Abstract
Host defense peptides and proteins are important components of the innate host defense against pathogenic microorganisms. They target negatively charged bacterial surfaces and disrupt microbial cytoplasmic membranes, which ultimately leads to bacterial destruction. Throughout evolution, pathogens devised several mechanisms to protect themselves from deleterious damage of host defense peptides. These strategies include (a) inactivation and cleavage of host defense peptides by production of host defense binding proteins and proteases, (b) repulsion of the peptides by alteration of pathogen’s surface charge employing modifications by amino acids or amino sugars of anionic molecules (e.g., teichoic acids, lipid A and phospholipids), (c) alteration of bacterial membrane fluidity, and (d) expulsion of the peptides using multi drug pumps. Together with bacterial regulatory network(s) that regulate expression and activity of these mechanisms, they represent attractive targets for development of novel antibacterials.
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Introduction
In order to survive on and within the host, bacterial pathogens have evolved numerous mechanisms to combat host immune system. Among these are secreted molecules that interfere with recognition of bacterial pathogens by host immune system and pathogen-associated components that make the invader more resistant to the arsenal of host antimicrobial molecules such as lysoyzme, group IIA phospholipase A2, and small cationic antimicrobial peptides (CAMPs) [1]. Bacterial resistance mechanisms to the latter will be the focus of this brief.
Infection is typically initiated when a breach of the skin or mucosal barriers allows bacterial pathogen access to adjoining tissues or the bloodstream. Whether an infection is contained or spreads depends on a complex interplay between the pathogen’s virulence determinants and host defense mechanisms. The effective host immune response is based on quick recognition, isolation, and elimination of the pathogen. Innate immunity is the first line of defense against the invading microbial pathogens [1, 2]. The swift action of cellular (e.g., phagocytes) and secreted compounds (e.g., CAMPs) of innate immunity is in part due to the ability of innate immunity to recognize invariant structures on pathogens, so-called pathogen-associated molecular patterns or PAMPs [3]. While the killing by phagocytes occurs by combined action of reactive oxygen species and CAMPs, CAMPs are also secreted by various types of epithelial cells and therefore represent an important component of host defense on their own [1, 4, 5]. Many CAMPs have additional immunomodulatory roles that contribute to antimicrobial host defense [6].
Host defense peptides and their role in innate immune defense
CAMPs represent an evolutionary ancient part of immune response that is found in all kingdoms of life, from bacteria to plants and animals [5, 7]. These are small (12–50 amino acids) hydrophobic molecules that usually have a net positive charge of +2 or more and display a broad spectrum antimicrobial activity against bacteria as well as fungi [5, 8]. In humans, CAMPs can be placed in three distinct groups based on their peptide structure, amino acid composition, and number of disulfide bonds [5, 9]. Cathelicidins are CAMPs with α-helical structure, which do not contain cysteine residues. The only human representative of cathelicidin is LL-37 (see Fig. 1) [9]. Defensins have a β-sheet structure that is stabilized by three disulphide bridges. The pattern of disulphide bridges determines whether defensins belong to the group of α- or β-defensins. Defensins belonging to the two groups also differ by which cell type they are produced. While α-defensins [e.g., human neutrophil peptides (HNPs)] are mainly produced by neutrophil granulocytes and the intestinal Paneth cells, β-defensins are produced by epithelial tissues such as skin and epithelia of gastrointestinal, respiratory, and genitourinary tracts (Fig. 1) [5, 8, 10–12]. Kinocidins are the third group of CAMPs related to chemokines that are released from platelets upon contact with bacteria (thrombocidins) or other cells [13, 14]. Thrombocidins have been shown to participate in endovascular infections, e.g., in elimination of bacteria in endocardial vegetations [15–17]. In addition to the three above-mentioned major groups of CAMPs, further human antimicrobial peptides have been described including anionic dermcidins produced in human sweat, the iron-regulatory hormone hepcidin [18], and split products form human proteins such as α-melanocyte-stimulating hormone [19].
There are several proposed mechanisms of how CAMPs kill bacteria. Whether CAMPs induce pore formation or disrupt the membrane in any other way is still a matter of debate and may be different for different CAMPs. The modes of action may include cytoplasmic leakage and additional antibacterial mechanisms such as inhibition of membrane-bound cell wall biosynthetic steps (e.g., interaction of human-β defensin-3 (HBD3) and plectasin with lipidII, or binding of friulimicin B to bactoprenol phosphate carrier C55–P) [20–22], all ultimately leading to death of bacteria [7, 23, 24]. CAMPs mechanism of action can be described in at least three steps [5, 25, 26]. In order to reach the bacterial membrane, CAMPs need to initially bind to the bacterial cell surface and traverse the bacterial cell wall. The third step is insertion of CAMPs in the bacterial membrane that ultimately leads to cell membrane disruption and cell death [5, 8, 23]. The cationic nature of CAMP allows the peptides to distinguish between largely neutrally charged host membranes and mostly anionic bacterial membranes. Besides bacterial membrane damage, CAMPs can assist in killing of microorganisms by modulating host immune responses. To this end, CAMPs can bind lipopolysaccharides (LPS), the outer leaflet of the outer membrane of all Gram-negative bacteria with high proinflammatory capacity, to dampen host inflammatory response [27, 28]. In addition, many CAMPs can act as chemokines, can induce apoptosis, or play a role in activation of autoimmune response [7, 28, 29]. CAMP-derived synthetic peptides lacking antimicrobial activity have been shown to modulate immune responses and contribute to the clearance of infection [30], which underscores the importance of immunomodulatory CAMP activities.
How do bacteria defend against host defense peptides?
Extracellular mechanisms
Bacteria can defend themselves against host defense peptides using several different strategies. Some pathogens produce extracellular molecules that bind and trap CAMPs and thereby prevent their antimicrobial action. An example of such a protein is Staphylococcus aureus (S. aureus) staphylokinase. Besides binding of plasminogen that may facilitate S. aureus invasion of tissues, staphylokinase also forms complexes with α-defensins (HNP 1-3) resulting in over 80% reduced activity of these CAMPs against S. aureus [31]. Streptococcal inhibitor of complement (SIC) closely resembles the protective action of staphylokinase against the CAMPs. Even though SIC has been initially discovered as a complement-binding protein, Pence et al. [32] recently found that SIC-deficient Streptococcus pyogenes is more sensitive to LL-37. It has been proposed that SIC protects S. pyogenes by direct binding and inactivation of CAMPs [33].
Alternatively, bacteria can intercept host defense peptides before they reach their target—the bacterial cytoplasmic membrane—using cell envelope-associated or secreted proteases. Several pathogenic microbes use proteases that cleave CAMPs, thereby abolishing their antibacterial activity. Examples of such secreted proteases can be found in S. aureus (e.g., V8 and aureolysin proteases [34]), Proteus mirabilis (ZapA [35]), S. pyogenes (streptopain SpeB [35]), or Pseudomonas aeruginosa (elastase [35]). Alternatively, CAMP-degrading proteases can also be associated with the bacterial cell envelope either anchored in the cell wall or in the outer membrane such as PgtE of Salmonella enterica [36] and E. coli OmpT [37]. It has been suggested that CAMP evolution led to introduction of multiple cysteine bridges, resulting in a form that is substantially more resistant to proteolysis [7].
Modifications of cell surface
Synthesis of capsular polysaccharides
To protect themselves from host immune response including host defense peptides, bacteria can encase themselves by producing elaborate extracellular matrices. S. epidermidis, for example, produces cationic exopolymer polysaccharide intercellular adhesin (PIA) and anionic poly-γ-glutamic acid (PGA), which were both shown to play a role in resistance to cationic LL-37 and HBD3 as well as anionic defense peptide dermcidin [38–40]. Because these polysaccharide matrices protect bacteria from both cationic and anionic host defense peptides, the resistance mechanism probably involves both electrostatic and mechanical (charge-independent) sequestration of host defense peptides, far from their ultimate target—bacterial cytoplasmic membrane.
Role of teichoic acids (TA) in resistance to host defense peptides
Host evolution of defense peptides led to selection of mostly positively charged compounds. For measuring bactericidal effects of defense peptides, it seems that the rule of ‘the more cationic the better’ may be applied [5, 41, 42]. Net positive charge may be beneficial in targeting of bacterial surfaces that are generally negatively charged. Molecules that contribute to the net negativity of bacterial surface include peptidoglycan, anionic polymers such as TA, teichuronic acid, lipopolysaccharide (LPS) and anionic phospholipids in the bacterial cytoplasmic membrane. In the last decade, substantial advances in uncovering novel bacterial resistance mechanism that reduce the net negative charge of bacterial surfaces have been shown to confer resistance to CAMPs. These mechanisms differ between Gram-positive and Gram-negative bacteria due to the different composition of their cell surfaces (see Fig. 2). The envelope of Gram-negative bacteria consists of a thin layer of peptidoglycan and an extra outer membrane with inner phospholipid layer and outer LPS layer [43]. In contrast, Gram-positive bacteria lack the outer membrane, but have a thick, multiple-peptidoglycan layer cell wall. Analogous to LPS, the cell envelope of Gram-positive bacteria also contains various glyco- or glycolipid polymers, namely TA, which are attached to either peptidoglycan (wall teichoic acid, WTA) or cell membrane (lipoteichoic acid, LTA) [44, 45]. Both LPS and teichoic acids are negatively charged and contribute to the net negativity of bacterial surface [43–46].
Structurally, WTA and LTA differ depending on bacterial species. For example in S. aureus, they consist of repeating units of ribitol and glycerol phosphates, respectively [47, 48]. The length of the glycerol phosphate polymers of LTA can exceed 40 U [49], extending through the peptidoglycan layers. Together, LTA and WTA form a negatively charged lattice that bridges bacterial cell membrane and cell wall. The biochemical role of TA in bacterial physiology has been a focus of many studies. TA have been implicated in maintaining proton gradient, cationic homeostasis in bacterial cell wall, regulating enzymatic activity of autolysins and assembly of division site [50]. The biochemical properties of TA have been tightly linked to the substitutions of TA by the amino acid d-alanine. d-alanine’s positively charged free amino group partially influences the net negativity of TA [44, 50, 51]. In turn, d-alanyl esters of TA modulate functions of TA that significantly increase resistance of Gram-positive bacteria to CAMPs and proteins [52–58].
d-Alanylation of TA
At least four proteins, encoded by a single operon dltABCD, are necessary for d-alanylation of both WTA and LTA. d-alanyl carrier protein ligase (Dcl; dltA) activates d-alanine using ATP. With assistance of DltD (dltD), this activated complex is delivered to the d-alanine carrier protein (Dcp) encoded by dltC. DltB (dltB) is predicted to be a transmembrane protein and is thought to be involved in passage of the d-alanyl-Dcp complex across the cytoplasmic membrane, where d-alanine is transferred to the glycerol phosphate backbone of LTA and WTA [44]. The glycerol phosphate backbone is derived from phosphatidyl-glycerol (PG), the major phospholipid on growing S. aureus. Transfer of d-alanine most likely occurs at the base of the growing chain of LTA and WTA [59]. An additional open reading frame (“dltX”) present within the 5′ end of the dlt operon of S. aureus [60] and other Gram-positive bacteria has been recently identified, however, its function is not yet fully understood.
Inactivation of any of dltA–D genes results in complete loss of d-alanine esters from TA [52, 61–63]. Such dlt mutants have been created in numerous Gram-positive organisms including S. aureus, Streptococcal species, Listeria, Bacillus, and Enterococcus [52–58, 64]. In vitro studies have shown that dltA S. aureus is more sensitive to host cationic peptides (defensins) and proteins, especially group IIA PLA2 [41, 52] and is defective in adherence to artificial surfaces and formation of biofilms [65]. In addition, this mutant strain is virulence-attenuated in several animal models [53, 54, 66, 67]. Similar phenotypes have been observed in other bacterial species (see Table 1 for details). Of note, d-alanylation may also regulate pro-inflammatory activity of LTA [68], which may be an important determinant in establishing persistent infections (e.g., biofilms) and colonization of special niches. d-alanylation of TA in Lactobacillus have been show to be crucial for establishing successful colonization of the gastrointestinal tract [69].
The mechanism of d-alanyl-TA resistance to cationic peptides
CAMPs killing action is accomplished in three steps: (1) binding of CAMPs to the cell wall that depends on electrostatic interaction between cationic surfaces of defense peptides and negatively charged bacterial surfaces, (2) penetration of the cell wall, and, (3) insertion of CAMPs in the cell membrane leading to membrane disruption [4, 5]. In the absence of d-alanine esters, increased binding of CAMPs has been observed, suggesting that initial binding of CAMPs may be the critical step controlled by d-alanine esters [52]. However, additional binding sites for CAMPs may be revealed secondary to CAMPs’ inflicted cell damage that may mask the additional effect of d-alanine esters on CAMP action. Additional insights on CAMPs action may be gained from studies of another cationic protein that is part of mammalian innate immune response, human group IIA phospholipase A2 (gIIA PLA2). Antibacterial action of gIIA PLA2 is besides enzymatic degradation of phospholipids closely similar to the smaller CAMPs, but each of the steps can be controlled by manipulation of the presence of Ca2+ in the medium [41, 70, 71]. The results of these studies suggest that lack of d-alanine esters in TA may promote the penetration and release of autolytic enzymes that enhance the activity of cell wall bound gIIA PLA2 [41].
Regulation of d-alanylation of TA
Environmental factors such as pH, temperature, and salt (e.g., NaCl) concentration [72–74] are known to affect the degree to which TA are substituted by d-alanines. For example, the degree of d-alanylation of LTA is 0.77 mol d-alanine/mol glycerol phosphate when bacteria are grown under low-salt conditions (0.2% NaCl), but decreased to only 0.3 mol d-alanine/mol glycerol phosphate when bacteria are grown in medium with much higher salt (7.5% NaCl) concentration [75]. d-alanylation of TA could be further modulated by regulating the activity of the proteins encoded by dlt operon [76] or their abundance by regulation of transcription of dlt genes.
In Bacillus subtilis, the dlt operon is part of the σx regulon and is regulated by the global transcriptional regulators AbrB and Spo0A [61]. In Streptococcus agalactiae, a two-component system, dltRS, that is part of the dlt operon, is presumably involved in transcriptional regulation of dlt expression [77]. In S. aureus, transcription profiling studies have demonstrated increased dlt mRNA in an accessory gene regulator (agr) mutant and decreased dlt mRNA in a rot (repressor of toxins) mutant suggesting a role for the agr and rot global regulators in negative and positive regulation of dlt, respectively [78, 79]. In addition, S. aureus represses dlt transcription in response to increased concentration of mono and divalent cations that is in part dependent on ArlRS two-component system [60]. Most recently, studies aimed to identify genome-wide responses of S. epidermidis to HBD3 revealed a novel three-component system ApsRSX also termed GraRSX in S. aureus [80–84]. ApsRSX is an unusual three components system, composed of the classical combination of sensor histidine kinase and response regulator plus an additional protein, the function of which has yet to be determined. Most significantly, ApsS sensor kinase has been shown to be activated in the presence of CAMPs, in turn upregulating expression of resistance genes against CAMPs: dlt operon, mprF for modification of phospholipids involved in resistance to CAMPs [85] (see Fig. 2 and below), and vraFG operon encoding a transporter conferring resistance to glycopeptide vancomycin [86].
Modification of bacterial cell membrane
Charge neutralization
The ultimate step in CAMPs is integration into bacterial membrane leading to bacterial destruction [5]. This final step has been shown to be driven by electrostatic interactions between cationic defense peptides and anionic lipids in bacterial cell membrane [24, 87]. Neutralization mechanisms similar to charge compensation of anionic TA by d-alanine described above have also been described for bacterial membrane lipids [53, 82, 85]. In many Gram-positive and Gram-negative pathogens, phosphatidylglycerol (PG), a major component of bacterial cell membrane, is modified by positively charged amino acid lysine. The synthesis of lysyl-PG is dependent on the MprF protein [85]. MprF is highly conserved in Gram-positive and Gram-negative pathogens and is a 97-kDa integral membrane protein composed of two functional domains. A C-terminal domain is responsible for synthesis of lysyl-PG at the inner leaflet of cytoplasmic membrane using PG and lysyl-tRNA as substrates [85, 88, 89], while the N-terminal—flippase domain—is responsible for translocation of lysyl-PG to the outer leaflet of cytoplasmic membrane [89]. In addition to increased sensitivity to CAMPs, mprF mutants are more sensitive to killing by neutrophils and virulence-attenuated in multiple animal models (Table 1) [53, 85]. MprF has also been shown to be under control of ApsRSX regulator [81, 83]. Increased lysyl-PG content and point mutation resulting in MprF gain of function have been recently reported to increase resistance to daptomycin in vitro and in vivo [90–92]. Daptomycin is an anionic antibiotic but has CAMP-like properties in the presence of calcium ions [93] and is used as one of the last resort drugs in treatment of multiple drug resistant S. aureus (MRSA). In addition, mprF affects sensitivity to several other antibiotics such as gentamicin, vancomycin, and moenomycin [94–96].
Gram-negative bacteria can repel CAMPs by regulating anionic surface charge. Lipid A, the anionic component of LPS, consists of two glucosamine units with free phosphate groups linked to four or more acyl chains [43]. Salmonella can modify lipid A and LPS core sugars by incorporation of aminoarabinose or phosphoethanol amine (Fig. 2), leading to reduction in the net negative charge of lipid A and increased resistance to the CAMP polymyxin B [97–99]. The genes responsible for decoration of lipid A by aminoarabinose (pmrEHFIJKL) and ethanolamine (pmrC) are under control of PmrAB two-component system. The activity of PmrA transcriptional regulator is controlled by two different systems: (a) a sensor kinase PmrB that senses iron, zinc and mild acidic conditions, and (b) PhoPQ two-component system that regulates PmrA activity through PmrD protein [97, 100].
Changes in membrane fluidity
Salmonella typhimurium PhoPQ is one of the most studied bacterial two-component systems that controls several groups of genes important for bacterial survival in the host [101–103]. PhoPQ is a Mg2+ sensor, responds to changes in pH [104–106] and to the presence of CAMPs [107]. Crystal structure of PhoQ sensor kinase has revealed a patch of acidic amino acids that are responsible for binding of divalent cations and CAMPs (Fig. 2) [107]. PhoP has been shown to directly or indirectly influence expression >100 genes. Among them are Mg2+ transporters (MgtA and MgtCB), SlyA, which regulates genes important for survival inside macrophages, RpoS that regulates genes important for resistance to oxidative stress, and PmrAB, a two-component system that regulates genes for modifications of LPS by amino arabinose [104, 108, 109]. One of the PhoPQ-mediated responses is also upregulation of pagP that is responsible for additional acylation of the lipid A and pagL that is involved in deacylation of lipid A (Fig. 2). Such modified acylation of lipid A in turn reduces fluidity and permeability of the bacterial outer membrane and renders bacteria more resistant to CAMPs [36, 110, 111]. PhoPQ homologs can be found in many Gram-negative pathogens including Shigella flexneri, Yersinia pestis, and P. aeruginosa, suggesting this two-component system and its function may be one of the evolutionary adaptations to pathogen life within the host [112].
Changes in membrane fluidity have also been shown to influence CAMP resistance of Gram-positive organisms. For example, incorporation of longer chain unsaturated fatty acids in membrane lipids results in an increased membrane fluidity and resistance of S. aureus to platelet-derived CAMP [113]. Several studies also suggested that pigment production in S. aureus may influence sensitivity to CAMPs. Just recently, Mishra et al. [114] discovered that an S. aureus carotenoid staphyloxanthin increases S. aureus resistance to human neutrophil defensin 1, platelet-derived CAMPs, and polymyxin B. However, the mechanism of resistance proposed in this study is that staphyloxanthin increases membrane rigidity [114]. Why extreme increase and extreme decrease in membrane fluidity lead to increased resistance to CAMPs is still subject of investigation, however, it appears that the mechanism of resistance may be specific for different CAMPs.
Expulsion of host defense peptides
CAMPs that after damaging the membrane end up inside the cytoplasm, can also be actively exported from the cell by certain multi drug resistance exporters (MDR). Such examples have been found in several bacterial species including Neisseria gonorrhoeae [115–117]. N. gonorrhoeae, utilizes the MtrCDE MDR exporter to expel diverse antibiotics and confer resistance to the CAMPs protegrin PG1 and LL-37. However, in a recent study using E. coli overexpressing certain MDR exporters s of S. aureus, P. aeruginosa, and E. coli failed to show increased resistance to several CAMPs, indicating that most MDR exporters do not mediate broad CAMP resistance and that only some MDR exporters can expel certain CAMPs [118]. Further more, resistance mediated by S. aureus MDR exporter QacA to rabbit tPMP has been suggested not to be associated with the ability of QacA active transport of CAMP but rather attributed to the secondary QacA mediated changes on membrane fluidity [119]
Conclusions
Several advances in bacterial host defenses against host antimicrobial peptides have been recently reported. They include secreted proteins or cell surface-associated proteins that irreversibly bind or cleave CAMPs or glycopolymeric matrices that trap CAMPs to prevent their access to bacterial cytoplasmic membrane (see Table 1). Perhaps the biggest advances were made in identification of cell wall-associated mechanisms, the primary action of which appears to be electrostatic repulsion of CAMPs (Fig. 2). Whether it is modification of lipid A of Gram-negative LPS by aminoarabinose or ethanolamine, modification of phosphatidylglycerol by lysine, or esterification of TA by d-alanine, the common denominator to all is a reduction of bacterial net negative surface charge. In the era of increasing numbers of bacterial infections resistant to multiple antibiotics, novel antibiotics are in dire need. Even though CAMPs can be designed in such a way that they are more resistant to proteolytic degradation, high concentrations are needed for their effectiveness and their potentially harmful immunomodulatory effect may not make them suitable for further considerations [7]. In contrast, drugs that target bacterial responses to CAMPs that appear to be conserved among pathogens may be much more suitable. Inhibition of d-alanylation of TA offers a premium target due to the multiple effects that lack of d-alanine esters have on pathogenicity of several Gram-positive pathogens (see Table 1). To this end, inhibitors of DltA, a protein involved in the first step of d-alanylation have shown remarkable success as potential therapeutics in vitro and in vivo, especially when used in combination with other antibiotics [120]. MprF may be an even better target, since it is present in both Gram-positive as well as Gram-negative pathogens. Furthermore, because of involvement of MprF in resistance to daptomycin, inhibitors of MprF could be of even greater value.
Perhaps the newest notion to exploit is a discovery that bacteria can sense and respond to the presence of CAMPs by upregulation of genes responsible for their resistance. These bacterial sensing systems are well conserved among pathogens and may present attractive targets for developing new antimicrobials.
Abbreviations
- CAMPs:
-
Cationic antimicrobial peptides
- gIIA PLA2:
-
group IIA phospholipase A2
- HBD3:
-
Human β-defensin 3
- HNPs:
-
Human neutrophil peptides
- LPS:
-
Lipopolysaccharide
- LTA:
-
Lipoteichoic acid
- MDR:
-
Multidrug resistance
- PAMPs:
-
Pathogen-associated molecular patterns
- PGA:
-
Anionic poly-γ -glutamic acid
- TA:
-
Teichoic acids
- PG:
-
Phosphatidylglycerol
- PIA:
-
Polysaccharide intercellular adhesin
- SIC:
-
Streptococcal inhibitor of complement
- WTA:
-
Wall teichoic acid
References
Weiss J, Bayer AS, Yeaman M (2006) Cellular and extracellular defenses against Staphylococcal infections. In: Fischetti VA, Novick RP, Ferretti JJ, Portnoy DA, Rood JI (eds) Gram-positive pathogens. ASM Press, Washington, DC
DeLeo FR, Diep BA, Otto M (2009) Host defense and pathogenesis in Staphylococcus aureus infections. Infect Dis Clin North Am 23:17–34
Akira S, Uematsu S, Takeuchi O (2006) Pathogen recognition and innate immunity. Cell 124:783–801
Lehrer RI (2004) Primate defensins. Nat Rev Microbiol 2:727–738
Ganz T (2003) Defensins: antimicrobial peptides of innate immunity. Nat Rev Immunol 3:710–720
Easton DM, Nijnik A, Mayer ML, Hancock REW (2009) Potential of immunomodulatory host defense peptides as novel anti-infectives. Trends Biotechnol 27:582–590
Peschel A, Sahl H-G (2006) The co-evolution of host cationic antimicrobial peptides and microbial resistance. Nat Rev Micro 4:529–536
Izadpanah A, Gallo RL (2005) Antimicrobial peptides. J Am Acad Dermatol 52:381–390 quiz 391–382
Lehrer RI, Ganz T (2002) Cathelicidins: a family of endogenous antimicrobial peptides. Curr Opin Hematol 9:18–22
Ayabe T (2000) Secretion of microbicidal alpha-defensins by intestinal Paneth cells in response to bacteria. Nature Immunol 1:113–118
Bals R (1999) Mouse [beta]-defensin 3 is an inducible antimicrobial peptide expressed in the epithelia of multiple organs. Infect Immun 67:3542–3547
Bensch KW, Raida M, Magert HJ, Schulz-Knappe P, Forssmann WG (1995) hBD-1: a novel [beta]-defensin from human plasma. FEBS Lett 368:331–335
Yeaman MR (2010) Platelets in defense against bacterial pathogens. Cell Mol Life Sci 67:525–544
Fitzgerald JR, Foster TJ, Cox D (2006) The interaction of bacterial pathogens with platelets. Nat Rev Micro 4:445–457
Bayer AS, Cheng D, Yeaman MR, Corey GR, McClelland RS, Harrel LJ, Fowler VG Jr (1998) In vitro resistance to thrombin-induced platelet microbicidal protein among clinical bacteremic isolates of Staphylococcus aureus correlates with an endovascular infectious source. Antimicrob Agents Chemother 42:3169–3172
Kupferwasser LI, Yeaman MR, Shapiro SM, Nast CC, Bayer AS (2002) In vitro susceptibility to thrombin-induced platelet microbicidal protein is associated with reduced disease progression and complication rates in experimental Staphylococcus aureus endocarditis: microbiological, histopathologic, and echocardiographic analyses. Circulation 105:746–752
Dhawan VK, Bayer AS, Yeaman MR (1998) In vitro resistance to thrombin-induced platelet microbicidal protein is associated with enhanced progression and hematogenous dissemination in experimental Staphylococcus aureus infective endocarditis. Infect Immun 66:3476–3479
Ganz T (2006) Hepcidin–a peptide hormone at the interface of innate immunity and iron metabolism. Curr Top Microbiol Immunol 306:183–198
Madhuri ST, Venugopal SK, Ghosh D, Gadepalli R, Dhawan B, Mukhopadhyay K (2009) In vitro antimicrobial activity of alpha-melanocyte stimulating hormone against major human pathogen Staphylococcus aureus. Peptides 30:1627–1635
Schneider T, Gries K, Josten M, Wiedemann I, Pelzer S, Labischinski H, Sahl HG (2009) The lipopeptide antibiotic friulimicin B inhibits cell wall biosynthesis through complex formation with bactoprenol phosphate. Antimicrob Agents Chemother 53:1610–1618
Sass V, Schneider T, Wilmes M, Korner C, Tossi A, Novikova N, Shamova O, Sahl H-G (2010) Human beta-defensin 3 inhibits cell wall biosynthesis in Staphylococci. Infect Immun 78:2793–2800
Schneider T, Kruse T, Wimmer R, Wiedemann I, Sass V, Pag U, Jansen A, Nielsen AK, Mygind PH, Raventos DS, Neve S, Ravn B, Bonvin AMJJ, De Maria L, Andersen AS, Gammelgaard LK, Sahl H-G, Kristensen H-H (2010) Plectasin, a fungal defensin, targets the bacterial cell wall precursor lipid II. Science 328:1168–1172
Lai Y, Gallo RL (2009) AMPed Up immunity: how antimicrobial peptides have multiple roles in immune defense. Trends Immunol 30:131–141
Zasloff M (2002) Antimicrobial peptides of multicellular organisms. Nature 415:389–395
Shai Y (1999) Mechanism of the binding, insertion and destabilization of phospholipid bilayer membranes by alpha-helical antimicrobial and cell non-selective membrane-lytic peptides. Biochim Biophys Acta 1462:55–70
Wimley WC, Selsted ME, White SH (1994) Interactions between human defensins and lipid bilayers: evidence for formation of multimeric pores. Protein Sci 3:1362–1373
Scott MG, Vreugdenhil AC, Buurman WA, Hancock RE, Gold MR (2000) Cutting edge: cationic antimicrobial peptides block the binding of lipopolysaccharide (LPS) to LPS binding protein. J Immunol (Baltimore, MD, 1950) 164:549–553
Durr M, Peschel A (2002) Chemokines meet defensins: the merging concepts of chemoattractants and antimicrobial peptides in host defense. Infect Immun 70:6515–6517
Mader JS, Marcet-Palacios M, Hancock REW, Bleackley RC (2011) The human cathelicidin, LL-37, induces granzyme-mediated apoptosis in cytotoxic T lymphocytes. Exp Cell Res 317:531–538
Nijnik A, Madera L, Ma S, Waldbrook M, Elliott MR, Easton DM, Mayer ML, Mullaly SC, Kindrachuk J, Jenssen Hv, Hancock REW (2010) Synthetic cationic peptide IDR-1002 provides protection against bacterial infections through chemokine induction and enhanced leukocyte recruitment. J Immunol 184:2539–2550
Jin T, Bokarewa M, Foster T, Mitchell J, Higgins J, Tarkowski A (2004) Staphylococcus aureus resists human defensins by production of staphylokinase, a novel bacterial evasion mechanism. J Immunol 172:1169–1176
Pence MA, Rooijakkers SHM, Cogen AL, Cole JN, Hollands A, Gallo RL, Nizet V (2010) Streptococcal inhibitor of complement promotes innate immune resistance phenotypes of invasive M1T1 Group A Streptococcus. J Innate Immun 2:587–595
Frick I-M, Akesson P, Rasmussen M, Schmidtchen A, Bjorck L (2003) SIC, a secreted protein of Streptococcus pyogenes that inactivates antibacterial peptides. J Biol Chem 278:16561–16566
Sieprawska-Lupa M, Mydel P, Krawczyk K, Wojcik K, Puklo M, Lupa B, Suder P, Silberring J, Reed M, Pohl J, Shafer W, McAleese F, Foster T, Travis J, Potempa J (2004) Degradation of human antimicrobial p LL-37 by Staphylococcus aureus-derived proteinases. Antimicrob Agents Chemother 48:4673–4679
Schmidtchen A, Frick I-M, Andersson E, Tapper H, Björck L (2002) Proteinases of common pathogenic bacteria degrade and inactivate the antibacterial peptide LL-37. Mol Microbiol 46:157–168
Guina T, Yi EC, Wang H, Hackett M, Miller SI (2000) A PhoP-regulated outer membrane protease of Salmonella enterica serovar typhimurium promotes resistance to [alpha]-helical antimicrobial peptides. J Bacteriol 182:4077–4086
Hui C-Y, Guo Y, He Q-S, Peng L, Wu S-C, Cao H, Huang S-H (2010) Escherichia coli outer membrane protease OmpT confers resistance to urinary cationic peptides. Microbiol Immunol 54:452–459
Vuong C, Voyich JM, Fischer ER, Braughton KR, Whitney AR, DeLeo FR, Otto M (2004) Polysaccharide intercellular adhesin (PIA) protects Staphylococcus epidermidis against major components of the human innate immune system. Cell Microbiol 6:269–275
Kocianova S, Vuong C, Yao Y, Voyich JM, Fischer ER, DeLeo FR, Otto M (2005) Key role of poly-γ-dl-glutamic acid in immune evasion and virulence of Staphylococcus epidermidis. J Clin Invest 115:688–694
Vuong C, Kocianova S, Voyich JM, Yao Y, Fischer ER, De Leo FR, Otto M (2004) A crucial role for exopolysaccharide modification in bacterial biofilm formation, immune evasion, and virulence. J Biol Chem 279:54881–54886
Koprivnjak T, Peschel A, Gelb MH, Liang NS, Weiss JP (2002) Role of charge properties of bacterial envelope in bactericidal action of human group IIA phospholipase A2 against Staphylococcus aureus. J Biol Chem 277:47636–47644
Koprivnjak T, Weidenmaier C, Peschel A, Weiss JP (2008) Wall teichoic acid deficiency in Staphylococcus aureus confers selective resistance to mammalian group IIA phospholipase A2 and human beta-defensin -3. Infect Immun 76:2169–2176
Raetz CRH, Reynolds CM, Trent MS, Bishop RE (2007) Lipid A modification systems in Gram-negative bacteria. Annu Rev Biochem 76:295–329
Neuhaus FC, Baddiley J (2003) A continuum of anionic charge: structures and functions of d-alanyl-teichoic acids in Gram-positive bacteria. Microbiol Mol Biol Rev 67:686–723
Weidenmaier C, Peschel A (2008) Teichoic acids and related cell-wall glycopolymers in Gram-positive physiology and host interactions. Nat Rev Microbiol 6:276–287
Trent MS, Stead CM, Tran AX, Hankins JV (2006) Invited review: diversity of endotoxin and its impact on pathogenesis. J Endotxin Res 12:205–223
Weidenmaier C, Kokai-Kun JF, Kulauzovic E, Kohler T, Thumm G, Stoll H, Gotz F, Peschel A (2008) Differential roles of sortase-anchored surface proteins and wall teichoic acid in Staphylococcus aureus nasal colonization. Int J Med Microbiol 298:505–513
Baddiley J, Buchanan JG, Hardy FE, Martin RO, Rajbhandary UL, Sanderson AR (1961) The structure of the ribitol teichoic acid of Staphylococcus aureus H. Biochim Biophys Acta 52:406–407
Morath S, Geyer A, Hartung T (2001) Structure-function relationship of cytokine induction by lipoteichoic acid from Staphylococcus aureus. J Exp Med 193:393–397
Fischer W (1994) Lipoteichoic acid and lipids in the membrane of Staphylococcus aureus. Med Microbiol Immunol (Berl) 183:61–76
Xia G, Kohler T, Peschel A (2009) The wall teichoic acid and lipoteichoic acid polymers of Staphylococcus aureus. Int J Med Microbiol 300:148–154
Peschel A, Otto M, Jack RW, Kalbacher H, Jung G, Gotz F (1999) Inactivation of the dlt operon in Staphylococcus aureus confers sensitivity to defensins, protegrins, and other antimicrobial peptides. J Biol Chem 274:8405–8410
Weidenmaier C, Peschel A, Kempf VAJ, Lucindo N, Yeaman MR, Bayer AS (2005) 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 73:8033–8038
Collins LV, Kristian SA, Weidenmaier C, Faigle M, Van Kessel KP, Van Strijp JA, Gotz F, Neumeister B, Peschel A (2002) Staphylococcus aureus strains lacking d-alanine modifications of teichoic acids are highly susceptible to human neutrophil killing and are virulence attenuated in mice. J Infect Dis 186:214–219
Abachin E, Poyart C, Pellegrini E, Milohanic E, Fiedler F, Berche P, Trieu-Cuot P (2002) Formation of d-alanyl-lipoteichoic acid is required for adhesion and virulence of Listeria monocytogenes. Mol Microbiol 43:1–14
Kovacs M, Halfmann A, Fedtke I, Heintz M, Peschel A, Vollmer W, Hakenbeck R, Bruckner R (2006) A Functional dlt operon, encoding proteins required for incorporation of d-alanine in teichoic acids in Gram-positive bacteria, confers resistance to cationic antimicrobial peptides in Streptococcus pneumoniae. J Bacteriol 188:5797–5805
Abi Khattar Z, Rejasse A, Destoumieux-Garzon D, Escoubas JM, Sanchis V, Lereclus D, Givaudan A, Kallassy M, Nielsen-Leroux C, Gaudriault S (2009) The dlt operon of Bacillus cereus is required for resistance to cationic antimicrobial peptides and for virulence in insects. J Bacteriol 191:7063–7073
Cox KH, Ruiz-Bustos E, Courtney HS, Dale JB, Pence MA, Nizet V, Aziz RK, Gerling I, Price SM, Hasty DL (2009) Inactivation of DltA modulates virulence factor expression in Streptococcus pyogenes. PLoS ONE 4:e5366
Haas R, Koch HU, Fischer W (1984) Alanyl turnover from lipoteichoic acid to teichoic acid in Staphylococcus aureus. FEMS Microbiol Lett 21:27–31
Koprivnjak T, Mlakar V, Swanson L, Fournier B, Peschel A, Weiss JP (2006) Cation-induced transcriptional regulation of the dlt operon of Staphylococcus aureus. J Bacteriol 188:3622–3630
Perego M, Glaser P, Minutello A, Strauch MA, Leopold K, Fischer W (1995) Incorporation of d-alanine into lipoteichoic acid and wall teichoic acid in Bacillus subtilis. Identification of genes and regulation. J Biol Chem 270:15598–15606
Grundling A, Missiakas DM, Schneewind O (2006) Staphylococcus aureus mutants with increased lysostaphin resistance. J Bacteriol 188:6286–6297
Grundling A, Schneewind O (2006) Cross-linked peptidoglycan mediates lysostaphin binding to the cell wall envelope of Staphylococcus aureus. J Bacteriol 188:2463–2472
Fabretti F, Theilacker C, Baldassarri L, Kaczynski Z, Kropec A, Holst O, Huebner J (2006) Alanine esters of enterococcal lipoteichoic acid play a role in biofilm formation and resistance to antimicrobial peptides. Infect Immun 74:4164–4171
Gross M, Cramton SE, Gotz F, Peschel A (2001) Key role of teichoic acid net charge in Staphylococcus aureus colonization of artificial surfaces. Infect Immun 69:3423–3426
Weidenmaier C, Peschel A, Xiong YQ, Kristian SA, Dietz K, Yeaman MR, Bayer AS (2005) Lack of wall teichoic acids in Staphylococcus aureus leads to reduced interactions with endothelial cells and to attenuated virulence in a rabbit model of endocarditis. J Infect Dis 191:1771–1777
Kristian SA, Lauth X, Nizet V, Goetz F, Neumeister B, Peschel A, Landmann R (2003) Alanylation of teichoic acids protects Staphylococcus aureus against Toll-like receptor 2-dependent host defense in a mouse tissue cage infection model. J Infect Dis 188:414–423
Deininger S, Stadelmaier A, von Aulock S, Morath S, Schmidt RR, Hartung T (2003) Definition of structural prerequisites for lipoteichoic acid-inducible cytokine induction by synthetic derivatives. J Immunol 170:4134–4138
Walter J, Loach DM, Alqumber M, Rockel C, Hermann C, Pfitzenmaier M, Tannock GW (2007) D-alanyl ester depletion of teichoic acids in Lactobacillus reuteri 100–23 results in impaired colonization of the mouse gastrointestinal tract. Environ Microbiol 9:1750–1760
Foreman-Wykert AK, Weinrauch Y, Elsbach P, Weiss J (1999) Cell-wall determinants of the bactericidal action of group IIA phospholipase A2 against Gram-positive bacteria. J Clin Invest 103:715–721
Weiss J, Inada M, Elsbach P, Crowl RM (1994) Structural determinants of the action against Escherichia coli of a human inflammatory fluid phospholipase A2 in concert with polymorphonuclear leukocytes. J Biol Chem 269:26331–26337
Heptinstall S, Archibald AR, Baddiley J (1970) Teichoic acids and membrane function in bacteria. Nature 225:519–521
MacArthur AE, Archibald AR (1984) Effect of culture pH on the d-alanine ester content of lipoteichoic acid in Staphylococcus aureus. J Bacteriol 160:792–793
Hurst A, Hughes A, Duckworth M, Baddiley J (1975) Loss of d-alanine during sublethal heating of Staphylococcus aureus S6 and magnesium binding during repair. J Gen Microbiol 89:277–284
Koch HU, Doker R, Fischer W (1985) Maintenance of d-alanine ester substitution of lipoteichoic acid by reesterification in Staphylococcus aureus. J Bacteriol 164:1211–1217
Kiriukhin MY, Neuhaus FC (2001) d-alanylation of lipoteichoic acid: role of the d-alanyl carrier protein in acylation. J Bacteriol 183:2051–2058
Poyart C, Lamy MC, Boumaila C, Fiedler F, Trieu-Cuot P (2001) Regulation of d-alanyl-lipoteichoic acid biosynthesis in Streptococcus agalactiae involves a novel two-component regulatory system. J Bacteriol 183:6324–6334
Dunman PM, Murphy E, Haney S, Palacios D, Tucker-Kellogg G, Wu S, Brown EL, Zagursky RJ, Shlaes D, Projan SJ (2001) Transcription profiling-based identification of Staphylococcus aureus genes regulated by the agr and/or sarA loci. J Bacteriol 183:7341–7353
Said-Salim B, Dunman PM, McAleese FM, Macapagal D, Murphy E, McNamara PJ, Arvidson S, Foster TJ, Projan SJ, Kreiswirth BN (2003) Global regulation of Staphylococcus aureus genes by Rot. J Bacteriol 185:610–619
Herbert S, Bera A, Nerz C, Kraus D, Peschel A, Goerke C, Meehl M, Cheung A, Gotz F (2007) Molecular basis of resistance to muramidase and cationic antimicrobial peptide activity of lysozyme in Staphylococci. PLoS Pathogens 3:e102
Meehl M, Herbert S, Gotz F, Cheung A (2007) Interaction of the GraRS two-component system with the VraFG ABC transporter to support vancomycin-intermediate resistance in Staphylococcus aureus. Antimicrob Agents Chemother 51:2679–2689
Kraus D, Herbert S, Kristian SA, Khosravi A, Nizet V, Gotz F, Peschel A (2008) The GraRS regulatory system controls Staphylococcus aureus susceptibility to antimicrobial host defenses. BMC Microbiol 8:85
Li M, Cha DJ, Lai Y, Villaruz AE, Sturdevant DE, Otto M (2007) The antimicrobial peptide-sensing system aps of Staphylococcus aureus. Mol Microbiol 66:1136–1147
Otto M (2009) Bacterial sensing of antimicrobial peptides. Contrib Microbiol 16:136–149
Peschel A (2001) 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 193:1067–1076
Kuroda M, Kuwahara-Arai K, Hiramatsu K (2000) Identification of the up- and down-regulated genes in vancomycin-resistant Staphylococcus aureus strains Mu3 and Mu50 by cDNA differential hybridization method. Biochem Biophys Res Commun 269:485–490
Matsuzaki K (2009) Control of cell selectivity of antimicrobial peptides. Biochim Biophys Acta Biomembranes 1788:1687–1692
Staubitz P, Neumann H, Schneider T, Wiedemann I, Peschel A (2004) MprF-mediated biosynthesis of lysylphosphatidylglycerol, an important determinant in staphylococcal defensin resistance. FEMS Microbiol Lett 231:67–71
Ernst CM, Staubitz P, Mishra NN, Yang S-J, Hornig G, Kalbacher H, Bayer AS, Kraus D, Peschel A (2009) The Bacterial defensin resistance protein MprF consists of separable domains for lipid lysinylation and antimicrobial peptide repulsion. PLoS Pathog 5:e1000660
Friedman L, Alder JD, Silverman JA (2006) Genetic changes that correlate with reduced susceptibility to daptomycin in Staphylococcus aureus. Antimicrob Agents Chemother 50:2137–2145
Rubio A, Conrad M, Haselbeck R, Kedar GC, Driver V, Finn J, Silverman J (2011) Regulation of mprF by antisense restores daptomycin susceptibility to daptomycin-resistant isolates of Staphylococcus aureus. Antimicrob Agents Chemother 55:364–367
Yang S-J, Xiong YQ, Dunman PM, Schrenzel J, Francois P, Peschel A, Bayer AS (2009) Regulation of mprF in daptomycin-nonsusceptible Staphylococcus aureus strains. Antimicrob Agents Chemother 53:2636–2637
Straus SK, Hancock REW (2006) Mode of action of the new antibiotic for Gram-positive pathogens daptomycin: comparison with cationic antimicrobial peptides and lipopeptides. Biophys Biochim Acta Biomembranes 1758:1215–1223
Camargo ILBdC, Neoh H-M, Cui L, Hiramatsu K (2008) Serial daptomycin selection generates daptomycin-nonsusceptible Staphylococcus aureus strains with a heterogeneous vancomycin-intermediate phenotype. Antimicrob Agents Chemother 52:4289–4299
Ruzin A, Severin A, Moghazeh SL, Etienne J, Bradford PA, Projan SJ, Shlaes DM (2003) Inactivation of mprF affects vancomycin susceptibility in Staphylococcus aureus. Biochim Biophys Acta 1621:117–121
Nishi H, Komatsuzawa H, Fujiwara T, McCallum N, Sugai M (2004) 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 48:4800–4807
Gunn JS, Miller SI (1996) PhoP–PhoQ activates transcription of pmrAB, encoding a two-component regulatory system involved in Salmonella typhimurium antimicrobial peptide resistance. J Bacteriol 178:6857–6864
Gunn JS, Lim KB, Krueger J, Kim K, Guo L, Hackett M, Miller SI (1998) PmrA–PmrB-regulated genes necessary for 4-aminoarabinose lipid A modification and polymyxin resistance. Mol Microbiol 27:1171–1182
Gunn JS (2001) Bacterial modification of LPS and resistance to antimicrobial peptides. J Endotxin Res 7:57–62
Lee H, Hsu F-F, Turk J, Groisman EA (2004) The PmrA-regulated pmrC gene mediates phosphoethanolamine modification of lipid A and polymyxin resistance in Salmonella enterica. J Bacteriol 186:4124–4133
Groisman EA, Heffron F, Solomon F (1992) Molecular genetic analysis of the Escherichia coli phoP locus. J Bacteriol 174:486–491
Groisman EA, Chiao E, Lipps CJ, Heffron F (1989) Salmonella typhimurium phoP virulence gene is a transcriptional regulator. Proc Natl Acad Sci USA 86:7077–7081
Garcia Vescovi E, Soncini FC, Groisman EA, Vescovi EG, Ayala YM, Di Cera E (1996) Mg2+ as an extracellular signal: environmental regulation of Salmonella virulence. Cell 84:165–174
Chamnongpol S, Cromie M, Groisman EA (2003) Mg2+ sensing by the Mg2+ sensor PhoQ of Salmonella enterica. J Mol Biol 325:795–807
Vescovi EG, Ayala YM, Di Cera E, Groisman EA (1997) Characterization of the bacterial sensor protein PhoQ. Evidence for distinct binding sites for Mg2+ and Ca2+. J Biol Chem 272:1440–1443
Alpuche Aranda CM, Swanson JA, Loomis WP, Miller SI (1992) Salmonella typhimurium activates virulence gene transcription within acidified macrophage phagosomes. Proc Natl Acad Sci USA 89:10079–10083
Bader MW, Sanowar S, Daley ME, Schneider AR, Cho U, Xu W, Klevit RE, Le Moual H, Miller SI (2005) Recognition of antimicrobial peptides by a bacterial sensor kinase. Cell 122:461–472
Charles RC, Harris JB, Chase MR, Lebrun LM, Sheikh A, LaRocque RC, Logvinenko T, Rollins SM, Tarique A, Hohmann EL, Rosenberg I, Krastins B, Sarracino DA, Qadri F, Calderwood SB, Ryan ET (2009) Comparative proteomic analysis of the phoP regulon in Salmonella enterica serovar typhi versus typhimurium. PLoS ONE 4:e6994
Prost LR, Sanowar S, Miller SI (2007) Salmonella sensing of anti-microbial mechanisms to promote survival within macrophages. Immunol Rev 219:55–65
Guo L, Lim KB, Gunn JS, Bainbridge B, Darveau RP, Hackett M, Miller SI (1997) Regulation of lipid A modifications by Salmonella typhimurium virulence genes phoP–phoQ. Science 276:250–253
Miller SI, Pulkkinen WS, Selsted ME, Mekalanos JJ (1990) Characterization of defensin resistance phenotypes associated with mutations in the phoP virulence regulon of Salmonella typhimurium. Infect Immun 58:3706–3710
Prost LR, Daley ME, Bader MW, Klevit RE, Miller SI (2008) The PhoQ histidine kinases of Salmonella and Pseudomonas spp. are structurally and functionally different: evidence that pH and antimicrobial peptide sensing contribute to mammalian pathogenesis. Mol Microbiol 69:503–519
Bayer AS, Prasad R, Chandra J, Koul A, Smriti M, Varma A, Skurray RA, Firth N, Brown MH, Koo S-P, Yeaman MR (2000) In Vitro resistance of Staphylococcus aureus to thrombin-induced platelet microbicidal protein is associated with alterations in cytoplasmic membrane fluidity. Infect Immun 68:3548–3553
Mishra NN, Liu GY, Yeaman MR, Nast CC, Proctor RA, McKinnell J, Bayer AS (2011) Carotenoid-related alteration of cell membrane fluidity impacts Staphylococcus aureus susceptibility to host defense peptides. Antimicrob Agents Chemother 55:526–531
Bengoechea JA, Skurnik M (2000) Temperature-regulated efflux pump/potassium antiporter system mediates resistance to cationic antimicrobial peptides in Yersinia. Mol Microbiol 37:67–80
Tzeng Y-L, Ambrose KD, Zughaier S, Zhou X, Miller YK, Shafer WM, Stephens DS (2005) Cationic antimicrobial peptide resistance in Neisseria meningitidis. J Bacteriol 187:5387–5396
Shafer WM, Qu X, Waring AJ, Lehrer RI (1998) Modulation of Neisseria gonorrhoeae susceptibility to vertebrate antibacterial peptides due to a member of the resistance/nodulation/division efflux pump family. Proc Natl Acad Sci USA 95:1829–1833
Rieg S, Huth A, Kalbacher H, Kern WV (2008) Resistance against antimicrobial peptides is independent of Escherichia coli AcrAB, Pseudomonas aeruginosa MexAB and Staphylococcus aureus NorA efflux pumps. Int J Antimicrob Ag 33:174–176
Bayer AS, Kupferwasser LI, Brown MH, Skurray RA, Grkovic S, Jones T, Mukhopadhay K, Yeaman MR (2006) Low-level resistance of Staphylococcus aureus to thrombin-induced platelet microbicidal protein 1 In Vitro associated with qacA gene carriage is independent of multidrug efflux pump activity. Antimicrob Agents Chemother 50:2448–2454
May JJ, Finking R, Wiegeshoff F, Weber TT, Bandur N, Koert U, Marahiel MA (2005) Inhibition of the d-alanine:d-alanyl carrier protein ligase from Bacillus subtilis increases the bacterium’s susceptibility to antibiotics that target the cell wall. FEBS 272:2993–3003
Peschel A (2002) How do bacteria resist human antimicrobial peptides? Trend Microbiol 10:179–186
Jin T, Bokarewa M, McIntyre L, Tarkowski A, Corey GR, Reller LB, Fowler VG Jr (2003) Fatal outcome of bacteraemic patients caused by infection with staphylokinase-deficient Staphylococcus aureus strains. J Med Microbiol 52:919–923
Kwiecinski J, Josefsson E, Mitchell J, Higgins J, Magnusson M, Foster T, Jin T, Bokarewa M (2010) Activation of plasminogen by staphylokinase reduces the severity of Staphylococcus aureus systemic infection. J Infect Dis 202:1041–1049
Belas R, Manos J, Suvanasuthi R (2004) Proteus mirabilis ZapA metalloprotease degrades a broad spectrum of substrates, including antimicrobial peptides. Infect Immun 72:5159–5167
Johansson L, Thulin P, Sendi P, Hertzen E, Linder A, Akesson P, Low DE, Agerberth B, Norrby-Teglund A (2008) Cathelicidin LL-37 in severe Streptococcus pyogenes soft tissue infections in humans. Infect Immun 76:3399–3404
Woods DE, Cryz SJ, Friedman RL, Iglewski BH (1982) Contribution of toxin A and elastase to virulence of Pseudomonas aeruginosa in chronic lung infections of rats. Infect Immun 36:1223–1228
Blackwood LL, Stone RM, Iglewski BH, Pennington JE (1983) Evaluation of Pseudomonas aeruginosa exotoxin A and elastase as virulence factors in acute lung infection. Infect Immun 39:198–201
Rupp ME, Ulphani JS, Fey PD, Bartscht K, Mack D (1999) Characterization of the importance of polysaccharide intercellular adhesin/Hemagglutinin of Staphylococcus epidermidis in the pathogenesis of biomaterial-based infection in a mouse foreign body infection model. Infect Immun 67:2627–2632
Rupp ME, Fey PD, Heilmann C, Gotz F (2001) Characterization of the importance of Staphylococcus epidermidis autolysin and polysaccharide intercellular adhesin in the pathogenesis of intravascular catheter-associated infection in a rat model. J Infect Dis 183:1038–1042
Jones A, Georg M, Maudsdotter L, Jonsson A-B (2009) Endotoxin, capsule, and bacterial attachment contribute to Neisseria meningitidis resistance to the human antimicrobial peptide LL-37. J Bacteriol 191:3861–3868
Yi K, Stephens DS, Stojiljkovic I (2003) Development and evaluation of an improved mouse model of meningococcal colonization. Infect Immun 71:1849–1855
Spinosa MR, Progida C, Tala A, Cogli L, Alifano P, Bucci C (2007) The Neisseria meningitidis capsule is important for intracellular survival in human cells. Infect Immun 75:3594–3603
Cortes G, Borrell N, de Astorza B, Gomez C, Sauleda J, Alberti S (2002) Molecular analysis of the contribution of the capsular polysaccharide and the lipopolysaccharide O side chain to the virulence of Klebsiella pneumoniae in a murine model of pneumonia. Infect Immun 70:2583–2590
Campos MA, Vargas MA, Regueiro V, Llompart CM, Alberti S, Bengoechea JA (2004) Capsule polysaccharide mediates bacterial resistance to antimicrobial peptides. Infect Immun 72:7107–7114
Llobet E, Tomas JM, Bengoechea JA (2008) Capsule polysaccharide is a bacterial decoy for antimicrobial peptides. Microbiology 154:3877–3886
Weidenmaier C, Kokai-Kun JF, Kristian SA, Chanturiya T, Kalbacher H, Gross M, Nicholson G, Neumeister B, Mond JJ, Peschel A (2004) Role of teichoic acids in Staphylococcus aureus nasal colonization, a major risk factor in nosocomial infections. Nat Med 10:243–245
Kristian SA, Datta V, Weidenmaier C, Kansal R, Fedtke I, Peschel A, Gallo RL, Nizet V (2005) d-alanylation of teichoic acids promotes group a streptococcus antimicrobial peptide resistance, neutrophil survival, and epithelial cell invasion. J Bacteriol 187:6719–6725
Kristian SA, Durr M, Van Strijp JAG, Neumeister B, Peschel A (2003) MprF-mediated lysinylation of phospholipids in Staphylococcus aureus leads to protection against oxygen-independent neutrophil killing. Infect Immun 71:546–549
Gunn JS, Ryan SS, Van Velkinburgh JC, Ernst RK, Miller SI (2000) Genetic and functional analysis of a PmrA–PmrB-regulated locus necessary for lipopolysaccharide modification, antimicrobial peptide resistance, and Oral virulence of Salmonella enterica serovar typhimurium. Infect Immun 68:6139–6146
Zhou Z, Ribeiro AA, Lin S, Cotter RJ, Miller SI, Raetz CRH (2001) Lipid A modifications in polymyxin-resistant Salmonella typhimurium. J Biol Chem 276:43111–43121
Belden WJ, Miller SI (1994) Further characterization of the PhoP regulon: identification of new PhoP-activated virulence loci. Infect Immun 62:5095–5101
Guo L (1998) Lipid A acylation and bacterial resistance against vertebrate antimicrobial peptides. Cell 95:189–198
Pelz A, Wieland K-P, Putzbach K, Hentschel P, Albert K, Gotz F (2005) Structure and biosynthesis of staphyloxanthin from Staphylococcus aureus. J Biol Chem 280:32493–32498
Clauditz A, Resch A, Wieland K-P, Peschel A, Gotz F (2006) Staphyloxanthin plays a role in the fitness of Staphylococcus aureus and its ability to cope with oxidative stress. Infect Immun 74:4950–4953
Liu GY, Essex A, Buchanan JT, Datta V, Hoffman HM, Bastian JF, Fierer J, Nizet V (2005) Staphylococcus aureus golden pigment impairs neutrophil killing and promotes virulence through its antioxidant activity. J Exp Med 202:209–215
Kupferwasser LI, Skurray RA, Brown MH, Firth N, Yeaman MR, Bayer AS (1999) Plasmid-mediated resistance to thrombin-induced platelet microbicidal protein in Staphylococci: role of the qacA locus. Antimicrob Agents Chemother 43:2395–2399
Warner DM, Folster JP, Shafer WM, Jerse AE (2007) Regulation of the MtrC–MtrD–MtrE efflux-pump system modulates the in vivo fitness of Neisseria gonorrhoeae. J Infect Dis 196:1804–1812
Jerse AE, Sharma ND, Simms AN, Crow ET, Snyder LA, Shafer WM (2003) A gonococcal efflux pump system enhances bacterial survival in a female mouse model of genital tract infection. Infect Immun 71:5576–5582
Acknowledgments
Our work is supported in part by Grants from Marie Curie International Grant 249285 to TK, the German Research Foundation (SFB766, TRR34) and the German Ministry of Education and Research (SkinStaph, MENAGE) to AP.
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Koprivnjak, T., Peschel, A. Bacterial resistance mechanisms against host defense peptides. Cell. Mol. Life Sci. 68, 2243–2254 (2011). https://doi.org/10.1007/s00018-011-0716-4
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DOI: https://doi.org/10.1007/s00018-011-0716-4