, and characterizing these interactions is the current focus of our laboratory. We are also using these approaches to unravel the mechanism of induction by HNPs. Because these new lipids and proteins are important for Sae TCS activity and virulence, they are potentially new anti-virulence targets. Answering these questions would provide insights into how S. aureus upregulates virulence factor production. Moreover, since other Gram-positive bacteria utilize IM-HKs, using SaeS as a model will provide insights into how these systems function generally. We are actively pursuing these questions in our laboratory.

MATERIALS AND METHODS

Bacterial strains and growth conditions

S. aureus strains used in this study are derivatives of USA300 S. aureus LAC, or N315 (Table S1). S. aureus strains were cultured either in tryptic soy broth (TSB, Becton Dickenson formulation containing 2.5 g liter⁻¹ dextrose) or chemically defined medium (CDM) (83). Briefly, CDM was formulated with alanine (672 µM), arginine (287 µM), aspartic acid (684 µM), cysteine (166 µM), glutamic acid (680 µM), glycine (670 µM), histidine (129 µM), isoleucine (228 µM), leucine (684 µM), lysine (342 µM), methionine (20 µM), phenylalanine (240 µM), proline (690 µM), serine (285 µM), threonine (260 µM), tryptophan (50 µM), tyrosine (275 µM), valine (684 µM), thiamine (56 µM), nicotinic acid (10 µM), biotin (0.04 µM), pantothenic acid (2.3 µM), MgCl₂ (1,000 µM), CaCl₂ (100 µM), monopotassium phosphate (40,000 µM), dipotassium phosphate (14,700 µM), sodium citrate dehydrate (1,400 µM), magnesium sulfate (400 µM), ammonium sulfate (7,600 µM), and glucose (27,753 µM). All strains were grown at 37°C. When necessary, media were solidified with agar (1.5% [wt/vol]) and supplemented with the following antibiotics at the indicated concentrations: chloramphenicol (Cm), 5 to 10 µg mL⁻¹; erythromycin (Erm), 5 to 10 µg mL⁻¹; or tetracycline (Tc) at 1.5 µg mL⁻¹. When indicated, media were supplemented with 0.5 mM branched-chain carboxylic acids (aC₅ [2-methylbutyric acid], iC₅ [isovaleric acid], and iC₄ [isobutyric acid]). Except for modifications pertinent to specific experiments described later, cultures utilized in experiments were grown overnight in test tubes from colonies picked from freshly streaked TSB-agar plates and then diluted 1:100 into flasks with the indicated medium (5:1 flask:medium volume). After 5–6 hours, cells were collected by centrifugation and used in experiments. Growth prior to the use of bacteria in individual experiments was monitored by the increase in absorbance at 600 nm (OD₆₀₀) using an Amersham Ultraspec 2100 Pro UV–visible spectrophotometer.

Genetic techniques

Oligonucleotides used in this study were synthesized by Integrated DNA Technologies (IDT; Coralville, IA) and are listed in Table S2. The plasmids used in this study are listed in Table S3. Restriction enzymes, T4 DNA ligase, and Q5 DNA polymerase were purchased from New England Biolabs (NEB). Plasmid and genomic DNA (gDNA miniprep kits were purchased from Promega. Unless indicated, plasmid constructions were performed using Gibson Assembly, as described (84, 85). E. coli NEB 5α (NEB) was used as a host for plasmids, which were then transferred when necessary into S. aureus strain RN4220 by electroporation, as previously described (86). Plasmid and marked mutations were moved between S. aureus strains via Φ85- or Φ11-mediated transduction (87). All plasmids and chromosomal alleles were verified by primer-independent sequencing (PlasmidSaurus) or Sanger Sequencing (Azenta/GeneWiz).

To construct an in-frame ilvD deletion, ~500 bp up- and downstream of the ilvD coding sequence were amplified using S. aureus chromosomal DNA. A third, overlapping PCR was used to fuse the two DNA fragments. PCR reactions used oSRB368–371. This DNA fragment was cut with NcoI and BamHI and ligated to the same sites of pMAD, resulting in pSRB72. Allelic exchange was performed as described previously (88), screening for white, Em^S colonies on TSB XG medium and ILV auxotrophy on CDM .

pDD12 plasmid construction

pOS-saeP1-gfp (a gift from Dr. Taeok Bae, Indiana University, 6,644 bp) harboring the saeP1 promoter region fused to GFP was linearized and amplified using primers oDD171 and oDD172. The tetracycline resistance gene (tetM, 2,265 bp) was amplified from S. aureus integration vector pWY53 [pCT3; (61)] using primers oDD173 and oDD174. Each primer was designed to add 25 bp overlapping for later Gibson assembly. The linearized vector and tetM insert were treated with DpnI (NEB) for 1 hour at 37°C and cleaned up using a PCR purification kit (NEB). Purified PCR products were then subjected to Gibson assembly and then transformed into E. coli DH5α.

Reporter fusion assays

Single colonies of S. aureus strains were freshly streaked out on plates from frozen stocks and were used to inoculate the appropriate medium. For experiments utilizing the ΔlpdA mbcS1 ΔcodY mutant strain, carboxylic acids were added to the medium in each corresponding tube, and cultures were grown overnight before being diluted 1:5 in 100 µL of the total volume per well of 96-well plates. Both OD₆₀₀ and GFP fluorescence (485 nm excitation; 535 nm emission) were read in a computer-controlled BioTek Synergy H1 Multimode plate reader (Agilent) using Gen5 IVD control software (version 1.11). Relative fluorescence units (RFU values) were calculated by dividing the 535 nm emission intensity values by the OD₆₀₀ values. For experiments assaying the effect of branched-chain amino acid biosynthesis or import on Sae TCS activity, inoculated cultures were grown overnight and then diluted into fresh medium (with or without BCCAs, depending on the experiment) to an optical density (OD₆₀₀) of 0.05, before 200-µL aliquots were transferred into each well of a 96-well plate. Both OD₆₀₀ and GFP fluorescence were read every 20 minutes over 16 hours using the same BioTek Synergy H1 Multimode plate reader, and RFU was calculated for each time point. For experiments involving the overexpression of brnQ2⁺, the inoculated cultures were grown overnight in test tubes and then diluted 1:100 into media flasks containing various concentrations of anhydrotetracycline. After 5–6 hours of incubation at 37°C, 100-µL aliquots were transferred to wells of a 96-well plate and then read using the abovementioned plate reader to determine the RFU. Finally, for experiments involving N315 and the evolved daptomycin-resistant strain D8, single colonies were used to inoculate tubes containing TSB +/-fatty acid-free bovine serum albumin (BSA, 10 mg mL^-1). Inoculated cultures were grown overnight (16 hours) and then diluted 1:5 in 1X PBS in a 96-well plate. The RFU was determined using the plate reader as described above.

Nuclease activity quantification via FRET

Secreted nuclease activity was quantified using a fluorescence resonance energy transfer (FRET) assay described previously (89). In brief, cells were grown to the exponential phase in CDM. These cultures were then serially diluted and grown overnight. Cultures that reached an OD₆₀₀ of ~1 after overnight incubation were double back-diluted to an OD₆₀₀ of 0.05 and allowed to grow back to the exponential phase (OD₆₀₀ of 0.5). To obtain secreted nuclease, 0.7 mL of the experimental culture was transferred to 0.22-µm Spin-X centrifuge tube filters (Corning Life Sciences) and centrifuged at 21,000 x g for 3 minutes. Sterilized culture supernatants were frozen at −20°C until use. The N315 strain supernatants were prepared in a slightly modified way. Single colonies were inoculated in tubes containing TSB and were grown overnight (16 hours). To obtain secreted nuclease, 0.5 mL of the resulting overnight culture was collected and stored as described above. The single-stranded oligonucleotide FRET substrate was diluted to a concentration of 2 mM in buffer A (20 mM Tris [pH 8.0], 0.5 M CaCl₂) and then mixed 1:1 with thawed nuclease-containing supernatants that were diluted as described below. Fluorescence, indicative of the substrate cleavage by nuclease (535 nm excitation; 590 nm emission), was measured at 30°C using the BioTek Synergy H1 plate reader described above. The relative fluorescence units were converted to units of nuclease activity by interpolation using a standard curve generated with purified micrococcal nuclease enzyme (Worthington Biochemicals). To ensure that the readouts from the assay of the sterilized culture supernatants fell within the linear regression range of the standard curve samples, the nuclease-containing supernatants were diluted 1:10 in water and then serially diluted twofold in water up to 14 times before being mixed with the FRET substrate.

Membrane fatty acid analysis

Strains were grown in CDM to the exponential phase, and then a 15-mL sample of culture was pelleted (between 10 and 100 mg wet cell pellet) and washed twice with phosphate-buffered saline (PBS) before being stored at −80°C. The fatty acids in each pellet were saponified and methylated before being analyzed using gas chromatographic analysis of fatty acid methyl esters (GC-FAME) as a fee for service at the Center for Microbial Identification and Taxonomy (Norman, OK).

Analysis of SaeS kinase activity

In vivo separation of SaeR and SaeR ~P was performed as described previously using 12% polyacrylamide gels containing 100 µM manganese and 50 µM of the acrylamide-pendant Phos-tag ligand (24, 63). Cells were grown in CDM to exponential phase (OD₆₀₀ 0.5–1) at 37°C with agitation. Cell pellets (OD₆₀₀ of 10) were collected at 13,000 x g and stored at −80°C prior to analysis. Whole-cell extracts were obtained by resuspending cell pellets in 300 µL cell extract buffer (20 mM Tris-HCl [pH 7.0], 1X Protease Inhibitor Cocktail Set I (Sigma-Aldrich)) and transferred to sterile screw cap tubes containing approximately 100 µL of 0.1 mm silica beads. The cells were homogenized at room temperature using a Precellys 24 bead beater (Bertin Technologies) for three cycles of 6,500 rpm for 30 seconds each, followed by 3-minute pauses on ice. The tubes were then centrifuged at 8,500 x g for 15 seconds to settle the beads, and the supernatant was transferred into new microcentrifuge tubes. Whole-cell extracts were normalized by protein concentration (A280) to 100 µg, mixed with 5X SDS loading dye, and electrophoresed on Phos-tag gels with standard running buffer (0.1% [wt/vol] SDS, 25  mM Tris-HCl, 192  mM glycine) at 4°C under constant voltage (150 V) for 2 hours. The gels were washed for 15 minutes with transfer buffer (25 mM Tris [pH 8.3], 192 mM glycine, and 20% methanol) with 1 mM EDTA, followed by a second wash without EDTA to remove manganese ions. Proteins were then transferred to 0.45-µM PVDF membranes (Cytiva). Membranes were incubated in blocking buffer (5% [wt/vol] skim milk in tris-buffered saline with Tween-20 (TBST) (20 mM Tris-HCl, and 150 mM NaCl with 0.1% [wt/vol] Tween 20 pH 7.6) for 1 hour. Membranes were then subjected to three brief washes in TBST and incubated with polyclonal rabbit antibodies to SaeR (1.5:1,000) for 1 hour. Membranes were then washed three times with and incubated with StarBright Blue 700 goat anti-rabbit IgG (1:3,500; Bio-Rad) for 1 hour. Membranes were subjected to three brief washes in TBST, and signals were visualized using an Amersham ImageQuant800. The densities of the SaeR ~P relative to the total SaeR signal were determined by quantification with Multi Gauge software (FujiFilm). The data are representative of three independent experiments, and a representative image is shown.

Analysis of SaeR and SrtA

The whole-cell extracts and protein concentrations described above were used to visualize total SaeR levels. Briefly, 100 µg of whole-cell extracts mixed with 5X SDS loading dye was subjected to 12% SDS-PAGE, and proteins were transferred to 0.45-µM PVDF membranes (Cytiva). After transfer, membranes were incubated in blocking buffer for 1 hour. Membranes were then washed three times with TBST and incubated with either polyclonal rabbit antibodies to SaeR (1.5:1,000) or SrtA (1:1,000) for 1 hour. Membranes were then washed three times and incubated with StarBright Blue 700 goat anti-rabbit IgG (1:3,500; Bio-Rad) for 1 hour. Membranes were subjected to three brief washes in TBST, and signals were visualized using an Amersham ImageQuant800. The densities of the SaeR and SrtA signal were determined by quantification with Multi Gauge software (FujiFilm). The data are representative of three different independent experiments, and a representative image is shown.

RNA extraction

RNA extraction was performed as previously described (58). Briefly, cells were grown as described above to the exponential phase (OD₆₀₀ of 0.5), and then a 4-mL sample was quenched with an equal volume of a 1:1 (vol/vol) mixture of ethanol–acetone prechilled to −20°C. The quenched samples were immediately frozen on dry ice and then transferred for long-term storage to a −80°C freezer. Once thawed, cells were pelleted, washed twice with TE buffer (10 mM Tris-Cl [pH 8], 1 mM EDTA), resuspended in TRIzol, and then mechanically disrupted using a Precellys 24 homogenizer (Bertin Technologies) with three 30-second pulses at 6,800 rpm. Samples were incubated for 1 minute on wet ice between pulses. Nucleic acids from each sample were extracted using a Direct-Zol kit (Zymo Research Corporation).

Quantitative reverse transcription-polymerase chain reaction (qRT-PCR)

Reverse transcription followed by quantitative PCR was performed essentially as previously described (58). The High-Capacity cDNA Reverse Transcription kit (Applied Biosystems) and random primers were used to reverse transcribe 250 ng of total RNA from each sample, as per the manufacturer’s instructions. The abundances of specific transcripts were determined using a Applied Biosystems QuantStudio Real-Time PCR system and Universal SYBR Green Supermix (Applied Biosystems). Reaction mixtures were incubated at 98°C for 2 minutes and then cycled between 98°C and 60°C. Threshold fluorescence measurements were converted to transcript abundance using standard curves generated from serial dilutions of genomic DNA spanning at least seven orders of magnitude. Target transcript abundance was normalized to the level of the gyrB transcript as the internal control since the abundance of this transcript remains constant (<twofold change) across all strains and conditions analyzed.

SUPPLEMENTAL MATERIAL

The following material is available online at https://doi.org/10.1128/jb.00191-24.

Supplemental figures and tables. jb.00191-24-s0001.pdf.

Fig. S1 to S4; Tables S1 to S3.

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