In order to identify factors that Gram-positive bacteria use to conceal their PGN present at the bacterial cell surface from host recognition, we have constructed S. aureus null mutants lacking non-essential genes involved in PGN metabolism and determined the ability of a fluorescent derivative of PGRP-SA (mCherry_PGRP-SA) to bind to their surfaces (Figure 1). Specifically, we tested mutants constructed in NCTC8325-4 S. aureus strain, expressing altered levels of autolysins (ΔarlR), lacking the major autolysin (Δatl) or factors that modulate autolysin activity (ΔfmtA), producing non-O-acetylated peptidoglycan with increased susceptibility to lysozyme (Δoat), or producing altered WTAs lacking attached β-O-GlcNAc (ΔtarS) or D-alanyl residues (ΔdltA). A mutant lacking the major autolysin Atl, a PGN hydrolase, was identified as the most severely impaired in its ability to avoid recognition by mCherry_PGRP-SA.

Figure 1

Download asset Open asset

PGN modifications, namely the attachment of teichoic acids or the degree of PGN polymerization (defined as the ratio between the amount of polymerized muropeptides and the amount of monomeric muropeptides present in the PGN macromolecule), have been previously associated with changes in the recognition of S. aureus PGN by host receptors (Filipe et al., 2005; Atilano et al., 2011). However, an S. aureus atl null mutant produces a PGN with a muropeptide composition similar to the parental strain (Figure 2A), that is, it shows no increase in the amount of polymerized muropeptides, which were previously reported to be better inducers of an innate immune response than monomeric muropeptides (Filipe et al., 2005). Moreover, the S. aureus atl null mutant produces WTAs (Figure 2B), indicating that the mechanism to conceal PGN dependent on the presence of atl is new, and different from that previously reported for a WTA mutant (Atilano et al., 2011). In agreement, the simultaneous deletion of tagO and atl had a synergistic effect on exposing the surface PGN to host recognition (Figure 2C).

Figure 2

Download asset Open asset

The atl gene encodes a polypeptide that is cleaved into two proteins capable of digesting PGN: an amidase (AM), which cleaves the amide bond between peptides and glycans, and a glucosaminidase (GL), which cleaves glycan strands (Figure 3A). To determine which one of these activities was required for the role of Atl in PGN concealment, we constructed S. aureus strains expressing, from the atl native chromosomal locus, Atl mutants with impaired amidase activity (AM^H265A [Bose et al., 2012]), glucosaminidase activity (GL^E1128A [Bose et al., 2012]), or both (Atl^H265A/E1128A double mutant) and confirmed the absence of the expected hydrolytic activity in a zymogram (Figure 3B).

Figure 3

Download asset Open asset

When the atl mutants were incubated with mCherry-PGRP-SA, both the mutant having only glucosaminidase activity (AM^H265A mutant) and the mutant having only amidase activity (GL^E1128A mutant) were still able to avoid strong PGRP-SA binding, indicating that either amidase or glucosaminidase enzymatic activity was sufficient to impair recognition of S. aureus by PGRP-SA (Figure 3C). Only when both activities were absent (Atl^H265A/E1128A double mutant) was mCherry-PGRP-SA capable of easily recognizing the bacterial cell surface (Figure 3C,D), albeit to lower levels than those observed for the atl null mutant. This difference may be due to residual amidase or glucosaminidase activity in the S. aureus Atl^H265A/E1128A double mutant, not detectable in a zymogram.

The proposed role for the amidase and glucosaminidase activities in concealing bacteria was confirmed using purified enzymes. Pre-incubation of S. aureus atl null mutant cells with either AM or GL (approximately 0.4 μM) completely abolished mCherry_PGRP-SA binding to the bacterial surface (Figure 4A). When both enzymes were added simultaneously to the atl null mutant cells, lower concentrations of each enzyme (<0.02 μM) were sufficient to conceal bacteria from the host immune receptor (Figure 4B). This is in agreement with the synergy in PGRP-SA binding observed for the S. aureus Atl^H265A/E1128 double mutant (Figure 2C).

Figure 4

Download asset Open asset

We then questioned if the mechanism used by autolysins to conceal PGN at the bacterial surface was dependent on their lytic activity. An alternative hypothesis could be that amidases, or other proteins capable of binding PGN, prevented binding of PGRP-SA by competing for the same PGN substrate, given that the structural fold of PGRPs closely resembles the fold of amidases (Zoll et al., 2010). To distinguish between the two hypotheses, we first incubated S. aureus atl null mutant cells with purified AM or with GL, and then washed them thoroughly to ensure complete removal of the proteins (Figure 5). This procedure did not increase recognition by PGRP-SA, showing that the activity of the autolysins, and not their physical presence, was required for protection.

Figure 5

Download asset Open asset

The results described above show that amidase or glucosaminidase activities can efficiently protect S. aureus cells from direct recognition by the Drosophila PGN receptor PGRP-SA. Does this protection confer an advantage to S. aureus cells during infection in the Drosophila model? Indeed, Tabuchi et al. have previously shown that S. aureus atl mutants are impaired in their virulence (Tabuchi et al., 2010). We have confirmed that the atl mutant used in this work, when compared to the parental S. aureus strain NCTC8325-4, was also severely impaired in its ability to kill wild type Drosophila, both the 25174 strain, recently established from wild caught flies (Mackay et al., 2012), as well as the yw strain, which is the parental lineage used to generate the PGRP seml mutant (Michel et al., 2001; Figure 6A,B). Furthermore, the atl mutant triggered a stronger induction of drosomycin expression (frequently used as a read-out for Toll activity) when normalized for the same number of infecting bacteria (Figure 6C,D). Importantly, our data showed that survival of Drosophila infected with an S. aureus atl mutant was dependent upon the presence of a functional PGRP-SA (Figure 6E). Furthermore, these mutant bacteria were capable of propagating similarly to the parental S. aureus strain in flies that lacked a functional PGRP-SA (Figure 6F), showing that impaired virulence is not due to a lower proliferation rate of the atl mutant.

Figure 6

Download asset Open asset

Two, non-mutually exclusive, hypotheses can explain the decreased virulence of the atl mutant. (1) S. aureus wild type bacteria produce amidases that cleave the stem peptide moiety of soluble PGN fragments. Given that stem peptides are required for binding of PGRPs to PGN, amidase activity would therefore reduce the inflammatory activity of the soluble PGN. In contrast, the atl mutant would release intact muropeptides, easily detected by PGRP-SA, and therefore would be unable to evade detection by the innate immune system. (2) Amidases produced by S. aureus wild type bacteria would shave and remove the most accessible PGN fragments at the surface of bacteria, eliminating the binding sites for PGRPs. As the atl mutant does not produce the Atl amidase, it would have extending PGN fragments at its surface, which would be easily detected by PGRP-SA, inducing an immune response.

If the first hypothesis is correct, inactivation of the amidase activity, but not of the glucosaminidase (which cleaves the glycans and therefore releases intact, inflammatory PGN fragments), should result in decreased virulence. Alternatively, if the second hypothesis is mainly correct, inactivation of either the amidase or the glucosaminidase activity should result in decreased virulence, as both activities can remove fragments of detectable PGN (containing peptides) from the bacterial surface. We therefore tested the virulence of bacteria with either impaired amidase activity (AM^H265A) or impaired glucosaminidase activity (GL^E1128A) and observed that both had a reduced ability to kill flies (Figure 7A) in accordance with the second hypothesis. Moreover, injection of seml Drosophila with atl mutant bacteria pre-coated with mCherry_PGRP-SA resulted in increased fly survival (Figure 7B), again favoring a role for PGRP-SA in the recognition of PGN directly at the bacterial surface. Nevertheless, the amount of PGRP-SA present at the surface of the injected bacteria may not have been sufficient to result in full complementation of the ability of seml flies to survive infection. This is in accordance with the hypothesis that a continuous and high expression of PGRP-SA during the entire infection process is required to ensure survival of the infected host (De Gregorio et al., 2001).

Figure 7

Download asset Open asset

Secreted autolysins not only bind to bacterial cell surfaces but they can also be found in the growth medium (Pasztor et al., 2010). This raises an interesting hypothesis: can secreted bacterial autolysins protect neighboring bacteria? If this was true, autolysin producers could in theory protect an entire bacterial population from recognition by the host. To test this hypothesis we collected and filtered the supernatant from a culture of S. aureus parental strain NCTC8325-4 (Atl producer) and incubated S. aureus atl null mutant cells in this medium for 30 min. The atl-encoded proteins secreted into the supernatant by the parental strain were capable of modifying the surface of the atl null mutant cells, consequently abolishing their recognition by PGRP-SA (Figure 8A,B). This was not observed when supernatant from a culture of S. aureus atl null mutant was used (Figure 8A,B), showing that atl products (and not other secreted PGN hydrolases) are responsible for PGN concealment.

Figure 8

Download asset Open asset

Importantly, we observed that atl mutants regained the ability to kill flies if pre-treated with a supernatant from a culture of the Atl producer NCTC8325-4 but not if pre-treated with supernatant from a culture of the atl mutant (Figure 8C).

Moreover, previously bound PGRP-SA could be removed by autolysin activity present in the medium, as seen by time-lapse microscopy of S. aureus atl null mutant cells initially covered by bound mCherry_PGRP-SA and then incubated with supernatant containing amidase and glucosaminidase enzymes (Figure 9).

Figure 9

Download asset Open asset

Video 1

Download asset

Video 2

Download asset

Together, these data suggest that the autolysins encoded by atl can protect not only the producer cells but also a population of invading bacteria from detection by the immune system during infection, enabling it to kill the infected host.

The S. aureus strain used in this work, NCTC8325-4, is susceptible to most antibiotics and has limited interest from a clinical point of view. We therefore tested if the role of atl-encoded enzymes in making PGN inaccessible to the host was conserved in other strains, namely in methicillin-resistant S. aureus (MRSA) strains. MRSA strains are a leading cause of bacterial infections in hospitals and an important cause of community-acquired (CA) bacterial infections in the United States. MW2 is a particularly virulent MRSA strain that caused the earliest reported cases of CA-MRSA infection in the United States (Chambers and Deleo, 2009). Deletion of the atl gene in MW2 led to higher binding of mCherry_PGRP-SA, that is, to improved recognition of the bacterial surface (Figure 10A). More importantly, MW2Δatl cells were severely impaired in virulence, as demonstrated by the decreased capacity to kill Drosophila (Figure 10B).

Figure 10

Download asset Open asset

The fact that PGN sensing is an evolutionarily conserved recognition mechanism in innate immunity, suggests that the strategy used by S. aureus to conceal PGN from the host could be effective across different animal phyla and even used by other bacteria. We therefore tested if we could observe this phenomenon in another Gram-positive pathogenic bacteria, namely Streptococcus pneumoniae, a frequent cause of pneumonia in developed countries and a major cause of infant mortality by septicemia in developing countries (Kadioglu et al., 2008). Lack of LytA, the major amidase of S. pneumoniae, resulted in bacteria that were better recognized by mCherry_PGRP-SA, a phenotype that was reversed when LytA was expressed from a plasmid (Figure 11A). Interestingly, it is possible that autolysins can participate in inter-species protection, as S. pneumoniae LytA, an amidase that releases PGN stem peptides, and Streptomyces globisporus mutanolysin, a commercially available muramidase that releases muropeptides, were capable of protecting S. aureus atl null mutant cells from PGRP-SA recognition (Figure 11B).

Figure 11

Download asset Open asset

These results suggest a general role for autolysins, such as amidases, in protecting bacteria from host recognition.

All bacterial strains used in this study are listed in Supplementary file 1. Escherichia coli strains were grown in Luria–Bertani broth (LB; Difco, France) or Luria–Bertani agar (LA; Difco) medium at 37°C with aeration. When needed, the antibiotics ampicillin (Amp; Sigma-Aldrich, Germany) and erythromycin (Ery; Sigma-Aldrich) were added at a final concentration of 100 µg/ml, and kanamycin (Km; Sigma-Aldrich) was added at a final concentration of 30 µg/ml. Lactococcus lactis LL108 strain was grown in M17 broth (Difco), supplemented with sucrose (0.5 M) and glucose (0.5% wt/vol) at 30°C without aeration. Erythromycin was used when required at 100 µg/ml. S. aureus strains were grown at 30°C with aeration in tryptic soy broth (TSB; Difco) or on tryptic soy agar (TSA; Difco). Medium was supplemented when required with Ery at 10 µg/ml and/or 5-bromo-4-chloro-3-indolyl β-D-galactopyranoside (X-Gal; Apollo Scientific, UK) at 100 µg/ml. S. pneumoniae was grown in C + Y medium at 37°C, without aeration, or on TSA plates supplemented with 5% vol/vol sheep blood (Probiológica, Portugal). Tetracycline (Sigma-Aldrich) and Ery were used, when required, at 1 µg/ml and 0.25 µg/ml, respectively.

Isogenic Drosophila Bloomington #25174 and #6599 (yw) were used as wild type flies. PGRP_SA^seml flies were used as a PGRP-SA mutant background (Michel et al., 2001). Stocks were raised on standard cornmeal-agar medium at 25°C.

All plasmids used in this study are listed in Supplementary file 1 and the sequences of the primers used are listed in Supplementary file 2.

The S. aureus oatA and fmtA null mutants were constructed using the integrative vector pORI280 (Leenhouts et al., 1996). To delete the oatA gene from the genome of S. aureus NCTC8325-4 we amplified two 1.3 Kb DNA fragments from NCTC8325-4 chromosomal DNA, corresponding to the upstream (primers P1_oatA and P2_oatA) and downstream (primers P3_oatA and P4_oatA) regions of the oatA gene. The upstream fragment was digested with BamHI and NcoI and cloned into pORI280. The downstream PCR fragment was digested with NcoI and BglII and then cloned into the previous plasmid, which already contained the upstream fragment, producing the plasmid pΔoatA. To delete the fmtA gene, we amplified two DNA fragments of approximately 0.9 Kb from S. aureus NCTC8325-4 chromosomal DNA, corresponding to the upstream (primers P5_fmtA and P6_fmtA) and downstream (primers P7_fmtA and P8_fmtA) regions of the fmtA gene. These fragments were joined by overlap PCR using primers P5_fmtA and P8_fmtA. The resulting PCR product was digested with BamHI and BglII, and cloned in the pORI280 plasmid, producing the plasmid pΔfmtA. Plasmids pΔoatA and pΔfmtA were sequenced and electroporated into the transformable RN4220 strain at 30°C (using Ery selection) as previously described (Veiga and Pinho, 2009). The plasmids were then transduced to NCTC8325-4 using the phage 80α. The resulting strains, carrying the plasmids integrated into the chromosome, were grown in liquid media, without antibiotic and at 37°C, for several generations to allow for loss of the plasmid and, consequently, of the lacZ and erm genes. Different dilutions of the cultures were plated on TSA plates containing X-Gal (without Ery) and incubated at 37°C overnight. White colonies were isolated and their sensitivity to erythromycin was confirmed. Absence of the oatA and fmtA genes (NCTCΔoatA and NCTCΔfmtA) was shown by PCR and confirmed by sequence analysis of the amplified DNA fragment.

All other S. aureus null mutants described were constructed using the thermosensitive vector pMAD (Arnaud et al., 2004). In order to generate the arlR null mutant, 0.8 Kb PCR fragments of the upstream and downstream regions of the arlR gene were amplified from chromosomal DNA of NCTC8325-4, using primer pairs P9_arlR/P10_arlR and P11_arlR/P12_arlR. The two PCR products were joined in an overlap PCR reaction using the primers P9_arlR/P12_arlR. The resulting product was digested with NcoI and BglII and cloned into pMAD, giving rise to the pΔarlR plasmid.

To delete the atl gene, the pΔatl plasmid was constructed by amplifying 1 Kb fragments from the downstream and upstream regions of the atl gene, using the primer pairs P13_atl/P14_atl and P15_atl/P16_atl. These two fragments were joined by overlap PCR using the primers P13_atl and P16_atl, and the obtained PCR product was digested with BglII and EcoRI and cloned into the pMAD vector, giving rise to the pΔatl plasmid.

To delete the tarS gene, we amplified by PCR two DNA fragments of approximately 0.6 Kb, corresponding to the upstream (primers P17_tarS/P18_tarS) and downstream (primers P19_tarS/P20_tarS) regions of the tarS gene. The two fragments were joined by overlap PCR, using primers P17_tarS and P20_tarS. The resulting PCR product was digested with BglII and NcoI and cloned into pMAD vector, producing the plasmid pΔtarS.

To delete the dltA gene, two 0.55 Kb DNA fragments were amplified by PCR from the genome of S. aureus NCTC8325-4, corresponding to the upstream (primers P25_dltA/P26_dltA) and downstream (primers P27_dltA/P28_dltA) regions of the dltA gene. The two fragments were joined by overlap PCR using primers P25_dltA and P28_dltA, and the resulting PCR product was digested with BglII and EcoRI and cloned into the pMAD vector, producing the plasmid pΔdltA.

The pΔarlR, pΔatl, pΔtarS, and pΔdltA plasmids were sequenced and electroporated into S. aureus RN4220 strain at 30°C (using Ery selection) and then transduced to NCTC8325-4 using phage 80α. Insertion and excision of the plasmids for deletion of the genes (arlR, atl, tarS, and dltA) from the NCTC8325-4 chromosome was completed as previously described (Arnaud et al., 2004), with the exception of the NCTCΔdltA strain where the last step was performed at 30°C due to the thermosensitive nature of cells lacking the dltA gene. All gene deletions were confirmed by PCR and the resulting strains were named NCTCΔarlR, NCTCΔatl, NCTCΔtarS, and NCTCΔdltA.

To make a NCTCΔatlΔtagO double mutant, the pΔtagO plasmid (Atilano et al., 2010) was transduced into the NCTCΔatl strain using phage 80α, at 30°C due to the thermosensitive nature of cells lacking the tagO gene. The double mutant was identified among the white colonies sensitive to Ery, by PCR and DNA sequencing, as described above.

The point mutants in the Atl amidase (AM) and Atl glucosaminidase (GL) domains were constructed by site-directed mutagenesis. To inactivate the AM domain, the conserved amino acid H265 was exchanged for an alanine (H265A mutation). To inactivate the GL domain, the amino acid residue E1128 was mutated to an alanine (E1128A mutation). To generate the H265A mutation, a 794 bp region upstream from the codon encoding H265 (primers P29_AtlAM and P30_AtlAM) and a 1070 bp region downstream from the codon encoding H265 (primers P31_AtlAM and P32_AtlAM) were amplified from NCTC8325-4 genomic DNA by PCR. Joining of the up and downstream regions by overlap PCR (primers P29_AtlAM and P32_AtlAM) resulted in the amplification of the DNA fragment encoding the mutated Atl amidase domain, which was cloned into the pMAD vector, using BamHI and EcoRI restriction enzymes, to construct plasmid pAM^H265A. A similar approach was used to generate the Atl point mutation in the Atl glucosaminidase (GL) domain. Briefly, a 659 bp fragment upstream of the codon encoding E1128 (primers P33_AtlGL and P34_AtlGL) and a 680 bp fragment downstream of the codon encoding E1128 (primers P35_AtlGL and P36_AtlGL) were amplified by PCR. The two fragments were joined by overlap PCR using primers P33_AtlGL and P36_AtlGL, digested and cloned into the EcoRI/BamHI sites of the pMAD vector, yielding plasmid pGL^E1128A. After sequencing, plasmids pAM^H265A and pGL^E1128A were electroporated into RN4220 and then transduced into NCTC8325-4. Insertion into and excision from the chromosome generated S. aureus point mutants in the amidase and glucosaminidase domains. The strains carrying the Atl mutations were verified by sequencing and were named AM^H265A and GL^E1128A. A double mutant, named Atl^H265A/E1128A, mutated in both the amidase and glucosaminidase domains, was constructed by phage transduction of plasmid pGL¹¹²⁸ into the AM^H265A strain, followed by insertion into and excision from the chromosome, as described above.

The S. pneumoniae lytA null mutant strain was constructed in the background of R36A strain, using the integrative vector pORI280. To delete the lytA gene, we amplified two 1.0 Kb DNA fragments from R36A chromosomal DNA, corresponding to the upstream (primers P41_LytA and P42_LytA) and downstream (primers P43_LytA and P44_LytA) regions of the lytA gene. The two fragments were then joined by overlap PCR using primers P41_LytA and P44_LytA. The PCR product was digested with BamHI and EcoRI and cloned into plasmid pORI280, producing the plasmid pΔlytA. This plasmid was propagated in L. lactis and purified before being transformed into the unencapsulated S. pneumoniae R36A strain. The R36AΔlytA strain was obtained by insertion and excision of the plasmid into the chromosome, as previously described (Leenhouts et al., 1996). For the construction of a plasmid expressing LytA (plytA), the lytA gene from S. pneumoniae R36A genome was amplified with primers P45_LytA and P46_LytA, digested with NheI and BglII, and cloned into plasmid pBCSLF001 (Henriques et al., 2011).

A 3177 bp fragment of the atl gene, missing the pro-peptide sequence, was amplified by PCR using genomic DNA of S. aureus strain COL with primers Pexp1 and Pexp4, which included BamHI and SalI restriction sites. The DNA fragment was purified, digested with the respective restriction enzymes, and cloned into pET28a(+) plasmid (Novagen, Merck Millipore, UK) using E. coli DH5α, yielding pET-AMR₁R₂R₃GL.

To construct a plasmid expressing the AM recombinant protein, plasmid pET-AMR₁R₂R₃GL was used as the template for site-directed mutagenesis using primers Pexp_stop1 and Pexp_stop2, to change codon 775 from AAA (Lys) to TAA (stop codon) resulting in pET-AMR₁R₂.

To construct a plasmid expressing the GL recombinant protein, primers Pexp2 and Pexp4 with BamHI and SalI restriction sites, were used to amplify a 1446 bp sequence encoding for the GL domain of Atl, using COL genomic DNA as the template. The DNA fragment was purified, digested with the respective restriction enzymes, and cloned into pET28a(+) plasmid using E. coli DH5α, yielding pET-R₃GL.

To construct a plasmid expressing protein AM^H265A, pET-AMR₁R₂ was used as the template for site-directed mutagenesis using primers Pexp_H265A1 and Pexp_H265A2, to change codon 265 from GTA (His) to GCA (Ala). A plasmid expressing protein GL^E1128A was constructed similarly, using pET-R₃GL as the template for directed mutagenesis with primers Pexp_E1228A1 and Pexp_E1228A2, changing codon GAA (Glu) to GCA (Ala). The amplified plasmids were digested with DpnI prior to transformation into E. coli DH5α, resulting in pET-AM^H265A and pET-GL^E1228A.

The recombinant plasmids (Supplementary file 1) were confirmed by restriction analysis and sequencing and used to transform E. coli BL21(DE3) strain for protein expression.

A functional mCherry-tagged derivative of PGRP-SA protein, mCherry-PGRP-SA, was expressed in E. coli and purified using a cobalt affinity resin (Clontech) as previously described (Atilano et al., 2011).

The AM and GL proteins were expressed as N-terminal 6×His-tag fusion proteins. Expression of the recombinant proteins was performed using an auto-induction based expression method. Cells were grown for 18 h at 37°C with shaking, harvested, and resuspended in 1/10 vol of purification lysis buffer (50 mM NaH₂PO₄, 10 mM imidazole, 300 mM NaCl, pH 8.0) containing 10 U/ml of benzonase nuclease (Novagen, Merck Millipore) and Complete-Mini Protease Inhibitor Cocktail (Roche, Switzerland). After cell disruption, the lysates were cleared. The recombinant proteins were soluble and therefore present in the supernatant.

Protein purification was performed using Ni-NTA agarose columns (Qiagen, Germany) under native conditions, according to the manufacturer’s instructions. The recombinant proteins were eluted with purification elution buffer (50 mM NaH₂PO₄, 250 mM imidazole, 300 mM NaCl, pH 8.0). The expression and purification yields were monitored by SDS–PAGE. The most concentrated elution fractions were dialyzed for 16 h in a 3500 MWCO SnakeSkin (Pierce Biotechnology, Rockford, IL) at 4°C, against 100 mM Tris, pH 7.5.

For the purification of S. pneumoniae LytA amidase, E. coli BL21(DE3) cells were transformed with the plasmid pGL100 (Garcia et al., 1987) and incubated overnight at 37°C, with vigorous shaking, in LB supplemented with 100 μg/ml ampicillin and 2% lactose. Cells were harvested by centrifugation, resuspended in 20 mM sodium phosphate buffer, pH 6.9, and broken by sonication. Clarified lysate was applied to DEAE-cellulose resin and incubated at 4°C for 1 h with stirring. Bound protein was washed five times with 20 mM sodium phosphate buffer containing 1.5 M NaCl, then eluted in the same buffer containing 2% choline. Protein was dialyzed against 20 mM sodium phosphate buffer, pH 6.9 to remove the choline and aliquots were stored at −20°C.

To quantify mCherry_PGRP-SA binding to bacterial cells, cultures were grown to mid-exponential phase (OD_{600 nm} 0.5), centrifuged, and resuspended in TSB to an OD of 2.5.

Approximately 100 µl of each culture was placed into wells of a 96-well plate and harvested at room temperature (RT) for 5 min at 3600×g. Cells were washed and then resuspended in 100 µl of sterile PBS (pH 6). mCherry_PGRP-SA was added to each well at a final concentration of 60 µg/ml, followed by incubation for 5 min at RT. Cells were then harvested and washed with PBS. The fluorescent signal of the mCherry_PGRP-SA was detected using a 532 nm laser in a Fuji FLA-5100 reader (FUJIFILM Life Science) and the fluorescence intensity of each well was determined using ImageJ software (Abràmoff et al., 2004).

To image the mCherry_PGRP-SA binding to bacterial cell surface by fluorescence microscopy, cells were treated as previously described (Atilano et al., 2011), except that mCherry_PGRP-SA was added to a final concentration of ≈300 µg/ml, in a reaction volume of 200 µl. After labeling, bacterial cells were centrifuged, washed twice with 200 µl of PBS, resuspended in 20 µl of PBS, and observed by fluorescence microscopy.

Crude autolytic enzyme extracts were prepared from S. aureus cultures grown in TSB to OD₆₀₀ 0.3 at 30°C. The cells were harvested by centrifugation (7000 rpm, 15 min at 4°C), and the cell pellet was washed in 15 ml of ice-cold 50 mM Tris–HCl (pH 7.5), 150 mM NaCl. Cells were resuspended in SDS 4% (500 ml) and incubated for 30 min at 25°C with stirring. Zymographic analysis of the proteins’ peptidoglycan hydrolytic activities was performed as described earlier (Heilmann et al., 1997). Briefly, crude cell lysates or pure proteins isolated as described above were separated by SDS–PAGE on a polyacrylamide gel (10% acrylamide/0.2% bisacrylamide) containing inactivated Micrococcus luteus or S. aureus cells (recovered at mid-exponential phase, washed with water, and heat inactivated by autoclaving at 121°C for 15 min) as a substrate in the resolving gel. Equal amounts of crude extracts (50 μg) or purified proteins (200 ng), quantified in a Nanodrop ND-1000 spectrophotometer (Thermo Scientific, Wilmington, NC), were loaded in the gel. After electrophoresis, gels were washed in deionized H₂O for 15 min at RT, and then incubated in renaturation buffer (0.1% Triton X-100, 10 mM CaCl₂, 10 mM MgCl₂, 50 mM Tris–HCl, pH 7.5) at 37°C with gentle agitation. The zymogram was stained in methylene blue solution (1% methylene blue in KOH 0.01%) for 3 min and destained in water until bands with peptidoglycan hydrolytic activity were observed as clear zones in the opaque gel.

WTAs were extracted by alkaline hydrolysis from overnight cultures, analyzed by native PAGE, and visualized by combined alcian blue/silver staining, as previously described (Atilano et al., 2010).

Mutanolysin (Sigma)-digested peptidoglycan was prepared from NCTC8325-4 and NCTCΔatl S. aureus strains as previously described (Filipe et al., 2005). The released muropeptides were reduced with sodium borohydride and separated by HPLC on a C₁₈ column (ODS-Hypersyl, 5 μm, 4.6×250 mm; Thermo Scientific) at a flow rate of 0.5 ml/min for 160 min with a gradient from 5% to 30% (vol/vol) methanol in 100 mM NaH₂PO₄, pH 2.0. The eluted muropeptides were detected by their UV absorption at 206 nm, and their abundance was estimated by calculating the percentage of the integrated area of each peak relative to the total area of the muropeptide peaks, using Shimadzu LC Solution software.

Cell-free supernatants from the S. aureus NCTCΔatl mutant and its parental strain NCTC8325-4 were obtained by harvesting overnight cultures at 10,000×g for 10 min at 4°C, and filtering the supernatants through a 0.22 µm filter (Merck Millipore). Sterility was confirmed by plating the supernatants on TSA plates at 30°C. To image the effect of the secreted Atl products on the PGRP-SA binding, exponentially growing NCTCΔatl and NCTC8325-4 cells (500 µl at OD 0.5) were incubated with the previously isolated cell-free supernatants (500 μl) for 30 min at 30°C. Cells were then washed twice with cold PBS, incubated with mCherry_PGRP-SA, and imaged by fluorescence microscopy.

To quantify mCherry_PGRP-SA binding to the parental and the atl null mutant S. aureus cells pre-treated with secreted Atl products, the cultures were grown to mid-exponential phase (OD ∼0.5), centrifuged, and resuspended in TSB to an OD of ∼2.5. A 100 µl volume of each culture was placed into the wells of a 96-well plate and harvested at RT, 5 min at 3600×g. Cells were washed once with 100 µl of PBS and then resuspended in 100 µl of cell-free supernatants (prepared as described above). After 30 min of incubation at 30°C, the cells were washed twice with 100 µl of PBS. Quantification of mCherry_PGRP-SA binding was done as described above.

AM (15.6 and 390.1 nM), GL (18.7 and 467.4 nM), AM plus GL (15.6 and 18.7 nM, respectively), LytA (3.5 nM), and mutanolysin (108.7 nM) were added to 200 µl of PBS-washed NCTCΔatl cells collected during exponential growth (500 µl at OD 0.5). After 30 min incubation at 30°C, cells were washed twice with cold PBS, incubated with mCherry_PGRP-SA as described before, and imaged by fluorescent microscopy.

To determine the relative amount of AM and GL still associated with the bacterial cells following the treatment with these two enzymes, cells were treated with AM and GL as described above and washed three times with PBS (control pellets without the enzymes, without washing, or without cells were used). Samples were serially diluted 1:2 to concentrations ranging from 6 µg/ml (undiluted) to 0.37 µg/ml (lowest dilution), and 2 µl were spotted onto a polyvinylidene difluoride (PVDF) membrane (GE Healthcare Life Sciences) and immunostained with specific anti-His mouse antibodies. Washed, unwashed, and control bacterial cells were also labeled with mCherry_PGRP-SA as described above and observed by fluorescence microscopy.

Bacterial cells samples (2 µl) incubated with fluorescent tagged mCherry_PGRP-SA were placed on a thin layer of 1% agarose (Bio-Rad) in PBS. For time-lapse microscopy experiments, 2 µl of NCTCΔatl culture, pre-labeled with mCherry-PGRP-SA, were placed on a thin layer of 0.6% agarose containing filtered supernatants of either NCTC8325-4 or NCTCΔatl cells. After 2.5 min, phase contrast and fluorescence images were taken every 30 s for 20.5 min and a final image was acquired after 50 min of incubation. Imaging was performed using a Leica DM 6000B microscope equipped with an iXon^EM + EMCCD camera (Andor Technologies), using Metamorph software (Meta Imaging series 7.5). Images were analyzed using ImageJ software (Abràmoff et al., 2004).

Overnight cultures of bacteria (10 ml) were washed and resuspended in PBS so that their final OD₆₀₀ was 0.350. Healthy looking adult female flies (n = 25), 2–4 days old, were injected in the thorax with 32 nl of a bacterial cell suspension (approximately 100 CFUs) or with PBS, using a nanoinjector (Nanoject II; Drummond Scientific, Broomall, PA), and fly survival was determined as previously described (Atilano et al., 2011). In the fly survival assay for the pathogenicity rescue of atl null mutants, 100 μl of each bacterial cell suspension were incubated with cell-free supernatants of wild type or Δatl mutants (prepared as described above) for 30 min at 30°C. Treated bacterial cells were washed twice, resuspended in an equal volume of PBS and further diluted so that approximately 100 CFUs would be injected per fly. Each experiment was performed in triplicate, on different days. Following injection, flies were kept at 30°C and survival assessed every 12 h over a period of 3 days. Since the trends in survival were the same for each sample of a triplicate (i.e., survival curves were positioned similarly, relative to one another), the three independent biological repeats were combined (n = 75) and estimates of survival curves were plotted.

Fly survival was also tested using PGRP-SA pre-coated S. aureus cells. For that purpose, overnight NCTCΔatl bacterial cells were washed and resuspended in PBS (200 μl; OD₆₀₀ 0.350) and pre-coated with mCherry_PGRP-SA at 300 μg/ml for 5 min at 4°C. Cells were washed once and resuspended in an equal volume of ice-cold PBS. PBS suspensions with non-coated atl cells or containing only mCherry_PGRP-SA (300 μg/ml) were used as experiment controls. Controls and samples were further diluted in PBS (1:250) and used to infect PGRP-SA deficient flies as described above.

The bacterial load per insect during infection was assessed as previously described (Atilano et al., 2011). Six female flies were homogenized in 1000 μl TSB with a micropestle device (VWR, Radnor, PA) until pieces of tissue were no longer visible. Homogenate samples (50 μl) underwent a 10-fold dilution series from 1× to 1/100,000× before being spread onto TSB agar plates and onto mannitol salt agar. The number of CFUs was scored after 24 h at 30°C. Triplicate assays were performed.

Total RNA was extracted from homogenized female flies (n = 6) using the Total RNA Purification Plus Kit (Norgen, Canada) according to the manufacturers’ instructions. A Maxima First Strand cDNA Synthesis Kit (Thermo Scientific) was used to produce cDNA. The total RNA (500 ng) was mixed with 5× reaction mix buffer (4 μl), Maxima enzyme mix (2 μl), and RNase-free dH₂O to a final volume of 20 μl. cDNA was synthesized in a T100 Thermal Cycler (Bio-Rad) and stored at −20°C. Drosomycin (FlyBase annotation symbol: CG10810) and tbp (FlyBase annotation symbol: CG9874) expression levels were measured using primer pairs: Drs (+) and Drs (−); Tbp (+) and Tbp (−). The housekeeping gene tbp (Matta et al., 2011) was used as a control to normalize the expression of the Drs gene. qPCR reactions were performed using the SensiFast SYBR No-ROX Kit (Bioline, UK) as outlined in the manufacturer’s instructions, in a Rotor-Gene Q real-time PCR cycler with a 72-well rotor (Qiagen). Negative controls had an equivalent amount of total RNA or no cDNA template. Three biological repeats were performed per time point. Gene expression was calculated on the basis of the comparative threshold cycle (C_T) value (Schmittgen and Livak, 2008). Levels of gene expression in all groups were shown as a ratio to the control group value.

Data for the mCherry_PGRP-SA binding to bacteria assays (n = 10–50) was non-normal but with equal variance, and therefore a non-parametric Kruskal–Wallis test followed by Dunn’s multiple comparison was applied. Estimated survival curves were constructed from the raw data sets and the log-rank (Mantel–Cox) test used to determine statistical significance between the curves. For clarity, 95% confidence intervals have been omitted from the graphs.

When the bacterial CFU numbers in fly survival assays did not present a normal distribution (Lilliefors test, p<0.05), log10 transformation was used. Repeated measures two-way ANOVA was used to analyze significant differences over time and between bacterial strains. Bonferroni post-tests were used to locate the time points where mean values were statistically separable between the two bacterial strains.

Analysis for qPCR data showed that the data followed a normal distribution (Lilliefors test, p>0.1) and had equal variance (Levene’s test, p>0.1). Thus, mean values were plotted with 95% confidence and repeated measures two-way ANOVA was used to look for significant differences between bacterial wild type strain and atl null mutant over time. Bonferroni post-tests were used to locate the time points where mean values were statistically separable between the two bacterial strains.

All data were plotted and analyzed using GraphPad Prism 6 (GraphPad Software).

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

This study was funded by Fundação para a Ciência e Tecnologia (FCT), Lisbon, Portugal, through research grants PTDC/SAU-IMU/111806/2009 and PTDC/BIA-MIC/111817/2009 (SRF), PTDC/BIA-BCM/099152/2008 (MGP), PTDC/BIA-MIC/101375/2008 (RGS) and PEst-OE/EQB/LA0004/2011. Wellcome Trust funded grant WT087680 (PL). ERC funded grant ERC-2012-StG-310987 (MGP). MLA, PMP, FV, MJC, PR, and IRG were supported by FCT fellowships SFRH/BD/28440/2006, SFRH/BD/41119/2007, SFRH/BD/78748/2011, SFRH/BPD/77758/2011, SFRH/BPD/23812/2005, and SFRH/BD/70162/2010, respectively.

© 2014, Atilano et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.