The bacterial strains and plasmids used in this study are described in Supplementary Table 1. Escherichia coli were grown in Luria broth (LB) medium (Oxoid). S. aureus and its derivative strains were grown in tryptic soy broth (TSB) medium (BD) at 37°C with shaking at 220 rpm. When needed, appropriate antibiotics were used for plasmid selection and maintenance at the following concentrations: for E. coli, ampicillin at 150 μg/mL and kanamycin at 50 μg/mL; for S. aureus, chloramphenicol at 15 μg/mL. Constructed plasmids were purified from E. coli Trans1-T1 and transformed into S. aureus RN4220 as the initial recipient and then S. aureus strain ST59 by electroporation. The media were solidified with 1.5% (w/v) agar when required.

S. aureus genomic DNA was prepared by a standard protocol for Gram-positive bacteria. Plasmid DNA was extracted with a plasmid purification kit (Sangon Biotech) according to the manufacturer’s instructions. PrimeSTAR HS DNA polymerase (TaKaRa) and Phanta Max Super-Fidelity DNA polymerase (Vazyme) were used for PCR amplification, respectively. All plasmids transformed into the target S. aureus strain were first introduced into S. aureus strain RN4220 by electroporation at 2.5 kV for modification. The primers used in this study are listed in Supplementary Table 2.

To construct the lcpB mutants, the upstream and downstream fragments of lcpB were amplified from S. aureus strain ST59 genomic DNA using the lcpB-up-F/lcpB-up-R, lcpB-down-F/lcpB-down-R sets of primers (Supplementary Table 2). The upstream and downstream regions of each gene were ligated to form an up-down fragment. The resultant fragment was digested with restriction enzymes and then cloned into pBTs. The resulting plasmids, pBTs-lcpB, were first electroporated into S. aureus strain RN4220 for modification and subsequently transformed into S. aureus strain ST59. The allelic replacement mutants were selected using a previously described method and were further confirmed by PCR and sequencing. N315 and Newman knockout strains were constructed in the same way.

For complementation, the plasmid pLI50 was used to construct pLI50-lcpB complemented plasmid. First, amplified the fragment carrying the ORF and its native promoter from ST59 genomic DNA. and digested with restriction enzymes and cloned into the shuttle plasmid pLI50 to derive the plasmid pLI50-lcpB. The recombinant plasmid was transformed into S. aureus RN4220 by electroporation and then into the ST59 lcpB mutant to derive the complemented strain. The wild-type, lcpB mutant strains were transformed with the plasmid pLI50 as the control strains, resulting in the ST59-pLI50, and ΔlcpB-pLI50 strains respectively. The complementation strains were selected using the same method described above and were further confirmed by PCR and sequencing.

Triton X-100-stimulated autolysis was measured as described previously (Liu et al., 2016). Briefly, overnight-grown bacterial cells were diluted with TSB to 0.05 at the absorbance 600 and then allowed to grow to the early exponential phase (OD₆₀₀ ≤ 0.8) at 37°C with shaking at 220 rpm. Cells were harvested, washed three times with PBS, resuspended in the original volume of Tris-HCl (0.05 M; pH 7.5) containing 0.05% (v/v) Triton X-100, incubated at 37°C with shaking. The autolysis level was checked by measuring the progressive decrease in absorbance (OD₆₀₀) each hour using a microplate reader (Elx800; Bio-Tek) and analyzed by comparing the percentage of reduction. The experiment was repeated at least three times, with similar results.

To detect morphological changes, strains were cultivated in TSB adding 15 μg/mL chloramphenicol and allowed to grow to the early exponential phase (OD₆₀₀ = 0.5) at 37°C with shaking at 220 rpm. Samples were prepared as previously described (Wang and Sun, 2021) and sent to the Core Facility Center for Life Science (USTC, China). Specimens were examined with a transmission electron microscopy which is operated at an accelerating voltage of 120 kV.

Antibiotic susceptibility testing was performed by the broth microdilution method, as described previously (Wang and Sun, 2021).

Normally, the overnight cultures of S. aureus were diluted 1:100 in TSB with 15 μg/mL chloramphenicol, grown to the early exponential (OD₆₀₀ = 0.5), and collected. When the stress response of S. aureus to antibiotics was detected, 10-fold MIC level were added at the early exponential (OD₆₀₀ = 0.5) and then cultured for 30 min. The collected cells were processed with 1 mL of RNAiso Plus (TaKaRa) in combination with 0.1-mm-diameter-silica beads in a FastPrep-24 automated system (MP biomedicals Solon, OH, United States), and then used RNase-free DNase I (TaKaRa) to remove the residual DNA. The concentration of total RNA was adjusted to 200 ng/μL. Reverse transcription was carried out with the PrimeScript 1st Strand cDNA synthesis kit (Takara) and real-time quantitative reverse transcription-PCR (RT-qPCR) was performed with SYBR Premix Ex Taq (TaKaRa) using a StepOne realtime system (Applied Biosystems). The relative quantity of cDNA measured by real-time PCR was normalized to the average abundance of wild-type strain samples using housekeeping gene hu as the reference gene (Valihrach and Demnerova, 2012).

To detect the cellular localization of LcpB, we fused LcpB with GFP at its C-terminal tail with its promoter using pALC, and FM4-64 was used to determine the location of cell membrane. For confocal microscopy, an inverted confocal laser scanning microscope (FV1000, Olympus) was used. For the observation of fluorescent signals of GFP, an argon ion laser (Ex = 488 nm, Em = 515–530 nm) was used. For the observation of FM4-64, fluorescent signals were acquired using a He-Ne laser (Ex = 559 nm, Em = 570–670 nm). Finally, all the confocal images were captured with FV10-ASW 4.2 Viewer software (Olympus).

The Gluathion S-transferases (GST)-tagged LcpB was expressed and purified using standard procedures. The fragment of the extracellular region lcpB ORF (30–408 residues) was amplified by PCR with the primer pair ΔTM-lcpB-F/ΔTM-lcpB-R from S. aureus strain ST59 genomic DNA, cloned into the expression vector pGEX-4T-2 to generate the plasmid pGEX-ΔTM-LcpB, and transformed into E. coli BL21 (DE3). The transformant was grown in LB at 37°C to an OD₆₀₀ of 0.6 and induced with 0.5 mM isopropyl-β-D-1-thiogalactopyranoside (IPTG) at 16°C for additional 12 h. The cells were harvested and lysed by sonication in a lysis buffer (50 mM Tris-HCl, pH 8.0). The bound protein was eluted with an elution buffer (10 mM reduced glutathione, 50 mM Tris-HCl, pH 8.0). The purity of the protein was analyzed using SDS-PAGE, and the protein concentration was determined using the Bradford Assay Kit (Beyotime).

The pyrophosphatase activity of ΔTM-LcpB was determined according to a previously published protocol (Siegel et al., 2019). ΔTM-LcpB, single mutated protein, and GST (4 μg) was incubated with 2 μmol FPP Farnesyl pyrophosphate (FPP) in 50 mM Tris-HCl (pH 8.0) for 1 h at 30°C. Inorganic phosphate released from these reactions was detected by a Phosphate Assay Kit (ab270004; abcam) according to the manufacturer’s instructions. Phosphate signal was measured with a microplate reader (Elx800; Bio-Tek) at the wavelength in 600 nm. Phosphate standards were used to generate a standard curve, with samples without phosphate used as control. The phosphate concentrations in the test samples were determined by linear regression analysis of the standard curve. The results were presented as an average from three independent experiments.

Overnight-grown bacterial cells were diluted with TSB to 0.05 at the absorbance 600 and then allowed to grow to the early exponential phase (OD₆₀₀ = 0.5). Harvested 200 mL cultures of the S. aureus ST59 strain and extracted WTA. The extraction and analysis method strictly followed the protocol of the work of Covas et al. (2016). The quantification of WTA was applied by the measure of the phosphate group from WTA using the Phosphate Assay Kit with a microplate reader (Elx800; Bio-Tek) at the wavelength in 600 nm. The extraction of WTA were analyzed by native PAGE and detected by Alcian blue–silver staining. The results were presented as an average from three independent experiments.

Hemolytic activity was determined by incubating samples with sheep red blood cells. Overnight cultures were collected by centrifugation, and supernatants (100 μL) were mixed with 900 μL phosphate-buffered saline (PBS) buffer containing 10% sheep red blood cells, and the mixtures were incubated at 37°C for proper time. It takes 1 h for ST59 strains, 2.5 h for N315 and Newman strains. The absorption of supernatant at 543 nm was measured after centrifugation. A mixture with 1 mL ddH₂O containing 10% sheep red blood cells was used as the positive control, and a mixture with 1 mL PBS containing 10% sheep red blood cells was used as the negative control. The percentage of hemolytic activity was calculated relative to the positive control, which was regarded as 100% hemolytic activity.

Outbred, immunocompetent female BALB/c mice between 5 and 6 weeks of age were purchased from Gem Pharmatech Technology Company and raised them to 6–8 weeks old. The hair on the back was removed by an animal shaver. Overnight-grown bacterial cells were diluted with TSB to 0.05 at the absorbance 600 and then allowed to grow to the early exponential phase (OD₆₀₀ = 0.5) at 37°C with shaking at 220 rpm. Bacteria were collected, washed twice, and diluted in sterile PBS. Viable cells were counted via colony forming units (CFU) counting on TSB agar plates in order to quantify the infectious dose. Mice were inoculated with 2.5 × 10⁸ live S. aureus cells or PBS alone in both flanks of the back by subcutaneous injection. Abscess areas, assessed as the maximal length times width of the developing ulcers, were measured daily. Seven days after infection, the mice were sacrificed. The skin lesions were excised and homogenized in PBS. The number of CFU recovered from each individual lesion was counted by serial dilution and plated onto LB agar plates. For histopathological analyses, the skin lesions were placed in 10% formalin. Paraffin embedding and hematoxylin and eosin (H&E) staining were performed by Wuhan Servicebio Technology.

The use and care of mice in the present study followed strictly the guidelines adopted by the Ministry of Health of the People’s Republic of China in June 2004. The protocol was approved by the Institutional Animal Care and Use Committee of the University of Science and Technology of China (USTCACUC182301015).

Statistical analysis was performed using Origin 2019 and GraphPad Prism 5. Data were analyzed using unpaired t-tests to compare two different conditions and analysis of variance for more conditions. All error bars show the standard errors of the means (SEM). All experiments were performed in biological triplicates.

LcpB is considered to be a transmembrane protein that plays an important role in maintaining cell morphology in S. aureus (Chan et al., 2013). Therefore, in order to get a better understanding of the effects of LcpB, we knocked out lcpB in the clinic isolates S. aureus ST59 (NCBI number: CP076823, Supplementary Table 3) and constructed the complemented strain. The function of LcpB was then assessed by examining several phenotypes related to the cell wall, including change in growth, rate of autolysis, and levels of MIC (Supplementary Table 4). Notably, we first evaluated the growth of the wild-type, lcpB mutant, and complemented strains. The findings showed that there was a decrease in the growth rate of the lcpB mutant strain (Figure 1A). Thereafter, the autolytic activity of these three strains was examined. The results revealed that there was an increase in the rate of autolysis rate in the lcpB mutant, compared to the wild-type. The phenotype of the complemented strain was, however, same as that of the wild-type (Figure 1B). Given that autolysis is related to cell wall synthesis (Templin et al., 1999; Wang and Sun, 2021; Zamakhaeva et al., 2021), these results suggested that deficiency of lcpB can affect cell wall synthesis. Therefore, we used transmission electron microscopy to observe the morphology of the cell wall in these three strains. As expected, the results clearly showed that wild-type bacteria revealed regular-shaped cells, while the lcpB mutant strain generated deformed cells with irregular envelopes in the early stage of growth (Figure 1C middle) while the wild-type and complemented strains were regular-shaped with smooth cell walls (Figure 1C left and right). These findings further revealed that the absence of LcpB resulted in defects in the cell wall.

As previous studies have shown, LCP proteins can bind to the promoter region of target genes to regulate their transcription (Lazarevic et al., 1992; Cieslewicz et al., 2001; Hanson et al., 2011). In order to explore the regulatory mechanism of LcpB, we first analyzed the change of gene transcription levels in the wild-type and lcpB mutant strains using RNA-seq. Interestingly, the results showed that deficiency of lcpB had minimal effect on the transcription of genes related to cell wall synthesis or degradation (Supplementary Table 5). We next verified the results of RNA-seq. RT-qPCR results showed no significant changes in the expression of cell wall related genes (Supplementary Figure 1), including autolysin genes lytM and atl (Sugai et al., 1995; Singh et al., 2010), cell wall synthesis related genes pbps, femX (Schneider et al., 2004; Sobral and Tomasz, 2019), and cell wall hydrolysis gene fmtA (Komatsuzawa et al., 1997). Since LcpB is involved in WTA synthesis (Chan et al., 2013; Srisuknimit et al., 2017), we further assessed the transcription levels of genes associated with the process, including tarG, tarH, tarO, tarL, tarS, tarA, and tarB. The results revealed little to no differences among the wild-type, lcpB mutant, and complemented strains (Figure 2A). These findings suggested that LcpB may not be a transcription regulator that affects the transcription of cell wall related genes, but to be involved in cell wall synthesis.

Moreover, members of the LCP family are putative transmembrane proteins (Hubscher et al., 2008). Therefore, we assessed transmembrane segment using TMHMM, a tool that predicts the topology of proteins (Krogh et al., 2001). Analysis of the LcpB amino acid sequence suggested the existence of a short intracellular part (1–6 residues), a single transmembrane region at the N-terminal region of the protein (7–28 residues), and that the main part of LcpB was located outside the cell (29–408 residues, Figure 2B). In order to further confirm the subcellular localization of the protein, we fused it with GFP at its C-terminal tail and the fusion gene was controlled by the native promoter of lcpB. Confocal microscopy observation showed that LcpB exhibites plasma membrane localization which is indicated by FM4-64 (Figure 2C). These results confirmed that LcpB is indeed a transmembrane protein.

Existing literature suggests that LCP proteins may have pyrophosphatase activity (Siegel et al., 2019). Therefore, in order to verify whether LcpB is a pyrophosphatase, we attempted to express the full-length protein but the efforts were unsuccessful. Consequently, the protein was truncated and its extracellular region was expressed (ΔTM-LcpB, 30–408 residues). It is also important to note that the members of the LCP family studied to date possess pyrophosphatase activity and can hydrolyze diphosphate phosphoanhydride bonds (Siegel et al., 2019). In order to ascertain whether LcpB has the same function, a diphosphate mimetic substrate was used and farnesyl pyrophosphate (FPP) was incubated with ΔTM-LcpB, which was expressed in and purified from Escherichia coli (Supplementary Figure 2). The pyrophosphatase activity of the ΔTM-LcpB proteins was then determined by quantitatively measuring the amount of inorganic phosphate (Pi) released from FPP. Compared to glutathione S-transferase (GST), the negative control, there was a significant increase in the amount of phosphate produced, after incubation with FPP (Figure 2D). These results confirmed that LcpB is a pyrophosphatase. Moreover, previous research showed that mutants lacking the LCP family can release cell wall teichoic acids into the extracellular medium (Chan et al., 2013). Therefore, we assessed the level of WTA content in the wild-type, lcpB mutant and complemented strains, in early growth stage. The findings revealed that there was a decrease in the amount of WTA in the cell wall of the lcpB mutant strain (Figure 2E and Supplementary Figure 3). These results suggested that LcpB does not regulate cell wall synthesis as a transcription factor and it may affect WTA synthesis through its pyrophosphatase activity.

Members of the LCP family contain conserved arginine residues which are thought to play a key role in enzyme activity (Kawai et al., 2011; Siegel et al., 2019). In addition, there are 15 arginine sites in LcpB, all of which are located in the extracellular region (Supplementary Figure 4A). In order to determine which arginine site was key site to the pyrophosphatase activity of LcpB, we constructed single-mutated complemented strains in which these arginines were mutated to alanines. The growth rate was then tested for preliminary screening. The results showed that, compared to the non-mutated strain, arginine mutations at position 86, 109, 207, 209, 217, and 220 led to a decrease in growth rate (Figure 3A). However, the other mutations had not effect on growth (Supplementary Figure 4B). Next, we assessed the amount of WTA in these six mutants. As expected, the amount of WTA in the six mutants decreased by 40–65%, compared to the non-mutated complementary strain (Figure 3B). These results therefore indicated that these six arginine sites are important for the activity of LcpB. The findings also showed that these six arginine sites were all located in the putative LCP domain. This suggested that the LCP domain plays a crucial role in the activity of LcpB. Consequently, these six proteins with single arginine mutations were expressed (Supplementary Figure 2), after which we evaluated their pyrophosphatase activity. Corresponding to results on the amount of WTA, ΔTM-LcpB with a single arginine mutation had a decrease in pyrophosphatase activity by 70–90% (Figure 3C). These results suggested that LcpB affected the synthesis of WTA through pyrophosphatase activity and the key arginine sites in the LCP domain regulated its enzyme activity.

We further analyzed and compared the RNA-seq data of the wild-type and lcpB mutant strains. The findings showed that several virulence-related genes were up-regulated (Supplementary Table 5), suggesting that LcpB may be involved in regulating virulence. In order to explore the relationship between LcpB and virulence, the RNA-seq results were first verified. Analysis through RT-qPCR showed that psmα, psmβ, scnL, and ehp were significantly up-regulated (Figure 4A). However, the transcription of agrA and RNAIII showed no visible alteration, suggesting that the regulation of virulence by LcpB may be agr-independent. Notably, PSMs are well-characterized toxins that play a significant role in skin infection by S. aureus (Kennedy et al., 2010). Moreover, SCN (SCIN) is an efficient modulator of neutrophil chemotaxis, phagocytosis, and killing, whose early expression is necessary for efficient modulation of early immune responses (Rooijakkers et al., 2006). Protein encoded by orf1855 is predicted to be a staphylococcal complement inhibitor which has high homology with SCN, and belongs to the SCN family. Therefore, the present study named orf1855 as scnL. Additionally, Ehp is a secreted protein which can bind to C3 to inhibit the alternative complement activation pathway (Hammel et al., 2007; Jongerius et al., 2007). Based on the RT-qPCR results, we assessed the interaction between the bacteria and the host through the levels of hemolytic activity, and the ability of form abscesses in the wild-type, lcpB mutant, and complemented strains. The hemolytic assay was firstly performed using the sheep red blood cells and the percentage of hemolytic activity was calculated relative to the positive control (100% hemolytic activity) by measuring the optical density at 543 nm. The lcpB mutant strain displayed increased hemolytic activity, compared to the wild-type strain after incubation for 2 h at 37°C and these changes could be restored by the complemented strain (Figures 4B,C). Since our above study showed that LcpB had six key arginine sites, we also tested the hemolytic ability of these single point mutant strains. The results showed that compared to the non-mutated strain, arginine mutations at position 109, 207, 209, and 217 led to an increase in hemolytic activity (Figure 4D). These results suggest that LcpB may regulate virulence through pyrophosphatase activity. In addition, we used a mouse model of subcutaneous abscess to investigate the contribution of LcpB to the pathogenicity of S. aureus. The findings showed that the ability of the lcpB mutant strain to cause skin abscesses in mice was significantly enhanced, compared to the wild-type and lcpB complemented strains. This was further demonstrated by photographs of the skin lesions (Figures 4E,F). Next, we examined bacterial colonization of the skin lesions. The results revealed that these three strains had a similar level of colonization (Figure 4G). It was therefore speculated that the increase in abscess area was not caused by the difference in bacterial quantity but by the enhanced virulence of the lcpB mutant strains. Moreover, histological examination of the lcpB mutant showed more extensive inflammation with leukocyte infiltration, destruction of the skin structure (Figure 4H). Overall, these findings showed that LcpB can promote the expression of virulence genes and affect the pathogenicity of S. aureus.

Wanner et al. (2017) reported before that agr system participates in WTA synthesis and virulence through regulating tarH. Our study showed that lcpB knockout affected WTA synthesis and virulence, but the transcriptional level of agrA, RNAIII, and tarH showed no significant difference. In order to examine whether the regulation of virulence by LcpB is agr-independent or not, we tried to knock out agr system in ST59, but failed. Therefore, another MRSA strain N315, an agr system deficient strain, was used for further study. Interestingly, lcpB deficiency greatly slowed down the growth (Figure 5A) and enhanced the hemolytic activity (Figures 5B,C). This is consistent with the phenotype of ST59 strain. This suggested that the regulation of WTA synthesis and virulence by LcpB is agr-independent.

In order to examine whether the mechanism of LcpB is common in Staphylococcus aureus or not, we knocked out lcpB in MSSA strain Newman. The results showed that lcpB deletion slightly affected the growth rate (Figure 5D) and had no visible alteration on hemolysis (Figures 5E,F). These results suggest that the effect of LcpB was strain specific and it may play a more important role in MRSA strains.