Present address: Division of Dermatology, University of California at San Diego, 3350 La Jolla Village Drive, San Diego, CA 92161, USA.
The antimicrobial peptide-sensing system aps of Staphylococcus aureus
Article first published online: 24 OCT 2007
Volume 66, Issue 5, pages 1136–1147, December 2007
How to Cite
Li, M., Cha, D. J., Lai, Y., Villaruz, A. E., Sturdevant, D. E. and Otto, M. (2007), The antimicrobial peptide-sensing system aps of Staphylococcus aureus. Molecular Microbiology, 66: 1136–1147. doi: 10.1111/j.1365-2958.2007.05986.x
- Issue published online: 24 OCT 2007
- Article first published online: 24 OCT 2007
- Accepted 30 September, 2007.
Staphylococcus aureus is a leading cause of hospital-associated and, more recently, community-associated infections caused by highly virulent methicillin-resistant strains (CA-MRSA). S. aureus survival in the human host is largely defined by the ability to evade attacks by antimicrobial peptides (AMPs) and other mechanisms of innate host defence. Here we show that AMPs induce resistance mechanisms in CA-MRSA via the aps AMP sensor/regulator system, including (i) the d-alanylation of teichoic acids, (ii) the incorporation of lysyl-phosphatidylglycerol in the bacterial membrane and a concomitant increase in lysine biosynthesis, and (iii) putative AMP transport systems such as the vraFG transporter, for which we demonstrate a function in AMP resistance. In contrast to the aps system of S. epidermidis, induction of the aps response in S. aureus was AMP-selective due to structural differences in the AMP binding loop of the ApsS sensor protein. Finally, using a murine infection model, we demonstrate the importance of the aps regulatory system in S. aureus infection. This study shows that while significant interspecies differences exist in the AMP–aps interaction, the AMP sensor system aps is functional and efficient in promoting resistance to a variety of AMPs in a clinically relevant strain of the important human pathogen S. aureus.
Staphylococcus aureus is a dangerous human pathogen that may cause severe diseases such as bacteraemia, toxic shock syndrome, scalded skin syndrome and endocarditis (Lowy, 1998). Traditionally, infections with S. aureus occurred exclusively in hospitalized and predisposed, e.g. immunocompromised, patients. However hyper-virulent strains of S. aureus have recently emerged that are resistant to antibiotics and may infect healthy adults outside the hospital setting (community-associated methicillin-resistant S. aureus, CA-MRSA), thus posing an even more severe threat to the public health system (Chambers, 2005). These strains are now by far the most common cause of skin and soft tissue infections reporting to the emergency department in the USA (Moran et al., 2006).
While the role of acquired immunity in prolonged staphylococcal infection is not well understood, the innate immune system clearly plays a major part in determining the outcome of an S. aureus infection (Komatsuzawa et al., 2006). Neutrophils (or polymorphonuclear leucocytes) form the central part of innate host defence, and their key role in combating S. aureus can be seen, for example in the significantly impaired defence to S. aureus infection in patients with chronic granulomatous disease (Lekstrom-Himes and Gallin, 2000). Neutrophils ingest invading bacteria and subsequently kill them by the combined activity of reactive oxygen species and antimicrobial proteins and peptides (Faurschou and Borregaard, 2003). Antimicrobial peptides (AMPs) form an evolutionarily conserved mechanism of innate host defence and are found in many groups of organisms, including amphibians, insects and other invertebrates. More recent research indicates that human AMPs have a much more important role in preventing infection than previously suspected (Hancock and Diamond, 2000). Notably, AMPs are also secreted by human epithelia and thus play a key role not only in eliminating invading S. aureus, but also in controlling colonization on the human skin and mucosal surfaces (Harder and Schroder, 2005).
The importance of AMPs in defending the human body from bacterial infection is reflected by the many mechanisms of resistance that human pathogenic bacteria have developed to cope with AMPs (Peschel and Sahl, 2006). These mechanisms include: alterations of the bacterial cell surface charge, which is aimed to decrease binding of the commonly cationic AMPs; proteolytic inactivation by secreted exoproteases; and AMP transporters that remove the AMPs from the bacterial cell and membrane.
Most likely because the constitutive production of molecules involved in resistance mechanisms represents a significant energetic burden to the bacteria, their expression is limited to times when AMPs are present. To that end, bacteria have regulatory mechanisms that sense the presence of AMPs and control the expression of resistance genes. In Gram-negative bacteria, a two-component system named PhoP/PhoQ senses cationic AMPs and, for example, controls the incorporation of aminoarabinose in lipid A, thereby preventing attachment of cationic AMPs (Bader et al., 2005). Very recently, we have discovered that the corresponding AMP sensing and gene regulatory task in the Gram-positive pathogen Staphylococcus epidermidis is performed by an unrelated three-component sensor/regulator, composed of a classical two-component histidine kinase/response regulator (ApsR, ApsS) and a third protein of unknown function (ApsX) (Li et al., 2007). The presence of similar genetic loci in other Gram-positive bacteria, including S. aureus, suggested that the aps system represents a widespread mechanism for AMP sensing and the regulation of AMP-resistance mechanisms. Therefore, here we investigated the role of the aps system in the induction of AMP-resistance mechanisms in S. aureus. We provide evidence for aps functionality in this important human pathogen in vitro and in vivo and describe significant differences compared with the prototype system of S. epidermidis.
Induction of AMP-resistance mechanisms in S. aureus MW2 is AMP-selective
To test whether AMPs induce resistance mechanisms in S. aureus, we used the interaction between human beta defensin 3 (hBD3) (Harder et al., 2001) and strain MW2, a prototype community-associated MRSA strain (Anonymous, 1999), as example. The AMP hBD3 was selected because it is a key component of innate host defence on human epithelia and a strong inducer of AMP-resistance mechanisms in S. epidermidis (Harder and Schroder, 2005; Li et al., 2007). We determined, by quantitative reverse transcription polymerase chain reaction (RT-PCR), the induction of three genes, dltB, vraF and mprF, representing the three main AMP-resistance mechanisms found to be under hBD3-inducible control in S. epidermidis (Li et al., 2007). The dlt operon is responsible for the d-alanylation of teichoic acids, a mechanism aimed to decrease the overall negative charge of the bacterial cell surface and thus the attraction of cationic AMPs (Peschel et al., 1999; Peschel, 2002); the MprF enzyme catalyses the lysinylation of phosphatidylglycerol, preventing the binding of cationic AMPs to the cytoplasmic membrane (Peschel et al., 2001). The vraFG genes code for a transporter also likely involved in AMP resistance (Li et al., 2007). Surprisingly, we did not detect induction of the dltB gene with hBD3, even when we increased the concentration to 100 μg ml−1, corresponding to 10 times that previously used for S. epidermidis (Fig. 1A). Further, there was no significant increase of mprF expression with increasing concentrations of hBD3, and significantly increased expression of vraF only at the high concentrations of 50 and 100 μg ml−1 hBD3. To rule out that this lack of induction was due to proteolytic degradation of hBD3 in our test system, we determined the stability of hBD3 under the tested conditions with S. aureus MW2. However, no differential induction of dltB with and without hBD3 was found even within short incubation times, at which no considerable degradation of hBD3 was observed (Fig. 1B and C). These findings indicated that hBD3 is a much weaker inducer of the aps response in S. aureus MW2 compared with S. epidermidis.
Prompted by the severely impaired induction of the dltB, vraF and mprF genes with hBD3 in S. aureus compared with S. epidermidis, we determined dltB expression under the influence of a series of AMPs at 2 μmol l−1 (which corresponds to ∼10 μg ml−1 for hBD3) (Fig. 1D). In addition to hBD3, the uncharged gramicidin and the anionic dermcidin and, notably, the cationic peptides histadin and magainin II failed to induce dltB expression. Among the peptides that we tested, only indolicidin, melittin, nisin and the human AMP LL-37 induced a significant increase in dltB expression. These results demonstrate that induction of the aps system in S. aureus MW2 is different from S. epidermidis and much more selective than described for that bacterium (Li et al., 2007).
Cationic AMPs induce AMP-resistance mechanisms and a concomitant metabolic adaptation in S. aureus MW2
To determine the gene regulatory response to cationic AMPs in S. aureus MW2, we selected the AMP indolicidin, an AMP of the cathelicidin family, because it showed the most significant influence on the expression of the dltB gene and in contrast to melittin, which also led to strong dltB induction, is of mammalian origin (Selsted et al., 1992) (Fig. 1D). We used whole-genome microarrays specific for the MW2 genome and analysed changes in gene expression with and without addition of 5 μg ml−1 (∼2 μMol l−1) indolicidin during mid-logarithmic growth (Table 1).
|Gene number||Gene product name||Wild-type with/without indolicidin||apsS with/without indolicidin||Wild-type/apsS (without indolicidin)||Wild-type/apsS (with indolicidin)|
|MW0814||d-alanylation of teichoic acids|
|d-alanine-activating enzyme (DltA)||2.16a||1.15||0.86||1.61|
|MW0816||d-alanyl carrier protein (DltC)||3.03||1.14||0.99||2.33|
|MW0817||Protein DltD precursor||2.73||1.21||0.85||1.92|
|Lysinylation of phosphatidylglycerol|
|MW1247||Lysyltransferase (MprF or FmtC)||4.90||1.28||1.28||5.00|
|Lysine biosynthesis (from aspartate)|
|MW1282||Aspartate-semialdehyde dehydrogenase (Asd)||2.40||1.76||1.27||1.72|
|MW1283||Dihydrodipicolinate synthase (DapA)||2.30||1.86||1.20||1.49|
|MW1284||Dihydrodipicolinate reductase (DapB)||2.35||1.84||1.20||1.54|
|MW1285||Tetrahydrodipicolinate N-acetyltransferase (DapD)||2.45||1.75||1.06||1.49|
|ABC transporter (VraFG)|
|MW0623||Bacitracin transport ATP binding protein VraF||14.17||1.06||1.85||25.00|
|MW0624||ABC transporter permease protein (VraG)||13.51||1.02||1.45||20.00|
|VraRS regulator and adjacent transporter|
|MW1824||Two-component response regulator VraR||2.37||5.38||0.86||0.38|
|MW1825||Two-component sensor protein VraS||2.88||5.92||0.89||0.43|
|MW1826||Transporter (similar to B. subtilis YvqF)||4.10||10.34||0.93||0.37|
|MW2620||Bacitracin transport ATP binding protein VraD||9.84||3.76||1.18||3.13|
|MW2621||ABC transporter permease protein VraE||12.02||5.32||1.16||2.63|
|MW1872||ABC transporter ATP binding protein||3.43||15.46||0.55||0.12|
|MW1874||ABC transporter ATP binding protein||4.14||26.81||0.68||0.10|
|MW1875||Transcriptional regulator, GntR family||3.24||23.21||0.71||0.10|
The most pronounced changes under the influence of indolicidin comprised the upregulation of established mechanisms of AMP resistance. All genes in the dlt operon were upregulated about 3-fold. The mprF gene was upregulated about 5-fold. In addition, increased activity of the MprF enzyme was confirmed on the translational level: the relative portion of lysyl-phosphatidylglycerol (L-PG) in phospholipids was significantly higher, and that of phosphatidylglycerol accordingly significantly lower, upon addition of indolicidin to cells of S. aureus MW2 (Fig. 2A). Furthermore, we observed upregulation of the transcription of genes encoding the biosynthesis of lysine from aspartic acid via the diaminopimelic acid pathway (Fig. 2B). This metabolic change is most likely caused by the increased requirement for lysine due to its increased incorporation in phospholipids by the MprF enzyme.
Further regulatory changes upon addition of indolicidin included the increased transcription of four transporter systems that may be involved in AMP resistance. The vraFG ABC transporter genes, which are situated next to the apsRSX regulatory system, and the vraDE ABC transporter genes, were strongly upregulated (about 10- to 15-fold). Two other ABC transporters systems were upregulated to a lower degree. The microarray results were confirmed by quantitative RT-PCR experiments with the dltB, vraF and mprF genes (Fig. 3). Taken together, these findings demonstrate that the gene regulatory response to the cationic AMP indolicidin in S. aureus comprises the upregulation of known and putative AMP-resistance mechanisms and, simultaneously, a metabolic adaptation to respond to the increased need for structural components of molecules involved in AMP resistance.
The vraFG transporter is involved in AMP resistance
The VraDE and VraFG transporters are similar to bacitracin-resistance transporters, underpinning their likely involvement in the resistance to AMPs. Specifically, our previous and current data on aps control in S. epidermidis and S. aureus suggest that the VraFG transport system contributes to AMP resistance. To investigate this hypothesis, we produced an isogenic deletion mutant of the vraFG genes in strain MW2 and determined minimal inhibitory concentration (MIC) values to a series of AMPs (Table 2). Our results indicate an important role of the vraFG transporter in promoting resistance to cationic AMPs, but not the anionic dermcidin.
|MW2||apsS||apsS (pTXapsS)||vraFG||vraFG (pRBvraFG)|
The S. aureus MW2 aps system controls AMP resistance
To investigate whether the aps system of S. aureus is involved in mediating the gene regulatory response to cationic AMPs and determining altered resistance to AMPs, we constructed an isogenic deletion mutant of the apsS sensor in the CA-MRSA strain MW2. Using a variety of cationic AMPs and the anionic AMP dermcidin, MIC values (Table 2) and survival of the wild-type and isogenic apsS mutant strains in killing assays were determined (data not shown). We found significantly reduced resistance of the apsS mutant strain compared with the wild-type strain towards all cationic peptides, but not the anionic dermcidin, indicating that the aps system of S. aureus MW2 is involved exclusively in regulating resistance to cationic AMPs. Of note, we observed aps-dependent killing with all tested cationic AMPs, including hBD3, demonstrating that induction of and susceptibility to AMP-resistance mechanisms are clearly distinguished features of the AMP–bacteria interaction, inasmuch as the spectrum of AMPs to which aps-controlled target mechanisms provide resistance is broader than that of aps inducers.
Most of the AMP-induced gene regulatory response is mediated by aps
To determine the role of the aps system in the induction of gene regulatory changes in S. aureus to indolicidin and define the S. aureus aps regulon, we performed whole-genome microarray analysis with the MW2 apsS deletion and wild-type strain with or without addition of indolicidin (Table 1). As a main result of these experiments, we were able to distinguish two sets of cationic AMP (indolicidin)-regulated genes in S. aureus: those under aps control and those under the control of a yet-undefined regulatory mechanism. The first group contains the dlt genes, mprF, vraFG, and the lysine biosynthesis operon. Analysis of phospholipids by thin-layer chromatography with the apsS mutant compared with the isogenic S. aureus MW2 wild-type strain confirmed that the MprF-catalysed incorporation of L-PG is regulated by aps (Fig. 2A). The second group contains the aforementioned ABC transporter operons. The ABC transporter vraDE genes appear to be under the control of both aps and an unidentified regulator. Further, the MW1871–1875 ABC transporter operon is under positive control by cationic AMPs via an unknown mechanism, but under negative control by aps. Microarray results were confirmed by quantitative RT-PCR of wild-type and apsS mutant, and apsS-complemented and control strains (Fig. 3).
Differences in induction by cationic AMPs between S. aureus and S. epidermidis are due to structural changes in the ApsS sensor
The aps system is composed of three components, ApsS, ApsR and ApsX (Li et al., 2007). The protein that interacts with AMPs is ApsS. Specifically, a single, 9-amino-acid extracellular loop of ApsS with a high density of negative charges is responsible for AMP binding and induction. Interestingly, the amino acid composition of this loop differs significantly between S. aureus and S. epidermidis. Whereas the overall similarity between the ApsS proteins of S. aureus and S. epidermidis is 70%, this loop only shows 33% similarity (Fig. 4A). For example, a serine residue found in the ApsS extracellular loop of S. epidermidis is replaced by a proline in S. aureus. As proline residues cause bends in polypeptides, this probably leads to significant changes in the structure of the S. aureus versus the S. epidermidis AMP binding loop. To confirm the hypothesis that the differences in induction of dltB expression between S. aureus and S. epidermidis are due to differences in ApsS, we complemented the apsS mutant of S. aureus MW2 with apsS of S. epidermidis and determined expression under the influence of different concentrations of hBD3. The S. aureus apsS mutant with heterologously expressed S. epidermidis apsS showed increased dltB expression upon addition of hBD3 (Fig. 4B) similar to the S. epidermidis wild type (Li et al., 2007), demonstrating that ApsS is responsible for the differences in induction seen between S. aureus and S. epidermidis. In a control experiment, complementation with the S. aureus apsS gene did not result in significant changes in hBD3 inducibility (data not shown). Moreover, to pinpoint the differences in induction to the AMP interaction loop of ApsS, we constructed a plasmid that expressed a hybrid protein consisting of the ApsS protein of S. aureus with the extracellular loop of S. epidermidis. This plasmid restored apsS inducibility by hBD3 in the MW2 apsS mutant (Fig. 4B), demonstrating that the differences in AMP induction are due to the structural changes compared with S. epidermidis in this part of the S. aureus ApsS protein.
The aps system is important for S. aureus virulence in vivo
To investigate whether control of AMP-resistance mechanisms by the aps system is important for S. aureus survival during infection, we compared the wild-type and apsS-deletion strains in a peritoneal infection model. Progression of disease was measured by determining the bacterial load in the kidneys after death. We detected a significantly reduced infectivity of the apsS mutant strain (Fig. 5), indicating that the aps system contributes to pathogen survival during S. aureus infection.
Sensing AMPs and reacting with effective countermeasures is important for the survival especially of commensal microorganisms living on human mucosal and epithelial surfaces, where AMPs represent a key mechanism of host defence (Harder and Schroder, 2005). Therefore, AMP sensors that control the expression of target AMP-resistance mechanisms, such as the Gram-negative PhoP/PhoQ system discovered in Salmonella typhimurium (Miller et al., 1989; Bader et al., 2005), and the Gram-positive aps system discovered in S. epidermidis (Li et al., 2007), have been proposed as targets for antimicrobial drug development due to their pivotal role in host–pathogen interaction (Brodsky and Gunn, 2005). An aps homologue of S. aureus has been described previously to be important for vancomycin resistance (Kuroda et al., 2000), partially via control of the vraFG transporter genes (Meehl et al., 2007). The system was named graRS, failing to recognize its three-component nature. In addition, a recent study has investigated the graRS regulon in S. aureus strain SA113 (Herbert et al., 2007). However, the detailed in vitro and in vivo roles of aps in Gram-positive bacteria other than the relatively innocuous S. epidermidis have not been analysed. Therefore, we determined here whether and to what extent aps controls AMP resistance and the expression of resistance mechanisms in S. aureus, specifically in the hyper-virulent community-associated MRSA strain MW2, and whether control by aps is crucial for pathogen success during infection.
First, we investigated whether AMP-resistance mechanisms in S. aureus are inducible by AMPs. To our surprise, we found that common AMP-resistance mechanisms, which are under inducible control in S. epidermidis (Li et al., 2007), were not inducible by the important human epithelial AMP hBD3 in S. aureus, or only to a very limited degree. Subsequently, we analysed a series of peptides and found that AMPs can be grouped into inducers and non-inducers of AMP-resistance mechanisms in S. aureus. There was no clear correlation of inducer function with the intensity of positive charge or secondary structure type. Rather, we found that functionality of an AMP as an inducer is determined by the ApsS extracellular AMP-sensing loop in a molecular interaction that certainly warrants further investigation.
The AMP-induced gene regulatory response in S. aureus was similar to that detected in S. epidermidis (Li et al., 2007), but included additional ABC transporter systems potentially involved in the removal of AMPs from the cell. Most AMP-inducible AMP-resistance mechanisms, including the dlt, mprF and vraFG genes, were found to be under aps control. However, the relative degree of upregulation was different compared with S. epidermidis. While the dlt operon was the most strongly upregulated target in S. epidermidis (Li et al., 2007), S. aureus MW2 controlled expression of the mprF and, especially, vraFG genes in response to AMPs in a more pronounced way, indicating significant differences in the aps regulons of the two species. Furthermore, the transcriptional profiling experiments revealed the presence of at least one additional yet-undefined mechanism of AMP sensing that regulates a series of ABC transporters probably involved in AMP resistance, including VraDE. Moreover, upregulation of lysine biosynthesis via the diaminopimelic acid pathway, the most common way of producing lysine in bacteria (Patte, 1994), indicated that the MprF-catalysed production of L-PG is accompanied by a metabolic shift to respond to the altered lysine requirements. The strong influence of aps in strain MW2 on the expression of mprF contrasts findings by Herbert et al. (2007), who did not discover regulation of mprF by aps (graRS) in S. aureus strain SA113. Thus, there appear to be strain-specific differences in the aps regulon among S. aureus strains. These may be linked to the known deficiency of strain SA113 in the important global regulator agr (Vuong et al., 2000) and the interaction of aps (graRS) with a series of global regulators described in that study (Herbert et al., 2007).
Initially described as important for resistance to the antibiotic vancomycin (Kuroda et al., 2000), the role of the VraFG transporter in AMP resistance has remained speculative. We demonstrate here for the first time that the VraFG transporter is involved in promoting resistance to AMPs. Interestingly, VraFG only conferred resistance to cationic AMPs, similar to the other main AMP-resistance mechanisms regulated by aps, thus underpinning the specificity of the aps gene regulatory response and target mechanisms for cationic AMPs.
Finally, we demonstrate efficiency of the S. aureus MW2 aps system in facilitating resistance to a variety of cationic AMPs and pathogen survival during infection. Notably, the aps system mediated increased protection also from those peptides that were not, or only weak inducers of the system, indicating that induction and susceptibility to resistance mechanisms are unrelated features of the AMP–pathogen interaction. Taken together, while our study indicates significant interspecies differences in AMP induction and regulation of target mechanisms, it shows that the aps system is functional in S. aureus and particularly CA-MRSA, and has a crucial role during infection, thus establishing physiological importance in a key Gram-positive pathogen.
Bacterial strains, growth conditions and chemicals
The bacteria and plasmids used are listed in Table 3. Escherichia coli DH5α and S. aureus RN4220 were used for cloning experiments. RN4220 was also used as a gateway strain prior to propagation of plasmids into target S. aureus strains. Bacteria were routinely grown in tryptic soy broth (TSB, Oxoid) medium with 0.25% glucose broth or agar plates at 37°C, unless otherwise indicated. Antibiotics were used at the following concentrations: ampicillin, 100 μg ml−1; chloramphenicol, 10 μg ml−1; and tetracycline, 12.5 μg ml−1. For microarray experiments, overnight cultures were diluted 1:100 into 50 ml of TSB and incubated at 37°C with shaking at 180 r.p.m. until grown to an OD600 of 2.5–3.0 (mid-logarithmic growth phase). Cultures were then split into two equal volumes, and cells were washed twice in 10 mM sodium phosphate buffer (pH 6.5) with 100 mM NaCl and resuspended in the same buffer. To one sample, the AMP indolicidin was added to a final concentration of 5 μg ml−1. Samples were incubated at 37°C for 1 h. Dermcidin (in its active, processed form, DCD-1 l) and hBD3 were synthesized by the Peptide Synthesis Unit, Research Technologies Branch, NIAID. Gramicidin D and nisin were purchased from Sigma. All other AMPs were purchased from GenScript Corporation.
|Strains/plasmids||Relevant genotype and property||Source/reference|
|RN4220||Derived from NCTC8325-4; r- m+||Kreiswirth et al. (1983)|
|MW2||CA-MRSA standard wild-type strain, clinical isolate||Anonymous (1999)|
|MW2 apsS||apsS gene-deletion mutant of MW2||This study|
|MW2 vraFG||vraFG gene-deletion mutant of MW2||This study|
|DH5α||endA1 recA1 gyrA96 thi-1 hsdR17(rK- mK+) relA1 supE44 (lacZYA-argF)U169 F-φ80 dlacZ M15 deoR phoAλ-||Commercial (Invitrogen)|
|pKOR1||cmR and ampR, temperature-sensitive vector for allelic replacement via lambda recombination and ccdB selection||Bae and Schneewind (2006)|
|pKOR1apsS||Vector for allelic replacement of apsS||This study|
|pKOR1vraFG||Vector for allelic replacement of vraFG||This study|
|pTX15||A staphylococcal expression vector that harbours a xylose-inducible promoter||Peschel et al. (1996)|
|pTX16||Derived from pTX15 by deletion of the lipase gene downstream of the xylose-inducible promoter.||Peschel et al. (1996)|
|pTXapsS||Derived from pTX15 by insertion of the MW2 apsS gene between the BamHI and MluI sites in place of the lipase gene fragment||This study|
|pTXapsS(Se loop)||Derived from pTX15 by insertion of the hybrid apsS gene between the BamHI and MluI sites||This study|
|pT181mcs||pT181 (Khan et al., 1981) with multiple cloning site of pUC19 (PvuII fragment) cloned in NdeI site||Seeber et al. (1990)|
|pTapsS(Se)||pT181mcs with S. epidermidis apsS gene (under control of its natural promoter)||Li et al. (2007)|
|pRB473||E. coli/Staphylococcus shuttle cloning plasmid, cmR and ampR||Bruckner (1997)|
|pRBvraFG||Derived from pRB473 by insertion of the MW2 vraF and vraG genes||This study|
Total RNA was isolated using an RNeasy Mini Kit (Qiagen) as recommended in a standard protocol. In brief, cell pellets were washed with RNase-free water, resuspended in 700 μl of RLT buffer supplemented with,-mercaptoethanol (10 μl β-mercaptoethanol per 1 ml RLT). The bacterial suspension was transferred to a 2 ml FastPrep lysing tube (Q-BioGene). The cells were lysed in a Bio101 high-speed homogenizer (Savant Instruments), at the following setting: speed, 6.0; time, 20 s. The lysate was incubated on ice for 5 min and centrifuged at 15 000 r.p.m. at 4°C for 15 min. The supernatant was collected and diluted with 500 μl of 100% ethanol. Samples were mixed and transferred to an RNeasy mini column. RNA isolation was performed according to the manufacturer's instructions. Remaining DNA was removed using RNase-free DNase I (Amersham Biosciences). Removal of contaminant DNA was confirmed by PCR. The reaction product was cleaned up with an RNeasy mini column. cDNA was synthesized and labelled according to the manufacturer's suggestions for Affymetrix antisense genome arrays (Affymetrix) as described (Li et al., 2007). A gel shift assay with NeutrAvidin (Pierce Biotechnology) was performed to estimate the labelling efficiency based on the instructions from Affymetrix. Biotinylated S. aureus cDNA was hybridized to custom Affymetrix GeneChips (RMLChip 3) with 96% coverage of genes from MW2 (2534 probe sets of 2632 ORFs) and scanned according to standard GeneChip protocols (Affymetrix). Each experiment was replicated at least three times. Affymetrix GeneChip Operating Software (GCOS v1.4, http://www.affymetrix.com) was used to perform the preliminary analysis of the custom chips at the probe-set level. Subsequent data analysis was performed as described (Li et al., 2007). To be included in the final gene list, gene expression must have been changed at least 2-fold for one of the treatments. The complete set of microarray data was deposited in NCBIs Gene Expression Omnibus (GEO, http://www.ncbi.nlm.nih.gov/geo/) and is accessible through GEO Series accession number GSE8400.
Construction of isogenic apsS, vraFG-deletion mutant strains and complementation plasmids
To delete the apsS or vraFG genes in S. aureus MW2, ∼1 kb DNA fragments, upstream and downstream of apsS or vraFG respectively, were PCR amplified from the chromosomal DNA of S. aureus MW2 using the primers listed in Table 4. PCR products were digested with BamHI, and then mixed together and ligated by T4 DNA ligase (Roche). The ligation product with attB sites at the two ends was used for recombination with plasmid pKOR1, yielding plasmid pKOR1apsS or pKOR1vraFG respectively. The resulting plasmid was transferred via electroporation first to S. aureus RN4220 and then to S. aureus MW2. Allelic replacement mutations were selected as described by Bae and Schneewind (2006). The proper integration was verified by analytical PCR and sequencing of the genomic DNA at the borders of the PCR-derived regions. For genetic complementation of the apsS mutant, the apsS gene was PCR amplified from MW2 genomic DNA using the primers listed in Table 4 and cloned into the BamHI and MluI sites of BamHI/MluI-digested pTX15. For heterologous complementation with the S. epidermidis apsS gene, the previously described plasmid pTapsS (S. epidermidis) was used (Li et al., 2007), in which S. epidermidis apsS is cloned in the plasmid pT181mcs with its natural promoter. For construction of the apsS hybrid expression plasmid, the primer pairs p1–p2 and p3–p4 (Table 4) were used to generate 238 and 1003 bp PCR fragments respectively from MW2 genomic DNA, resulting in the incorporation of 27 bp corresponding to the S. epidermidis apsS extracellular loop. Purified PCR products of p1–p2 and p3–p4 were hybridized by overlap extension PCR using primer pair p5–p6 and cloned into the BamHI and MluI sites of BamHI/MluI-digested pTX15. For genetic complementation of the vraFG mutant, the vraFG genes were PCR amplified with the primers listed in Table 4; the PCR product was digested with EcoRI and BamHI, and cloned into EcoRI/BamHI-digested plasmid pRB473.
|Name||DNA sequence (5′−3′)|
|For the construction of the apsS-deletion mutant|
|For the construction of the apsS complementation and hybrid plasmids|
|For the construction of the vraFG-deletion mutant and complementation plasmid|
|For quantitative RT-PCR|
Oligonucleotide primers and probes (Table 4) were synthesized by Sigma. Probes for quantitative RT-PCR were used to continuously monitor formation of PCR products during PCR. cDNA was synthesized from total RNA using the SuperScript III first-strand synthesis system (Invitrogen) according to the manufacturer's instructions. The resulting cDNA and negative control samples were amplified with TaqMan Universal PCR Master Mix (Applied Biosystems). Reactions were performed in a MicroAmp Optical 96-well reaction plate using a 7700 Sequence Detector (Applied Biosystems). Standard curves were determined for each gene, by use of purified chromosomal template DNA at concentrations of 0.005–50 ng ml−1. All RT-PCR experiments were performed in triplicate, with gyrB RNA used as a control.
Killing assays were performed as described before (Lai et al., 2007). In brief, S. aureus cells were grown to exponential growth phase (OD600, 2.5–3.0), cultures were harvested, washed twice with 10 mM sodium phosphate buffer (pH 6.5) with 100 mM NaCl, and resuspended in the same buffer. The cells were diluted to a final concentration of 106 cells in each sample. The bacteria were exposed to a range of AMP concentrations and incubated at 37°C for 3 h, and appropriate dilution series of the samples were plated on TSB agar plates. The samples were incubated at 37°C for 24 h, and surviving S. aureus cells were counted.
Determination of MICs
Staphylococcus aureus cells were grown to mid-log phase; cultures were harvested, washed twice with 10 mM sodium phosphate buffer (pH 6.5) with 100 mM NaCl, and resuspended in LB medium. The cells were diluted using Luria–Bertani (LB) media to a final concentration of 105 ml−1 in each sample. The bacteria were exposed to a range of AMP concentrations and incubated at 37°C with shaking at 180 r.p.m. for at least 12 h, after which OD600 was measured. The MIC was defined as the concentration at which the OD600 was reduced by 50%.
Peptide degradation assay
Staphylococcus aureus MW2 cells were grown to exponential growth phase (OD600, 2.5–3.0); cultures were harvested, washed twice with 10 mM sodium phosphate buffer (pH 6.5) with 100 mM NaCl, and resuspended in the same buffer. The bacteria were exposed to 50 μg ml−1 hBD3 and incubated at 37°C. At different incubation time points, 1 ml of the culture was collected, and supernatants were lyophilized. Degradation was analysed on 15% Tricine SDS-PAGE gels.
Isolation and detection of membrane phospholipids
Staphylococcus aureus MW2 and apsS-deletion mutant overnight cultures were diluted 1:100 into 50 ml of TSB and incubated at 37°C with shaking at 180 r.p.m. until grown to an OD600 of 2.5–3.0 (mid-logarithmic growth phase). Cultures were then split into two equal volumes, and cells were washed twice in 10 mM sodium phosphate buffer (pH 6.5) with 100 mM NaCl and resuspended in the same buffer. To one sample, indolicidin was added to a final concentration of 5 μg ml−1. Samples were incubated at 37°C for 1 h, cultures were harvested, washed once in sodium acetate buffer (20 mM, pH 4.5), and cells were extracted as a wet pellet using a modified Bligh and Dyer method (1959). The cell pellets were resuspended in chloroform–methanol–water (10:20:8, v/v) at 4°C, and four times their weight of glass beads (Sigma) was added to the mixture. The cells were disrupted by vortexing three times for 2 min at 4°C. After centrifugation at 4000 r.p.m. for 15 min, the mixture was filtered and divided into two phases by adding chloroform and water to a final concentration of chloroform–methanol–water of 20:20:18. After settling (24 h), the lower phase was collected, roto-evaporated under reduced pressure, and dissolved in chloroform–methanol (2:1, by volume). Equal amounts of lipid extracts were spotted onto silica 60 F254 HPTLC plates (Merck) and developed with chloroform–methanol–water (65:25:4, by volume). Phospholipids were stained with 5% H2SO4.
CD1 Swiss female mice were obtained from Charles River Laboratories and were between 4 and 6 weeks of age at the time of use. All animals were housed and maintained under pathogen-free conditions at the Rocky Mountain Laboratory animal facility. The study protocols were reviewed and approved by the Animal Use Committee at Rocky Mountain Laboratories, NIAID, National Institutes of Health (NIH). S. aureus strains were grown to mid-exponential phase, washed twice with sterile PBS, then resuspended in PBS at 1 × 108 cfus per 100 μl, and each mouse was injected intraperitoneally with 2 × 108 cfus of live S. aureus in 0.2 ml PBS. Control animals received 0.2 ml PBS only. After inoculation, animal health and disease advancement were monitored every 3 h for the first 24 h, then every 8 h for up to 72 h. Animals were euthanized immediately if showing signs of respiratory distress, mobility loss, or inability to eat and drink. At the time of death, the kidneys were removed, weighed, homogenized and serially diluted in PBS. Dilution series of the samples were plated on TSB agar plates. The samples were incubated at 37°C for 24 h, and the cfu counts per 100 mg kidney were determined.
This work was supported by the Intramural Research Program of the NIAID, NIH.
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