• We are sorry, but NCBI web applications do not support your browser and may not function properly. More information
Logo of nihpaAbout Author manuscriptsSubmit a manuscriptNIH Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
IUBMB Life. Author manuscript; available in PMC Oct 1, 2010.
Published in final edited form as:
PMCID: PMC2757517
NIHMSID: NIHMS134010

Tuning the properties of the bacterial membrane with aminoacylated phosphatidylglycerol

Abstract

The bacterial envelope is a semi-permeable barrier that protects the cell from the hostilities of the environment. To survive the ever-changing conditions of their surroundings, bacteria need to rapidly adjust the biochemical properties of their cellular envelope. Amino acid (aa) addition to phosphatidylglycerol (PG) of the membrane is one of the mechanisms used by bacteria to lower the net negative charge of their cellular envelope, thereby decreasing its affinity for several antibacterial agents such as the cationic antimicrobial peptides (CAMPs) produced by the innate immune response during host infection. This process requires the activity of an integral membrane protein, called aa-PG synthase (aaPGS), to transfer the aa of aminoacyl-tRNA (aa-tRNA) onto the PG of the membrane. aaPGSs constitute a new family of virulence factors that are found in a wide range of microorganisms. aa-PGs not only provide resistance to cationic antimicrobial peptides, but also to other classes of antibacterial agents and to environmental stresses such as those encountered during extreme osmotic or acidic conditions. This review will describe the known biochemical properties of aa-PGSs, their specificity for aa-tRNAs and phospholipids, and the growing repertoire of aa used as substrates by these enzymes. Their prevalence in bacteria and the phenotypes and modulations of membrane properties associated with these molecules will be addressed, as well as their regulation as a component of the envelope stress response system in certain bacteria.

Introduction

Initially discovered in the early 1960's in a few Gram-positive bacteria, lysinylation and alanylation of PG uses aa-tRNAs, the substrates of protein biosynthesis, as aa donor molecules to synthesize aa-PG within the cell membrane (Figure 1B). The biological function of aa-PG was not realized for almost four decades and has recently drawn renewed interest. During the screening of staphylococci mutants for their sensitivity towards antibacterial peptides, Peschel et al., established a correlation between the gene mprF (multi peptide resistance factor, also named fmtC and lpiA) and the biosynthesis of Lys-PG in the bacterial membrane and resistance to components of the innate immune system (1). mprF is a virulence factor that allows evasion of neutrophil killing activity, and its deletion results in attenuation of S. aureus virulence. Identification of a lysyl-PG synthase (LysPGS) in staphylococci has provided new perspectives on the mechanisms of adaptation of the bacterial membrane to changing environmental conditions, and the different factors utilized by microorganisms to evade the host immune response.

Figure 1
Known mechanisms for decreasing the net negative charge of the bacterial envelope. Increasing the net positive charge of cell wall components confers resistance to multiple CAMPs. A. D-alanylation of lipo-teichoic acid chains. In Gram positive bacteria ...

In many microorganisms, aa-PGs are constitutively expressed but their biosynthesis can be enhanced under various environmental stress conditions. A series of studies showed that aminoacylation of PG in the bacterial membrane is one of a variety of methods that some species have evolved to counter the anti-microbial arsenal developed by other microorganisms inhabiting the same niche, or by the innate immune cells of a host organism (see (2) for review). Cationic antimicrobial peptides (CAMPs) are one of the most primitive weapons in this arsenal and are produced in all organisms. CAMPs preferentially target bacteria because of the high negative charge of the bacterial cell envelope, which is due in part to the elevated content of phospholipids. Other molecules, such as the teichoic acid chains of the Gram-positive cell wall, or the lipid-A found in the outer membrane of Gram-negative bacteria, also contribute to the net negative charge of the bacterial membrane. The diversity of CAMP structures reflects the multiplicity of their modes of action and cellular targets. CAMPs can disrupt normal membrane function using different mechanisms of pore formation or by interacting with specific components of the cell wall. Also, CAMPs not only target the cellular envelope, but some are capable of diffusing through the membrane to inhibit intracellular targets (3, 4). Addition of lysine to PG contributes to lowering of the net negative charge of the cellular membrane, and a decrease in the affinity for CAMPs. Addition of Lys to the membrane is one of several mechanisms that involve lowering of the net negative charge of the cellular envelope. For instance, addition of D-Ala to teichoic acid in Gram-positive bacteria, and addition of ethanolamine or 4-amino-4-deoxy-L-arabinose to lipid A in Gram-negative bacteria, are complementary strategies used by microorganisms to resist CAMPs (Figure 1) (2, 5).

Recent findings showed that aa-PGs not only enhance the resistance of microorganisms to CAMPs, but also provide protection against other classes of non-positively charged antimicrobial molecules that are not targeting the lipid bilayer. aa-PGs confer an advantage in other challenging growth conditions such as those encountered during osmotic or acidic stress. This review will describe the known biochemical properties of aaPGSs, their occurrence in bacteria, and the phenotypes and modulations of the membrane properties associated with these molecules. Recent progress regarding the regulation of aaPGSs expression will also be addressed.

Phylogenic distribution and structure of aaPGSs

Sequence comparisons with known aaPGS sequences allowed the identification of mprF homologs in a wide variety of microorganisms (Figure 2). 347 sequences of mprF homologs are distributed among 31 genera of Gram-positive bacteria (mostly firmicutes and actinobacteria), 59 genera of Gram-negative bacteria (mostly proteobacteria), and in three species of archaea in the Methanosarcina genus. mprF homologs are predominantly found in animal and plant pathogens, and in symbiotic and soil colonizing microorganisms. Interestingly, mprF homologs are heterogeneously distributed within a given genus. For example, among the 42 taxa of the Bacillus genus, mprF is not found in four species (i.e. Bacillus clausii KSM-K16, Bacillus coahuilensis m4-4, Bacillus halodurans C-125, and Bacillus selenitireducens MLS10).

Figure 2
Phylogenic distribution of mprF homologs in bacteria. mprF sequences were identified using Blast with the C-terminal hydrophilic domain of LysPGS from B. subtilis against the Reference Sequence database of proteins (RefSeq, http://www.ncbi.nlm.nih.gov ...

aaPGSs are composed of an N-terminal integral membrane domain and a C-terminal hydrophilic domain, and they do not share apparent similarity with any protein of known function. The membrane domain varies in size and in sequence, and is in fact absent from the aaPGS homologs of several species (e.g. Streptomyces, Burkholderia). Because of the structural variability found in the integral membrane domain, it is likely that the catalytic site is located in the C-terminal hydrophilic moiety of the enzyme. This idea is supported by results obtained through mutation analysis in which recurrent deletion of the N-terminal membrane domain of the B. subtilis protein did not disrupt activity of the enzyme in vitro (Roy, H., unpublished results). Based on the predicted secondary structure, the membrane domain is thought to consist of a variable number of α-helices (4 to 14) that orient the N-terminal extremity and the hydrophilic C-terminal moiety towards the cytoplasm where aa-tRNA, the substrate of aaPGSs, is located. The secondary structure of some aaPGS is predicted to consist of a helical domain with an odd number of helices that orient the hydrophilic domain towards the periplasm (1, 6). However, this topology is unlikely because of the absence of aa-tRNA in the periplasmic compartment. Interestingly, aaPGSs of several actinomycetes (e.g. Mycobacterium, Streptomyces) are fused by the C-terminal end to a lysyl-tRNA synthetase, suggesting that the flow of substrates in the PG lysylation pathway is channeled in these organisms (7).

Thirty-three of the available genome sequences encode two or more mprF paralogs. These paralogs are most often encountered in Gram-positive bacteria, particularly in members of the classes Actinobacteria (e.g. Mycobacterium, Streptomyces, etc.) and Clostridia. A recent investigation of the two aaPGS paralogs in Clostridium perfringens showed that each enzyme exhibits distinct substrate specificity. One enzyme is specific for the transfer of Lys, whereas the second is specific for the transfer of Ala onto PG; both enzymes use the corresponding aminoacylated tRNAs (i.e. Lys- tRNALys and Ala-tRNAAla) (7).

Substrate specificity of aaPGSs

aa-PGs were discovered at the same time as the essential components of the translation machinery, almost five decades ago. A wide variety of aa were determined to be associated with phospholipids in the membrane, and because of their abundance and diversity, it was proposed that these amino-lipids may participate in protein biosynthesis as aa donors (8, 9). This hypothesis was disproven and it was determined that although phospholipids have the propensity to associate with a variety of aa in a non-covalent manner, only a small subset of aa are utilized for tRNA-dependent covalent linkage to PG in the membrane (10) (11).

The covalent attachment of several types of aa to membrane phospholipids in a variety of bacterial species has been experimentally demonstrated by conventional chromatography and mass spectrometry techniques. Most of these organisms contain a homolog to mprF in their genomes, and the biochemical properties of the PG aminoacylation activity have been studied in some instances (Table 1).

Table 1
List of bacterial species in which L-aa-phospholipids have been detected, and in which aaPGS biochemical properties have been studied.

aa-tRNA specificity

To date, two aa have been directly shown to act as substrates for aaPGSs. Each PG aminoacylation activity corresponds to a distinct protein and specifically uses either Ala- tRNAAla or Lys- tRNALys. The mprF system and protein biosynthesis machinery compete equitably for the cellular pool of aa-tRNA, since the affinity of aaPGSs and elongation factor Tu (EF-Tu) for the aa-tRNA substrate is comparable (in the μM range for each) (7, 12, 13). The contribution of the tRNA moiety of aa-tRNA to the specificity of aaPGSs recognition seems to be rather low. Alanylated minihelix RNA, consisting of a 12 bp hairpin which mimics the acceptor stem and T-stem of the tRNA, displays unaltered kinetic parameters relative to full-length tRNA with AlaPGS from C. perfringens. Similarly, a heterologous aa-tRNA substrate such as Lys- tRNAAsp is also active indicating that the nature of the tRNA moiety is not a critical determinant for substrate recognition by LysPGS and AlaPGS (7). This moderate specificity for the tRNA moiety is also illustrated by the ease with which mprF pathways from various organisms (S. aureus, Rhizobium tropici, B. subtilis, C. perfringens, P. aeruginosa) can be successfully reconstituted in Escherichia coli (7, 13-17). Divergence in tRNA sequences often represents a species barrier preventing horizontal transfer of tRNA-dependent pathways. The lack of specificity of aaPGSs for the tRNA moiety of aa-tRNA provides a mechanistic basis (e.g. (18)) underlying the mprF pathway as a mobile virulence factor.

Since a limited number of aa-PGs are found in bacterial membranes, and the tRNA moiety is not a crucial determinant for aaPGS recognition, it is expected that the aa moiety of aa-tRNA is a more important determinant for substrate recognition. Specificity for the aa moiety has been investigated with LysPGS from S. aureus (12) and AlaPGS from C. perfringens (previously called C. welchii) (19). The nature of the atoms comprising the aliphatic chain of Lys is not important for LysPGS activity since β-aminoethylcyteinyl- tRNALys is a suitable substrate. However, the nature of the aa seems critical since Cys- tRNALys is not recognized by the enzyme (12). Also, the α-amino group of the aa is an important determinant for the transferase reaction since neither N-acetyl-Alanyl- tRNAAla nor lactyl- tRNAAla are substrates (19).

Interestingly, the phospholipid aminoacylation pathway described in this review resembles the pathway of formation of pentapeptides for cross-linking peptidoglycan chains, a mechanism involved in microbial resistance that is principally found in Gram-positive bacteria. Like aaPGSs, this system uses aa-tRNAs, membrane lipids, and transferase activities, which in this case are supported by enzymes belonging to the Fem family of proteins. To synthesize the pentapeptide bridge, these enzymes transfer the aa of aa-tRNAs to lipid II, the membrane lipid precursor located on the intracellular face of the membrane during the initial steps of peptidoglycan assembly (for review see (20)). Fem proteins display variable aa-tRNA specificities (i.e. Gly, Ser, Thr, Ala) depending on the bacterial species. Similar to aaPGSs, Fem proteins recognize aa-tRNAs features that are distinct from those recognized by components of the protein biosynthesis machinery and minimal aa-tRNA structures such as minihelices are efficient substrates (21, 22). If there is a mechanistic similarity between Fem and aaPGS proteins, there is no sequence similarity between these two families of proteins. It is also worth mentioning that addition of Gly to the peptidoglycan pentapeptide bridge in certain Staphylococci species uses a dedicated Gly-tRNAGly. This Gly- tRNAGly displays weak affinity for the elongation factor Tu, increasing the availability of this unusual aa-tRNA for peptidoglycan synthesis (23-25).

Phospholipid specificity

The specificity of LysPGS for PG has been investigated using chemically synthesized analogs of PG. Experiments with LysPGS from S. aureus revealed that the polar head of PG (i.e. the terminal glycerol phosphate group) is a crucial determinant for catalysis, and transfer of Lys from Lys- tRNALys occurs preferentially onto the 3′ hydroxyl group of PG. The 3′deoxy PG analog (phosphatidyl-(3′-deoxy)glycerol) is not a substrate for the reaction, whereas the 2′-deoxy PG is still active (26, 27). Isomerization of the lysyl group between the 2′ and 3′ hydroxyl group of PG is catalyzed under strong acidic conditions in vitro (28), and may also occur under physiological conditions since the 2′ and 3′ isomer forms have been detected in vivo (5, 29).

Cardiolipin (CL), a common lipid exhibiting a free 2′hydroxyl group, is not a substrate of the Staphylococcal enzyme (26, 27). However, this specificity does not seem to be conserved among aaPGSs since LysPGS of Listeria species is able to form Lys-PG as well as Lys-CL. In these organisms Lys-CL is a major constituent of the membrane and represents up to 30% of the total phospholipids (6, 30).

The nature of the phospholipid fatty acid chain is not a major specificity determinant since the aaPGS activity of several organisms has been successfully transplanted into E. coli. Additionally, the saturation level and length of fatty acid chains within a given organism can vary in response to changing environmental conditions (31). Aminoacylated phospholipids with such variations in their fatty acid composition have been observed in the same organism (e.g., (30, 32)). However, LysPGS from S. aureus exhibits a slight preference for PG extracted from the same organism over PG extracted from egg yolk. This distinction may be due to variations in the composition of the fatty acid chains, or may be attributed to differences in the chirality of the the PG glycerol moiety (a racemic mixture is found in egg PG) (27).

Other aa-tRNA specificities within the repertoire of aaPGSs

Some other aa have been also found associated with the membrane of certain bacteria. L-Ornithine containing lipids are phosphorus free lipids found in many bacteria. They are synthesized in phosphate limiting conditions in a tRNA-independent manner so that available phosphate can be rerouted to nucleic acid biosynthesis. The operon olsAB was recently reported as coding for the biosynthetic pathway of this amino acid containing lipid (33). A few other species of aminoacylated phospholipids were reported for which little is known. D-Ala-PG and D-Ala-CL have been identified in the membrane of Vagococcus fluvialis (29). Gly-PG has been reported in C. perfringens (9) and Ornithyl-PG found in B. cereus (34) and Mycobacterium (35). More investigations are required to determine whether these unusual aminoacylated lipids are formed through a tRNA-dependent or independent mechanism. Finally, L-Arg-PG in the Enterococcus faecium membrane has been found along with other aminoacylated phospholipids such as Ala-PG, and some PG acylated by two molecules of Lys (36, 37). Our preliminary investigation showed that arginylation of PG in this organism occurs through a dual specific aaPGS (Arg/LysPGS) that is able to use Arg- tRNAArg or Lys- tRNALys as an aa donor (Roy, unpublished data). This latter finding illustrates that more aa may be involved in the modification of membrane lipids. The functional divergence among aaPGSs is reflected in the structural diversity of this family of proteins.

Localization and properties of aa-PG in the lipid bilayer

Before giving an overview of the phenotypes linked to aa-PG and the regulation of its biosynthesis, we will review the biophysical properties of aa-PG derived from studies of monolayer membrane models reconstituted in vitro. Some of these observations have been corroborated by observations in vivo.

Biosynthesis of aa-PG in vivo

The bacterial membrane is a complex structure mainly composed of phospholipids. The principal phospholipids encountered in bacteria are the anionic phospholipid PG and the zwitterionic phosphatidylethanolamine (PE) which bears a neutral charge at physiological pH. Other phospholipids such as Cardiolipin (CL), the cationic Lys-PG, and the zwitterionic Ala-PG and PS are less prominent in the lipid composition of bacterial membranes. The identity of the phospholipid polar head is determined by a biosynthetic pathway that consists of two main branches, one for the biosynthesis of negatively charge phospholipids (i.e. PG, CL), and the other for synthesis of negative or zwitterionic neutral species (i.e. PS, PE) (Figure 3). By directing lipid biosynthesis through one branch or the other, bacteria can adjust the electrostatic properties of their membranes. The mprF pathway branches directly from the biosynthetic pathway that produces negatively charged phospholipids (i.e. PG, CL). This enables bacteria to adapt the electrostatic properties of their membrane quickly and efficiently through the modification of pre-existing negatively charged species without the need for de novo lipid synthesis. The position of phospholipid aminoacylation in membrane metabolism, and the fact that aaPGSs are not universally conserved among bacteria, suggest that this pathway arose late during the evolution of phospholipid metabolism. Interestingly, bacteria use a similar mechanism at the level of phospholipid fatty acid chain synthesis to adapt the membrane to environment requirements without having to synthesize new phospholipids. Bacteria have evolved systems to alter the structure of pre-existing fatty acids by decreasing saturation levels or incorporating cylopropane within the hydrocarbon chain, thereby influencing membrane fluidity and permeability (31).

Figure 3
Phospholipid biosynthesis and aminoacylation pathways in bacteria. The structure of the polar head groups of common phospholipids in the membrane of bacteria is detailed and the net charge at neutral pH of each is indicated in brackets. aa addition to ...

The relative amount of individual lipids varies greatly from one organism to another. Under typical growth conditions in rich media at neutral pH, the membrane of S. aureus is almost exclusively composed of PG (42%) and Lys-PG (42%) (38). In B. subtilis Lys-PG (22%), PE (30%), PG (36%) and CL (12%) are all well represented showing that, in both cases, Lys-PG biosynthesis is not only induced in response to stress, but also contributes to lipid homeostasis under regular growth conditions. In both of these organisms, Lys-PG is dispensable for cellular growth, and in B. subtilis, phospholipids can be removed individually from the composition of the membrane without impairing cell survival (39). Recent investigations into the distribution of phospholipids at the membrane surface revealed the existence of CL and PE rich domains located at the septal regions of B. subtilis, indicating a reduced amount of Lys-PG and PG in this region (40, 41). Interestingly, the proteins involved in phospholipid biosynthesis, including LysPGS, are all located in the septal region (40, 41). There is less data on the localization of aaPGS and aa-PG in the membranes of Gram-negative bacteria. However, it has been recently shown that Ala-PG is incorporated into the inner and outer membranes of E. coli expressing AlaPGS from P. aeruginosa. Since E. coli normally does not synthesize aa-PG, these results indicate that Ala-PG may diffuse to the outer membrane through a non-specific transport mechanism (17).

Properties of membranes containing aa-PG

Lipid membranes are a composite material, which display biophysical properties that differ from the sum of the properties of each of the individual components. This composite behavior is provided by the variety of inter- and intra- molecular interactions that lipids can form that are modulated by parameters such as temperature and type of counter ions present. Intermolecular lipid-lipid interactions have been demonstrated for several common head groups of phospholipids from bacteria. For instance, the ethanolamine group of PE can form intermolecular bonding with the phosphate groups of neighboring phospholipids, modulating the biophysical properties of the membrane containing this lipid (42). Several biophysical studies conducted on phospholipid monolayers have revealed some physical properties of membranes containing aa-PG. Lys and Ala of the polar head exist in a loop conformation allowing for the formation of salt bridges with phosphate groups within the same molecule or with those of neighboring phospholipids. These intra- or intermolecular interactions exist within a range of pH values (3 to 7) at which deprotonation of the phosphate and protonation of the amine group of aa-PG occurs. The molecular packing and fluidity of phospholipids can vary as a function of the counter ions present at the surface of the membrane, which can shield the salt bridge interactions of the lipids. This effect seems greatly attenuated by Ala-PG, since Ala-PG containing membranes display constant packing properties in a wide range of ionic conditions (43). Furthermore, results from these studies suggested that Lys-PG and Ala-PG, in the presence of certain divalent ions such as Ca2+, may raise the surface potential of the membrane conferring decreased permeability to cations and protons (43-45). Indeed, Lys-PG containing liposomes have been shown to exhibit decreased permeability to Rb+ and increased fluidity (38). Also, a recent study investigating the physical properties of the bacterial membrane in S. aureus demonstrated that an increasing amount of Lys-PG correlates with an increase in membrane fluidity and a decrease in permeability to daptomycin (46).

Major changes in lipid composition that affect the structural properties of cell membranes may also impact the activity of a number of membrane associated proteins. Several studies have reported an effect of Lys-PG on the activity of enzymes. For instance, one study demonstrated that Lys-PG inhibits the first step of genomic DNA replication by inhibiting the activity of the protein DnaA. Disruption of mprF in S. aureus increases the number of replication origins per cell, suggesting that Lys-PG may play a role in the regulation of cell cycle events (47). Another study has shown that the activity of phospholipase A2 (PLA2), a cationic host defense factor involved in bacterial clearing during infection, decreases when the overall electrostatic potential of B. subtilis protoplasts is increased by the presence of Lys-PG (48). It is worth noting that S. aureus utilizes a different mechanism in which resistance to PLA2 is not due to Lys-PG in the membrane, but is dependent on the D-alanylation of teichoic acid chains within the cell wall (49).

aa-PG within the bacterial membrane is linked to several resistance phenotypes

Cellular membranes are semi-permeable barriers that define and preserve biologically relevant compartments and have a central role in cellular life. The bacterial membrane is responsible for energy synthesis, maintenance of cell shape, motility, and cell aggregation as well as for sensing environmental stimuli and stresses. To survive the changing environmental conditions, living cells must quickly adapt their cell membranes by adjusting the levels and properties of the constituent lipids. The cell must choose from a variety of lipid components that vary in fatty acid chain length and saturation levels, and that bear a repertoire of polar head groups (31). Alteration of membrane composition leads to various phenotypical changes that are directly or indirectly related to membrane function. The presence of aa-PG in the membrane has been found to be associated with resistance to a variety of antimicrobial compounds and to several peculiar physicochemical conditions.

aa-PG as a factor of resistance to cationic antimicrobial molecules

mprF was discovered in Staphylococci by screening of transposon insertion libraries for mutants sensitive to gallidermin. Disruption of mprF in S. aureus leads to a loss of synthesis of Lys-PG and enhances sensitivity to a wide range of bacterial and mammalian CAMPs (Table 2) (1). The presence of Lys-PG in the membrane can provide up to a 30-fold increase in the minimum inhibitory concentration (MIC) of certain CAMPs, although this number is variable for individual compounds. For instance, the MIC of weakly positively charged CAMPs, such as gramicidin D or S, is not affected by Lys-PG (1). Since the initial discovery in S. aureus, the correlation between CAMP resistance and mprF has been demonstrated in various bacterial contexts for both Lys-PG and Ala-PG (Table 2). Depletion of positively charged phospholipids (i.e. PE and Lys-PG) from the cell membrane of B. subtilis decreases its resistance to CAMPs, underlining the effectiveness of positive charges against CAMPs (39). aa-PG exerts its effect towards CAMPs through electrostatic repulsion, but may also alter the properties of the lipid bilayer by increasing fluidity and diminishing permeability to CAMPs (50). This effect may also explain how zwitterionic Ala-PG which, in spite of its neutral net charge, increases the resistance of P. aeruginosa to the CAMP protamine (17). The evolutionary related Ala-PGS and Lys-PGS probably arose via duplication to better adapt the bacterial response to a specific set of CAMPs or antibacterial molecules in a given environment.

Table 2
Antimicrobial resistances linked to aa-PG

Interestingly, the protective effects of Lys-PG differ among isolates of B. anthracis. In B. anthracis Stern, mprF confers an increase in resistance to three different CAMPs, however, in B. anthracis ΔANR, increased resistance is observed for only one CAMP (32). These findings illustrate that CAMP resistance is multifactorial and that the variation of sensitivity to certain CAMPs results from the sum of the expression levels of multiple resistance mechanisms and to subtle differences in the membrane composition and components. It is worth noting that in B. subtilis, variations in the lipid membrane composition of other phospholipids, besides Lys-PG, strongly modulate the resistance to various CAMPs (39). Variability in the effects conferred by Lys-PG in different genetic contexts is not limited to B. anthracis, but is also observed for resistance of S. aureus to glycopeptides and lipopeptides.

Resistance to glycopeptides and lipopeptides

Vancomycin is a positively charged glycopeptide that inhibits peptidoglycan crosslinking in the periplasm. In S. aureus, vancomycin resistance mediated by Lys-PG varies between individual strains. Different studies have reported that mprF is a factor that enhances (51), diminishes (52) or has no effect on (53) the resistance of S. aureus to vancomycin. These differences are not due to different levels of mprF expression, which remain constant among 24 different strains of S. aureus that display various susceptibility patterns to vancomycin (54). This disparity in the vancomycin sensitivity profile seems dependent on the genetic background of each S. aureus isolate (52). In addition to vancomycin resistance, a possible link between mprF and daptomycin (an acidic pore forming lipopeptide) resistance has recently been suggested. Several clinical isolates of S. aureus developed daptomycin or vancomycin resistances during treatment of infected patients (55-57). In some of these isolates a mutation in mprF was identified, further implicating mprF as a potential factor for modulating resistance to these antibiotics (58-61). However, a direct correlation between mprF and daptomycin resistance has not been found, since the mutations in mprf were not always detected in these isolates (59, 60, 62-65). Instead, it is proposed that the genetic background and the regulatory network (see corresponding section below) of factors modulating the additive properties of the cell membrane affect the resistance of a given isolate to vancomycin and daptomaycin (64).

Resistance to β-lactams

Concurrent with the discovery of mprF by Peschel et al., as a factor for resistance to CAMPs, mprF was shown to be involved in resistance to several β-lactams (53). Initial experiments in S. aureus involved the screening of a transposon insertion library for mutants sensitive to oxacillin. Further characterization of mprF disruptants showed that mprF increases the MIC of four different β-lactams by 4 to 8 fold, and prompted the authors to name the new orf fmtC (factor which affects methicillin resistance in the presence and absence of Triton)(53). Interestingly, biosynthesis of Lys-PG is induced during the treatment of S. aureus with β-lactam (66, 67). Also, Ala-PG has been shown to increase the resistance of P. aeruginosa to cefsulodin (17). It is unclear how aa-PG increases resistance to β-lactams since these compounds are negatively charged and are not thought to directly interact with the lipid bilayer as they target the peptidoglycan biosynthesis machinery located in the periplasm (68). In the presence of β-lactams, the weakly cross-linked peptidoglycan makes the growing bacteria more susceptible to the osmotic-mechanical pressure of the cytoplasm. Thus, the increase of resistance to β-lactams is probably due to an indirect effect of aa-PG in the membrane. For instance, aa-PG may increase the mechanical resistance of the cytoplasmic membrane to osmotic pressure or may regulate the activity of key enzymes of the cell wall such as autolysins in S. aureus that condition cell wall turnover and cellular cycle (for review (69)), or porins in P. aeruginosa that are involved in the diffusion of β-lactams into the periplasmic space.

Resistance to challenging acidic and osmotic conditions

In symbiotic or pathogenic bacteria of plants (i.e. Rhizobiales) mprF (also called lpiA: for low pH-inducible protein) is induced in acidic growth conditions (70). In Rhizobiales, lpiA is organized in an operon that also encodes acvB (71), and both genes are required for acid tolerance in Rhizobium tropici and Sinorhizobium medicae (71, 72). acvB encodes for a protein of unknown function that contains a lipase signature motif with a catalytic Ser residue (Prosite number PS00120) (71). While Lys-PG has been detected in R. tropici, this modification has not been detected in in the membrane of S. medicae during adaptation to acid conditions (72). Outside of the rhizobia, the possible role of Lys-PG in acid tolerance mechanisms has not been directly addressed. However, biosynthesis of Lys-PG is a cellular response to acid conditions that has been observed in numerous Bacillus, Streptococcus, and Staphylococcus species (73-75). The role of Lys-PG in resistance to low pH is not entirely understood. However, it has been proposed that Lys-PG may diminish the permeability of the lipid bilayer to protons, which are known to diffuse through lipid bilayers (see previous section) (43, 44). Tuning of membrane permeability by Ala-PG has recently been illustrated in P. aeruginosa, in which resistance to high concentrations of lactate is linked to the presence of this aa-PG in the membrane (17). Lactate is a fermentation product that is used as a food preservative. At high concentrations, this osmolyte can act as proton carrier via passive diffusion of the protonated form through the lipid bilayer. Ala-PG may reduce diffusion of this osmolyte through the membrane, thereby lowering accumulation of protons and lactate inside the cell that could otherwise lead to cell death (76).

Resistance to host immunity

mprF is an essential factor for bacterial virulence and for colonization of host organisms. mprF allows evasion of neutrophil activity and enhances the virulence of S. aureus in mice (1) and in endovascular infection of rabbits (77). The amount of Lys-PG present in staphylococcal membranes correlates with the invasiveness of different Staphylococcus species (78). mprF provides a specific resistance to defensins produced by neutrophils, but does not attenuate the oxidative burst within phagolysosomes. Therefore, mprF does not provide any advantage to S. aureus in human monocytes, a type of cell lacking defensins (79). Similar observations were made for L. monocytogenes for which mprF confers an advantage for infection of epithelial cells and macrophages in mice (6). Finally, in the context of rhizobiales (i.e. R. tropici) the mprF homolog, lpiA, provides a competitive advantage for the colonization of the roots of plants (nodulation), a process in which defensin like peptides also play an important role (71).

mprF as a factor of the cell envelope stress response

Bacteria have developed a range of complex and sensitive signal transduction systems with which they adjust gene expression to adapt to their changing surroundings. The principal components and the regulatory mechanisms that coordinate the cell envelope stress response are now well explored in Gram positive as well as Gram negative bacteria (for review: (80, 81)). These systems play an integral role in the cellular stress response, but are also involved in regulation during normal growth. Regulation involves the activity of a stress-sensor anchored in the cell membrane and a cytoplasmic transcriptional regulator that activates specific genes. aa-PG biosynthesis is induced during various stress conditions such as acidification of the medium or by antibiotics targeting the cell envelope. Recent investigations into the regulation of the envelope stress response in bacteria revealed several regulatory pathways that control the expression of mprF.

Stimuli inducing mprF expression

In numerous microorganisms, acidification of the cellular environment induces the biosynthesis of Lys-PG. This response occurs in S. aureus (73, 74), E. faecium (74, 82), B. subtilis (11, 83) and B. megaterium, in which the Lys-PG content can reach up to 80% of the total lipids present in the membrane (84). Induction of the mprF pathway has also been observed in several Gram-negative Rhizobiaceae such as S. medicae and R. tropici (15, 70-72). In early studies, it was determined that the increase of Lys-PG levels in the membrane of S. aureus under acidic conditions results from an increase in Lys-PG biosynthesis and a decrease in Lys-PG catabolism. This occurs in spite of decreased activity of LysPGS at acidic pH (pH optimal=7) (12, 75). In Staphylococci, numerous cell envelope inhibitors are also able to trigger Lys-PG synthesis. For instance, in S. aureus, methicillin, vancomycin, penicillin-G, D-cycloserine and bacitracin, which target the cell envelope, all trigger Lys-PG expression. However, chloramphenicol and puromycin, which target the protein biosynthesis machinery, do not induce Lys-PG production (66, 67, 85)

Antimicrobial peptide sensor in Staphylococci

It has been recently shown in S. epidermidis that mprF is regulated by a CAMP sensing three-component system apsXRS (for antimicrobial peptide sensor; also named graRS) (86, 87). This system resembles a classical two-component histidine kinase response regulator and is composed of a sensor (ApsS), a regulator (ApsR), and a third protein (ApsX) of unknown function. ApsS is an integral membrane protein that binds CAMPs with a nine aa extracellular loop that is negatively charged. The aspXRS system responds to a wide variety of structurally unrelated CAMPs, but certain CAMPs act as better activators than others with no obvious correlation to charge density or structure. apsXRS activates three systems involved in CAMP resistance including mprF, the dlt operon responsible for the D-alanylation of teichoic acid, and vraFG which codes for an ABC transporter involved in CAMP resistance in S. aureus (88). Interestingly, apsXRS also enhances transcription of lysC, which encodes an enzyme involved in Lys biosynthesis, suggesting that this system not only induces the principal mechanisms of CAMP resistance, but also induces expression of substrates for these systems (i.e. mprF) (86). apsXRS is well conserved in Staphylococci, however the set of genes that are regulated by this system varies between organisms. This can perhaps explain the variability of CAMP resistance phenotypes observed among Staphylococcus species (see previous section). This variability in the regulation system of mprF is also encountered in other strains. For instance, S. aureus SG511-Berlin exhibits a mutation in the sensor apsS resulting in an increased susceptibility to CAMPs (89). A thorough analysis in S. aureus SA113, revealed that the apsXRS system is in fact directly or indirectly responsible for the regulation of 248 genes including some that encode colonization factors, exotoxins, and CAMP resistance factors, excluding mprF which is not under the control of the aps system in this peculiar isolate(90, 91).

MprF regulation in L. monocytogenes

mprF is encoded outside of the main virulence regulon of L. monocytogenes, which is controlled by the themoinducible transcriptional activator PrfA (6, 92). Instead, mprF is under the control of a two component system called virSR, which consists of a sensor protein, VirS, and a regulator protein, VirR. It is still unclear what signal VirS responds to, but it has been demonstrated that this system activates several genes involved in cell envelope stress response and virulence of L. monocytogenes. The disruption of virS results in decreased entry of L. monocytogenes into host cells (93). As with the apsXRS system of S. aureus, VirR induces expression of mprF, the dlt operon and an ABC transporter. In addition to these genes, VirR enhances expression of VanZ, which is involved in resistance (by an unknown mechanism) to vancomycin and teichoplanin (94). It is worth noting that the regulator VirR can be activated by other sensor proteins besides VirS, indicating that multiple types of stimuli may trigger the mprF pathway in L. monocytogenes (93).

MprF regulation in S. medicae

Little is known about the regulation of mprF in Gram-negative bacteria. In several Rhizobiaceae, mprF (lpiA) is essential for survival at low pH and is induced under acidic conditions. In S. medicae, lpiA is not under the control of phrR or actSR, which are signal transduction components required for growth in acidic conditions (70). Alternatively, expression of lpiA may be regulated by the sensor component, TcsA, and two regulator components, FsrR and TcrA. The genes encoding these regulatory components are located upstream of lpiA and this system does not share any sequence similarity with the aps or vir systems that regulate expression of mprF in gram-positive bacteria (72). However, the organization of the locus is well conserved within several species of the Rhizobiaceae group, and it encodes the genes responsible for phospholipid aminoacylation as well as the genes regulating its expression.

Conclusion and outlook

Pathways that participate in the remodeling of the bacterial envelope, such as the mprF-catalyzed modifications, are central to adapting the biophysical properties of the bacterial membrane to the ever-changing conditions of the environment. During infection, these factors and their regulatory systems are essential for the virulence of certain pathogenic organisms, and allow for the evasion of antibiotics and innate host defenses. For this reason, these mechanisms represent interesting targets for new therapeutic strategies to combat microbial infections. Understanding the mechanisms of cell membrane permeation by antibiotics and the strategies of counteraction used by bacteria is crucial for the development of new bactericidal agents. Increased knowledge would help in the design of new inhibitors targeting enzymatic activities, their regulatory mechanisms, or specific components of the remodeled membrane such as Lys-PG. In spite of recent progress, many questions remain unanswered concerning phospholipid aminoacylation in bacteria. Topics of interest include the structure-function relationship of aaPGSs, the regulation of their expression, and how addition of aa to the membrane mediates bacterial resistance to different classes of antimicrobial molecules.

The repertoire of aa used by bacteria to modify their phospholipids needs to be explored by examining the specificity of aaPGS homologs from a wide variety of microorganisms, especially those containing several mprF paralogs. The 3D structure of aaPGSs, or at least of the hydrophilic domain bearing the active site, would provide insight into the structural features required for enzymatic activity and reveal what defines the specificity of the reaction. Also unclear is the exact role of the membrane domain of aaPGSs. More work is required to determine whether this domain simply serves to anchor the protein to the membrane, or if it supports additional functions. For instance, this domain may regulate the active site, or it could be involved in the transfer of aa-PGs from the inner leaflet of the membrane, where it is synthesized, to the outer leaflet where aa-PGs probably exert their role in CAMP resistance.

PG-lysylation was recently identified as a cell envelope stress response induced by CAMPs in Gram-positive bacteria; however, comparabale data for Gram-negative organisms are not available. Also, it is unknown if the regulatory mechanisms that induce Lys-PG expression in response to CAMPs in Gram-positive bacteria, also respond to other stress stimuli such as acid stress or alternative antibiotics (e.g. β-lactams). In addition, the regulation of aa-PG synthesis outside of the stress response, and the role of aa-PG in phospholipid homeostasis are not yet known. Several clues indicate that aa-PG may be implicated in many more general processes that affect bacterial life, and may also play a larger role than is currently realized in modulating resistance to a range of cell stimuli.

Acknowledgments

I would like to thank M. Ibba, K. Dare, and A.M. Smith for critical review of the manuscript. The author's work on this topic is supported a grant from NIH to Michael Ibba (GM65183).

References

1. Peschel A, Jack RW, Otto M, Collins LV, Staubitz P, Nicholson G, Kalbacher H, Nieuwenhuizen WF, Jung G, Tarkowski A, van Kessel KP, van Strijp JA. Staphylococcus aureus resistance to human defensins and evasion of neutrophil killing via the novel virulence factor MprF is based on modification of membrane lipids with l-lysine. J Exp Med. 2001;193:1067–1076. [PMC free article] [PubMed]
2. Peschel A, Sahl HG. The co-evolution of host cationic antimicrobial peptides and microbial resistance. Nat Rev Microbiol. 2006;4:529–536. [PubMed]
3. Hechard Y, Sahl HG. Mode of action of modified and unmodified bacteriocins from Gram-positive bacteria. Biochimie. 2002;84:545–557. [PubMed]
4. Zasloff M. Antimicrobial peptides of multicellular organisms. Nature. 2002;415:389–395. [PubMed]
5. Peschel A. How do bacteria resist human antimicrobial peptides? Trends Microbiol. 2002;10:179–186. [PubMed]
6. Thedieck K, Hain T, Mohamed W, Tindall BJ, Nimtz M, Chakraborty T, Wehland J, Jansch L. The MprF protein is required for lysinylation of phospholipids in listerial membranes and confers resistance to cationic antimicrobial peptides (CAMPs) on Listeria monocytogenes. Mol Microbiol. 2006;62:1325–1339. [PubMed]
7. Roy H, Ibba M. RNA-dependent lipid remodeling by bacterial multiple peptide resistance factors. Proc Natl Acad Sci U S A. 2008;105:4667–4672. [PMC free article] [PubMed]
8. Hunter GD, Goodsall RA. Lipo-amino acid complexes from Bacillus megaterium and their possible role in protein synthesis. Biochem J. 1961;78:564–570. [PMC free article] [PubMed]
9. Macfarlane MG. Characterization of Lipoamino-Acids as O-Amino-Acid Esters of Phosphatidyl-Glycerol. Nature. 1962;196:136–138.
10. Lennarz WJ, Nesbitt JA, 3rd, Reiss J. The participation of sRNA in the enzymatic synthesis of O-L-lysyl phosphatidylgylcerol in Staphylococcus aureus. Proc Natl Acad Sci U S A. 1966;55:934–941. [PMC free article] [PubMed]
11. den Kamp JA, Redai I, van Deenen LL. Phospholipid composition of Bacillus subtilis. J Bacteriol. 1969;99:298–303. [PMC free article] [PubMed]
12. Nesbitt JA, 3rd, Lennarz WJ. Participation of aminoacyl transfer ribonucleic acid in aminoacyl phosphatidylglycerol synthesis. I. Specificity of lysyl phosphatidylglycerol synthetase. J Biol Chem. 1968;243:3088–3095. [PubMed]
13. Oku Y, Kurokawa K, Ichihashi N, Sekimizu K. Characterization of the Staphylococcus aureus mprF gene, involved in lysinylation of phosphatidylglycerol. Microbiology. 2004;150:45–51. [PubMed]
14. Staubitz P, Neumann H, Schneider T, Wiedemann I, Peschel A. MprF-mediated biosynthesis of lysylphosphatidylglycerol, an important determinant in staphylococcal defensin resistance. FEMS Microbiol Lett. 2004;231:67–71. [PubMed]
15. Sohlenkamp C, Galindo-Lagunas KA, Guan Z, Vinuesa P, Robinson S, Thomas-Oates J, Raetz CR, Geiger O. The lipid lysyl-phosphatidylglycerol is present in membranes of Rhizobium tropici CIAT899 and confers increased resistance to polymyxin B under acidic growth conditions. Mol Plant Microbe Interact. 2007;20:1421–1430. [PubMed]
16. Roy H, Ibba M. Monitoring Lys-tRNA (Lys) phosphatidylglycerol transferase activity. Methods. 2008;44:164–169. [PMC free article] [PubMed]
17. Klein S, Lorenzo C, Hoffmann S, Walther JM, Storbeck S, Piekarski T, Tindall BJ, Wray V, Nimtz M, Moser J. Adaptation of Pseudomonas aeruginosa to various conditions includes tRNA-dependent formation of alanyl-phosphatidylglycerol. Mol Microbiol. 2009;71:551–565. [PubMed]
18. Mazauric MH, Roy H, Kern D. tRNA glycylation system from Thermus thermophilus, tRNAGly identity and functional interrelation with the glycylation systems from other phylae. Biochemistry. 1999;38:13094–13105. [PubMed]
19. Gould RM, Thornton MP, Liepkalns V, Lennarz WJ. Participation of aminoacyl transfer ribonucleic acid in aminoacyl phosphatidylglycerol synthesis. II. Specificity of alanyl phosphatidylglycerol synthetase. J Biol Chem. 1968;243:3096–3104. [PubMed]
20. Rohrer S, Berger-Bachi B. FemABX peptidyl transferases: a link between branched-chain cell wall peptide formation and beta-lactam resistance in gram-positive cocci. Antimicrob Agents Chemother. 2003;47:837–846. [PMC free article] [PubMed]
21. Maillard AP, Biarrotte-Sorin S, Villet R, Mesnage S, Bouhss A, Sougakoff W, Mayer C, Arthur M. Structure-based site-directed mutagenesis of the UDP-MurNAc-pentapeptide-binding cavity of the FemX alanyl transferase from Weissella viridescens. J Bacteriol. 2005;187:3833–3838. [PMC free article] [PubMed]
22. Fonvielle M, Chemama M, Villet R, Lecerf M, Bouhss A, Valery JM, Etheve-Quelquejeu M, Arthur M. Aminoacyl-tRNA recognition by the FemXWv transferase for bacterial cell wall synthesis. Nucleic Acids Res. 2009;37:1589–1601. [PMC free article] [PubMed]
23. Bumsted RM, Dahl JL, Söll D, Strominger JL. Biosynthesis of peptidoglycan of bacterial cell walls. X. Further study of the glycyl transfer ribonucleic acids active in peptidoglycan synthesis in Staphylococcus aureus. J Biol Chem. 1968;243:779–782. [PubMed]
24. Stewart TS, Roberts RJ, Strominger JL. Novel species of tRNA. Nature. 1971;230:36–38. [PubMed]
25. Giannouli S, Kyritsis A, Malissovas N, Becker HD, Stathopoulos C. On the role of an unusual tRNAGly isoacceptor in Staphylococcus aureus. Biochimie. 2009;91:344–351. [PubMed]
26. Lennarz WJ, Bonsen PP, van Deenen LL. Substrate specificity of O-L-lysylphosphatidylglycerol synthetase. Enzymatic studies on the structure of O-L-lysylphosphatidylglycerol. Biochemistry. 1967;6:2307–2312. [PubMed]
27. Bonsen PP, de Haas GH, van Deenen LL. Synthetic and structural investigations on 3-phosphatidyl-1′-(3′-O-L-lysyl)glycerol. Biochemistry. 1967;6:1114–1120. [PubMed]
28. Tocanne JF, Verheij HM, den Kamp JA, van Deenen LL. Chemical and physicochemical studies of lysylphosphatidylglycerol derivatives. Occurrence of a 2′ -> 3′ lysyl migration. Chem Phys Lipids. 1974;13:389–403. [PubMed]
29. Fischer W, Arneth-Seifert D. D-Alanylcardiolipin, a major component of the unique lipid pattern of Vagococcus fluvialis. J Bacteriol. 1998;180:2950–2957. [PMC free article] [PubMed]
30. Fischer W, Leopold K. Polar lipids of four Listeria species containing L-lysylcardiolipin, a novel lipid structure, and other unique phospholipids. Int J Syst Bacteriol. 1999;49(Pt 2):653–662. [PubMed]
31. Zhang YM, Rock CO. Membrane lipid homeostasis in bacteria. Nat Rev Microbiol. 2008;6:222–233. [PubMed]
32. Samant S, Hsu FF, Neyfakh AA, Lee H. The Bacillus anthracis protein MprF is required for synthesis of lysylphosphatidylglycerols and for resistance to cationic antimicrobial peptides. J Bacteriol. 2009;191:1311–1319. [PMC free article] [PubMed]
33. Gao JL, Weissenmayer B, Taylor AM, Thomas-Oates J, Lopez-Lara IM, Geiger O. Identification of a gene required for the formation of lyso-ornithine lipid, an intermediate in the biosynthesis of ornithine-containing lipids. Mol Microbiol. 2004;53:1757–1770. [PubMed]
34. Houtsmuller UM, van DL. Identification of a bacterial phospholipid as an O-ornithine ester of phosphatidyl glycerol. Biochim Biophys Acta. 1963;70:211–213. [PubMed]
35. Khuller GK, Subrahmanyam D. On the ornithinyl ester of phosphatidylglycerol of Mycobacterium 607. J Bacteriol. 1970;101:654–656. [PMC free article] [PubMed]
36. Gould RM, Lennarz WJ. Biosynthesis of aminoacyl derivatives of phosphatidylglycerol. Biochem Biophys Res Commun. 1967;26:512–515. [PubMed]
37. dos Santos Mota JM, den Kamp JA, Verheij HM, van Deenen LL. Phospholipids of Streptococcus faecalis. J Bacteriol. 1970;104:611–619. [PMC free article] [PubMed]
38. Haest CW, de Gier J, den Kamp JO, Bartels P, van Deenen LL. Chages in permeability of Staphylococcus aureus and derived liposomes with varying lipid composition. Biochim Biophys Acta. 1972;255:720–733. [PubMed]
39. Salzberg LI, Helmann JD. Phenotypic and Transcriptomic Characterization of Bacillus subtilis Mutants with Grossly Altered Membrane Composition. J Bacteriol 2008 [PMC free article] [PubMed]
40. Nishibori A, Kusaka J, Hara H, Umeda M, Matsumoto K. Phosphatidylethanolamine domains and localization of phospholipid synthases in Bacillus subtilis membranes. J Bacteriol. 2005;187:2163–2174. [PMC free article] [PubMed]
41. Matsumoto K, Kusaka J, Nishibori A, Hara H. Lipid domains in bacterial membranes. Mol Microbiol. 2006;61:1110–1117. [PubMed]
42. Boggs JM. Lipid intermolecular hydrogen bonding: influence on structural organization and membrane function. Biochim Biophys Acta. 1987;906:353–404. [PubMed]
43. Sacre MM, El Mashak EM, Tocanne JF. A monolayer (pi,deltaV) study of the ionic properties of alanylphosphatidylglycerol: effects of pH and ions. Chem Phys Lipids. 1977;20:305–318. [PubMed]
44. Tocanne JF, Ververgaert PH, Verkleij AJ, van Deenen LL. A monolayer and freeze-etching study of charged phospholipids. I. Effects of ions and pH on the ionic properties of phosphatidylglycerol and lysylphosphatidylglycerol. Chem Phys Lipids. 1974;12:201–219. [PubMed]
45. Tocanne JF, Ververgaert PH, Verkleij AJ, van Deenen LL. A monolayer and freeze-etching study of charged phospholipids. II. Ionic properties of mixtures of phosphatidylglycerol and lysylphosphatidylglycerol. Chem Phys Lipids. 1974;12:220–231. [PubMed]
46. Jones T, Yeaman MR, Sakoulas G, Yang SJ, Proctor RA, Sahl HG, Schrenzel J, Xiong YQ, Bayer AS. Failures in clinical treatment of Staphylococcus aureus Infection with daptomycin are associated with alterations in surface charge, membrane phospholipid asymmetry, and drug binding. Antimicrob Agents Chemother. 2008;52:269–278. [PMC free article] [PubMed]
47. Ichihashi N, Kurokawa K, Matsuo M, Kaito C, Sekimizu K. Inhibitory effects of basic or neutral phospholipid on acidic phospholipid-mediated dissociation of adenine nucleotide bound to DnaA protein, the initiator of chromosomal DNA replication. J Biol Chem. 2003;278:28778–28786. [PubMed]
48. Op den Kamp JA, Kauerz MT, van Deenen LL. Action of phospholipase A 2 and phospholipase C on Bacillus subtilis protoplasts. J Bacteriol. 1972;112:1090–1098. [PMC free article] [PubMed]
49. Koprivnjak T, Peschel A, Gelb MH, Liang NS, Weiss JP. Role of charge properties of bacterial envelope in bactericidal action of human group IIA phospholipase A2 against Staphylococcus aureus. J Biol Chem. 2002;277:47636–47644. [PubMed]
50. Xiong YQ, Mukhopadhyay K, Yeaman MR, Adler-Moore J, Bayer AS. Functional interrelationships between cell membrane and cell wall in antimicrobial peptide-mediated killing of Staphylococcus aureus. Antimicrob Agents Chemother. 2005;49:3114–3121. [PMC free article] [PubMed]
51. Ruzin A, Severin A, Moghazeh SL, Etienne J, Bradford PA, Projan SJ, Shlaes DM. Inactivation of mprF affects vancomycin susceptibility in Staphylococcus aureus. Biochim Biophys Acta. 2003;1621:117–121. [PubMed]
52. Nishi H, Komatsuzawa H, Fujiwara T, McCallum N, Sugai M. Reduced content of lysyl-phosphatidylglycerol in the cytoplasmic membrane affects susceptibility to moenomycin, as well as vancomycin, gentamicin, and antimicrobial peptides, in Staphylococcus aureus. Antimicrob Agents Chemother. 2004;48:4800–4807. [PMC free article] [PubMed]
53. Komatsuzawa H, Ohta K, Fujiwara T, Choi GH, Labischinski H, Sugai M. Cloning and sequencing of the gene, fmtC, which affects oxacillin resistance in methicillin-resistant Staphylococcus aureus. FEMS Microbiol Lett. 2001;203:49–54. [PubMed]
54. Wootton M, Macgowan AP, Walsh TR. Expression of tcaA and mprF and glycopeptide resistance in clinical glycopeptide-intermediate Staphylococcus aureus (GISA) and heteroGISA strains. Biochim Biophys Acta. 2005;1726:326–327. [PubMed]
55. Cui L, Tominaga E, Neoh HM, Hiramatsu K. Correlation between Reduced Daptomycin Susceptibility and Vancomycin Resistance in Vancomycin-Intermediate Staphylococcus aureus. Antimicrob Agents Chemother. 2006;50:1079–1082. [PMC free article] [PubMed]
56. Patel JB, Jevitt LA, Hageman J, McDonald LC, Tenover FC. An association between reduced susceptibility to daptomycin and reduced susceptibility to vancomycin in Staphylococcus aureus. Clin Infect Dis. 2006;42:1652–1653. [PubMed]
57. Sakoulas G, Alder J, Thauvin-Eliopoulos C, Moellering RC, Jr, Eliopoulos GM. Induction of daptomycin heterogeneous susceptibility in Staphylococcus aureus by exposure to vancomycin. Antimicrob Agents Chemother. 2006;50:1581–1585. [PMC free article] [PubMed]
58. Friedman L, Alder JD, Silverman JA. Genetic changes that correlate with reduced susceptibility to daptomycin in Staphylococcus aureus. Antimicrob Agents Chemother. 2006;50:2137–2145. [PMC free article] [PubMed]
59. Julian K, Kosowska-Shick K, Whitener C, Roos M, Labischinski H, Rubio A, Parent L, Ednie L, Koeth L, Bogdanovich T, Appelbaum PC. Characterization of a daptomycin-nonsusceptible vancomycin-intermediate Staphylococcus aureus strain in a patient with endocarditis. Antimicrob Agents Chemother. 2007;51:3445–3448. [PMC free article] [PubMed]
60. Pillai SK, Gold HS, Sakoulas G, Wennersten C, Moellering RC, Jr, Eliopoulos GM. Daptomycin nonsusceptibility in Staphylococcus aureus with reduced vancomycin susceptibility is independent of alterations in MprF. Antimicrob Agents Chemother. 2007;51:2223–2225. [PMC free article] [PubMed]
61. Murthy MH, Olson ME, Wickert RW, Fey PD, Jalali Z. Daptomycin non-susceptible meticillin-resistant Staphylococcus aureus USA 300 isolate. J Med Microbiol. 2008;57:1036–1038. [PubMed]
62. Rose WE, Leonard SN, Sakoulas G, Kaatz GW, Zervos MM, Sheth A, Carpenter CF, Rybak MJ. Evaluation of daptomycin activity against Staphylococcus aureus following vancomycin exposure in an in vitro pharmacodynamic model with simulated endocardial vegetations. Antimicrob Agents Chemother 2007 [PMC free article] [PubMed]
63. Boucher HW, Sakoulas G. Perspectives on Daptomycin resistance, with emphasis on resistance in Staphylococcus aureus. Clin Infect Dis. 2007;45:601–608. [PubMed]
64. Camargo IL, Neoh HM, Cui L, Hiramatsu K. Serial daptomycin selection generates daptomycin-nonsusceptible Staphylococcus aureus strains with a heterogeneous vancomycin-intermediate phenotype. Antimicrob Agents Chemother. 2008;52:4289–4299. [PMC free article] [PubMed]
65. Rose WE, Leonard SN, Sakoulas G, Kaatz GW, Zervos MJ, Sheth A, Carpenter CF, Rybak MJ. daptomycin activity against Staphylococcus aureus following vancomycin exposure in an in vitro pharmacodynamic model with simulated endocardial vegetations. Antimicrob Agents Chemother. 2008;52:831–836. [PMC free article] [PubMed]
66. Hebeler BH, Chatterjee AN, Young FE. Regulation of the bacterial cell wall: effect of antibiotics on lipid biosynthesis. Antimicrob Agents Chemother. 1973;4:346–353. [PMC free article] [PubMed]
67. Rozgonyi F, Biacs P, Szitha K, Kiss J. Effect of methicillin on the fatty acid composition of phospholipids in methicillin sensitive Staphylococcus aureus. Acta Microbiol Acad Sci Hung. 1981;28:97–110. [PubMed]
68. Wilke MS, Lovering AL, Strynadka NC. Beta-lactam antibiotic resistance: a current structural perspective. Curr Opin Microbiol. 2005;8:525–533. [PubMed]
69. Smith TJ, Blackman SA, Foster SJ. Autolysins of Bacillus subtilis: multiple enzymes with multiple functions. Microbiology. 2000;146(Pt 2):249–262. [PubMed]
70. Glenn AR, Reeve WG, Tiwari RP, Dilworth MJ. Acid tolerance in root nodule bacteria. Novartis Found Symp. 1999;221:112–126. discussion 126-130. [PubMed]
71. Vinuesa P, Neumann-Silkow F, Pacios-Bras C, Spaink HP, Martinez-Romero E, Werner D. Genetic analysis of a pH-regulated operon from Rhizobium tropici CIAT899 involved in acid tolerance and nodulation competitiveness. Mol Plant Microbe Interact. 2003;16:159–168. [PubMed]
72. Reeve WG, Brau L, Castelli J, Garau G, Sohlenkamp C, Geiger O, Dilworth MJ, Glenn AR, Howieson JG, Tiwari RP. The Sinorhizobium medicae WSM419 lpiA gene is transcriptionally activated by FsrR and required to enhance survival in lethal acid conditions. Microbiology. 2006;152:3049–3059. [PubMed]
73. Houtsmuller UM, Van D. On the Accumulation of Amino Acid Derivatives of Phosphatidylglycerol in Bacteria. Biochim Biophys Acta. 1964;84:96–98. [PubMed]
74. Houtsmuller UM, van Deenen LL. On the amino acid esters of phosphatidyl glycerol from bacteria. Biochim Biophys Acta. 1965;106:564–576. [PubMed]
75. Gould RM, Lennarz WJ. Metabolism of Phosphatidylglycerol and Lysyl Phosphatidylglycerol in Staphylococcus aureus. J Bacteriol. 1970;104:1135–1144. [PMC free article] [PubMed]
76. Rubin HE, Nerad T, Vaughan F. Lactate acid inhibition of Salmonella typhimurium in yogurt. J Dairy Sci. 1982;65:197–203. [PubMed]
77. Weidenmaier C, Peschel A, Kempf VA, Lucindo N, Yeaman MR, Bayer AS. DltABCD- and MprF-mediated cell envelope modifications of Staphylococcus aureus confer resistance to platelet microbicidal proteins and contribute to virulence in a rabbit endocarditis model. Infect Immun. 2005;73:8033–8038. [PMC free article] [PubMed]
78. Nahaie MR, Goodfellow M, Minnikin DE, Hajek V. Polar lipid and isoprenoid quinone composition in the classification of Staphylococcus. J Gen Microbiol. 1984;130:2427–2437. [PubMed]
79. Kristian SA, Durr M, Van Strijp JA, Neumeister B, Peschel A. MprF-mediated lysinylation of phospholipids in Staphylococcus aureus leads to protection against oxygen-independent neutrophil killing. Infect Immun. 2003;71:546–549. [PMC free article] [PubMed]
80. Rowley G, Spector M, Kormanec J, Roberts M. Pushing the envelope: extracytoplasmic stress responses in bacterial pathogens. Nat Rev Microbiol. 2006;4:383–394. [PubMed]
81. Jordan S, Hutchings MI, Mascher T. Cell envelope stress response in Gram-positive bacteria. FEMS Microbiol Rev. 2008;32:107–146. [PubMed]
82. Kocun FJ. Amino acid containing phospholipids as major components of the phospholipids of Streptococcus faecalis 10C1. Biochim Biophys Acta. 1970;202:277–282. [PubMed]
83. van Iterson W, den Kamp JA. Bacteria-shaped gymnoplasts (protoplasts) of Bacillus subtilis. J Bacteriol. 1969;99:304–315. [PMC free article] [PubMed]
84. den Kamp JO, Houtsmuller UM, van Deenen LL. On the phospholipids of Bacillus megaterium. Biochim Biophys Acta. 1965;106:438–441. [PubMed]
85. Rozgonyi F, Kiss J, Jekel P, Vaczi L. Effect of methicillin on the phospholipid content of methicillin sensitive Staphylococcus aureus. Acta Microbiol Acad Sci Hung. 1980;27:31–40. [PubMed]
86. Li M, Cha DJ, Lai Y, Villaruz AE, Sturdevant DE, Otto M. The antimicrobial peptide-sensing system aps of Staphylococcus aureus. Mol Microbiol. 2007;66:1136–1147. [PubMed]
87. Li M, Lai Y, Villaruz AE, Cha DJ, Sturdevant DE, Otto M. Gram-positive three-component antimicrobial peptide-sensing system. Proc Natl Acad Sci U S A. 2007;104:9469–9474. [PMC free article] [PubMed]
88. Meehl M, Herbert S, Gotz F, Cheung A. Interaction of the GraRS two-component system with the VraFG ABC transporter to support vancomycin-intermediate resistance in Staphylococcus aureus. Antimicrob Agents Chemother. 2007;51:2679–2689. [PMC free article] [PubMed]
89. Sass P, Bierbaum G. Native graS mutation supports the susceptibility of Staphylococcus aureus strain SG511 to antimicrobial peptides. Int J Med Microbiol 2009 [PubMed]
90. Herbert S, Bera A, Nerz C, Kraus D, Peschel A, Goerke C, Meehl M, Cheung A, Gotz F. Molecular basis of resistance to muramidase and cationic antimicrobial peptide activity of lysozyme in staphylococci. PLoS Pathog. 2007;3:e102. [PMC free article] [PubMed]
91. Kraus D, Herbert S, Kristian SA, Khosravi A, Nizet V, Gotz F, Peschel A. The GraRS regulatory system controls Staphylococcus aureus susceptibility to antimicrobial host defenses. BMC Microbiol. 2008;8:85. [PMC free article] [PubMed]
92. Johansson J, Mandin P, Renzoni A, Chiaruttini C, Springer M, Cossart P. An RNA thermosensor controls expression of virulence genes in Listeria monocytogenes. Cell. 2002;110:551–561. [PubMed]
93. Mandin P, Fsihi H, Dussurget O, Vergassola M, Milohanic E, Toledo-Arana A, Lasa I, Johansson J, Cossart P. VirR, a response regulator critical for Listeria monocytogenes virulence. Mol Microbiol. 2005;57:1367–1380. [PubMed]
94. Arthur M, Depardieu F, Molinas C, Reynolds P, Courvalin P. The vanZ gene of Tn1546 from Enterococcus faecium BM4147 confers resistance to teicoplanin. Gene. 1995;154:87–92. [PubMed]
95. Jones DE, Smith JD. Phospholipids of the differentiating bacterium Caulobacter crescentus. Can J Biochem. 1979;57:424–428. [PubMed]
96. Kampfer P, Rossello-Mora R, Falsen E, Busse HJ, Tindall BJ. Cohnella thermotolerans gen. nov., sp. nov., and classification of ‘Paenibacillus hongkongensis’ as Cohnella hongkongensis sp. nov. Int J Syst Evol Microbiol. 2006;56:781–786. [PubMed]
97. Fischer W. The polar lipids of group B Streptococci. II. Composition and positional distribution of fatty acids. Biochim Biophys Acta. 1977;487:89–104. [PubMed]
98. Exterkate FA, Otten BJ, Wassenberg HW, Veerkamp JH. Comparison of the phospholipid composition of Bifidobacterium and Lactobacillus strains. J Bacteriol. 1971;106:824–829. [PMC free article] [PubMed]
99. Smith MW, Steim JM. A phosphatidylinositol-containing derivative of Lactobacillus casei ATCC 7469. Biochim Biophys Acta. 1980;619:515–521. [PubMed]
100. Kalin JR, Allen CM. Lipid activation of undecaprenol kinase from Lactobacillus plantarum. Biochim Biophys Acta. 1980;619:76–89. [PubMed]
101. Koyama T, Yoshida I, Ogura K. Undecaprenyl diphosphate synthase from Micrococcus luteus B-P 26: essential factors for the enzymatic activity. J Biochem. 1988;103:867–871. [PubMed]
102. Short SA, White DC. Metabolism of phosphatidylglycerol, lysylphosphatidylglycerol, and cardiolipin of Staphylococcus aureus. J Bacteriol. 1971;108:219–226. [PMC free article] [PubMed]
103. Gould RM, Dawson RM. The trypsin-catalyzed hydrolysis of monomolecular films of lysylphosphatidylglycerol. Biochim Biophys Acta. 1972;288:1–11. [PubMed]
104. Sinha DB, Gaby WL. Structural Composition of Polar Lipid-Amino Acid Complex in Pseudomonas Aeruginosa. J Biol Chem. 1964;239:3668–3673. [PubMed]
105. Hachmann AB, Angert ER, Helmann JD. Genetic Analysis of Factors Affecting Susceptibility of Bacillus subtilis to Daptomycin. Antimicrob Agents Chemother 2009 [PMC free article] [PubMed]
106. Trent MS, Ribeiro AA, Lin S, Cotter RJ, Raetz CR. An inner membrane enzyme in Salmonella and Escherichia coli that transfers 4-amino-4-deoxy-L-arabinose to lipid A: induction on polymyxin-resistant mutants and role of a novel lipid-linked donor. J Biol Chem. 2001;276:43122–43131. [PubMed]
107. Ciccarelli FD, Doerks T, von Mering C, Creevey CJ, Snel B, Bork P. Toward automatic reconstruction of a highly resolved tree of life. Science. 2006;311:1283–1287. [PubMed]
PubReader format: click here to try

Formats:

Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...

Links

Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...