# Peptidoglycan Crosslinking Relaxation Plays an Important Role in Staphylococcus aureus WalKR-Dependent Cell Viability

Aurelia Delauné, Olivier Poupel, Adeline Mallet, Yves-Marie Coïc, Tarek Msadek, Sarah Dubrac

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Aurelia Delauné, Olivier Poupel, Adeline Mallet, Yves-Marie Coïc, Tarek Msadek, et al., Peptidoglycan Crosslinking Relaxation Plays an Important Role in Staphylococcus aureus WalKR-Dependent Cell Viability. PLoS ONE, 2011, 6 (2), pp.e17054. 10.1371/journal.pone.0017054. pasteur-02870016

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# Peptidoglycan Crosslinking Relaxation Plays an Important Role in *Staphylococcus aureus* WalKR-Dependent Cell Viability

Aurelia Delaune¹,², Olivier Poupel¹,², Adeline Mallet³, Yves-Marie Coic⁴,⁵, Tarek Msadek¹,²*, Sarah Dubrac¹,²

¹Institut Pasteur. Biology of Gram-Positive Pathogens, Department of Microbiology, Paris, France, ²CNRS, URA 2172, Paris, France, ³Institut Pasteur, Ultrastructural Microscopy Platform, Imagopole, Paris, France, ⁴Institut Pasteur, Chemistry of Biomolecules, Department of Structural Biology and Chemistry, Paris, France, ⁵CNRS, URA 2128, Paris, France

## Abstract

The WalKR two-component system is essential for viability of *Staphylococcus aureus*, a major pathogen. We have shown that WalKR acts as the master controller of peptidoglycan metabolism, yet none of the identified regulon genes explain its requirement for cell viability. Transmission electron micrographs revealed cell wall thickening and aberrant division septa in the absence of WalKR, suggesting its requirement may be linked to its role in coordinating cell wall metabolism and cell division. We therefore tested whether uncoupling autolysin gene expression from WalKR-dependent regulation could compensate for its essential nature. Uncoupled expression of genes encoding lytic transglycosylases or amidases did not restore growth to a WalKR-depleted strain. We identified only two WalKR-regulon genes whose expression restored cell viability in the absence of WalKR: *lytM* and *ssaA*. Neither of these two genes are essential under our conditions and a *ΔlytM* *ΔssaA* mutant does not present any growth defect. LytM is a glycyl-glycyl endopeptidase, hydrolyzing the pentaglycine interpeptide crossbridge, and SsaA belongs to the CHAP amidase family, members of which such as LysK and LytA have been shown to have D-alanyl-glycyl endopeptidase activity, cleaving between the crossbridge and the stem peptide. Taken together, our results strongly suggest that peptidoglycan crosslinking relaxation through crossbridge hydrolysis plays a crucial role in the essential requirement of the WalKR system for cell viability.

**Citation:** Delaune A, Poupel O, Mallet A, Coic Y-M, Msadek T, et al. (2011) Peptidoglycan Crosslinking Relaxation Plays an Important Role in *Staphylococcus aureus* WalKR-Dependent Cell Viability. PLoS ONE 6(2): e17054. doi:10.1371/journal.pone.0017054

**Editor:** Malcolm Horsburgh, University of Liverpool, United Kingdom

**Received:** October 21, 2010; **Accepted:** January 14, 2011; **Published:** February 28, 2011

**Copyright:** © 2011 Delaune et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

**Funding:** This work was supported by research funds from the European Commission (StaphDynamics [LHSM-CT-2006-019064] and BaSysBio [LSHG-CT-2006-037469] grants), the Centre National de la Recherche Scientifique (CNRS URA 2172), Agence Nationale de la Recherche (ANR Grablron and NaBab) and the Institut Pasteur (PTR N°256 and PTR N°336). Aurelia Delaune received a Young Scientist Fellowship from the Conseil Pasteur-Weizmann. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

**Competing Interests:** The authors have declared that no competing interests exist.

* E-mail: tmsadek@pasteur.fr

## Introduction

*Staphylococcus aureus* is the most common Gram-positive bacterium causing both nosocomial and community-acquired infections, which extend from superficial skin lesions to life-threatening deep-tissue invasive diseases such as endocarditis, osteomyelitis and pneumonia [1]. Asymptomatic carriage of this pathogen, usually within the nares, is widespread and often linked to *S. aureus* bacteremia [2,3].

Despite its status as a major human pathogen, many of the mechanisms involved in *S. aureus* virulence and fitness remain poorly understood. Two-component systems (TCSs) are key regulatory pathways allowing bacteria to adapt their genetic expression to environmental changes. Composed of a sensor histidine kinase, generally bound to the cell membrane, and its cognate response regulator, they act to control expression of specific sets of genes in answer to defined signals. Most *S. aureus* strains have 16 TCSs, with an additional one present in MRSA [4,5]. Although many of these systems remain to be characterized, several have been shown to play a role in virulence (AgrC/AgrA, SaeS/SaeR) [6], antibiotic resistance (VraS/VraR, GraS/GraR)

[7–9], or adaptation to environmental changes such as oxygen pressure (SrrA/SrrB) [10]. Among the 16 TCSs common to all *S. aureus* strains, only one is essential for bacterial survival, the WalK/WalR system (aka YycG/YycF), highly conserved among low G+C % Gram-positive bacteria [11–13].

In previous work, we highlighted the major role the WalKR system plays in *S. aureus* peptidoglycan metabolism by controlling expression of most of the cell wall hydrolase genes [14]. Nevertheless, none of the identified WalKR-regulated genes appear to be essential, and the reason this signal transduction pathway is required for cell viability remains to be established. Cell wall metabolism is vital for bacterial fitness, playing an important role in cell shape and division, resistance to external stresses such as osmotic pressure, as well as host-pathogen interactions. While the role of cell wall plasticity in bacterial fitness is well documented, none of the *S. aureus* cell wall hydrolases have been characterized as essential for bacterial growth, presumably due to the high level of genomic redundancy in this pathogen, which produces several autolytic enzymes with similar activities (see MiST Database: http://genomics.ornl.gov/mist) [15,16]. The WalKR system has also been extensively studied in *B. subtilis* and *S. pneumoniae*, and

<figure><img src="image_3.png" /></figure>
shown to be involved in regulation of cell wall metabolism in these
bacteria [13,17–19]. Furthermore, in *S. pneumoniae*, the essentiality
of the WalKR system has been linked to the regulation of a single
gene, *pcsB*, encoding a putative cell wall amidase [17,18].

In this study, we tested whether overexpression of WalKR-
regulated autolysin genes in a WalKR-independent manner could
compensate for the essential nature of the system. Indeed, since the
WalKR system controls the synthesis of most of the identified cell wall
hydrolases, we reasoned that the essentiality of this system could be
linked to global regulation of one type of autolytic activity rather than
that of a single gene. This allowed us to show that only increased
production of LytM or SsaA can restore cell viability in the absence of
WalKR. Although the *lytM* and *ssaA* genes are not essential when
WalKR is present, our results indicate that peptidoglycan crosslinking
relaxation through crossbridge hydrolysis must play a crucial role in
the essential nature of the WalKR system.

Results

WalKR depletion leads to cell wall thickening and a
defect in cell division

We have previously shown that the WalKR system is a major
regulator of cell wall degradation, and is also necessary for
peptidoglycan biosynthesis and turn-over [14]. Since none of the
identified WalKR regulon genes appear to be essential, this
suggested that the WalKR requirement for cell viability could be
specifically linked to its role in controlling cell wall metabolism.
Cell wall plasticity is involved in daughter cell separation and
mutant strains deleted for two of the major *Staphylococcus aureus* cell

wall hydrolase genes (*atlA* and *sle1*) harbor an abnormal rough cell
surface and a pronounced defect in cell separation [20,21].

In order to identify morphological changes associated with
WalKR-depletion, we used electron microscopy to examine S.  
aureus cells where the *walRK* operon is placed under the control of  
an IPTG-dependent inducible promoter (strain ST1000, *Pspac-  
*walRK*) [12], grown in the presence or absence of IPTG. Although  
no gross morphological differences were observed by scanning  
electron microscopy (data not shown), ultrastructural analysis by  
transmission electron microscopy (TEM) showed significant  
changes. Indeed, when WalKR is produced, the ST1000 strain  
displays the typical diplococcal S. aureus morphology, with a single  
central division septum (Fig. 1A). However, as shown in Fig. 1B,  
WalKR-depleted cells exhibit abnormal and misplaced division  
septa, with the formation of several new septa before separation of  
the daughter cells (indicated by arrows, Fig. 1B). WalKR-depleted  
cells also displayed a rougher cell surface, with amorphous “fuzzy”  
extracellular material, and increased cell wall thickness, almost  
twice that of cells producing WalKR (58.56 +/- 12.05 nm vs.  
32.64 +/- 4.54 nm, respectively, Student *t* test P-value <0.05)  
(Fig. 1A and 1B). Interestingly, all of these phenotypes are  
strikingly similar to those described for S. aureus *atlA sle1* mutants  
[21] and VISA (vancomycin intermediate S. aureus) strains with  
reduced susceptibility to vancomycin [22].

Uncoupling cell wall hydrolase gene expression from
WalKR-dependent regulation

Given the dramatic impact of WalKR-depletion on septum
placement and cell separation, we reasoned that WalKR-

<figure><img src="image_4.png" /></figure>

**Figure 1.** *S. aureus* starved for WalK/WalR display thickened cell walls and aberrant division septa. Strain ST1000 carrying a chromosomal Pspac-walRK fusion was grown in TSB +/- 1 mM IPTG, harvested at OD<sub>600nm</sub> = 1 (corresponding to exponential phase for the culture with IPTG and cessation of growth for the culture without IPTG) and embedded in thin sections for ultrastructure examination by transmission electron microscopy. Panel A: ST1000 grown with 1 mM IPTG. Panel B: ST1000 grown without IPTG. Bars represent 0.5 µm for the upper panels and 2 µm for the lower panels. Black arrows indicate aberrant division septa.
doi:10.1371/journal.pone.0017054.g001

<figure><img src="image_5.png" /></figure>
independent expression of one or more cell wall hydrolase genes might restore cell viability in the absence of WalKR. We therefore tested the effect of uncoupling expression of autolysin-encoding genes from WalKR-dependent regulation.

Peptidoglycan hydrolases include lytic transglycosylases cleav-
ing the glycan strand, amidases that hydrolyze the amide bonds
linking the stem peptides to the glycan strand, and glycyl-glycyl
endopeptidases cleaving the *S. aureus*-specific pentaglycine
interpeptide crossbridges [16]. As shown in Fig. 2, we chose
to overexpress WalKR-regulated genes shown or presumed to
be involved in cell wall degradation. We have recently shown
that *sle1* [21], encoding a CHAP-domain amidase (Cysteine,
Histidine-dependent Amidohydrolases/Peptidases) [23,24], is a
novel member of the WalKR regulon (A. Delaune *et al*., in
preparation). The major autolysin gene, *atlA*, which encodes a
bifunctional enzyme with amidase and glucosaminidase domains
[25] was not included in this approach since we were unable to
obtain an expression plasmid carrying this gene, presumably
because its expression is deleterious for *E. coli*. Genes were
cloned into plasmid pCN51, placing them under control of the
Pcad cadmium chloride-inducible promoter [26] (see Materials
and Methods). The resulting overexpression plasmids were then
introduced into strain ST1000 (*Pspac-walRK*), whose growth is
IPTG-dependent [12]. Growth of strain ST1000 remained
IPTG-dependent in the presence of the control vector pCN51
and cadmium chloride (Fig. 3A and Fig. 4A, open symbols),
indicating that neither the pCN51 vector nor the presence of
cadmium chloride interfere with expression from the *Pspac*
promoter.

<figure><img src="image_6.png" /><figcaption>**Figure 2. Diagram of S. aureus peptidoglycan and cleavage sites for WalKR-regulon encoded cell wall hydrolases.** Peptidoglycan cleavage sites for WalKR-regulon encoded cell wall hydrolases are indicated: A) glycan strand glycosidic bond (glucosaminidase: AtlA). B) glycan strand glycosidic bond (lytic transglycosylases: SceD and IsaA). C) N-Acetylmuramyl-L-alanyl amide bond (AtlA amidase domain, and the Sle1, SsaA, SAOUHSC_00671, _00773, _02576 and _02883 CHAP domain hydrolases). D) pentaglycine bridge glycyl-glycyl bond (LytM endopeptidase).</figcaption></figure>

doi:10.1371/journal.pone.0017054.g002

<figure><img src="image_7.png" /></figure>

<figure><img src="image_8.png" /></figure>

<figure><img src="image_9.png" /><figcaption>**Figure 3. Overexpression of *lytM* restores viability to cells starved for WalKR.** Cells were grown in TSB +/− 1 mM IPTG and gene overexpression was performed using the multicopy pCN51 plasmid carrying a CdCl₂ inducible promoter whose expression was induced using 0.25 µM CdCl₂. A) Growth curves are depicted for derivatives of S. aureus strain ST1000 carrying a *Pspac-walRK* chromosomal fusion and the pCN51 control vector (ST1012, open symbols) in TSB-CdCl₂ in the presence (○) or absence (◊) of 1 mM IPTG. Growth of the ST1000 derivatives overproducing the LytM glycyl-glycyl endopeptidase (ST1002, ▲) or the inactive LytM H291A enzyme (ST1128, ■) were carried out in TSB-CdCl₂ without IPTG. In the presence of IPTG, strains overproducing cell wall hydrolases grew similarly to the control ST1012 strain (data not shown). B) Relative levels of *lytM* transcripts were measured by qRT-PCR during growth of strains ST1012 (pCN51 control) or ST1002 (overexpressing *lytM*), in TSB-CdCl₂. Expression levels were normalized using the 16S rRNA as an internal standard and are indicated as n-fold change, expressed as the means and standard deviations of quadruplicate experiments. C) Western blot showing LytM and LytM H291A overproduction. Cells were grown in TSB-CdCl₂ with or without 1 mM IPTG, and the LytM and LytM H291A proteins were detected by Western blotting using purified polyclonal rabbit antibodies. The arrow indicates the position of the mature enzyme, with a predicted molecular mass of 32 kDa. Lanes: 1, ST1012; 2, ST1012 +IPTG; 3, ST1002 (*lytM*); 4, ST1002 +IPTG; 5, ST1128 (*lytM*\* H291A); 6, ST1128 +IPTG; 7, Purified mature his-tagged 'LytM.</figcaption></figure>

doi:10.1371/journal.pone.0017054.g003

<figure><img src="image_10.png" /></figure>
<figure><img src="image_11.png" /></figure>

<figure><img src="image_12.png" /><figcaption>**Figure 4.** Overexpression of *ssaA* restores viability to cells starved for WalKR. Cells were grown in TSB +/− 1 mM IPTG and gene overexpression was performed using the multicopy pCN51 plasmid in the presence of 0.25 µM CdCl₂. A) Growth curves are depicted for derivatives of S. aureus strain ST1000 carrying a Pspac-walRK chromosomal fusion and the pCN51 control vector (ST1012, open symbols) in TSB-CdCl₂ in the presence (○) or absence (◇) of 1 mM IPTG. Growth of the ST1000 derivatives overproducing SsaA (ST1123, ▲) or inactive SsaA C171S (ST1124, ■) was carried out in TSB-CdCl₂ without IPTG. In the presence of IPTG, strains overproducing cell wall hydrolases grew similarly to the control ST1012 strain (data not shown). B) Relative levels of *ssaA* transcripts were measured by qRT-PCR during growth of strains ST1012 (pCN51 control) or ST1123 (overexpressing *ssaA*), in TSB-CdCl₂. Expression levels were normalized using the 16S rRNA as an internal standard and are indicated as n-fold change, expressed as the means and standard deviations of quadruplicate experiments.</figcaption></figure>

doi:10.1371/journal.pone.0017054.g004

## Regulation of lytic transglycosylase genes is not responsible for WalKR essentiality

Lytic transglycosylases cleave the cell wall glycan strand. Lytic activity of IsaA and SceD has been demonstrated and we have shown that the corresponding genes are controlled by the WalKR system [14,27]. We cloned the *isaA* and *sceD* genes in the pCN51 vector, placing them under the control of the P*ead* promoter. The resulting plasmids, pAD13 and pAD01, were introduced into strain ST1000 and IPTG-dependent growth was examined in the

presence of CdCl₂. WalKR-independent overexpression of *isaA* or *sceD* was verified by qRT-PCR, and did not allow growth of the Pspac-walRK strain in the absence of IPTG (data not shown).

## Production of the LytM glycyl-glycyl endopeptidase restores growth to WalKR-depleted cells

The S. aureus NCTC 8325 core genome encodes three M23 peptidase domain proteins with putative glycyl-glycyl endopeptidase activity. Among them is the previously characterized LytM enzyme [28,29], whereas the other two, SAOUHSC_00174 and _02464, have not been studied. Transcription of *lytM* is under the control of the WalKR system, and we have shown that WalR binds directly to its target operator sequence upstream from the *lytM* promoter [12,14]. The SAOUHSC_00174 and _02464 genes are not preceded by a WalR binding site, and we have shown that their expression is not controlled by the WalKR system (data not shown).

As shown in Fig. 3A, a Pspac-walRK strain overexpressing *lytM* (strain ST1002 carrying plasmid pSD3-13) grew as well in the absence of IPTG, i.e. without expression of *walRK*, as the control strain (ST1000/pCN51) in the presence of IPTG, indicating that *lytM* overexpression is able to fully restore growth and viability to cells starved for the WalKR system. We verified by quantitative real-time PCR (qRT-PCR) analysis that *walRK* expression remained inducible by IPTG, and unexpressed in the absence of IPTG in the ST1012 (ST1000/pCN51) and ST1002 (ST1000/pSD3-13) strains grown in the presence of CdCl₂ (data not shown).

Overexpression of *lytM* and corresponding LytM overproduction were verified by qRT-PCR and Western blot analysis. As shown in Fig. 3B, pSD3-13 increases *lytM* transcription approximately 50-fold, and this increased transcription is directly correlated to significant LytM protein accumulation (Fig. 3C, lane 1 compared to lane 3). We note that *in vivo* levels of LytM protein are undetectable in the ST1000/pCN51 strain with or without WalKR production (Fig. 3C, lanes 1 and 2), although basal levels of the protein could be seen after overexposing the Western blot (data not shown). Although the genes encoding the other two M23 peptidase proteins (SAOUHSC_00174 and _02464) are not regulated by the WalKR system, we also tested their ability to compensate WalKR depletion, and showed that neither of the two proteins allowed growth of WalKR-depleted cells when overproduced (data not shown).

In order to determine whether the glycyl-glycyl endopeptidase enzymatic activity of LytM was specifically required for its capacity to restore viability to WalKR-depleted cells, we used site-directed mutagenesis to inactivate its catalytic site. The M23 peptidase domain of LytM extends from residues 208 to 309, and it has previously been shown that histidine residue 291 is crucial for LytM activity [30]. We placed the *lytM*** mutant allele (H291A) under control of P*ead* in pCN51, generating plasmid pSD3-24. As shown in Fig. 3A, expression of *lytM*** cannot restore growth to cells lacking WalKR. In order to compare LytM and LytM H291A production *in vivo* and to rule out the possibility that the inactive LytM H291A protein might be specifically targeted by proteases, we performed Western blot experiments with polyclonal LytM antibodies. As mentioned above, native levels of LytM are undetectable in the ST1000/pCN51 strain, however as shown in Fig. 3C introduction of pSD3-13 or pSD3-24 into strain ST1000 clearly led to significant overproduction of LytM (lanes 3 & 4) or LytM H291A (lanes 5 & 6), respectively.

LytM belongs to the lysostaphin-type peptidase family, whose prototype is the S. simulans-produced lysostaphin, which also acts as a glycyl-glycyl endopeptidase, cleaving the pentaglycine cross-

<figure><img src="image_13.png" /></figure>
bridge [31]. Since active recombinant lysostaphin is readily available [32], we tested the effect of adding sub-lethal concentrations (0.01 and 0.025 µg/ml; higher amounts lead to cell lysis) to the extracellular medium on growth of the ST1000 IPTG-dependent *Pspac-walRK* strain. There was no effect on the growth profile of the WalKR-depleted culture, indicating that extracellular glycyl-glycyl activity of lysostaphin cannot compensate for the absence of WalKR (data not shown).

We also tested the effect of adding purified LytM extracellularly, by purifying a His-tagged recombinant form of mature LytM, which was also unable to restore growth in the absence of WalKR, suggesting localization of the enzyme may be important (data not shown). In a reciprocal experiment, we cloned a DNA fragment encoding a recombinant active and secreted form of lysostaphin (*lss7* allele, partially deleted in the propeptide encoding region) [33] under the control of the CdCl₂-inducible promoter of pCN51, generating the lysostaphin production plasmid pAD08. The induction conditions used (0.25 µM CdCl₂) allowed normal growth of strain ST1000/pAD08 in the presence of IPTG. However, unlike LytM, intracellular production of lysostaphin at sub-lethal levels under conditions of WalKR depletion did not restore cell growth, suggesting that LytM and lysostaphin may differ with respect to their role in cell wall metabolism (data not shown).

### ssaA is the only WalKR-dependent CHAP domain amidase gene able to restore growth to a WalKR-depleted strain

CHAP domains are conserved in several peptidoglycan hydrolases, often associated with amidase activity [23, 24, 34, 35]. Careful examination of the *S. aureus* NCTC 8325 genome [5] allowed us to predict 14 genes encoding CHAP domain proteins (SAOUHSC_00256, _00427 or sle1, _00671, _00773, _01219, _01515, _02019, _02023, _02173, _02571 or ssaA, _02576, _02855, _02883, _02979). In agreement with the presence of our defined consensus WalR binding site DNA sequence, we have shown that six of these are controlled by the WalKR system: ssaA, SAOUHSC_00671, _00773, _02576, _02883 [14] and sle1 (A. Delaune et al., in preparation).

We tested the effect of overexpressing each of these six genes on growth of the ST1000 (*Pspac-walRK*) strain in the presence or absence of IPTG. Whereas WalKR-independent overexpression of *sle1*, SAOUHSC_00671, _00773, _02576 or _02883, as verified by qRT-PCR, had no effect on IPTG-dependent growth (data not shown), we showed that expression of *ssaA* from the *Pcad* promoter (plasmid pAD12) fully restores growth of *S. aureus* in the absence of WalKR (Fig. 4A). While this work was in progress, a similar finding was independently observed by the group of Simon Foster (S. Foster, University of Sheffield, personal communication).

Growth of the ST1000 strain, starved for the WalKR system but overexpressing *ssaA*, was identical to that of the strain expressing *walRK*, and we measured *ssaA* expression from the *Pcad* promoter and showed that this overexpression does not interfere with regulated expression of *walRK* from the *Pspac* promoter which remained unexpressed in the absence of IPTG (Fig. 4B and data not shown respectively). As shown in Fig. 4B, *ssaA* transcription was increased approximately 30-fold in strain ST1123, carrying the pAD12 plasmid with the *Pcad-ssaA* fusion, as compared to the strain carrying the control pCN51 vector under the same culture conditions (TSB with 0.25 µM CdCl₂), similar to the *lytM* overexpression levels shown in Fig. 3B.

CHAP domains are characterized by two highly conserved motifs containing cysteine, histidine and asparagine residues, all of which have been shown to be critical for enzymatic activity [35,36]. In order to verify whether the ability of SsaA to restore

viability to WalKR-starved cells was linked to its enzymatic activity, we used site-directed mutagenesis to change cysteine residue 171 to serine, a mutation that has previously been shown to abolish the activity of PlyC, a bacteriophage lysin with a CHAP domain [37]. The mutated *ssaA*\* (C171S) allele was placed under the control of the *Pcad* promoter and introduced into strain ST1000. As shown in Fig. 4A, overexpression of the *ssaA*\* allele is not able to restore growth to a WalKR-depleted strain. We verified by qRT-PCR that the mutated *ssaA*\* allele was overexpressed from the *Pcad* promoter at levels comparable to those of the native *ssaA* allele (data not shown). Taken together, these data indicate that SsaA is the only CHAP domain protein encoded by WalKR regulon genes able to restore growth to WalKR-depleted cells and that this is dependent on its enzymatic activity.

### SsaA and LytM are not sufficient to completely compensate for the absence of WalKR

As shown above, WalKR-depleted cells exhibit abnormal division septa and a rougher cell surface. We therefore examined the morphology of cells overproducing LytM or SsaA in the absence of WalKR, comparing strains ST1012 (pCN51), ST1002 (pSD3-13) and ST1123 (pAD12) by TEM. As shown in Fig. 5A, overexpression of *lytM* or *ssaA* only partially restored cell morphology: although cell wall thickness was diminished, cells presented a separation defect and some abnormal and misplaced division septa remained present in WalKR-depleted LytM- or SsaA-overproducing strains even though growth was indistinguishable from that of cells producing WalKR. In order to test the viability of these abnormal cells, we used fluorescence microscopy and the Live/Dead BacLight™ bacterial viability assay on stationary phase cultures of the ST1000 strain carrying either the pCN51 control plasmid or the pSD3-13 and pAD12 plasmids allowing overexpression of *lytM* or *ssaA*, respectively. As expected, viability of strain ST1000 carrying the pCN51 vector remains strictly dependent on WalKR, and almost all of the cells died when starved for WalKR (red staining, Fig. 5B upper panel). As shown in Fig. 5B, the ST1000/pSD3-13 or pAD12 strains were still viable in the absence of WalKR (green stain) but formed aggregates characteristic of bacteria with a cell separation defect (upper panel). This defect is not linked to *lytM* or *ssaA* overexpression since when the WalKR system was produced, the cells were well separated (lower panel).

## Discussion

We report here the first data establishing a direct link between WalKR-dependent regulation of cell wall metabolism and WalKR essentiality in *S. aureus*. The strategy of uncoupling expression of genes encoding cell wall hydrolases from WalKR-dependent activation allowed us to show that only two WalKR-regulon autolysin genes are able to compensate for the absence of this essential TCS, *lytM* and *ssaA*. Our results suggest that loss of cell viability following WalKR depletion can be compensated through peptidoglycan crosslinking relaxation.

LytM is produced with a signal peptide, suggesting that it is secreted, and carries a peptidase M23 domain similar to that of *S. simulans* lysostaphin [28]. LytM is a glycyl-glycyl endopeptidase, cleaving the pentaglycine interpeptide crossbridges of the *S. aureus* cell wall [29,30,38]. Cross-bridges stabilize the *S. aureus* cell wall and are essential since mutants impaired in their biosynthesis (*fmhB*, *femA*, *femB*) are non- or barely viable [39–41]. Furthermore, the pentaglycine interpeptide bridges are involved in exposure of cell surface proteins that are covalently anchored to them by

<figure><img src="image_14.png" /></figure>
A

<figure><img src="image_15.png" /></figure>

B

<figure><img src="image_16.png" /></figure>

**Figure 5.** Overexpression of *lytM* or *ssaA* restores cell viability but not morphology to cells lacking the WalK/WalR TCS.
Bacteria were grown in TSB CdCl₂ 0.25 µM. To produce the WalKR system in derivatives of S. aureus strain ST1000 carrying a Pspac-walRK chromosomal fusion and the indicated plasmids, 1 mM IPTG was added to the culture (walRK on). In the absence of IPTG, cells were depleted for WalKR (walRK off). A) Transmission electron microscopy ultrastructure of strains overproducing SsaA or LytM. Cells were grown in TSB-CdCl₂ +/− 1 mM IPTG, harvested at an OD₆₀₀nm = 1 (corresponding to exponential phase for the culture with IPTG and cessation of growth for the culture without IPTG) and embedded in thin sections for ultrastructure examination by transmission electron microscopy. Bars represent 1 µm for the upper panels and 2 µm for the lower panels. B) Viability assay of strains overproducing SsaA or LytM using the LIVE/DEAD BacLight™ assay. Two hours after reaching stationary phase, 10 ml of culture were sampled for fluorescent staining. For the control ST1012 strain without IPTG, the sample was taken 4 hours after cessation of growth following WalKR-depletion. Live bacteria appear green, following SYTO-9 staining, and dead cells are stained in red due to penetration of propidium iodide through compromised membranes. doi:10.1371/journal.pone.0017054.g005

sortase-dependent cross-linking [42]. Several studies have established a link between the degree of crosslinking of peptidoglycan and methicillin resistance in a PBP2'-independent manner, as well as resistance to other unrelated classes of antibiotics such as glycopeptides [39,43]. Pentaglycine cross-bridges thus play key roles in bacterial fitness, antibiotic resistance and in virulence, through the control of surface protein display, so it is not surprising that their biosynthesis genes are essential.

SsaA carries a carboxy-terminal CHAP domain, and its ortholog in S. epidermidis is highly antigenic and was recovered from whole cells and culture supernatant, suggesting it is also secreted [44]. SsaA has been linked to the hypersensitivity to macrolide-lincosamide-streptogramin B antibiotics of a thermosensitive walRK mutant in S. aureus, though the underlying molecular mechanism is unknown [45]. CHAP domains are conserved in a subset of peptidoglycan hydrolases [23, 24, 34, 35] and we have identified 14 genes encoding CHAP-domain proteins in the S. aureus NCTC 8325 genome [5], and shown that six of these are regulated by the WalKR system [14] and (A. Delaune et al., in preparation). Of these, only ssaA is able to compensate for

the absence of WalKR when it is overexpressed (Fig. 4). Among the bacterial CHAP proteins of S. aureus, only Sle1, which acts as a N-acetylmuramyl-L-alanyl amidase, has been characterized [21].

Degradation of peptidoglycan is critical, since cell wall plasticity is required for cell shape and cellular division. AtlA, one of the major S. aureus autolysins, is a bifunctional enzyme with amidase and glucosaminidase domains, and localizes in rings associated with division septa, consistent with its requirement for daughter cell separation by splitting the crosswall [46-48]. The Sle1 amidase also appears to play a crucial role in splitting the septum during cell division, since a sle1 mutant displays a higher number of septa per cell but without subsequent cell separation [21]. Interestingly, strains with mutations inactivating either atlA or sle1 are viable and their growth rates are normal, with only an atlA sle1 mutant presenting impaired growth [21].

We have shown here that WalKR depletion leads to a rougher cell surface, abnormal division septa and a defect in cell separation, but that cell viability is restored when LytM or SsaA are overproduced. Little is known about the role of these two autolytic enzymes in bacterial fitness. Unlike AtlA, LytM is not localized around the division septa, but distributed over the cell surface, suggesting that its role could be more general, involved in cell wall plasticity rather than cell division [29]. SsaA localization has never been studied. A ΔlytM mutant had not been described in the literature, however we constructed a mutant strain in which the entire lytM coding sequence was deleted and have shown that this mutant strain has no growth defect in rich medium, and behaves as the parental strain with respect to lysostaphin sensitivity and resistance to high osmolarity (data not shown). Essentiality of ssaA was initially controversial since although a mutant strain had been generated some time ago [45] a recent report on a genome-wide screening for essential genes in S. aureus using transposon-mediated differential hybridization suggested that it might be essential [49]. We constructed a ΔssaA mutant as well as a ΔlytM ΔssaA mutant (Strains ST1158 & ST1164; Table 1), and both grew as well as the parental strain, and did not display increased sensitivity to Triton X-100-induced lysis (data not shown).

Although LytM is the only glycyl-glycyl endopeptidase characterized so far in S. aureus [28,29], genome scanning allowed us to predict two more chromosomal genes encoding potential glycyl-glycyl endopeptidases as they share the LytM Pfam M23 peptidase domain (SAOUHSC_00174, and _02464). This likely functional redundancy could explain why the ΔlytM mutant has no obvious phenotype. The SAOUHSC_00174 and _02464 genes do not have a WalR binding site in their upstream region, their expression does not appear to be controlled by WalKR and their overexpression did not restore growth to cells depleted for WalKR (data not shown). The two genes appear to be expressed at a very low basal level and their roles in cell wall metabolism, if any, remains to be established.

Another well-known glycyl-glycyl endopeptidase cleaving the pentaglycine interpeptide crossbridge is lysostaphin, produced by S. simulans [50]. LytM and lysostaphin are described as having similar enzymatic activities, but we have shown that lysostaphin, added extracellularly or produced within the cells, was not able to restore viability to WalKR-depleted bacteria. This could suggest that location/targeting of LytM may be crucial or that LytM enzymatic activity could have a mild effect, allowing the cell wall to gain enough plasticity to restore growth, whereas the effect of lysostaphin has been shown to be drastic as it leads to cell lysis by forming holes in the cell wall [51]. Since distribution over the cell surface has been observed for LytM, it seems unlikely that localization is crucial for its activity [29]. The main enzymatic activities of LytM and lysostaphin are the same, i.e. glycyl-glycyl endopeptidase, but while they share a

<figure><img src="image_17.png" /></figure>
<table><caption>**Table 1.** Strains and plasmids used in this study.</caption><thead><tr><th>Strain or plasmid</th><th>Description</th><th>Source or reference*</th></tr></thead><tbody><tr><td colspan="3"><strong>Strains</strong></td></tr><tr><td>RN4220</td><td>Restriction-deficient transformation recipient strain</td><td>[68]</td></tr><tr><td>ST1000</td><td>RN4220 Pspac-walRK</td><td>[12]</td></tr><tr><td>ST1002</td><td>Pspac-walRK pSD3-13</td><td>pSD3-13ST1000</td></tr><tr><td>ST1012</td><td>Pspac-walRK pCN51</td><td>pCN51ST1000</td></tr><tr><td>ST1020</td><td>Pspac-walRK pAD01</td><td>pAD01ST1000</td></tr><tr><td>ST1031</td><td>ΔlytM</td><td>pAD07RN4220</td></tr><tr><td>ST1057</td><td>Pspac-walRK pAD04</td><td>pAD01ST1000</td></tr><tr><td>ST1069</td><td>Pspac-walRK pAD06</td><td>pAD01ST1000</td></tr><tr><td>ST1081</td><td>Pspac-walRK pAD08</td><td>pAD01ST1000</td></tr><tr><td>ST1083</td><td>Pspac-walRK pAD09</td><td>pAD01ST1000</td></tr><tr><td>ST1123</td><td>Pspac-walRK pAD12</td><td>pAD01ST1000</td></tr><tr><td>ST1124</td><td>Pspac-walRK pSD3-25</td><td>pSD3-25ST1000</td></tr><tr><td>ST1128</td><td>Pspac-walRK pSD3-24</td><td>pSD3-24ST1000</td></tr><tr><td>ST1133</td><td>Pspac-walRK pAD13</td><td>pAD01ST1000</td></tr><tr><td>ST1134</td><td>Pspac-walRK pAD14</td><td>pAD01ST1000</td></tr><tr><td>ST1135</td><td>Pspac-walRK pAD15</td><td>pAD01ST1000</td></tr><tr><td>ST1158</td><td>ΔssaA</td><td>pAD18RN4220</td></tr><tr><td>ST1164</td><td>ΔlytM ΔssaA</td><td>pAD18ST1031</td></tr><tr><td colspan="3"><strong>Plasmids</strong></td></tr><tr><td>pCN51</td><td>Vector for CdCl<sub>2</sub>-dependent gene expression</td><td>[26]</td></tr><tr><td>pET28/16</td><td>Vector for protein overproduction in E. coli</td><td>[69]</td></tr><tr><td>pMAD</td><td>Allelic exchange vector</td><td>[63]</td></tr><tr><td>pCXlss7</td><td>Plasmid allowing lysostaphin production</td><td>[33]</td></tr><tr><td>pSD3-13</td><td>pCN51-lytM</td><td>This study</td></tr><tr><td>pSD3-24</td><td>pCN51-lytM*</td><td>This study</td></tr><tr><td>pSD3-25</td><td>pCN51-ssaA*</td><td>This study</td></tr><tr><td>pAD01</td><td>pCN51-sceD</td><td>This study</td></tr><tr><td>pAD04</td><td>pCN51-sle1</td><td>This study</td></tr><tr><td>pAD05</td><td>pET28/16-'lytM</td><td>This study</td></tr><tr><td>pAD06</td><td>pCN51-SAOUHSC_00773</td><td>This study</td></tr><tr><td>pAD07</td><td>pMADΔlytM</td><td>This study</td></tr><tr><td>pAD08</td><td>pCN51-Iss7</td><td>This study</td></tr><tr><td>pAD09</td><td>pCN51-SAOUHSC_02883</td><td>This study</td></tr><tr><td>pAD12</td><td>pCN51-ssaA</td><td>This study</td></tr><tr><td>pAD13</td><td>pCN51-isaA</td><td>This study</td></tr><tr><td>pAD14</td><td>pCN51-SAOUHSC_00671</td><td>This study</td></tr><tr><td>pAD15</td><td>pCN51-SAOUHSC_02576</td><td>This study</td></tr><tr><td>pAD18</td><td>pMADΔssaA</td><td>This study</td></tr></tbody></table>

catalytic domain, they also contain other domains as well, in particular the amino-terminal half of LytM (residues 26 to 212) whose specific role remains to be established. We and others have shown that LytM is not efficient in lysing S. aureus cells (data not shown) while it can cleave pentaglycine bridges and modify cell wall thickness without disrupting it [28,30,38].

As we have shown here, overexpression of either lytM or ssaA restored cell viability, but only partially compensated cell

morphology in the absence of WalKR, with a cell separation defect and abnormal, incomplete or misplaced division septa still present (Fig. 5). This partial restoration is very likely due to the fact that WalKR controls expression of both atla [14] and sle1 (A. Delaune *et al.*, in preparation), since their expression is lowered in cells starved for WalKR and these remaining phenotypes are highly similar to those of a sle1 atla mutant [21]. The initial finding that both LytM and SsaA can compensate for the absence of WalKR when they are overproduced was puzzling at first, since their enzymatic activities were thought to be distinct. Indeed, SsaA, with a CHAP domain, was annotated as an amidase, cleaving the N-acetylmuramoyl-L-alanyl amide bond, whereas LytM is a glycyl-glycyl endopeptidase. However, CHAP domains are in fact associated with two different types of peptidoglycan cleavage activities: N-acetylmuramoyl-L-alanyl amidase as well as D-alanyl-glycyl endopeptidase activity [23,24,34,35]. This latter activity, effectively cleaving between the pentaglycine crossbridge and the stem peptide, is formally equivalent to that of LytM, since the end result of hydrolysing either the D-alanyl-glycyl or glycyl-glycyl bonds will lead to peptidoglycan crosslinking relaxation and release of a polyglycine extremity. Whereas bacterial CHAP domains are usually located at the carboxy-terminus, many S. aureus bacteriophage-encoded endolysins also have CHAP domains but at the amino-terminus [35]. Two of these endolysins have been well characterized: Ø11 LytA, and phage K LysK, sharing a tripartite organization: an amino-terminal CHAP domain, a central amidase2 domain, and a SH3b cell wall-binding domain. Both LytA and LysK have two distinct demonstrated enzymatic activities: a N-acetylmuramyl-L-alanyl amidase activity conferred by the amidase2 central domain and a D-alanyl-glycyl endopeptidase activity involving the CHAP domain [52,53]. Our results, showing that SsaA behaves differently from the other WalKR-regulon encoded CHAP domain proteins, strongly suggest that the SsaA CHAP domain is also endowed with D-alanyl-glycyl endopeptidase activity, which could explain why it can act as well as LytM in restoring cell viability in the absence of WalKR.

The only other bacterium in which essentiality of the WalKR system can be bypassed by expression of a regulon gene is *Streptococcus pneumoniae*. In Streptococci, WalKR essentiality is slightly different than in S. aureus or B. subtilis since only the gene encoding the response regulator is essential [13,54,55]. Transcriptome analyses have shown that WalKR regulates expression of several genes involved in cell wall degradation [17]. Among them is *pcsB*, which encodes a CHAP domain protein, thought to have amidase activity, and involved in cell septation during division [17]. Interestingly, the group of Malcolm Winkler (Indiana University) was able to show that constitutive expression of the essential *pcsB* gene is sufficient to bypass the requirement of WalR for cell viability in S. pneumoniae [17]. In agreement with our results, WalKR essentiality in S. pneumoniae as in S. aureus appears linked to regulation of a gene involved in cell wall hydrolysis, and in both cases a gene encoding a CHAP domain protein (*pcsB* or SsaA, respectively) is involved.

The essential nature of the WalKR system has made it a highly attractive target for novel antimicrobial compounds [56,57] and recent results indicate that this TCS is specifically activated in S. aureus during nasal colonization [58,59], suggesting it may play an important role in host-pathogen adaptation and virulence and emphasizing the importance of understanding its requirement for cell viability.

We have shown here that the genes encoding LytM and SsaA are not essential in S. aureus, but that peptidoglycan crosslinking relaxation through pentaglycine cross-bridge cleavage is important

<figure><img src="image_18.png" /></figure>

<footnote>* Arrows indicate plasmid introduction by electroporation.
doi:10.1371/journal.pone.0017054.t001</footnote>
in restoring cell viability in the absence of WalKR. Cell wall
degradation is an essential process, with cross wall splitting
required during cell division and peptidoglycan expansion
requiring the release of free extremities to add new cell wall
monomers through the activity of penicillin binding proteins. As
shown in this paper, overproduction of either LytM or SsaA
restores the viability of WalKR-depleted cells but not normal cell
separation. We can speculate that the enzymatic activities of SsaA
and LytM allow normal cell wall enlargement by increasing free
polyglycine extremities, allowing cell wall biosynthesis through
transpeptidase reactions. Future work will aim at understanding
why this is so important in a context in which most of the cell wall
hydrolases are down-regulated.

Materials and Methods

Bacterial strains and growth media

*Escherichia coli* K12 strain DH5α™ [F^− (φ80dlacZΔM15) Δ(lacZYA-argF) U169 recA1 endA1 hsdR17 (r_k^−, m_k^+) phoA supE44 λ^− thi-1 gyrA96 relA1] (Invitrogen) was used for cloning experiments, and *E. coli* strain BL21 λ DE3 [60] (Novagen) for protein overproduction and purification. *E. coli* strains were grown in LB medium and ampicillin (100 µg/ml) was added when required. *Staphylococcus aureus* strains and plasmids used in this study are listed in Table 1. *S. aureus* strain ST1000 (RN4220 *Pspac-walRK*) [12] and derivatives described in Table 1 were grown in Trypticase Soy Broth (TSB; Difco) supplemented with chloramphenicol (10 µg ml⁻¹), erythromycin (1 µg ml⁻¹) and IPTG (1 mM) when required. Cadmium chloride (CdCl₂) was added at a final concentration of 0.25 µM for expression from the *Pcad* promoter. Bacterial growth experiments were carried out in microtiter plates (100 µl culture volume) and incubated in a Synergy 2 thermoregulated spectrophotometer plate reader (BioTek) using the Gen5™ Microplate Software (BioTek). *E. coli* and *S. aureus* strains were transformed by electroporation using standard protocols [61] and transformants were selected on LB or Trypticase Soy Agar (TSA; Difco) plates, respectively, with the appropriate antibiotics.

DNA manipulations

Oligonucleotides used in this study were synthesized by Sigma-
Proligo and their sequences are listed in Table S1. *S. aureus*
chromosomal DNA was isolated using the MasterPure™ Gram-
positive DNA purification Kit (Epicentre Biotechnologies). Plasmid
DNA was isolated using a QIAprep Spin Miniprep kit (Qiagen) and
PCR fragments were purified using the Qiaquick PCR purification
kit (Qiagen). T4 DNA ligase and restriction enzymes (New England
Biolabs), PCR reagents and Pwo thermostable DNA polymerase
(Roche) were used according to the manufacturer's recommenda-
tions. Nucleotide sequencing of plasmid constructs was carried out
by Beckman Coulter Genomics.

Plasmid and mutant construction

Plasmid pCN51 carrying the *Pcad-cadC* promoter module (cadmium chloride-inducible promoter and the CadC repressor gene), was used for gene expression in *S. aureus* [26]. Plasmid pET28/16, a derivative of plasmid pET28a (Novagen), was used for protein overexpression and purification [62]. The pMAD allelic replacement vector [63] was used to generate the ΔlytM and ΔssaA mutant strains.

To uncouple gene expression from WalKR-dependent activa-
tion, DNA fragments corresponding to the coding sequences of
cell wall hydrolase genes with their associated ribosome-binding
sites were generated by PCR using *S. aureus* RN4220 chromosomal

DNA and the oligonucleotide pairs listed in Table S1, introducing BamHI and EcoRI sites at the 5' and 3' ends of the DNA fragments, respectively. The *lytM*\* (encoding the LytM H291A protein) and *ssaA*\* (encoding the SsaA C171S protein) mutant alleles were generated by site-directed mutagenesis using strand-overlap extension PCRs (SOE-PCR) [64]. Briefly, two DNA fragments overlapping at their ends were synthesized by PCR using oligonucleotide pairs OSA215/OSA259 and OSA260/OSA217 for *lytM*\* and OAD042/OSA265 and OSA266/OAD043 for *ssaA*\* and fused by SOE-PCR using the external oligonucleotides, (OSA215/OSA217 for *lytM*\*, OAD042/ OAD043 for *ssaA*\*) introducing BamHI and EcoRI sites at the 5' and 3' ends of the DNA fragments. The DNA fragments were cloned between the corresponding restriction sites of plasmid pCN51 in order to place gene transcription under the control of the Pcad promoter. The resulting plasmids are listed in Table 1. DNA ligations were transformed into *E. coli* DH5α™ and the nucleotide sequences of the DNA inserts were determined before introducing the plasmids into the *S. aureus* ST1000 strain.

For intracellular production of lysostaphin in *S. aureus*, a 897-bp DNA fragment encoding an active recombinant form of lysostaphin (*lys7*) was generated by PCR using plasmid pCXlss7 [33], and oligonucleotides OP274 and OSA252 (see Table S1), and cloned between the BamHI and EcoRI sites of the pCN51 vector, resulting in plasmid pAD08.

Mature LytM (without its signal peptide, residues 26 to 316) was
overproduced in *E. coli* using plasmid pAD05, constructed by
cloning a 887-bp PCR-generated NcoI/XhoI DNA fragment
corresponding to ‘*lytM* (oligonucleotide pair OAD027/OAD018)
between the corresponding restriction sites of plasmid pET28/16,
replacing the stop codon with an *XhoI* restriction site. This allows
the creation of a translational fusion adding six histidine residues
to the carboxy-terminus of the corresponding proteins, placing
expression of the genes under the control of a T7 bacteriophage
promoter.

For generation of a ΔlytM mutant strain, two DNA fragments, of 615 and 804 bp, were generated by PCR using oligonucleotides OAD003/OAD004 and OAD013/OAD014, respectively (see Table S1), corresponding to the DNA regions located immediately upstream and downstream from the *lytM* gene. These DNA fragments were cloned in tandem in two consecutive steps, between the *BamHI* and *BglII* restriction sites of the pMAD vector, resulting in plasmid pAD07. The plasmid was introduced by electroporation into *S. aureus* strain RN4220, and transformants were selected at 30°C on TSA plates containing erythromycin and 5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside (X-Gal) (100 µg/ml). Integration and excision of pAD07 were then performed as previously described [63], yielding strain ST1031 (ΔlytM mutant).

The same approach was used to obtain the Δ*ssaA* mutant. Oligonucleotides OAD064/OAD065 and OAD066/OAD067 were used to generate by PCR two DNA fragments of 559 bp and 607 bp respectively (Table S1). These fragments, corresponding to the upstream and downstream regions of the *ssaA* gene, were cloned between the *BamHI* and *NcoI* restrictions sites of the pMAD vector to construct pAD18. The plasmid was then introduced into *S. aureus* strain RN4220 or ST1031 (*ΔlytM*). Integration and excision of pAD18 were performed as previously described [63], yielding strains ST1158 (RN4220 *ΔssaA*) and ST1164 (*ΔlytM* *ΔssaA*). The markerless gene deletions were confirmed by PCR amplification.

Total RNA extraction

Strains were grown in TSB- CdCl2 (0.25 µM) +/- 1 mM
IPTG at 37°C, with aeration until OD₆₀₀ = 1. Cells were pelleted
by centrifugation (2 min, 20,800×g) and immediately frozen at

<figure><img src="image_19.png" /></figure>
−20°C. RNA extractions were then performed as previously described [65], followed by a DNase I treatment with the TURBO DNA-free reagent (Ambion, Austin, TX) in order to eliminate residual contaminating genomic DNA.

## cDNA synthesis and quantitative real time PCRs (qRT-PCR)

cDNA synthesis was carried out as previously described [14]. Oligonucleotides were designed with the BEACON Designer 4.02 software (Premier Biosoft International, Palo Alto, CA) in order to synthesize 100–200 bp amplicons (see Table S1). Quantitative real-time PCRs (qRT-PCRs), critical threshold cycles (CT) and n-fold changes in transcript levels were performed and determined as previously described using iQ™ SYBR Green Supermix (Bio-Rad, Hercules, CA) and normalized with respect to 16S rRNA whose levels did not vary under our experimental conditions [14].

## Protein overproduction and purification

The plasmid carrying the *LytM* gene (pAD05) was introduced into the BL21 λ DE3/pREP4 strain, allowing coproduction of the GroESL chaperonin in order to optimize recombinant protein solubility [66]. The resulting strains were grown in 2 liters of LB medium at room temperature, expression was induced during the mid-exponential growth phase by addition of 1 mM IPTG and incubation pursued for 4 hours. Protein purification by immobilized metal affinity chromatography (IMAC) was then performed as previously described [12]. Purified proteins were analyzed by SDS-PAGE on a 4–15% polyacrylamide gel (Bio-Rad, Hercules, CA).

## Western blotting experiments

Purified *LytM* was used to raise polyclonal rabbit antibodies, provided by Covalab (Villeurbanne, France) which were purified against *LytM* using the AminoLink® Kit from Thermo Scientific. Bacterial lysates were prepared from 45 ml cultures grown at 37°C with aeration in TSB + CdCl₂ 0.25 µM +/− 1 mM IPTG until approximately OD₆₀₀ nm = 1. Cells were harvested by centrifugation (10 min, 5200×g), resuspended in 500 µl of 2X Laemmli SDS sample buffer and heated for 5 min at 99°C [67]. After centrifugation (10 min, 20,800×g), 20 µl of SDS-soluble proteins or purified *LytM* (0.5 µg) were separated on a 12.5% SDS-PAGE gel, and transferred to a nitrocellulose membrane using a semi-dry blotter (Bio-Rad) and the following buffer: 25 mM Tris, 192 mM glycine, 20% ethanol. *LytM*, *LytM* H291A and purified *LytM* proteins were detected using the purified polyclonal rabbit antibodies (1:50,000), horseradish peroxidase-coupled anti-rabbit secondary antibodies (Zymed), and the Pico chemiluminescence Western blot kit (Pierce).

## References

1. Lowy FD (1998) *Staphylococcus aureus* infections. N Engl J Med 339: 520-532.

2. von Eiff C, Becker K, Machka K, Stammer H, Peters G (2001) Nasal carriage as a source of *Staphylococcus aureus* bacteremia. Study Group. N Engl J Med 344: 11-16.

3. Wertheim HF, Melles DC, Vos MC, van Leeuwen W, van Belkum A, et al. (2005) The role of nasal carriage in *Staphylococcus aureus* infections. Lancet Infect Dis 5: 751-762.

4. Kuroda M, Ohta T, Uchiyama I, Baba T, Yuzawa H, et al. (2001) Whole genome sequencing of meticillin-resistant *Staphylococcus aureus*. Lancet 357: 1225-1240.

5. Gillaspy AF, Worrell V, Orvis J, Roe BA, Dyer DW, et al. (2006) The *Staphylococcus aureus* NCTC 8325 Genome. In: Fischetti VA, Novick RP, Ferretti JJ, Portnoy DA, Rood JI, eds. Gram-Positive Pathogens. 2nd ed. Washington, DC: ASM Press. pp 381-412.

6. Novick RP (2003) Autoinduction and signal transduction in the regulation of staphylococcal virulence. Mol Microbiol 48: 1429-1449.

7. Gardete S, Wu SW, Gill S, Tomasz A (2006) Role of VraSR in antibiotic resistance and antibiotic-induced stress response in *Staphylococcus aureus*. Antimicrob Agents Chemother 50: 3424-3434.

## Viability testing

Bacteria were stained using the LIVE/DEAD BacLight™ viability assay L-7012 (Molecular Probes, Invitrogen, Carlsbad, CA). This stain distinguishes live cells from dead bacteria, based on membrane integrity and two nucleic acid stains. The green fluorochrome (SYTO 9) can penetrate intact membranes while the larger red fluorochrome (propidium iodide) only penetrates compromised membranes of dead bacteria. Bacterial cultures were grown in TSB CdCl₂ (0.25 µM) +/− IPTG (1 mM). At appropriate optical densities, cultures were washed 3 times in 0.9% NaCl and concentrated in 0.9% NaCl between 2 to 10 times depending on the initial optical densities. Staining and fluorescent microscopic observations were then carried out as specified by the manufacturer.

## Electron microscopy

For ultrastructural analyses, bacteria were grown in TSB +/− 1 mM IPTG and 0.25 µM CdCl₂ when indicated until OD₆₀₀ = 1. Bacteria were fixed overnight at 4°C with 2.5% glutaraldehyde in 0.1 M cacodylate buffer, pH 7.4. Fixed samples were treated with 1% osmium tetroxide in 0.1 M cacodylate buffer, pH 7.4, for 1 h, dehydrated in ethanol and embedded in epoxy resin. Ultrathin sections were stained with 2% uranyl acetate and lead citrate and examined using a Jeol JEM1010 transmission electron microscope (Jeol, Tokyo, Japan) at 80 kV and an Eloise MegaView III camera (Eloise SARL, Roissy, France).

## Supporting Information

<caption>**Table S1** Oligonucleotides used in this study. (DOC)</caption>

## Acknowledgments

We would like to thank Fritz Götz for the kind gift of plasmid pCXlss7 and many helpful suggestions, as well as Marie-Christine Prévost (Institut Pasteur, Ultrastructural Microscopy Platform), Françoise Baleux (Institut Pasteur, Chemistry of Biomolecules), Ivo Gomperts-Boneca and Simon J. Foster for helpful discussion.

## Author Contributions

Conceived and designed the experiments: AD SD TM. Performed the experiments: AD OP AM Y-MC SD. Analyzed the data: AD OP AM SD TM. Contributed reagents/materials/analysis tools: AD OP AM Y-MC SD. Wrote the paper: AD SD TM.

8. Herbert S, Bera A, Nerz C, Kraus D, Peschel A, et al. (2007) Molecular basis of resistance to muramidase and cationic antimicrobial peptide activity of lysozyme in staphylococci. PLoS Pathog 3: e102.

9. Meehl M, Herbert S, Gotz F, Cheung A (2007) Interaction of the GraRS two-component system with the VraFG ABC transporter to support vancomycin-intermediate resistance in *Staphylococcus aureus*. Antimicrob Agents Chemother 51: 2679-2689.

10. Pragman AA, Yarwood JM, Tripp TJ, Schlievert PM (2004) Characterization of virulence factor regulation by SrrAB, a two-component system in *Staphylococcus aureus*. J Bacteriol 186: 2430-2438.

11. Martin PK, Li T, Sun D, Biek DP, Schmid MB (1999) Role in cell permeability of an essential two-component system in *Staphylococcus aureus*. J Bacteriol 181: 3666-3673.

12. Dubrac S, Msadek T (2004) Identification of genes controlled by the essential YycG/YycF two-component system of *Staphylococcus aureus*. J Bacteriol 186: 1175-1181.

13. Dubrac S, Bisicchia P, Devine KM, Msadek T (2008) A matter of life and death: cell wall homeostasis and the WalKR (YycGF) essential signal transduction pathway. Mol Microbiol 70: 1307-1322.

<figure><img src="image_20.png" /></figure>
14. Dubrac S, Boneca IG, Poupel O, Msadek T (2007) New insights into the WalK/WalR (YycG/YycF) essential signal transduction pathway reveal a major role in controlling cell wall metabolism and biofilm formation in *Staphylococcus aureus*. *J Bacteriol* 189: 8257-8269.

15. Ulrich LE, Zhulin IB (2007) MiST: a microbial signal transduction database. *Nucleic Acids Res* 35: D386–390.

16. Vollmer W, Joris B, Charlier P, Foster S (2008) Bacterial peptidoglycan (murein) hydrolases. *FEMS Microbiol Rev* 32: 259–286.

17. Ng WL, Robertson GT, Kazmierczak KM, Zhao J, Gilmour R, et al. (2003) Constitutive expression of PcsB suppresses the requirement for the essential VirC (YycF) response regulator in *Streptococcus pneumoniae* R6. *Mol Microbiol* 50: 1647–1663.

18. Ng WL, Kazmierczak KM, Winkler ME (2004) Defective cell wall synthesis in *Streptococcus pneumoniae* R6 depleted for the essential PcsB putative murein hydrolase or the VirC (YycF) response regulator. *Mol Microbiol* 53: 1161–1175.

19. Bisicchia P, Noone D, Lioliou E, Howell A, Quigley S, et al. (2007) The essential YycFG two-component system controls cell wall metabolism in *Bacillus subtilis*. *Mol Microbiol* 65: 180–200.

20. Foster SJ (1995) Molecular characterization and functional analysis of the major autolysin of *Staphylococcus aureus* 8325/4. *J Bacteriol* 177: 5723-5725.

21. Kajimura J, Fujiwara T, Yamada S, Suzaki Y, Nishida T, et al. (2005) Identification and molecular characterization of an N-acetylmuramyl-L-alanine amidase Sle1 involved in cell separation of *Staphylococcus aureus*. *Mol Microbiol* 58: 1087–1101.

22. Sieradzki K, Tomasz A (2003) Alterations of cell wall structure and metabolism accompany reduced susceptibility to vancomycin in an isogenic series of clinical isolates of *Staphylococcus aureus*. *J Bacteriol* 185: 7103–7110.

23. Bateman A, Rawlings ND (2003) The CHAP domain: a large family of amidases including GSP amidase and peptidoglycan hydrolases. *Trends Biochem Sci* 28: 234–237.

24. Rigden DJ, Jedrzejes MJ, Galperin MY (2003) Amidase domains from bacterial and phage autolysins define a family of gamma-D,L-glutamate-specific amidohydrolases. *Trends Biochem Sci* 28: 230–234.

25. Oshida T, Sugai M, Komatsuzawa H, Hong YM, Suginaka H, et al. (1995) A *Staphylococcus aureus* autolysin that has an N-acetylmuramoyl-L-alanine amidase domain and an endo-beta-N-acetylglucosaminidase domain: cloning, sequence analysis, and characterization. *Proc Natl Acad Sci U S A* 92: 285–289.

26. Charpentier E, Anton AI, Barry P, Alfonso B, Fang Y, et al. (2004) Novel cassette-based shuttle vector system for gram-positive bacteria. *Appl Environ Microbiol* 70: 6076–6085.

27. Stapleton MR, Horsburgh MJ, Hayhurst EJ, Wright L, Jonsson IM, et al. (2007) Characterization of IsaA and SceD, two putative lytic transglycosylases of *Staphylococcus aureus*. *J Bacteriol* 189: 7316–7325.

28. Ramadurai L, Jayaswal RK (1997) Molecular cloning, sequencing, and expression of *lytM*, a unique autolytic gene of *Staphylococcus aureus*. *J Bacteriol* 179: 3625–3631.

29. Ramadurai L, Lockwood KJ, Nadakavukaren MJ, Jayaswal RK (1999) Characterization of a chromosomally encoded glycylglycine endopeptidase of *Staphylococcus aureus*. *Microbiology* 145(Pt 4): 801–808.

30. Odintsov SG, Sabala I, Marcyjaniak M, Bochtler M (2004) Latent LytM at 1.3A resolution. *J Mol Biol* 335: 775–785.

31. Bochtler M, Odintsov SG, Marcyjaniak M, Sabala I (2004) Similar active sites in lysostaphins and D-Ala-D-Ala metallopeptidases. *Protein Sci* 13: 854–861.

32. Mierau I, Leij P, van Swam I, Blommestein B, Floris E, et al. (2005) Industrial-scale production and purification of a heterologous protein in *Lactococcus lactis* using the nisin-controlled gene expression system NICE: the case of lysostaphin. *Microb Cell Fact* 4: 15.

33. Thumm G, Gotz F (1997) Studies on prolysostaphin processing and characterization of the lysostaphin immunity factor (Lif) of *Staphylococcus simulans* biovar *staphyolyticus*. *Mol Microbiol* 23: 1251–1265.

34. Layec S, Decaris B, Leblond-Bourget N (2008) Characterization of proteins belonging to the CHAP-related superfamily within the Firmicutes. *J Mol Microbiol Biotechnol* 14: 31–40.

35. Zou Y, Hou C (2010) Systematic analysis of an amidase domain CHAP in 12 *Staphylococcus aureus* genomes and 44 staphylococcal phage genomes. *Comput Biol Chem*.

36. Yokoi KJ, Kawahigashi N, Uchida M, Sugahara K, Shinohara M, et al. (2005) The two-component cell lysis genes *holWMT* and *lysWMT* of the *Staphylococcus warneri* M phage φWMY: cloning, sequencing, expression, and mutational analysis in *Escherichia coli*. *Gene* 351: 97–108.

37. Nelson D, Schuch R, Chahales P, Zhu S, Fischetti VA (2006) PlyC: a multimeric bacteriophage lysin. *Proc Natl Acad Sci U S A* 103: 10765–10770.

38. Firczuk M, Mucha A, Bochtler M (2005) Crystal structures of active LytM. *J Mol Biol* 354: 578–590.

39. Ling B, Berger-Bachi B (1998) Increased overall antibiotic susceptibility in *Staphylococcus aureus femAB* null mutants. *Antimicrob Agents Chemother* 42: 936–938.

40. Rohrer S, Ehlert K, Tschierske M, Labischinski H, Berger-Bachi B (1999) The essential *Staphylococcus aureus* gene *fmhB* is involved in the first step of peptidoglycan pentaglycine interpeptide formation. *Proc Natl Acad Sci U S A* 96: 9351–9356.

41. Tschierske M, Mori C, Rohrer S, Ehlert K, Shaw KJ, et al. (1999) Identification of three additional *femAB*-like open reading frames in *Staphylococcus aureus*. *FEMS Microbiol Lett* 171: 97–102.

42. Schneewind O, Fowler A, Faull KF (1995) Structure of the cell wall anchor of surface proteins in *Staphylococcus aureus*. *Science* 268: 103–106.

43. Maidhof H, Reinicke B, Blumel P, Berger-Bachi B, Labischinski H (1991). *femA*, which encodes a factor essential for expression of methicillin resistance, affects glycine content of peptidoglycan in methicillin-resistant and methicillin-susceptible *Staphylococcus aureus* strains. *J Bacteriol* 173: 3507–3513.

44. Lang S, Livesley MA, Lambert PA, Littler WA, Elliott TS (2000) Identification of a novel antigen from *Staphylococcus epidermidis*. *FEMS Immunol Med Microbiol* 29: 213–220.

45. Martin PK, Bao Y, Boyer E, Winterberg KM, McDowell L, et al. (2002) Novel locus required for expression of high-level macrolide-lincosamide-streptogramin B resistance in *Staphylococcus aureus*. *J Bacteriol* 184: 5810–5813.

46. Baba T, Schneewind O (1998) Targeting of muralytic enzymes to the cell division site of Gram-positive bacteria: repeat domains direct autolysin to the equatorial surface ring of *Staphylococcus aureus*. *Embo J* 17: 4639–4646.

47. Sugai M, Komatsuzawa H, Akiyama T, Hong YM, Oshida T, et al. (1995) Identification of endo-beta-N-acetylglucosaminidase and N-acetylmuramyl-L-alanine amidase as cluster-dispersing enzymes in *Staphylococcus aureus*. *J Bacteriol* 177: 1491–1496.

48. Yamada S, Sugai M, Komatsuzawa H, Nakashima S, Oshida T, et al. (1996) An autolysin ring associated with cell separation of *Staphylococcus aureus*. *J Bacteriol* 178: 1565–1571.

49. Chaudhuri RR, Allen AG, Owen PJ, Shalom G, Stone K, et al. (2009) Comprehensive identification of essential *Staphylococcus aureus* genes using Transposon-Mediated Differential Hybridisation (TMDH). *BMC Genomics* 10: 291.

50. Schindler CA, Schuhardt VT (1964) Lyostaphin: A new bacteriolytic agent for the *Staphylococcus aureus*. *Proc Natl Acad Sci U S A* 51: 414–421.

51. Francius G, Domenech O, Mingeot-Leclercq MP, Dufrene YF (2008) Direct observation of *Staphylococcus aureus* cell wall digestion by lysostaphin. *J Bacteriol* 190: 7904–7909.

52. Becker SC, Dong S, Baker JR, Foster-Frey J, Pritchard DG, et al. (2009) LysK CHAP endopeptidase domain is required for lysis of live staphylococcal cells. *FEMS Microbiol Lett* 294: 52–60.

53. Navarre WW, Ton-That H, Faull KF, Schneewind O (1999) Multiple enzymatic activities of the murein hydrolase from staphylococcal phage φhi1. Identification of a D-alanyl-glycine endopeptidase activity. *J Biol Chem* 274: 15847–15856.

54. Throup JP, Koretke KK, Bryant AP, Ingraham KA, Chalker AF, et al. (2000) A genomic analysis of two-component signal transduction in *Streptococcus pneumoniae*. *Mol Microbiol* 35: 566–576.

55. Wagner C, Saizieu Ad A, Schonfeld HJ, Kamber M, Lange R, et al. (2002) Genetic analysis and functional characterization of the *Streptococcus pneumoniae* vic operon. *Infect Immun* 70: 6121–6128.

56. Okada A, Igarashi M, Okajima T, Kinoshita N, Umerita M, et al. (2010) *Walkmycin B* targets *WalK* (YycG), a histidine kinase essential for bacterial cell growth. *J Antibiot* 63: 89–94.

57. Gotoh Y, Doi A, Furuta E, Dubrac S, Ishizaki Y, et al. (2010) Novel antibacterial compounds specifically targeting the essential WalR response regulator. *J Antibiot* 63: 127–134.

58. Burian M, Wolz C, Goerke C (2010) Regulatory adaptation of *Staphylococcus aureus* during nasal colonization of humans. *PLoS One* 5: e10040.

59. Burian M, Rautenberg M, Kohler T, Fritz M, Krismer B, et al. (2010) Temporal expression of adhesion factors and activity of global regulators during establishment of *Staphylococcus aureus* nasal colonization. *J Infect Dis* 201: 1414–1421.

60. Studier FW, Moffatt BA (1986) Use of bacteriophage T7 RNA polymerase to direct selective high-level expression of cloned genes. *J Mol Biol* 189: 113–130.

61. Sambrook J, Fritsch EF, Maniatis T (1989) Molecular cloning: a laboratory manual, second edition. Cold Spring Harbor, N. Y.: Cold Spring Harbor Laboratory.

62. Chastanet A, Fert J, Msadek T (2003) Comparative genomics reveal novel heat shock regulatory mechanisms in *Staphylococcus aureus* and other Gram-positive bacteria. *Mol Microbiol* 47: 1061–1073.

63. Arnaud M, Chastanet A, Debarbouille M (2004) New vector for efficient allelic replacement in naturally nontransformable, low-GC-content, gram-positive bacteria. *Appl Environ Microbiol* 70: 6887–6891.

64. Ho SN, Hunt HD, Horton RM, Pullen JK, Pease LR (1989) Site-directed mutagenesis by overlap extension using the polymerase chain reaction. *Gene* 77: 51–59.

65. Even S, Burguire P, Auger S, Soutourina O, Danchin A, et al. (2006) Global control of cysteine metabolism by CymR in *Bacillus subtilis*. *J Bacteriol* 188: 2184–2197.

66. Amrein KE, Takacs B, Stieger M, Molnos J, Flint NA, et al. (1995) Purification and characterization of recombinant human p50csk protein-tyrosine kinase from an *Escherichia coli* expression system overproducing the bacterial chaperones GroES and GroEL. *Proc Natl Acad Sci U S A* 92: 1048–1052.

67. Laemmli UK (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. *Nature* 227: 680–685.

68. Kreiswirth BN, Lofdahl S, Betley MJ, O'Reilly M, Schlievert PM, et al. (1983) The toxic shock syndrome exotoxin structural gene is not detectably transmitted by a prophage. *Nature* 305: 709–712.

69. Chastanet A, Msadek T (2003) ClpP of *Streptococcus salivarius* is a novel member of the dually regulated class of stress response genes in gram-positive bacteria. *J Bacteriol* 185: 683–687.

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