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Journal of Antimicrobial Chemotherapy logoLink to Journal of Antimicrobial Chemotherapy
. 2025 Sep 23;80(11):2978–2988. doi: 10.1093/jac/dkaf325

The toxin–antitoxin system SavRS contributes to vancomycin resistance in vancomycin-intermediate Staphylococcus aureus by mediating cell wall thickening

Weifeng Xu 1,2, Ping Yan 3,4, Yujie Li 5,6,7,✉, Baolin Sun 8,9,✉
PMCID: PMC12596048  PMID: 40985144

Abstract

Background

The emergence of vancomycin-intermediate Staphylococcus aureus (VISA) has significantly challenged the treatment of S. aureus infection. Toxin–antitoxin (TA) systems have been reported to mediate bacterial stress adaptation and virulence, but their role in vancomycin resistance remains elusive. This study investigated the vancomycin resistance mechanism regulated by the TA system SavRS in VISA.

Methods

savRS mutants in Mu50 and XN108 were generated via homologous recombination. To investigate the regulatory mechanism of vancomycin resistance mediated by savRS in VISA, phenotypic analyses including MICs, growth kinetics and cell wall thickness measurements were performed. Expression of cell wall synthesis-related genes was analysed using quantitative RT–PCR (RT–qPCR) and promoter-lacZ reporter assay. Electrophoretic mobility shift assay (EMSA) was performed to assess the binding of SavRS to the promoters of the cell wall synthesis-related genes. Pull-down assay identified an upstream regulatory element of savRS associated with vancomycin resistance. Quantitative assessment of bacterial burden in murine organ systems following vancomycin administration revealed the critical regulatory role of savRS in mediating vancomycin resistance in vivo.

Results

Compared with the WT, the savRS mutant exhibited enhanced vancomycin sensitivity, accelerated growth and reduced cell wall thickness. Correspondingly, RT–qPCR revealed marked down-regulation of the cell wall synthesis-related genes (glyS, dltA, scdA, pbp2, ddl). EMSA and promoter-lacZ reporter assay confirmed direct binding of SavRS to a conserved promoter motif, MGHYYTCCTCA. Pull-down assay identified UspA as an upstream regulator of SavRS, demonstrating that UspA directly controls savRS transcription and modulates VISA resistance. Mouse infection experiments showed that savRS promotes VISA to vancomycin resistance in vivo.

Conclusions

SavRS critically regulates vancomycin resistance in VISA.

Introduction

Staphylococcus aureus is a major human pathogen characterized by high prevalence, adaptability and virulence.1–3 The emergence of MRSA has established vancomycin as the primary therapeutic option for severe infections.4,5 This glycopeptide antibiotic targets peptidoglycan biosynthesis in Gram-positive bacteria by binding to D-alanyl-D-alanine termini of cell wall precursors.6,7 Clinically, S. aureus is classified into three phenotypes based on MICs: vancomycin-resistant S. aureus (VRSA; MICs of ≥16 mg/L), vancomycin-intermediate S. aureus (VISA; MICs of 4–8 mg/L) and vancomycin-susceptible S. aureus (VSSA; MICs of ≤2 mg/L).8 VRSA resistance originates from horizontal acquisition of the vanA transposon, which encodes enzymes that replace D-alanyl-D-alanine with low-affinity D-alanyl-D-lactate in peptidoglycan precursors.9,10 In contrast, VISA lacks vanA and develops resistance through peptidoglycan accumulation, which increases D-alanyl-D-alanine availability and competitively inhibits vancomycin binding.9,11 This mechanism correlates with characteristic cell wall thickening and reduced autolytic activity.12–14 Notably, VISA exhibits higher clinical prevalence than VRSA.15–17 Given the critical role of antibiotic efficacy in public health, further investigation into VISA resistance mechanisms remains imperative.

Toxin–antitoxin (TA) systems comprise two components: the toxins capable of inhibiting bacterial growth or bactericidal activity; and the antitoxins that neutralize the effects of the toxins in the TA systems.18 Most toxins are proteinaceous; antitoxins typically exist as RNA molecules or proteins.19,20 Although their roles were initially identified as maintaining plasmid stability via a post-segregational killing mechanism involving antitoxin degradation and subsequent toxin-mediated elimination of plasmid-free cells, recent research has revealed broader physiological functions, including bacterial growth regulation, persister cell formation, programmed cell death simulation, stress response adaptation and phage resistance.21–25 SavRS, a type II TA system originally identified in S. aureus NCTC 8325, has been demonstrated to regulate virulence gene transcription (hla and efb).26 Clinical isolates of antibiotic-resistant bacteria frequently harbour TA systems, with current understanding linking their antibiotic resistance association to growth inhibition properties and plasmid stabilization of resistance genes.27–29 Given the strong correlation between TA systems and antimicrobial resistance, we propose that novel regulatory mechanisms remain to be elucidated.

This investigation employed a previously identified TA system SavRS to examine its regulatory role in vancomycin resistance among VISA strains. Our findings demonstrate that the type II TA system SavRS enhances vancomycin resistance in VISA both in vitro and in vivo through mechanisms extending beyond conventional plasmid maintenance functions. This study elucidates the regulatory role of TA systems in S. aureus vancomycin resistance.

Materials and methods

Ethics

Mice care and experimentation in this study were conducted in strict compliance with the guidelines adopted by the Ministry of Health of the People’s Republic of China in June 2004. The study protocol received formal approval from the Institutional Animal Care and Use Committee at the University of Science and Technology of China (USTCACUC24080124026).

Strains and growth conditions

The bacterial strains utilized in this study are catalogued in Table S1 (available as Supplementary data at JAC Online). Plasmids are listed in Table S2. S. aureus isolates were maintained in tryptic soy broth (TSB; Difco) or on tryptic soy agar (TSA; Difco), with media supplementation of 15 mg/L chloramphenicol for strains harbouring chloramphenicol-resistant plasmids. Escherichia coli cultures were grown in lysogeny broth (LB; Oxoid) or on lysogeny broth agar (LA; Oxoid), supplemented with 50 mg/L kanamycin or 15 mg/L ampicillin for plasmid-bearing strains. All bacterial cultures were incubated at 37°C with shaking (220 rpm) unless otherwise specified.

Construction of gene knockout mutants and complementation strains

The shuttle plasmid pBTs was employed for targeted mutagenesis of savRS and uspA using homologous recombination. Briefly, 600–800 bp flanking regions of target genes were PCR-amplified using primers encoding 25 bp homology arms. These fragments were fused through overlap extension PCR and subsequently cloned into pBTs via enzymatic recombination. Recombinant plasmids were initially transformed into E. coli TOP10 for propagation, then electroporated into S. aureus RN4220 for methylation modification prior to final transfer into Mu50 and XN108 clinical strains. Mutant strains were generated through established scarless gene replacement protocols.30 For complementation assay, target genes with native promoters were cloned into pLI50 and reintroduced into respective mutant backgrounds.

Antibiotic sensitivity assay

MICs were determined by broth microdilution per CLSI guidelines using Mueller–Hinton II (MH II; Difco) medium with appropriate adjustment.31 Bacterial suspensions (1 × 10⁶ cfu/mL) were diluted 10-fold to the final concentration of 1 × 10³ cfu/mL and spot-plated on vancomycin-gradient TSA.30 The population characteristics were analysed via dilutions (10 μL aliquots) on vancomycin-concentration TSA and cfu were quantified after 48 h incubation at 37°C.32

Growth curves

Overnight cultures of S. aureus were subcultured into fresh TSB or vancomycin-supplemented MH II broth to achieve standardized inoculum density. Continuous growth monitoring was performed using a microplate reader (ELx800; Bio-Tek) with hourly OD600 measurements under standardized shaking conditions (37°C, 220 rpm) until cultures reached late-exponential phase.

RNA extraction, reverse transcription, quantitative RT–PCR (RT–qPCR)

Total RNA extraction and cDNA generation were performed as previously described.33 Briefly, mid-exponential phase S. aureus cells were harvested by centrifugation. Cell pellets were resuspended in RNAiso Plus (Takara Bio) and transferred to lysing matrix tubes containing zirconia/silica beads. Mechanical lysis was performed using a FastPrep-24 homogenizer (MP Biomedicals). Following chloroform extraction, samples were centrifuged. The aqueous phase was combined with isopropanol, incubated and centrifuged. RNA pellets were washed twice with 75% ethanol, air-dried and resuspended in DEPC-treated water. cDNA synthesis was performed with PrimeScript™ RT reagent Kit with gDNA Eraser (Takara Bio). Quantitative PCR analysis was conducted using 2× RealStar Green Fast Mixture (GenStar) on a QuantStudio 6 Pro system (Applied Biosystems), with the primer sequences detailed in Table S3.

Transmission electron microscopy (TEM) observation of the cell wall

Mid-logarithmic phase S. aureus cultures were processed using established protocols and analysed by the Core Facility Center for Life Science (USTC, China).34 TEM imaging was performed at 120 kV acceleration voltage. Cell wall thickness was quantified using ImageJ, with mean thickness values derived from four equidistant points per cellular cross-section.

Protein expression and purification

E. coli BL21 (DE3) transformants harbouring expression plasmids were grown at 37°C to an OD600 of 0.5, induced with 0.5 mM IPTG, and incubated overnight at 16°C with agitation. Protein purification was performed as previously described.30 Cells were harvested by centrifugation and lysed in lysis buffer containing lysozyme via ice incubation followed by ultrasonication. Clarified lysates obtained by centrifugation were subjected to Ni-NTA affinity chromatography (Beyotime, Shanghai). After three washes with native wash buffer, bound proteins were eluted with imidazole-containing buffer. Purified proteins underwent quantification with the Bradford Protein Assay Kit (Beyotime, Shanghai).

Electrophoretic mobility shift assay (EMSA)

The EMSA was adapted from previously described methods.35 Biotinylated DNA probes encompassing target promoter regions were generated via PCR using 5′-biotinylated primers. The DNA probes were heated to 95°C and annealed to room temperature to minimize secondary structure formation. Protein–DNA binding reactions (20 min, 25°C) contained serially diluted recombinant protein in 1× binding buffer. Reaction mixtures were resolved on 6% native polyacrylamide gels (0.5× TBE, 80 V, 50 min) and electrotransferred to nylon membranes (0.5× TBE, 200 mA, 150 min). Chemiluminescent detection employed streptavidin–HRP conjugates with ImageQuant LAS 4000 imaging. Non-specific competitor DNA served as negative controls.

Pull-down assay

The DNA pull-down assay was performed with modifications as previously described.36  S. aureus cultures across early-exponential, mid-logarithmic and stationary growth phases were harvested and lysed in lysis buffer containing 1 mg/mL lysostaphin and 60 μg of DNase I. Following enzymatic lysis, cells were sonicated on ice and clarified by centrifugation. Lysates were concentrated using 10 kDa centrifugal filters.

Biotinylated promoter probes were prepared via PCR with 5′-biotinylated primers. Streptavidin magnetic beads (Promega) were preconditioned with 0.5× saline sodium citrate (SSC) buffer and equilibrated in 2× binding/washing (B/W) buffer. Probe-bound beads were incubated with biotinylated DNA, followed by pre-clearing with salmon sperm DNA-treated lysates. DNA–protein complexes were captured, washed sequentially with lysis buffer containing/omitting competitor DNA, and processed for LC-MS/MS analysis (USTC Proteomics Core).37

β-Galactosidase activity assay

The β-galactosidase assay using o-nitrophenyl-β-D-galactopyranoside (ONPG) as substrate, including all buffers, was performed as previously described.38 Mid-log phase S. aureus cells were harvested by centrifugation. OD600 and volume (V, mL) were recorded. The pellets were resuspended in 100 µL of ABT LSA and incubated at 37°C until clear, then 100 µL of ABT buffer and 100 µL of ONPG were added, followed by incubation at 37°C until a yellow colour developed. Reaction times (T, min) were recorded. Reactions were terminated with Na2CO3. After centrifugation, supernatant OD420 values were measured. Miller units were calculated as: 1000 × OD420/(T × V × OD600).

RNA-Seq analysis

MU50 and ΔsavRS strains were harvested by centrifugation at mid-log phase (three biological replicates), flash-frozen in liquid nitrogen and stored at −80°C. RNA extraction, quality assessment and library construction were performed by Shanghai Personal Biotechnology Co. Ltd. Sequencing was subsequently conducted on their Illumina NovaSeq 6000 platform. The raw data of RNA-Seq have been uploaded to the NCBI database (PRJNA1298783).

Quantitative analysis of bacterial burden in murine visceral tissues

Eight-week-old BALB/c mice (GemPharmatech, Nanjing, China) were randomized into four experimental groups. Two groups received intraperitoneal challenge with 1 × 108 cfu of either WT or ΔsavRS, followed by vancomycin therapy (32 mg/kg) initiated 24 h post-infection. PBS-injected groups served as vehicle controls. Following 5 day antimicrobial treatment, euthanized mice underwent aseptic homogenization of heart, liver, spleen and kidney for quantitative bacteriological analysis.

Statistical analysis

All experiments were performed with more than three biological replicates. The Shapiro–Wilk test was used for testing the normality. Statistical significance was determined using unpaired or paired two-tailed Student’s t-tests, as appropriate for the experimental design.

Results

Deletion of savRS decreases the vancomycin resistance in VISA

To explore the molecular mechanism of the vancomycin resistance in VISA, we generated savRS mutants in VISA strains Mu50 (ΔsavRS), along with plasmid-complemented strains CsavRS. The plate susceptibility assay demonstrated complete growth inhibition of ΔsavRS on solid medium containing 1 mg/L vancomycin, contrasting with WT and CsavRS (Figure 1a). CLSI-guided antibiotic susceptibility testing demonstrated significantly reduced vancomycin MIC (2 mg/L) in ΔsavRS, while complemented strains maintained WT MIC, confirming the savRS-dependent resistance phenotype (Figure 1b). Population analysis profiling revealed significantly diminished vancomycin resistance in ΔsavRS compared with WT and CsavRS (Figure 1c). The liquid culture growth assay corroborated these findings, with ΔsavRS showing growth arrest at 2 mg/L vancomycin; WT and CsavRS strains maintained proliferation at these concentrations (Figure 1e–g). Notably, the accelerated growth rates observed in ΔsavRS via growth curve analysis align with its decreased vancomycin resistance, consistent with established correlations between slowed bacterial growth and vancomycin resistance acquisition in S. aureus.39

Figure 1.

Figure 1.

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Deletion of savRS decreases vancomycin resistance in S. aureus Mu50. (a) The model of savRS in S. aureus. (b) The growth of the strains was assessed by plate susceptibility assay on vancomycin-containing plates with gradient concentrations. (c) The vancomycin MICs were determined by the broth microdilution method according to CLSI. (d) The vancomycin resistance levels were evaluated by population analysis profiles and the colonies were counted after incubation at 37℃ for 48 h. Growth curves of WT (e), ΔsavRS (f) and CsavRS (g) in MH II broth with different concentrations of vancomycin. (h) RT–qPCR analysis of savRS expression under vancomycin pressure (n = 5). For (d), (e), (f), (g) and (h), data are expressed as the mean ± SD; statistical significance was calculated with a two-tailed unpaired Student’s t-test.

In addition, in another VISA strain XN108, ΔsavRS failed to grow on medium containing 2 mg/L vancomycin (Figure S1a). Antibiotic susceptibility testing showed that the vancomycin MIC for XN108 ΔsavRS decreased from 8 to 4 mg/L (Figure S1b). Population analysis revealed a significantly reduced survival rate of XN108 ΔsavRS on vancomycin plates (Figure S1c). In liquid medium with >2 mg/L vancomycin, XN108 ΔsavRS exhibited slower growth than WT and CsavRS (Figure S1d–f). These consistent phenotypic changes confirm the critical role of savRS in maintaining VISA vancomycin resistance.

To investigate savRS transcriptional regulation under vancomycin stress, VISA strains were exposed to vancomycin at 10× MIC during culture, followed by RT–qPCR analysis. The results demonstrated significant up-regulation of savRS transcription following high-concentration vancomycin challenge in Mu50 (Figure 1h). The same results as above also appeared in XN108 (Figure S1g), indicating that this TA system actively responds to vancomycin-induced stress.

SavRS regulates cell wall thickness

To investigate whether SavRS modulates vancomycin resistance through cell wall thickness regulation, we tested the cell wall thickness of the target strains via TEM. Quantitative measurements revealed significantly reduced cell wall thickness in ΔsavRS (23.54 ± 2.96 nm) compared with the WT (52.04 ± 7.24 nm) and complemented strain (55.58 ± 11.25 nm) (Figure 2a and b), establishing a direct correlation between SavRS function and cell wall morphology. Subsequent RT–qPCR analysis demonstrated the down-regulation of key cell wall biosynthesis genes (glyS, dltA, scdA, pbp2 and ddl) in ΔsavRS (Figure 2c), suggesting impaired peptidoglycan synthesis in ΔsavRS. We also found a significantly reduced cell wall thickness in XN108 ΔsavRS (33.39 ± 4.18 nm) compared with both XN108 WT (61.68 ± 7.46 nm) and XN108 CsavRS (61.58 ± 8.54 nm) strains (Figure S2a and b). Consistent with this morphological alteration, RT–qPCR analysis demonstrated significant down-regulation of key cell wall biosynthesis genes (glyS, dltA and pbp2) in the XN108 ΔsavRS (Figure S2c).

Figure 2.

Figure 2.

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savRS deletion disrupts cell wall biosynthesis in S. aureus Mu50. (a) Transmission electron micrographs of the cell walls of WT, ΔsavRS and CsavRS. Bar = 200 nm. (b) Cell wall thickness quantification using ImageJ (n = 10). (c) RT–qPCR analysis of cell wall biosynthesis gene expression (n = 3). Data are expressed as the mean ± SD; statistical significance was calculated with a two-tailed unpaired Student’s t-test.

Furthermore, the lipoglycopeptide antibiotic dalbavancin exhibits reduced susceptibility due to S. aureus cell wall thickening.40 MIC testing showed that ΔsavRS decreased dalbavancin MICs from 0.5 to 0.25 mg/L in Mu50 and from 1 to 0.25 mg/L in XN108 (Figure S3). This further supports that savRS regulates VISA strain resistance through cell wall thickening.

SavRS binds specific motifs in promoters of cell wall synthesis genes and promotes expression

To elucidate the molecular mechanism underlying SavRS-mediated regulation of the cell wall synthesis-related genes, competitive EMSA was performed using purified SavRS protein and biotin-labelled glyS, dltA and pbp2 promoter regions. The results demonstrated concentration-dependent DNA–protein complex formation, which was abolished by excess unlabelled competitor DNA but unaffected by random sequence controls (Figure 3a–c), indicating direct transcriptional regulation by SavRS. We also identified a conserved MGHYYTCCTCA motif in the promoters of affected genes through MEME suite analysis. Substitution of this motif with a scrambled sequence abolished the electrophoretic mobility shift between the dltA promoter probe and SavRS protein (Figure 3d), confirming the motif-specific interaction. β-Galactosidase activities from lacZ fusion reporter plasmids pOSpbp2, pOSdltA and pOSglyS were significantly higher in Mu50 than in ΔsavRS, supporting the EMSA results (Figure 3e–g). These findings collectively demonstrate that SavRS maintains vancomycin resistance through direct transcriptional activation of the cell wall biosynthesis genes via recognition of a conserved promoter motif.

Figure 3.

Figure 3.

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SavRS directly promotes the expression of pbp2, glyS and dltA. The purified SavRS protein was incubated with the pbp2 (a), dltA (b) and glyS (c) promoter regions that were amplified by PCR. (d) The MGHYYTCCTCA motif within the dltA promoter region was substituted with a synthetic randomized sequence of equivalent length for EMSA. Unlabelled probes served as specific competitors; non-specific DNA acted as a non-specific competitor. β-Galactosidase activity regulated by pbp2 (e), dltA (f) and glyS (g) promoters in WT and ΔsavRS (n = 3). Data are expressed as the mean ± SD; statistical significance was calculated with a two-tailed unpaired Student’s t-test.

savRS responds to vancomycin stress through the universal stress protein UspA

Given that SavRS functions as an intracellular protein incapable of direct interaction with vancomycin, we investigated its stress response mechanism by identifying upstream regulatory factors. The pull-down assay detected interaction between the savRS promoter and UspA (Figure S4; Tables S4 and S5), a conserved bacterial stress–response factor that is up-regulated during growth arrest and under environmental challenges.41 MIC analysis demonstrated that ΔuspA reduced vancomycin MIC to 4 mg/L, while the double mutant ΔsavRSΔuspA exhibited a further reduction to 1 mg/L (Figure 4a). Population analysis profiling revealed additive effects, with Mu50 ΔsavRSΔuspA showing greater vancomycin susceptibility than single mutants, both significantly exceeding WT sensitivity (Figure 4b). Liquid culture growth assay confirmed that Mu50 ΔuspA displayed growth arrest at 3 mg/L vancomycin, while Mu50 ΔsavRSΔuspA failed to proliferate even at 1 mg/L (Figure 4c).

Figure 4.

Figure 4.

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UspA regulates savRS transcription and influences vancomycin resistance in S. aureus Mu50. (a) The vancomycin MICs were determined by the broth microdilution method according to CLSI. (b) The vancomycin resistance levels were evaluated by population analysis profiles and the colonies were counted after incubation at 37℃ for 48 h. Growth curves of ΔuspA (c) and ΔuspAΔsavRS (d) in MH II broth with different concentrations of vancomycin. (e) RT–qPCR analysis of savRS expression (n = 3). (f) EMSA of UspA gradient concentration with the biotin-labelled promoters of savRS, unlabelled promoters of savRS and non-specific DNA competitive-binding UspA. For (b), (c), (d) and (e), data are expressed as the mean ± SD; statistical significance was calculated with a two-tailed unpaired Student’s t-test.

To investigate the precise regulatory mechanism by which UspA modulates savRS, we conducted RT–qPCR analysis, revealing significantly reduced savRS transcription in ΔuspA (Figure 4e). EMSA with recombinant UspA and biotin-labelled putative savRS promoter demonstrated concentration-dependent binding that was abolished by excess unlabelled competitor (100×), but unaffected by random sequence controls (Figure 4f). Combined with prior pull-down data, these results establish UspA as a direct transcriptional activator of savRS through promoter binding. These results consistently indicate that UspA can act as a savRS upstream element involved in the regulation of vancomycin resistance in VISA strains.

savRS enhances the vancomycin resistance of VISA strains in hosts

To validate the in vivo relevance of SavRS-mediated vancomycin resistance, we conducted murine infection studies using BALB/c mice challenged via intraperitoneal injection with either Mu50 or ΔsavRS, followed by a 5 day course of vancomycin treatment. Standard plate counts at 7 days post-infection revealed significantly reduced bacterial burdens in hearts, livers, spleens and kidneys of the mice infected with ΔsavRS and treated with vancomycin, compared with PBS-treated controls (Figure 5a–d). In contrast, vancomycin treatment did not significantly reduce organ colonization in mice infected with WT. These findings demonstrate that the SavRS-dependent vancomycin resistance observed in vitro critically determines the therapeutic outcomes in vivo, with ΔsavRS strain infections showing enhanced susceptibility to vancomycin.

Figure 5.

Figure 5.

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Disruption of savRS significantly reduces the survival rate of S. aureus in mouse organs. Bacterial loads in kidneys (a), spleens (b), livers (c) and hearts (d) of mice infected with Mu50 or ΔsavRS strains after vancomycin treatment (32 mg/kg) (n = 6). The black lines connecting each other indicate that they were infected by the same strain. Statistical significance was determined using a two-tailed paired Student’s t-test.

Discussion

VISA remains a critical therapeutic challenge due to its global prevalence and recalcitrant infections. Although TA systems are widely implicated in clinical antibiotic resistance through mechanisms including growth inhibition, plasmid stabilization and toxin-mediated persistence via stress adaptation, these conventional mechanisms inherently compromise bacterial fitness. However, based on the clinical characteristics of VISA strains, we believe that there may be a resistance pathway that does not completely sacrifice bacterial growth. To investigate this, we targeted the type II TA system savRS in VISA for genetic disruption. The savRS deletion significantly diminished vancomycin resistance in VISA strains. Specifically, the vancomycin MIC for Mu50 ΔsavRS decreased from 8 to 2 mg/L, while XN108 ΔsavRS exhibited an MIC reduction from 8 to 4 mg/L. Although the magnitude of resistance attenuation exhibited strain-dependent variation, this phenotypic consistency demonstrates the conserved regulatory role of savRS in VISA resistance development. Mechanistic investigations further demonstrated that SavRS directly binds to the conserved binding motif MGHYYTCCTCA located within promoters of cell wall biosynthesis genes, thereby enhancing their transcriptional activation and subsequently increasing peptidoglycan layer thickness. This observation aligns with the established VISA resistance paradigm, in which thickened cell walls prevent vancomycin from accessing its target. Furthermore, EMSA and the pull-down assay revealed that savRS expression is regulated by UspA through direct interaction with the savRS promoter region. Consistent with this interaction, uspA deletion increased vancomycin susceptibility in the Mu50. These findings position UspA as the master upstream regulator of the SavRS-mediated resistance pathway (Figure 6). Notably, the observed synergistic sensitivity in the double mutant suggested a degree of functional independence between these two regulatory systems. Previous reports indicate that UspA is present both on the surface and in the cytoplasm of S. aureus,42 and that cytoplasmic UspA is activated by the stringent response signalling nucleotide (p)ppGpp.41,43 The (p)ppGpp synthases RelP and RelQ are inducible by vancomycin and are part of the VraRS cell wall stress regulon in S. aureus.44,45 Therefore, we hypothesized that vancomycin induction might internalize surface UspA into the cytoplasm and promote savRS expression, or that vancomycin-induced cell wall stress might promote (p)ppGpp accumulation, triggering uspA transcription and thereby promoting savRS expression. Our findings reveal a distinct paradigm wherein the SavRS protein complex functions synergistically to directly target cell wall biosynthesis genes, rather than relying on conventional growth-arrest strategies for antibiotic evasion. This work reveals a non-canonical TA mechanism where coordinated regulation enables simultaneous antibiotic resistance and physiological fitness, posing enhanced clinical threats.

Figure 6.

Figure 6.

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The TA system SavRS regulates cell wall metabolism in VISA, directly mediating antibiotic resistance. UspA induces savRS transcription in response to vancomycin challenge. The SavRS subsequently induces expression of cell wall synthesis-related genes (glyS, pbp2 and dltA), thereby promoting peptidoglycan thickening and conferring vancomycin tolerance.

In addition to the aforementioned cell wall-related gene expression regulated by savRS, RNA-Seq revealed widespread down-regulation of additional cell wall-associated resistance genes in ΔsavRS (Figure S5a), correlating with increased susceptibility to cell wall-targeting antibiotics vancomycin and dalbavancin. Furthermore, the significant down-regulation of the membrane-related mprF gene in ΔsavRS coincided with enhanced daptomycin sensitivity (Figures S5a and S6).46 Transcriptional dysregulation of multiple resistance genes indicates the broad regulatory role of savRS in S. aureus resistance (Figure S5b). Concurrently, numerous virulence genes showed altered expression, with significant enrichment in S. aureus infection pathways (Figure S5c and d). In murine infection models, ΔsavRS exhibited heightened vancomycin susceptibility versus WT. Notably, even without antibiotics, differential bacterial loads were observed. Although RNA-Seq revealed both up- and down-regulated virulence genes, bacterial load analysis indicated that the savRS deletion increased the bacterial burden in the hearts and kidneys of mice; the enhanced virulence resulting from ΔsavRS is consistent with our previous report of enhanced RBC lysis and exacerbated abscesses in a mouse skin infection model.26 These data indicate that savRS enhances vancomycin resistance while simultaneously attenuating virulence during infection, resulting in a resistance–virulence ‘seesaw effect’. These findings collectively suggest that savRS plays a multifunctional role in host–pathogen interactions. Future studies will elucidate the specific molecular pathways by which savRS regulates the infection process, which may expand our understanding of the function of the TA systems in bacterial physiology and virulence regulation and provide new targets for the clinical treatment of VISA infections.

Supplementary Material

dkaf325_Supplementary_Data
dkaf325_supplementary_data.doc (8.2MB, doc)

Contributor Information

Weifeng Xu, State Key Laboratory of Immune Response and Immunotherapy, School of Basic Medical Sciences, Division of Life Sciences and Medicine, University of Science and Technology of China, Hefei 230027, China; Department of Oncology, the First Affiliated Hospital of USTC, Division of Life Sciences and Medicine, University of Science and Technology of China, Hefei, Anhui 230027, China.

Ping Yan, State Key Laboratory of Immune Response and Immunotherapy, School of Basic Medical Sciences, Division of Life Sciences and Medicine, University of Science and Technology of China, Hefei 230027, China; Department of Oncology, the First Affiliated Hospital of USTC, Division of Life Sciences and Medicine, University of Science and Technology of China, Hefei, Anhui 230027, China.

Yujie Li, State Key Laboratory of Immune Response and Immunotherapy, School of Basic Medical Sciences, Division of Life Sciences and Medicine, University of Science and Technology of China, Hefei 230027, China; Department of Oncology, the First Affiliated Hospital of USTC, Division of Life Sciences and Medicine, University of Science and Technology of China, Hefei, Anhui 230027, China; Anhui Province Key Laboratory of Embryo Development and Reproduction Regulation, Anhui Province Key Laboratory of Environmental Hormone and Reproduction, School of Biological and Food Engineering, Fuyang Normal University, Fuyang 236037, China.

Baolin Sun, State Key Laboratory of Immune Response and Immunotherapy, School of Basic Medical Sciences, Division of Life Sciences and Medicine, University of Science and Technology of China, Hefei 230027, China; Department of Oncology, the First Affiliated Hospital of USTC, Division of Life Sciences and Medicine, University of Science and Technology of China, Hefei, Anhui 230027, China.

Funding

This work was supported by the National Natural Science Foundation of China (32370184), the Natural Science Foundation of Anhui Province, China (2408085MC061), the Fundamental Research Funds for the Central Universities (WK9100000080) and the USTC Research Funds of the Double First-Class Initiative (YD9100002063).

Transparency declarations

None to declare.

Author contributions

Weifeng Xu: Conceptualization (Lead), Data curation (Lead), Formal analysis (Lead), Investigation (Lead), Methodology (Lead), Resources (Lead), Software (Lead), Validation (Lead), Visualization (Lead), Writing—original draft (Lead); Ping Yan: Writing—original draft (Supporting); Yujie Li: Funding acquisition (Equal), Project administration (Equal), Supervision (Equal), Writing—review & editing (Equal); Baolin Sun: Funding acquisition (Equal), Project administration (Equal), Supervision (Equal), Writing—review & editing (Equal).

Supplementary data

Figures S1 to S5 and Tables S1 to S3 are available as Supplementary data at JAC Online.

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