Defensive Remodeling: How bacterial surface properties and biofilm formation promote resistance to antimicrobial peptides Reut Nuri, Tal Shprung, Yechiel Shai PII: DOI: Reference:
S0005-2736(15)00179-0 doi: 10.1016/j.bbamem.2015.05.022 BBAMEM 81917
To appear in:
BBA - Biomembranes
Received date: Revised date: Accepted date:
10 February 2015 25 May 2015 26 May 2015
Please cite this article as: Reut Nuri, Tal Shprung, Yechiel Shai, Defensive Remodeling: How bacterial surface properties and biofilm formation promote resistance to antimicrobial peptides, BBA - Biomembranes (2015), doi: 10.1016/j.bbamem.2015.05.022
This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
PT
ACCEPTED MANUSCRIPT
SC
MA
Reut Nuri, Tal Shprung and Yechiel Shai*
NU
promote resistance to antimicrobial peptides
RI
Defensive Remodeling: How bacterial surface properties and biofilm formation
AC CE P
Rehovot, 76100 Israel.
TE
D
Department of Biological Chemistry, the Weizmann Institute of Science,
*To whom correspondence should be addressed, at the Department of Biological Chemistry, The Weizmann Institute of Science, Rehovot, 76100 Israel. Tel: 972-8-9342711; Fax: 972-89344112; E-mail: [email protected]
Key words Antimicrobial peptides, resistance, surface, biophysical properties, biochemical properties, biofilm, cross resistance.
1
ACCEPTED MANUSCRIPT Abstract .....................................................................................................................................................................................4
2.
Introduction ............................................................................................................................................................................4
3.
Bacterial resistance to AMPs as a result of changes in bacterial outer surface ..........................................6
PT
1.
Extracellular polysaccharides ................................................................................................................................7
3.2
Lipid modifications in bacterial membranes ...................................................................................................8
RI
3.1
Phosphatidylglycerol (PG) modifications ......................................................................................... 8
3.2.2
Changing the fatty acids composition ................................................................................................ 9
3.2.3
In-Vitro studies .......................................................................................................................................... 10
NU
SC
3.2.1
LTA D-alanylation of LTA in Gram-positive bacteria ................................................................................ 10
3.4
Surface modifications of Gram-negative bacteria....................................................................................... 11
MA
3.3
3.4.1
Acylation and deacylation of Lipid A ................................................................................................ 12
3.4.2
Addition of glycine to Lipid A .............................................................................................................. 13
D
3.4.3 De-phosphorylation, hydroxylation and addition of Phosphoethanolamine (PE) to Lipid A 14 Addition of amino-sugars to lipid A .................................................................................................. 15
3.4.5
Changing the length of O-antigen....................................................................................................... 16
TE
3.4.4
Modifications of the bacterial outer surface that influence peptide properties. ..................................... 17
5.
Mechanisms of bacterial AMP resistance in biofilm ............................................................................................ 17
6.
AC CE P
4.
5.1
Creation of sub-populations in the biofilm- .................................................................................................. 18
5.2
A higher mutation rate- ......................................................................................................................................... 19
5.3
Upregulation of efflux pumps - ........................................................................................................................... 19
5.4
Induction of TCSs within the biofilm- .............................................................................................................. 19
5.5
Diffusion in biofilm- ................................................................................................................................................ 20
Two component systems (TCSs) regulate AMP resistance .............................................................................. 20 6.1
TCS activation by AMPs ......................................................................................................................................... 21
6.2
Genes activated by TCS that are involved in AMP resistance ................................................................ 22
7.
Ways to overcome AMP resistance ............................................................................................................................ 23
8.
Summary ............................................................................................................................................................................... 25 2
ACCEPTED MANUSCRIPT
TE
D
MA
NU
SC
RI
PT
References............................................................................................................................................................................. 27
AC CE P
9.
3
ACCEPTED MANUSCRIPT
1. Abstract
PT
Multidrug resistance bacteria are a major concern worldwide. These pathogens cannot be treated
RI
with conventional antibiotics and thus alternative therapeutic agents are needed. Antimicrobial
SC
peptides (AMPs) are considered to be good candidates for this purpose. Most AMPs are short and positively charged amphipathic peptides, which are found in all forms of life. AMPs are
NU
known to kill bacteria by binding to the negatively charged bacterial surface, and in most cases cause membrane disruption. Resistance towards AMPs can be developed, by modification of
MA
bacterial surface molecules, secretion of protective material and up-regulation or elimination of specific proteins. Because of the general mechanisms of attachment and action of AMPs,
TE
D
bacterial resistance to AMPs often involves biophysical and biochemical changes such as surface rigidity, cell wall thickness, surface charge, as well as membrane and cell wall modification.
AC CE P
Here we focus on the biophysical, surface and surrounding changes that bacteria undergo in acquiring resistance to AMPs. In addition we discuss the question of whether bacterial resistance to administered AMPs might compromise our innate immunity to endogenous AMPs.
2. Introduction
The fight between bacteria and man, once thought to be at its end when antibiotics were discovered, has gained much attention in recent years. The enhanced and unregulated use of antimicrobials has increased the occurrence of bacteria resistant to multiple antibiotics [1, 2], thus urging the research community to develop new antibacterial agents. One of the most promising strategies explored is the use of antimicrobial peptides (AMPs) [3-6]. AMPs are hydrophobic and amphipathic short peptides, which in most cases have a net positive charge [4, 4
ACCEPTED MANUSCRIPT 7]. Initially AMPs were found in invertebrates [3] and later in all known forms of life including humans [8, 9], where they serve as part of our innate immune system [5, 10, 11]. AMPs rapidly
PT
neutralize a broad range of microbes, both Gram-positive and Gram-negative, mainly through
RI
binding and perturbing their cell envelope, which contains lipoteichoic acid (LTA) or lipopolysaccharide (LPS) respectively, as well as their cytoplasmic membranes [5, 6, 12]. This
SC
makes AMPs promising candidates in the quest to find new and better antimicrobial agents [13].
NU
It is believed that the chemical and physical characteristics of AMPs such as a positive net charge secondary structure and amphipathic nature are important for their antimicrobial
MA
activity [6, 14-16]. For example, in the case of Gram-negative bacteria, due to their net positive charge, AMPs have high affinity to the negatively charged LPS, and then can displace divalent
TE
D
cations that are naturally bound to it. However, it has been shown that some AMPs specifically bind oprI, a lipoprotein of Pseudomonas aeruginosa, located on the outer membrane and
AC CE P
suggested to serve as an AMP receptor. OprI is important for cell wall integrity and has homologs in other Gram-negative bacteria [17]. The binding of AMPs to the bacterial surface causes a local interference with the outer membrane and enables peptides to traverse it. When reaching the inner membrane, AMPs can function either without causing membrane disruption by penetrating the cell and affecting cytoplasmic components such as DNA and DNA-polymerase [18]. Alternatively/in addition, AMPs can function in membrane disruptive manner thus impairing cell integrity [18]. Various mechanisms were suggested for the membrane disruptive activity of AMPs [5, 6, 10, 19]. The main models include: (1) the barrel-stave model - peptides assemble to form pores with their hydrophobic surfaces facing the membrane core [20]; (2) the carpet model – peptides bind to the membrane in a carpet-like manner. When a threshold concentration has been reached, they act as
5
ACCEPTED MANUSCRIPT a detergent and disrupt the membrane [10, 21] and (3) the toroidal model - peptides form transient pores composed of peptides and lipids head groups [22, 23]. These pores were
PT
suggested to be as one of the stages in the carpet mechanism [5].
RI
Bacteria however, have evolved mechanisms to counteract AMPs. There are two main types of resistance, (i) induced resistance, caused by a transient induction of bacterial systems
SC
that help the bacteria cope with AMPs, and (ii) constitutive resistance as a result of genetic
NU
changes [24, 25]. Currently, there are several putative mechanisms known for bacterial resistance to AMPs [26-29]: remodeling of the bacterial surface by making it less negatively charged and
MA
less permeable [30-35], modulation of host immunity gene expression in order to suppress the release of native AMPs [36, 37], biofilm formation [38, 39], proteolytic degradation of AMPs
TE
D
by peptidases and proteases [40, 41], efflux pumps that expel the AMPs [27, 42, 43], and trapping molecules [44-46] and vesicles [47] that capture the peptides and prevent them from
AC CE P
reaching the bacteria.
In this review we focus on the mechanisms that affect the biophysical characteristics of the bacteria and bacterial surrounding such as capsid and biofilm, as well as biochemical changes of the outer surface, including the regulation of some of these processes by two component systems (TCSs).
3. Bacterial resistance to AMPs as a result of changes in bacterial outer surface The outer surface of Gram-positive and Gram-negative bacteria is negatively charged due to LTA and LPS, respectively. AMPs on the other hand, are hydrophobic, positively charged molecules, and therefore it is believed that AMPs are initially attracted to the bacterial surface 6
ACCEPTED MANUSCRIPT due to electrostatic interactions. In line with this, it is assumed that neutralization of the negative charge can reduce peptide binding to the bacterial surface, and will be further discussed. Note
Extracellular polysaccharides
RI
3.1
PT
that everal reviews were published on this subject [26, 48-51].
SC
Extracellular polysaccharides are polymers produced by some Gram-positive and Gram-negative
NU
bacteria. When covalently linked to the cell wall they are termed capsule [28]. The question of whether or not a capsule in this strain contributes to AMP resistance is still under debate [51].
MA
There are some reports indicating that capsules cause AMP-resistance [51-53]. For example, Klebsiella pneumoniae bacteria mutated in capsule production are more susceptible to AMPs and
D
bind better polymyxin B (PmB) compared to the wild-type (WT) [54]. Moreover, PmB was
TE
found to induce capsule production in these bacteria, and it was suggested that the capsule serves
AC CE P
as a shield towards AMPs, thus limiting their interaction with the bacterial surface [55]. In addition, it was shown that K. pneumonia, Streptococcus pneumonia and P. aeruginosa tend to naturally release capsule polysaccharides (CPS) fractions into the environment, which contributes to their resistance to AMPs. This secretion is enhanced in the presence of PmB. Supernatant containing CPS of these strains showed increased resistance in Escherichia coli strain [54]. Moreover, the authors demonstrated that anionic CPS but not cationic or neutral charged CPS of Streptococcus pneumonia, specifically bind dansyl-PmB [54]. In Group B Streptococcus it was demonstrated that bacteria mutated in capsule production did not demonstrate a higher susceptibility to several tested AMPs [35]. To conclude, some studies have demonstrated AMP resistance mediated by capsules and some show that capsules do not change the resistance. We suggest therefore that whether or not a capsule confers AMP resistance depends on the bacterial species, and perhaps on the AMP itself. 7
ACCEPTED MANUSCRIPT 3.2
Lipid modifications in bacterial membranes Phosphatidylglycerol (PG) modifications
PT
3.2.1
In Gram-positive bacteria, a gene encodes for a large membrane protein mprF responsible for the
RI
addition of lysine or alanine to phosphatidylglycerol (PG) to form lysyl-phosphatidylglycerol
SC
(LPG) and alanyl- phosphatidylglycerol (APG), respectively [56], as well as for the translocation of these products to the outer leaflet [56-58]. In several studies, mprF mutants of S. aureus were
NU
found to be more susceptible to AMPs, which might suggest that the addition of these amino
MA
acids to the membrane reduces susceptibility to AMPs [59] (A detailed review regarding AMP resistance mediated by mprF can be found in reference [60]). However, in another study it was
D
shown that there was no change in susceptibility of S. aureus mutants to mprF for the tested
TE
peptides [35].
AC CE P
The membrane of S. aureus consists mostly of unmodified PG (38-76% of the phospholipids), its dimer, diphosphatidylglycerol (DPG) also known as cardiolipin (5-30%) and about 14-38% of LPG, [58] (Figure 1). Some daptomycin (a lipopeptide) resistant S. aureus isolated strains, showed gain of function mutations in mprF and had a higher LPG content compared to the susceptible strains. These strains were found to be more resistant to thrombininduced platelet microbicidal proteins (tPMP), and showed a reduced net surface charge [61]. In line with that, less daptomycin was bound to their surface compared to the other strains, however the binding of classical AMPs was not examined in this study [61]. In the Gram-negative bacteria P. aeruginosa, a homolog for MprF was found, which is responsible for the addition of alanine to PG to form APG. This modification resulted in increased resistance to antimicrobial agents including the AMP protamine sulfate [62].
8
ACCEPTED MANUSCRIPT 3.2.2
Changing the fatty acids composition
It was suggested that membrane lipid composition can affect cell wall properties and change
PT
interactions with AMPs, and therefore the susceptibility to these agents [63]. S. aureus strains
RI
resistant to thrombin-induced platelet microbicidal protein 1 (tPMP-1), showed a higher
SC
proportion of longer and more unsaturated fatty acids in the membrane, as well as higher membrane fluidity compared to the control, but no differences in phospholipid composition [64].
NU
Daptomycin resistant S. aureus isolated strains also showed a higher membrane fluidity
MA
compared to the control [65-67]. Nevertheless, one study reports on a correlation between membrane fluidity and the level of resistance to AMPs [67]. Conversely, other AMP-resistant
D
bacteria showed a higher rigidity. Two S. aureus strains, one resistant to magainin 2 and the
TE
other to gramicidin D, had a higher proportion of LPG in their cell membrane compared to the control strain and both mutants showed higher rigidity and surface positive charge [68]. E.
AC CE P
faecalis pediocin-resistant strains showed a higher ratio of saturated and unsaturated fatty acids, as well as, reduction in branched fatty acids in these strains compared to the WT. These changes in fatty acid composition increase cell membrane rigidity, and are thought to reduce the insertion of pediocin (AMP) to the membrane [69]. In summary, membrane fluidity is not a direct indication for resistance to AMPs, and cell membrane can be more or less fluid in the context of the organism and the AMP. Comparison of nisin resistant and susceptible strains of Listeria monocytogenes revealed that the ratio of PG and DPG was higher than in the WT. An in-vitro nisin binding assay demonstrated that nisin binds better to a lipid-monolayer derived from the WT than the resistant strain [70]. Other studies reported that nisin resistant L. monocytogenes strains had a greater proportion of saturated fatty acids whereas the WT had more unsaturated ones [71]. In 9
ACCEPTED MANUSCRIPT Enterococcus faecalis, a Gram-positive bacterium, pediocin-resistant strains had more LPG and phosphoethanolamine (PEA) in their cell membrane compared to the WT [69]. AMP resistance
PT
in Salmonella enterica Typhimurium, a Gram-negative bacterium, was found to be mediated by
RI
increased levels of DPG and palmitoylated acyl-phosphatidylglyceroles in the inner leaflet of the outer membrane, which is mainly composed of glycerophospholipids. This palmitoylation is
In-Vitro studies
NU
3.2.3
SC
conducted by the protein pagP and regulated by the two-component system PhoPQ [72].
MA
An in- vitro study examined the effect of binding and pore formation ability of AMPs on large unilamellar vesicle (LUVs) composed of different proportion of lipids. LUVs were generated
D
containing 1-palmitoyl-2-oleoyl-phosphatidylglycerol (POPG) and different concentrations of
TE
either zwitterionic Gln-PE or positively charged Lys-PE. It was shown that while increasing the
AC CE P
concentration of the Gln-PE in the membrane does not change the level of bound cecropin A and mastoparan X (AMPs), the increase of the Lys-PE in the membrane reduced the binding. Pore formation assay revealed that increasing concentration of Lys-PE, starting with 10%, reduced the levels of pore formation by both cecropin A and mastoparan X, while 10% of Gln-PE increased pore formation. In both peptides however, higher concentrations resulted in reduced pore forming ability [73].
3.3
LTA D-alanylation of LTA in Gram-positive bacteria
The surface of Gram-positive bacteria consists of the cytoplasmic membrane, a thick peptidoglycan layer and lipoteichoic acid (LTA). It has been reported that D-alanylation of the LTA contributes to bacterial resistance to AMPs [74-76]. It was found that the dlt operon is responsible for the incorporation of D-alanine (D-ala) into teichoic acid, which results in an 10
ACCEPTED MANUSCRIPT increase in the positive charge of the bacterial surface. Mutants of dltA, the gene encoding a Dalanyl carrier protein ligase (that adds D-alanine to the Gro-P moiety of the LTA) showed
PT
increased susceptibility to AMPs, whereas overexpression of this gene confers increased
RI
resistance [77] (Figure 2, Table 1).
Reducing the net negative charge of the LTA is considered to contribute to the reduction of
SC
electrostatic interactions between the positively charged AMPs and the negatively charged
NU
bacterial surface, thus reducing susceptibility [34, 76, 78, 79]. However, recent experimental data on Group B Streptococus (GBS) indicate that there is no significant change in the amount of
MA
peptide bound to the dltA mutant bacteria compared to the WT. It was proposed that the higher resistance observed in the WT compared to the dltA mutant is due to an increase in cell wall
TE
D
density and reduced AMP penetration [35], i.e. the addition of D-alanine to Gro-P moiety of the LTA does not change the amount of bound peptide, but rather serves as a shield that prevents
AC CE P
peptides from entering the bacteria. No difference in cell wall thickness was found in this study [35] as well as in other studies [35, 61, 80]. However, several studies showed a thicker cell wall of different AMP- resistant bacteria [81, 82], which suggests that cell wall thickness might be, but is not necessarily, correlated with resistance. Importantly, the D-alanylation of the LTA contributes to the resistance of bacteria to only some AMPs [35] and so far, other known modifications of the LTA (e.g. the addition of phosphocholine and glycosylation) were not tested for their effect on resistance to antimicrobial peptides [83].
3.4
Surface modifications of Gram-negative bacteria
The surface of Gram-negative bacteria is complex and covered with several constituents such as LPS, proteinaceous curli fibers, cellulose and capsules [84-88]. LPS, which is thought to be the main surface element, consists of a conserved lipophilic component, lipid-A, linked with 11
ACCEPTED MANUSCRIPT different polysaccharides or oligosaccharides, giving it a net negative charge. The LPS composition is dynamic; it may be modified and is found to be diverse in length and composition
PT
amongst the different Gram-negative bacteria species [87]. This unique structure, besides helping
RI
to maintain cellular equilibrium, serves as a physical barrier and may provide protection against antibacterial agents [19, 89]. Some LPS modifications were shown to contribute to bacterial
SC
resistance to AMPs. The modifications of the LPS depend on bacterial species and
NU
environmental conditions, and it was shown that not all modifications are found in all bacteria species [71]. Here we will discuss some of the changes in the outer surface with respect to AMP
MA
resistance and susceptibility. Lipid A and inner core modifications that are known to affect AMP
Acylation and deacylation of Lipid A
TE
3.4.1
D
resistance are summarized in Figure 3 and Table 1.
AC CE P
Addition of an acyl chain to Lipid A was found to contribute to bacterial resistance to AMPs. PagP is a gene that encodes for an outer membrane palmitoyl transferase in S. enteric Typhimurium [72]. This enzyme adds additional palmitate (C16:0 acyl chain) to the lipid A moiety and this acylation is thought to increase the hydrophobic interactions between the acyl chains in the Lipid A, thus increasing outer membrane fluidity [90-92] (Figure 3). S. enterica Typhimurium mutants in pagP showed higher membrane permeability in response to AMPs compared to the control strain, as well as, Legionella pneumophila mutants in Rcp, a PagP homolog that showed a higher susceptibility to C18G and PmB [93] (Figure 3). To support the finding that lipid A acylation contributes to bacterial resistance to AMPs, deacylation was found to increase bacteria susceptibility to AMPs. PagL is an enzyme responsible for deacylation of lipid A. It is located on the outer membrane of several Gramnegative bacteria such as S. enterica Typhimurium, and removes R-3-hydroxymyristate from 12
ACCEPTED MANUSCRIPT position 3 of some lipid A precursors [94]. It was found that pagL is expressed upon PhoPQ activation but no deacylation of lipid A was detected under these conditions. Moreover, it was
PT
demonstrated that pagL is latent in the WT, and the regulation of the latency involves
RI
aminoarabinose modifications of the lipid A regulated by pmrA. Therefore, the pagL null mutation did not change cell permeability to ethidium compared to the WT [94]. However,
SC
permeability of pmrA-pagL double mutant was lower compared to pmrA mutant, that lacks
NU
aminoarabinose modifications on the lipid A, which indicates that deacylation by PagL increases the permeability of the membrane. In addition, it was shown that a WT strain containing a non-
MA
latent pagL vector possesses an increased susceptibility to various antimicrobial agents compared to the WT [95, 96]. However, investigating the effect of pmrA/pagL and pmrc/pagL double
TE
D
mutants on the susceptibility to PmB showed increased susceptibility to this AMP in the double mutants compared to pmrA and pmrC single mutants [97]. pagL mutant did not show increased
AC CE P
susceptibility compared to the WT, which might suggest that deacylation of lipid A is important to PmB resistance under certain conditions [97].
3.4.2
Addition of glycine to Lipid A
It was found that AlmEFG operon is involved in glycine and diglycine addition to the LPS of the Gram-negative bacteria Vibrio cholera. This modification confers polymyxin resistance in these species. The AlmEFG operon has some sequence homology with complexes involved in Dalanylation in Gram-positive bacteria [29, 98]. This modification process requires three proteins, AlmE activates glycine and transfers it to the carrier protein AlmF. The glycine is then transferred to AlmG which in turn transfers the glycine to the lipid A [29]. However, it was not yet reported how this modification confers Vibrio cholera resistance to polymyxin, and whether this system is present in other Gram-negative bacteria (Figure 3). 13
ACCEPTED MANUSCRIPT 3.4.3
De-phosphorylation, hydroxylation and addition of Phosphoethanolamine (PE) to Lipid A
PT
Studies have shown that de-phosphorylation of lipid A contributes to resistance against AMPs. It
RI
was shown that Helicobacter pylori possesses a protein called LpxE that is responsible for the
SC
de-phosphorylation of lipid A at position 1. This de-phosphorylation allows the addition of phosphoethanolamine (PEA) by EptA (PmrC), a PEA transferase. In addition, LpxF was found to
NU
dephosphorylate lipid A at position 4 leaving an unmodified hydroxyl group [99]. LpxE and
MA
LpxF mutants were shown to possess a higher susceptibility to PmB. In addition, a lipid A fully phosphorylated at positions 1 and 4 was shown to bind PmB more efficiently, rendering these
D
bacteria more susceptible to PmB [99]. It was initially found that the LPS of PmB resistant E.
TE
coli contain two to three fold higher amounts of PEA compared to the WT [100]. Addition of PEA to the LPS is also thought to change the net negative charge of the LPS and by that
AC CE P
contributes to the electrostatic repulsion of the positively charged AMPs [27]. PEA attachment can occur either on the phosphate group of the lipid A or of the inner core oligosaccharides (kdo). In E. coli and S. enterica Typhimurium, these modifications are conducted by two distinct enzymes. A pmrA-regulated PmrC (EptA) is responsible for the PEA incorporation into the lipid A [101], whilst the enzyme EptB couples the PEA moiety to the 3-deoxy-D-manno-octulosonic acid (Kdo) of the inner core polysaccharide [102]. However, EptB was not reported to be involved in antimicrobial resistance, and turned out to be regulated by other proteins rather than PmrA [103] (Figure 3). In other Gram-negative bacteria such as Neisseria gonorrhoeae and meningococcal bacteria, the gene lptA was found to encode for a protein with a similar activity as PmrC [104, 105]. Mutants in the pmrC of S. enterica Typhimurium and in lptA of N. gonorrhoeae were more 14
ACCEPTED MANUSCRIPT sensitive to PmB compared to the WT [105]. In addition, an in vivo study with N. gonorrhoeae showed that the addition of PEA to LPS confers a survival advantage compared to lptA mutant in
PT
human and rabbit [106]. It was found that pmrA mutants, that lack the aminoarabinose and PEA
RI
modifications on the lipid A, are more permeable to ethidium, which suggests that these modifications are involved in generating a robust permeation barrier [96]. However PmrA also
SC
regulates other genes that might affect membrane permeability. Hydroxylation of the myristate at
NU
position 2 on the lipid A in S. enterica Typhimurium by LpxO was shown to reduce permeability of the membrane and suggests that this hydroxylation can increase the hydrogen bond between
MA
neighboring molecules of LPS, which contributes to the outer membrane stabilization [107]
Addition of amino-sugars to lipid A
TE
3.4.4
D
(Figure 3).
AC CE P
There are several amino–sugars that are added to the lipid A moiety and are known to contribute to AMP resistance. These include (i) aminoarabinose, (ii) glucosamine, and (iii) galactoseamine (Figure 3).
(i) Addition of 4-amino-4-deoxy-L-arabinose (L-Ara4) to the phosphate group of the Lipid A correlated with acquired resistance to AMPs. L-Ara4 addition is done by ArnT, an enzyme regulated by PmrAB TCS [108]. The addition of L-Ara4 causes reduction of the net negative charge of the LPS, which is thought to reduce the attachment of positively charged AMPs, due to electrostatic repulsion [25, 94, 100]. In addition to its direct role in AMP resistance, the addition of L-ara4 to LPS has a regulatory effect on other processes related to LPS, such as the deacylation of lipid A by pagL (as described in section 3.4.1 above) [95], [96] and LPS export. It was found that L-ara4 synthesis is essential for bacteria viability since it is important for proper export of LPS molecules to the outer membrane of Burkholderia 15
ACCEPTED MANUSCRIPT cenocepacia [109]. It is possible that part of the AMP-resistance that is related to L-ara4 is due to its regulatory effects and not to direct effects of this modification.
PT
(ii) It has been reported that LPS undergoes a modification where glucosamine is added
RI
to the two phosphate groups of the lipid A in several species of Bordetella genus by ArnT homolog [110, 111]. This modification was found to confer resistance to AMPs, and reduce
SC
outer membrane perturbation by EDTA [112].
NU
(iii) Addition of galactosamine to lipid A was first discovered in Francisella tularensis Holarctica and Francisella novicida , and is done by ArnT homolog [113-115]. It was found later
MA
that in order for the galactosamine addition to occur, deacetylation of the lipid A needed to take place. This deacetylation is done by NaxD, and NaxD mutants were more susceptible to PmB
Changing the length of O-antigen
AC CE P
3.4.5
TE
D
compared to the parental strain [116 ].
To date, there is no conclusive evidence of whether the existence and length of the O-antigen moiety of the LPS affects AMP resistance. On the one hand, it was shown that a S. enterica Typhimurium mutant in O-antigen production is more sensitive to PmB compared to the WT strain. This mutant has a deletion in wbaP, a gene responsible for the addition of phosphogalactose to undecaprenyl-phosphate, a step necessary for O-antigen biosynthesis in this strain. However, this strain was still infectious and causes gut inflammation in mice models [117]. On the other hand, in F. tularensis it was shown that the absence of O-antigen did not cause a higher susceptibility to AMPs [118]. In addition, there is no conclusive answer of whether the length of the O-antigen in S. enterica Typhimurium affects resistance to AMPs. O-antigen length in these bacteria is defined by WZZst and fepE (WZZfepE) genes regulated by PmrAB TCS [119]. WZZst and fepE code 16
ACCEPTED MANUSCRIPT for a long O-antigen and a very long O-antigen, respectively [120, 121]. There is an ongoing debate on the question of whether the very long O-antigen is important for AMP resistance.
SC
RI
[119], while others showed no change in susceptibility [122] .
PT
Some claim that S. enterica Typhimurium mutants in FepE showed higher susceptibility to PmB
4. Modifications of the bacterial outer surface that influence peptide
NU
properties.
MA
Binding of AMPs to the LPS could change their chemical properties. It was shown [89] that a 12-mer peptide K5L7 has lower antibacterial activity compared to its diastereomer 4D-K5L7 in
D
which 4 L-amino acids were replaced with their enantiomers, D-amino acids. Nonetheless, when
TE
comparing the effect of these peptides on PG and phosphatidylethanolamine (PE) vesicles, an increase in the permeability of the corresponding vesicles was found for both. Furthermore, they
AC CE P
strongly bind LPS with a similar affinity; however each one of them was affected differently by the LPS. Both peptides have undefined structure in solution, but in LPS, K5L7 became aggregated whereas 4D-K5L7 did not. These results suggest that membrane binding is important for AMP activity, but it is not the only factor that affects the permeating activity of the peptides. It might also suggest that peptides can change their properties depending on the lipid they are bound to, and might explain the different activity of peptides on Gram-positive and negative bacteria. Later work also supports this hypothesis [63].
5. Mechanisms of bacterial AMP resistance in biofilms Bacterial biofilms are a colonization of bacteria on either a biotic or abiotic surface, where they form three dimensional multi-cell structures in which bacteria are embedded in an extracellular 17
ACCEPTED MANUSCRIPT matrix (ECM) composed of polysaccharides, DNA and proteins. Biofilms can be well structured and involve quorum sensing and formation of molecules that mediate migration of cells within
PT
the biofilm and formation of specific structures as seen in P. aeruginosa [123, 124]. Biofilms are
RI
important for bacterial survival in their natural environments [125], and protects the bacteria from the immune system and antibiotics including AMPs [126, 127]. Bacterial susceptibility to
SC
AMPs in biofilm was lower compared to the planktonic state [128, 129], however biofilm-
NU
resistance to AMPs was not extensively studied compared to other antimicrobial agents. Furthermore, no specific mechanisms for AMP-resistance in biofilm were discovered. It is
MA
assumed that mechanisms involved in the AMP-resistance in biofilms are similar to those known for other antimicrobial agents. Some of the putative mechanisms of resistance against
TE
D
antimicrobial agents in biofilms can theoretically be applied to AMPs and are therefore discussed in the following sections. A detailed review about resistance mechanisms to antibiotic in biofilm
5.1
AC CE P
has been published recently [130].
Creation of sub-populations in the biofilm-
P. aeruginosa biofilms that were grown in a flow chamber formed a mushroom shaped structure, and two well distinguished subpopulations were found in it [131]. When a biofilm was treated with colistin, there was a clear difference between the bacteria in the middle layer and those on the outer layer. The bacteria in the middle of the colony died while those on the outer layer were kept alive [131]. In addition, the bacteria on the outer surface and not in the middle were characterized with motility as was demonstrated after photo bleaching [131]. Resistant individuals were also found in bacteria with motility deficiencies. However these bacteria were not placed on the outer part of the biofilm, but were randomly incorporated in it. This might suggest that the motility is important for the arrangement of the different populations in the 18
ACCEPTED MANUSCRIPT biofilm, but not directly involved in AMP-resistance [131, 132]. Furthermore, pmr operon expression pattern as observed with GFP was correlated to the two bacterial subpopulations
PT
[131]. In addition, cells of the different subpopulations were characterized by different metabolic
RI
levels whereas cells in the outer surface were more metabolically active compared to the ones in
A higher mutation rate-
NU
5.2
SC
the middle. This suggests that metabolically active bacteria are more resistant to colistin [132].
It was found that bacteria in biofilm tend to have a higher mutation rate compared to the
MA
planktonic bacteria. The higher mutation rate can accelerate the natural occurring process of resistance to antimicrobial agents [130]. The accelerated mutation rate was found to involve a
D
reduced expression of DNA repair genes and mismatch repair system, as well as, elevation in
TE
oxidative stress [130]. In addition, several studies reported for a higher frequency of horizontal
5.3
AC CE P
gene transfer in biofilm. This is another aspect that contributes to a higher mutation rate [133].
Upregulation of efflux pumps -
Efflux pumps were shown to confer biofilm resistance in several bacterial strains in their planktonic state [27, 42, 43] and biofilm state [132, 134]. It was reported that efflux pumps undergo upregulation in biofilm compared to the planktonic state [135-137]. In addition, MexAB-OprM efflux pumps were found to be important for the development of tolerance to colistin in a sub-population of P. aeruginosa biofilm [132]. A detailed review about the role of efflux pumps in biofilm resistance to antibiotics can be found in reference [137].
5.4
Induction of TCSs within the biofilm-
19
ACCEPTED MANUSCRIPT P. aeruginosa resistant to colistin showed upregulation of pmr operon genes, which was suggested to contribute to bacterial biofilm tolerance due to LPS modifications [131, 132]. It was
PT
also suggested that the DNA that is found as part of the ECM reduces the level of divalent
Diffusion in biofilm-
SC
5.5
RI
cations in the environment thus activating PhoPQ TCS and conferring resistance to AMPs [38].
NU
It is not clear whether biofilm can reduce the diffusion rate of AMPs and in so doing help to resist them. It was shown that the diffusion rate of fluorescein, a small molecule that does not
MA
interact with the biofilm, is not affected by the presence of a biofilm. However, it was predicted that the diffusion of large molecules will be influenced by the biofilm [138]. Periplasmic glucan
D
was found to confer resistance to several antibiotics in P. aeruginosa, in a biofilm specific
TE
manner, where a mutant for ndvB, a gene necessary for periplasmic glucan, was shown to be
AC CE P
more susceptible to these agents [139]. In this work the authors suggested that glucans interact with the antibiotics, and therefore slow the diffusion of these molecules. However, resistance to AMPs was not checked in this respect. In addition, it was shown that P. aeruginosa and K. pneumonia biofilms reduce the diffusion rate of beta-lactam antibiotics only when betalactamase enzymes are present within the biofilms [130, 140]. For more details on diffusion of
antibiotics in P. aeruginosa biofilms the following review is recommended [141].
6. Two component systems (TCSs) regulate AMP resistance TCSs are commonly used in bacteria to sense and respond to environmental changes [142-145]. Many of the outer surface modifications that were discussed above are a result of activation of such systems via regulation of the expression of downstream genes as will be discussed in the
20
ACCEPTED MANUSCRIPT following section. TCSs consist of a membrane sensor that receive signals from the environment and transfers this information by activating a transcriptional response regulator through
PT
phosphorylation or de-phosphorylation [24, 143, 146-150]. In these systems, a receptor (usually
RI
histidine kinase) is situated in the cytoplasmic membrane, and is activated by an environmental signal. The sensor in turn, phosphorylates a cytoplasmic protein that acts as a transcription
SC
factor. The response involves activation of sets of genes such as virulence genes, ion transporters
NU
and membrane-remodeling genes that help the Gram-positive and Gram-negative bacteria to adjust better to the environment within hours. Some of these TCS are known to help bacteria
TCS activation by AMPs
D
6.1
MA
cope with AMPs and are activated by them.
TE
S. enterica PhoPQ TCS is known to be activated by low pH, low divalent cation concentration
AC CE P
and AMPs [151-153]. It has been shown recently that peptides with a higher positive charge were more potent in PhoPQ activation [15]. Studies on the mode of PhoQ activation by AMPs suggest that they displace divalent cations that bind to a highly acidic domain within the PhoQ [149, 153-158], which corroborates the finding that the more positively charged the AMP is, the more it tends to activate the PhoPQ [15]. Surprisingly, a histidine-rich AMP with a charge of only +1 under these conditions was highly potent in PhoPQ activation [15]. This supports a second mechanism for removing divalent cations and activation of PhoPQ by certain AMPs In that case, histidine, a known chelator for divalent cations, exerts this activity in the LPS environment, reducing their concentration in the environment leading to PhoPQ activation [15]. However, activation by different signals promotes different responses [159]. PmrAB TCS is known to be activated by high Fe+3 concentration [158]. In addition, there is a crosstalk between these two TCSs, and PhoPQ activated gene pmrD encodes for a protein activator that activates 21
ACCEPTED MANUSCRIPT pmrA, which in turn represses pmrD expression [151]. These systems have homologous systems in several Gram-negative bacteria, but the interaction between them in different species can be
PT
different. For example in E. coli another unknown bacterial system is involved in PmrAB
RI
activation in a PmrD dependent manner [160]. Both PhoPQ and PmrAB were found to be essential for the virulence of many bacteria species [143, 161, 162].
SC
GraRS (ApsRS) is a well-studied TCS with respect to AMPs resistance, that is found in
NU
several Gram-positive bacteria such as S. aureus and Staphylococcus epidermidis [163]. Similarly to PhoPQ activation in Gram-negative bacteria, it is assumed that AMPs bind a highly
MA
negative extracellular loop in the GraS (ApsS) sensor protein, and causes activation of this system [164]. GraRS was shown to crosstalk with another bacterial system Stk1/Stp1, where
TE
D
Stk1 phosphorylates the response regulator GraR, and thus enabling alanilation of the wall theichoic acid (WTA) by dlt operon, which is regulated by GraRS TCS. S. aureus mutants in the
AC CE P
phosphorylation site of Skt1on GraR showed a similar reduced level of alanine in the WTA [165]. However, Bacillus subtilis homolog for GraR, BceR, does not undergo Skp1 dependent phosphorylation [165]. Other TCS in S. aureus were also reported to play a role in vancomycin resistance. Such TCSs are WalKR, VraSR and AirSR (YhcSR) [166, 167].
6.2
Genes activated by TCS that are involved in AMP resistance
Upon activation, TCSs can activate tens and even hundreds of genes [168, 169]. Some of the genes activated by PhoPQ and PmrAB are able to change the composition of the LPS and other surface elements and so change their general surface thickness and charge, thus altering AMP’s effect on these bacteria [92, 100, 145, 170]. For example, PhoPQ induces pagP and pagL expression. pagP and pagL, as described before, encode for acyltransferases which catalyze 22
ACCEPTED MANUSCRIPT palmitoylation of lipid-A and glycerophospholipids of the inner leaflet of the outer membrane [72, 90]. On the other hand, pagL encodes a deacylase which removes a fatty acid from the
PT
lipid-A core [94]. This leads to increase or decrease of both the membrane thickness and the
RI
hydrophobicity of the LPS [90, 92]. Other PhoPQ and PmrAB activated genes are pmrE, pmrHJIFKLM operon that are involved in LPS charge modifications [3, 94, 100, 171-176].
SC
Because these systems induce genes that influence the hydrophobicity and charge of the LPS, it
NU
is highly logic to assume that they contribute to resistance toward AMPs. Indeed PhoPQ and PmrAB relevance to AMP-resistance was demonstrated by showing that phoP-KO or pmrA-KO
MA
bacteria displayed increased susceptibility to some AMPs (particularly PmB and colistin) compared to the WT [13, 177-183]. Despite extensive studies there are still genes induced by
TE
D
these systems that have an unknown function [184, 185]. In addition, most of the work done was limited to certain types of AMPs (particularly PmB, colistin and LL-37), which do not represent
AC CE P
the major classical AMPs (Figure 4).
GraRS TCS consists of GraS- a sensor protein, GraR (ApsR)- a response regulator and another accessory protein with an unknown function, GraX (ApsX). Mutations in each of these three genes increase susceptibility to AMPs [164, 186]. GraRS is known to regulate the expression of mprF, dlt operon and vraFG operon [57, 187], which are known to be involved in AMP resistance. Over expression of GraRS in S. aureus caused higher resistance to vancomycin [163]. As previously mentioned, MprF is responsible for the addition of lysine and alanine to PG on the plasma membrane, DltA is involved in the addition of D-alanine to the LTA, and VraFG is an AMP efflux pump. In Listeria monocytogenes a TCS named VirRS regulates dlt operon and mprF homolog, among other genes [169] (Figure 4).
7. Ways to overcome AMP resistance 23
ACCEPTED MANUSCRIPT Mammalian AMPs serve as part of the innate immune system, and consist of two main groups, cathelicidins and the cysteine rich defensins. Mutations in genes related to AMP production in
PT
humans and mice led to frequent bacterial infections, whilst transgenic mice expressing high
RI
amounts of human defensin 5 were protected from certain bacterial infections [188]. All known mechanisms of bacterial resistance to AMPs refer to general changes in bacteria properties that
SC
allegedly can be applied to all AMPs. Therefore, there is ongoing debate as to whether the use of
NU
AMPs in the clinic can lead to bacteria resistance to naturally occurring AMPs, and by that damage the activity of the innate immune system against various pathogens [13]. Indeed, a study
MA
showed that there is cross resistance between some AMPs, and thus pre-exposure to one peptide can lead to higher resistance to other peptides [189]. Lfcin B resistant S. aureus bacteria showed
TE
D
mild cross resistance with Indolicidin and a derivative of magainin (Ala3,13,18-magainin), while for the same strains no resistance was observed for LL-37 [190]. Induced S. enterica
AC CE P
Typhimurium resistant mutants to LL-37, CNY100HL and Wheat Germ Histones (WGH), showed cross resistance to the other peptides to a certain extent. Several lines of S. aureus that are resistant to daptomycin, a lipopeptide, showed a higher resistance to tPMPs and human neutrophil peptide-1 (hNP-1). However, resistance to PMB was not consistent in most of the lines [67].
Moreover, it has been also shown that there is no cross resistance between different AMPs. So far, it is suggested that peptides show cross resistance when they share common mechanisms of action [191], but this assumption should be further evaluated. In addition, in some cases the resistance to the AMPs was abolished after removing the AMP [129, 190, 192]. This kind of induced resistance is only transient and as a result of the activation of bacterial systems to environmental changes such as encountering an AMP. Therefore, it is indeed possible that the
24
ACCEPTED MANUSCRIPT use of AMPs for the treatment of bacterial infections will cause resistance to innate immune AMPs. However, a better understanding of the mechanisms that affect cross resistance between
PT
AMP, as well as the analysis of the nature of cross resistance, might allow this problem to be
RI
overcome, so that AMPs can be used as the first line treatment for infectious diseases. In addition, combined therapies using two different AMPs or a regular antibiotic and AMP might
NU
SC
also provide a solution to the cross resistance between AMPs.
8. Summary
MA
In summary, bacteria were found to acquire AMP resistance by two general mechanisms: (1) induced resistance, by induction of specific bacterial systems that contribute to AMP resistance.
TE
D
When the bacteria are no longer under these conditions the resistance will vanish. (2) Constitutive resistance which results from genetic changes. In this review we discussed briefly
AC CE P
these two general mechanisms which involve: (i) Extracellular polysaccharide secretion to form capsules that protect the bacteria from various antimicrobial agents including AMPs. (ii) Modifying bacteria surface molecules, such as LTA in Gram-positive bacteria with the addition of D-alanine. Modification of the LPS moieties: modification of the lipid A by the addition of glycine, amino sugars such as aminoarabinose, glucosamine and galactoseamine, dephosphorilation, hydroxylation , addition of phosphoethanolamine, addition and removal of acyl chains, and changing the length of O antigen moiety. (iii) Changes in bacterial surface which can cause alterations in peptide properties, and as such its effect on the bacteria.
25
ACCEPTED MANUSCRIPT (iv) Biofilm formation including the secretion of extra cellular matrix, which consists of polysaccharides, proteins and DNA, all of which form a barrier to AMPs.
PT
(v) Peptide activation of TCSs resulting in gene expression patterns, which in many cases
RI
regulate the above mechanisms, most of which induce biophysical changes to the bacteria, such as surface rigidity, cell wall thickness, surface charge and membrane fluidity.
SC
Importantly, despite the main concern that bacterial resistance to administered AMPs might
NU
cause resistance to natural AMPs and so sabotage the natural immune response, there are
MA
indications for only limited cross resistance between different AMPs.
D
Acknowledgements
TE
This investigation was supported in part by a grant from the Conseil Pasteur-Weizmann to PTC, TM and YS); the Israel Ministry of Health (IMOH), Grant No. 3-7291 and the European
AC CE P
Community's Seventh Framework Programme (FP7/2007-2013) under grant agreement n° 278998. Yechiel Shai is the incumbent of the Harold S. and Harriet B. Brady Professorial Chair in Cancer Research.
26
ACCEPTED MANUSCRIPT
B. Spellberg, R. Guidos, D. Gilbert, J. Bradley, H.W. Boucher, W.M. Scheld, J.G.
SC
[1]
RI
PT
9. References
Bartlett, J. Edwards, Jr., The epidemic of antibiotic-resistant infections: a call to action
NU
for the medical community from the Infectious Diseases Society of America, Clin Infect
[2]
MA
Dis 46 (2008) 155-164.
J.F. Acar, G. Moulin, Antimicrobial resistance: a complex issue, Rev Sci Tech 31 (2012)
H.G. Boman, Peptide antibiotics and their role in innate immunity, Annu Rev Immunol 13 (1995) 61-92.
T. Koprivnjak, A. Peschel, Bacterial resistance mechanisms against host defense
AC CE P
[4]
TE
[3]
D
23-31.
peptides, Cell Mol Life Sci 68 (2011) 2243-2254. [5]
M. Zasloff, Antimicrobial peptides of multicellular organisms, Nature 415 (2002) 389395.
[6]
Y. Shai, A. Makovitzky, D. Avrahami, Host defense peptides and lipopeptides: modes of action and potential candidates for the treatment of bacterial and fungal infections, Curr Protein Pept Sci 7 (2006) 479-486.
[7]
L.T. Nguyen, E.F. Haney, H.J. Vogel, The expanding scope of antimicrobial peptide structures and their modes of action, Trends Biotechnol 29 (2011) 464-472.
27
ACCEPTED MANUSCRIPT [8]
B. Agerberth, H. Gunne, J. Odeberg, P. Kogner, H.G. Boman, G.H. Gudmundsson, FALL-39, a putative human peptide antibiotic, is cysteine-free and expressed in bone
G.H. Gudmundsson, B. Agerberth, J. Odeberg, T. Bergman, B. Olsson, R. Salcedo, The
RI
[9]
PT
marrow and testis, Proc. Natl. Acad. Sci. U. S. A. 92 (1995) 195-199.
human gene FALL39 and processing of the cathelin precursor to the antibacterial peptide
Y. Shai, Mechanism of the binding, insertion and destabilization of phospholipid bilayer
NU
[10]
SC
LL-37 in granulocytes, Eur. J. Biochem. 238 (1996) 325-332.
membranes by alpha-helical antimicrobial and cell non-selective membrane-lytic
[11]
MA
peptides, Biochim Biophys Acta 1462 (1999) 55-70. U.H. Durr, U.S. Sudheendra, A. Ramamoorthy, LL-37, the only human member of the
TE
D
cathelicidin family of antimicrobial peptides, Biochim Biophys Acta 1758 (2006) 14081425.
H. Jenssen, P. Hamill, R.E. Hancock, Peptide antimicrobial agents, Clin Microbiol Rev
AC CE P
[12]
19 (2006) 491-511. [13]
A.K. Marr, W.J. Gooderham, R.E. Hancock, Antibacterial peptides for therapeutic use: obstacles and realistic outlook, Curr Opin Pharmacol 6 (2006) 468-472.
[14]
Z. Jiang, A.I. Vasil, J.D. Hale, R.E. Hancock, M.L. Vasil, R.S. Hodges, Effects of net charge and the number of positively charged residues on the biological activity of amphipathic alpha-helical cationic antimicrobial peptides, Biopolymers 90 (2008) 369383.
[15]
T. Shprung, A. Peleg, Y. Rosenfeld, P. Trieu-Cuot, Y. Shai, Effect of PhoP-PhoQ activation by broad repertoire of antimicrobial peptides on bacterial resistance, J Biol Chem 287 (2012) 4544-4551.
28
ACCEPTED MANUSCRIPT [16]
L.M. Yin, M.A. Edwards, J. Li, C.M. Yip, C.M. Deber, Roles of hydrophobicity and charge distribution of cationic antimicrobial peptides in peptide-membrane interactions, J
Y.M. Lin, S.J. Wu, T.W. Chang, C.F. Wang, C.S. Suen, M.J. Hwang, M.D. Chang, Y.T.
RI
[17]
PT
Biol Chem 287 (2012) 7738-7745.
Chen, Y.D. Liao, Outer membrane protein I of Pseudomonas aeruginosa is a target of
J.P. Powers, R.E. Hancock, The relationship between peptide structure and antibacterial activity, Peptides 24 (2003) 1681-1691.
R.E. Hancock, G. Diamond, The role of cationic antimicrobial peptides in innate host
MA
[19]
NU
[18]
SC
cationic antimicrobial peptide/protein, J Biol Chem 285 (2010) 8985-8994.
defences, Trends. Microbiol. 8 (2000) 402-410.
D
G.A. Woolley, B.A. Wallace, Model ion channels: gramicidin and alamethicin, J Membr
TE
[20]
Biol 129 (1992) 109-136.
Y. Shai, Mode of action of membrane active antimicrobial peptides, Biopolymers 66
AC CE P
[21]
(2002) 236-248. [22]
K. Matsuzaki, Why and how are peptide-lipid unteractions utilized for self defense? Magainins and tachyplesins as archyetypes, Biochim. Biophys. Acta. 1462 (1999) 1-10.
[23]
L. Yang, T.A. Harroun, T.M. Weiss, L. Ding, H.W. Huang, Barrel-stave model or toroidal model? A case study on melittin pores, Biophys J 81 (2001) 1475-1485.
[24]
E. Garcia Vescovi, F.C. Soncini, E.A. Groisman, The role of the PhoP/PhoQ regulon in Salmonella virulence, Res Microbiol 145 (1994) 473-480.
[25]
J.S. Gunn, K.B. Lim, J. Krueger, K. Kim, L. Guo, M. Hackett, S.I. Miller, PmrA-PmrBregulated genes necessary for 4-aminoarabinose lipid A modification and polymyxin resistance, Mol Microbiol 27 (1998) 1171-1182.
29
ACCEPTED MANUSCRIPT [26]
V. Nizet, Antimicrobial peptide resistance mechanisms of human bacterial pathogens, Curr Issues Mol Biol 8 (2006) 11-26. Y.L. Tzeng, K.D. Ambrose, S. Zughaier, X. Zhou, Y.K. Miller, W.M. Shafer, D.S.
PT
[27]
RI
Stephens, Cationic antimicrobial peptide resistance in Neisseria meningitidis, J Bacteriol 187 (2005) 5387-5396.
K.L. Nawrocki, E.K. Crispell, S.M. McBride, Antimicrobial Peptide Resistance
SC
[28]
[29]
NU
Mechanisms of Gram-Positive Bacteria, Antibiotics (Basel) 3 (2014) 461-492. J.C. Henderson, C.D. Fage, J.R. Cannon, J.S. Brodbelt, A.T. Keatinge-Clay, M.S. Trent,
MA
Antimicrobial peptide resistance of Vibrio cholerae results from an LPS modification pathway related to nonribosomal peptide synthetases, ACS Chem Biol 9 (2014) 2382-
TE
[30]
D
2392.
A.X. Tran, J.D. Whittimore, P.B. Wyrick, S.C. McGrath, R.J. Cotter, M.S. Trent, The
AC CE P
lipid A 1-phosphatase of Helicobacter pylori is required for resistance to the antimicrobial peptide polymyxin, J Bacteriol 188 (2006) 4531-4541. [31]
J.B. McPhee, M. Bains, G. Winsor, S. Lewenza, A. Kwasnicka, M.D. Brazas, F.S. Brinkman, R.E. Hancock, Contribution of the PhoP-PhoQ and PmrA-PmrB twocomponent regulatory systems to Mg2+-induced gene regulation in Pseudomonas aeruginosa, J Bacteriol 188 (2006) 3995-4006.
[32]
J.B. McPhee, S. Lewenza, R.E. Hancock, Cationic antimicrobial peptides activate a twocomponent regulatory system, PmrA-PmrB, that regulates resistance to polymyxin B and cationic antimicrobial peptides in Pseudomonas aeruginosa, Mol Microbiol 50 (2003) 205-217.
30
ACCEPTED MANUSCRIPT [33]
S.M. Moskowitz, R.K. Ernst, S.I. Miller, PmrAB, a two-component regulatory system of Pseudomonas aeruginosa that modulates resistance to cationic antimicrobial peptides and
A. Peschel, How do bacteria resist human antimicrobial peptides?, Trends Microbiol 10
RI
[34]
PT
addition of aminoarabinose to lipid A, J Bacteriol 186 (2004) 575-579.
(2002) 179-186.
R. Saar-Dover, A. Bitler, R. Nezer, L. Shmuel-Galia, A. Firon, E. Shimoni, P. Trieu-
SC
[35]
NU
Cuot, Y. Shai, D-alanylation of lipoteichoic acids confers resistance to cationic peptides in group B streptococcus by increasing the cell wall density, PLoS Pathog 8 (2012)
[36]
MA
e1002891.
G.H. Gudmundsson, P. Bergman, J. Andersson, R. Raqib, B. Agerberth, Battle and
TE
D
balance at mucosal surfaces--the story of Shigella and antimicrobial peptides, Biochem Biophys Res Commun 396 (2010) 116-119. B. Sperandio, B. Regnault, J. Guo, Z. Zhang, S.L. Stanley, Jr., P.J. Sansonetti, T. Pedron,
AC CE P
[37]
Virulent Shigella flexneri subverts the host innate immune response through manipulation of antimicrobial peptide gene expression, J Exp Med 205 (2008) 11211132. [38]
H. Mulcahy, L. Charron-Mazenod, S. Lewenza, Extracellular DNA chelates cations and induces antibiotic resistance in Pseudomonas aeruginosa biofilms, PLoS Pathog 4 (2008) e1000213.
[39]
C. Chan, L.L. Burrows, C.M. Deber, Helix induction in antimicrobial peptides by alginate in biofilms, J Biol Chem 279 (2004) 38749-38754.
[40]
M. Sieprawska-Lupa, P. Mydel, K. Krawczyk, K. Wojcik, M. Puklo, B. Lupa, P. Suder, J. Silberring, M. Reed, J. Pohl, W. Shafer, F. McAleese, T. Foster, J. Travis, J. Potempa,
31
ACCEPTED MANUSCRIPT Degradation of human antimicrobial peptide LL-37 by Staphylococcus aureus-derived proteinases, Antimicrob Agents Chemother 48 (2004) 4673-4679. A. Schmidtchen, I.M. Frick, E. Andersson, H. Tapper, L. Bjorck, Proteinases of common
PT
[41]
RI
pathogenic bacteria degrade and inactivate the antibacterial peptide LL-37, Mol Microbiol 46 (2002) 157-168.
D.M. Warner, W.M. Shafer, A.E. Jerse, Clinically relevant mutations that cause
SC
[42]
NU
derepression of the Neisseria gonorrhoeae MtrC-MtrD-MtrE Efflux pump system confer different levels of antimicrobial resistance and in vivo fitness, Mol Microbiol 70 (2008)
[43]
MA
462-478.
W.M. Shafer, X. Qu, A.J. Waring, R.I. Lehrer, Modulation of Neisseria gonorrhoeae
TE
D
susceptibility to vertebrate antibacterial peptides due to a member of the resistance/nodulation/division efflux pump family, Proc Natl Acad Sci U S A 95 (1998)
[44]
AC CE P
1829-1833.
H.C. Maisey, D. Quach, M.E. Hensler, G.Y. Liu, R.L. Gallo, V. Nizet, K.S. Doran, A group B streptococcal pilus protein promotes phagocyte resistance and systemic virulence, FASEB J 22 (2008) 1715-1724.
[45]
X. Lauth, M. von Kockritz-Blickwede, C.W. McNamara, S. Myskowski, A.S. Zinkernagel, B. Beall, P. Ghosh, R.L. Gallo, V. Nizet, M1 protein allows Group A streptococcal survival in phagocyte extracellular traps through cathelicidin inhibition, J Innate Immun 1 (2009) 202-214.
[46]
I.M. Frick, P. Akesson, M. Rasmussen, A. Schmidtchen, L. Bjorck, SIC, a secreted protein of Streptococcus pyogenes that inactivates antibacterial peptides, J Biol Chem 278 (2003) 16561-16566.
32
ACCEPTED MANUSCRIPT [47]
A.J. Manning, M.J. Kuehn, Contribution of bacterial outer membrane vesicles to innate bacterial defense, BMC Microbiol 11 (2011) 258. J.L. Anaya-Lopez, J.E. Lopez-Meza, A. Ochoa-Zarzosa, Bacterial resistance to cationic
PT
[48]
[49]
RI
antimicrobial peptides, Crit Rev Microbiol 39 (2013) 180-195.
J.S. Gunn, The Salmonella PmrAB regulon: lipopolysaccharide modifications,
M.R. Yeaman, N.Y. Yount, Mechanisms of antimicrobial peptide action and resistance,
NU
[50]
SC
antimicrobial peptide resistance and more, Trends Microbiol 16 (2008) 284-290.
Pharmacol Rev 55 (2003) 27-55.
A.O. Olaitan, S. Morand, J.M. Rolain, Mechanisms of polymyxin resistance: acquired
MA
[51]
and intrinsic resistance in bacteria, Front Microbiol 5 (2014) 643.
D
A. Jones, M. Georg, L. Maudsdotter, A.B. Jonsson, Endotoxin, capsule, and bacterial
TE
[52]
attachment contribute to Neisseria meningitidis resistance to the human antimicrobial
[53]
AC CE P
peptide LL-37, J Bacteriol 191 (2009) 3861-3868. H. Miajlovic, S.G. Smith, Bacterial self-defence: how Escherichia coli evades serum killing, FEMS Microbiol Lett 354 (2014) 1-9. [54]
E. Llobet, J.M. Tomas, J.A. Bengoechea, Capsule polysaccharide is a bacterial decoy for antimicrobial peptides, Microbiology 154 (2008) 3877-3886.
[55]
M.A. Campos, M.A. Vargas, V. Regueiro, C.M. Llompart, S. Alberti, J.A. Bengoechea, Capsule polysaccharide mediates bacterial resistance to antimicrobial peptides, Infect Immun 72 (2004) 7107-7114.
[56]
C.M. Ernst, S. Kuhn, C.J. Slavetinsky, B. Krismer, S. Heilbronner, C. Gekeler, D. Kraus, S. Wagner, A. Peschel, The lipid-modifying multiple peptide resistance factor is an oligomer consisting of distinct interacting synthase and flippase subunits, MBio 6 (2015).
33
ACCEPTED MANUSCRIPT [57]
C.M. Ernst, P. Staubitz, N.N. Mishra, S.J. Yang, G. Hornig, H. Kalbacher, A.S. Bayer, D. Kraus, A. Peschel, The bacterial defensin resistance protein MprF consists of separable
PT
domains for lipid lysinylation and antimicrobial peptide repulsion, PLoS Pathog 5 (2009)
[58]
RI
e1000660.
A. Peschel, R.W. Jack, M. Otto, L.V. Collins, P. Staubitz, G. Nicholson, H. Kalbacher,
SC
W.F. Nieuwenhuizen, G. Jung, A. Tarkowski, K.P. van Kessel, J.A. van Strijp,
NU
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-
[59]
MA
lysine, J Exp Med 193 (2001) 1067-1076.
S.J. Yang, N.N. Mishra, A. Rubio, A.S. Bayer, Causal role of single nucleotide
TE
D
polymorphisms within the mprF gene of Staphylococcus aureus in daptomycin resistance, Antimicrob Agents Chemother 57 (2013) 5658-5664. C.M. Ernst, A. Peschel, Broad-spectrum antimicrobial peptide resistance by MprF-
AC CE P
[60]
mediated aminoacylation and flipping of phospholipids, Mol Microbiol 80 (2011) 290299. [61]
A.S. Bayer, N.N. Mishra, G. Sakoulas, P. Nonejuie, C.C. Nast, J. Pogliano, K.T. Chen, S.N. Ellison, M.R. Yeaman, S.J. Yang, Heterogeneity of mprF sequences in methicillinresistant Staphylococcus aureus clinical isolates: role in cross-resistance between daptomycin and host defense antimicrobial peptides, Antimicrob Agents Chemother 58 (2014) 7462-7467.
[62]
S. Klein, C. Lorenzo, S. Hoffmann, J.M. Walther, S. Storbeck, T. Piekarski, B.J. Tindall, V. Wray, M. Nimtz, J. Moser, Adaptation of Pseudomonas aeruginosa to various
34
ACCEPTED MANUSCRIPT conditions includes tRNA-dependent formation of alanyl-phosphatidylglycerol, Mol Microbiol 71 (2009) 551-565. J.K. Muraih, J. Harris, S.D. Taylor, M. Palmer, Characterization of daptomycin
PT
[63]
RI
oligomerization with perylene excimer fluorescence: stoichiometric binding of phosphatidylglycerol triggers oligomer formation, Biochim Biophys Acta 1818 (2012)
A.S. Bayer, R. Prasad, J. Chandra, A. Koul, M. Smriti, A. Varma, R.A. Skurray, N. Firth,
NU
[64]
SC
673-678.
M.H. Brown, S.P. Koo, M.R. Yeaman, In vitro resistance of Staphylococcus aureus to
MA
thrombin-induced platelet microbicidal protein is associated with alterations in cytoplasmic membrane fluidity, Infect Immun 68 (2000) 3548-3553.
D
N.N. Mishra, A.S. Bayer, Correlation of cell membrane lipid profiles with daptomycin
TE
[65]
resistance in methicillin-resistant Staphylococcus aureus, Antimicrob Agents Chemother
[66]
AC CE P
57 (2013) 1082-1085.
T. Jones, M.R. Yeaman, G. Sakoulas, S.J. Yang, R.A. Proctor, H.G. Sahl, J. Schrenzel, Y.Q. Xiong, A.S. Bayer, 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 52 (2008) 269-278.
[67]
N.N. Mishra, J. McKinnell, M.R. Yeaman, A. Rubio, C.C. Nast, L. Chen, B.N. Kreiswirth, A.S. Bayer, In vitro cross-resistance to daptomycin and host defense cationic antimicrobial peptides in clinical methicillin-resistant Staphylococcus aureus isolates, Antimicrob Agents Chemother 55 (2011) 4012-4018.
35
ACCEPTED MANUSCRIPT [68]
T. Shireen, M. Singh, T. Das, K. Mukhopadhyay, Differential adaptive responses of Staphylococcus aureus to in vitro selection with different antimicrobial peptides,
R. Kumariya, S.K. Sood, Y.S. Rajput, N. Saini, A.K. Garsa, Increased membrane surface
RI
[69]
PT
Antimicrob Agents Chemother 57 (2013) 5134-5137.
positive charge and altered membrane fluidity leads to cationic antimicrobial peptide
A. Verheul, N.J. Russell, T.H.R. Van, F.M. Rombouts, T. Abee, Modifications of
NU
[70]
SC
resistance in Enterococcus faecalis, Biochim Biophys Acta 1848 (2015) 1367-1375.
membrane phospholipid composition in nisin-resistant Listeria monocytogenes Scott A,
[71]
MA
Appl Environ Microbiol 63 (1997) 3451-3457.
H. Zhou, J. Fang, Y. Tian, X.Y. Lu, Mechanisms of nisin resistance in Gram-positive
[72]
TE
D
bacteria, Annals of Microbiology 64 (2014) 413-420. Z.D. Dalebroux, S.I. Miller, Salmonellae PhoPQ regulation of the outer membrane to
[73]
AC CE P
resist innate immunity, Curr Opin Microbiol 17 (2014) 106-113. E. Cox, A. Michalak, S. Pagentine, P. Seaton, A. Pokorny, Lysylated phospholipids stabilize models of bacterial lipid bilayers and protect against antimicrobial peptides, Biochim Biophys Acta 1838 (2014) 2198-2204. [74]
S.M. McBride, A.L. Sonenshein, The dlt operon confers resistance to cationic antimicrobial peptides in Clostridium difficile, Microbiology 157 (2011) 1457-1465.
[75]
Z. Abi Khattar, A. Rejasse, D. Destoumieux-Garzon, J.M. Escoubas, V. Sanchis, D. Lereclus, A. Givaudan, M. Kallassy, C. Nielsen-Leroux, S. Gaudriault, The dlt operon of Bacillus cereus is required for resistance to cationic antimicrobial peptides and for virulence in insects, J Bacteriol 191 (2009) 7063-7073.
36
ACCEPTED MANUSCRIPT [76]
S.A. Kristian, V. Datta, C. Weidenmaier, R. Kansal, I. Fedtke, A. Peschel, R.L. Gallo, V. Nizet, D-alanylation of teichoic acids promotes group a streptococcus antimicrobial
PT
peptide resistance, neutrophil survival, and epithelial cell invasion, J Bacteriol 187 (2005)
[77]
RI
6719-6725.
F.C. Neuhaus, J. Baddiley, A continuum of anionic charge: structures and functions of D-
SC
alanyl-teichoic acids in gram-positive bacteria, Microbiol Mol Biol Rev 67 (2003) 686-
[78]
NU
723.
A. Peschel, H.G. Sahl, The co-evolution of host cationic antimicrobial peptides and
[79]
MA
microbial resistance, Nat Rev Microbiol 4 (2006) 529-536. A. Peschel, M. Otto, R.W. Jack, H. Kalbacher, G. Jung, F. Gotz, Inactivation of the dlt
TE
D
operon in Staphylococcus aureus confers sensitivity to defensins, protegrins, and other antimicrobial peptides, J Biol Chem 274 (1999) 8405-8410. S.J. Yang, C.C. Nast, N.N. Mishra, M.R. Yeaman, P.D. Fey, A.S. Bayer, Cell wall
AC CE P
[80]
thickening is not a universal accompaniment of the daptomycin nonsusceptibility phenotype in Staphylococcus aureus: evidence for multiple resistance mechanisms, Antimicrob Agents Chemother 54 (2010) 3079-3085. [81]
L. Cui, E. Tominaga, H.M. Neoh, K. Hiramatsu, Correlation between Reduced Daptomycin Susceptibility and Vancomycin Resistance in Vancomycin-Intermediate Staphylococcus aureus, Antimicrob Agents Chemother 50 (2006) 1079-1082.
[82]
S. Maisnier-Patin, J. Richard, Cell wall changes in nisin-resistant variants of Listeria innocua grown in the presence of high nisin concentrations, FEMS Microbiol Lett 140 (1996) 29-35.
37
ACCEPTED MANUSCRIPT [83]
M.G. Percy, A. Grundling, Lipoteichoic acid synthesis and function in gram-positive bacteria, Annu Rev Microbiol 68 (2014) 81-100. X. Zogaj, M. Nimtz, M. Rohde, W. Bokranz, U. Romling, The multicellular morphotypes
PT
[84]
RI
of Salmonella typhimurium and Escherichia coli produce cellulose as the second component of the extracellular matrix, Mol Microbiol 39 (2001) 1452-1463. A.P. White, D.L. Gibson, S.K. Collinson, P.A. Banser, W.W. Kay, Extracellular
SC
[85]
NU
polysaccharides associated with thin aggregative fimbriae of Salmonella enterica serovar enteritidis, J Bacteriol 185 (2003) 5398-5407.
C.E. de Rezende, Y. Anriany, L.E. Carr, S.W. Joseph, R.M. Weiner, Capsular
MA
[86]
polysaccharide surrounds smooth and rugose types of Salmonella enterica serovar
[87]
TE
D
Typhimurium DT104, Appl Environ Microbiol 71 (2005) 7345-7351. C. Alexander, E.T. Rietschel, Bacterial lipopolysaccharides and innate immunity, J
[88]
AC CE P
Endotoxin Res 7 (2001) 167-202. C.R. Raetz, C. Whitfield, Lipopolysaccharide endotoxins, Annu Rev Biochem 71 (2002) 635-700. [89]
N. Papo, Y. Shai, A molecular mechanism for lipopolysaccharide protection of Gramnegative bacteria from antimicrobial peptides, J. Biol. Chem. 280 (2005) 10378-10387.
[90]
R.E. Bishop, H.S. Gibbons, T. Guina, M.S. Trent, S.I. Miller, C.R. Raetz, Transfer of palmitate from phospholipids to lipid A in outer membranes of gram-negative bacteria, EMBO J 19 (2000) 5071-5080.
[91]
A.H. Delcour, Outer membrane permeability and antibiotic resistance, Biochim Biophys Acta 1794 (2009) 808-816.
38
ACCEPTED MANUSCRIPT [92]
L. Guo, K.B. Lim, C.M. Poduje, M. Daniel, J.S. Gunn, M. Hackett, S.I. Miller, Lipid A acylation and bacterial resistance against vertebrate antimicrobial peptides, Cell 95
M. Robey, W. O'Connell, N.P. Cianciotto, Identification of Legionella pneumophila rcp,
RI
[93]
PT
(1998) 189-198.
a pagP-like gene that confers resistance to cationic antimicrobial peptides and promotes
M.S. Trent, W. Pabich, C.R. Raetz, S.I. Miller, A PhoP/PhoQ-induced Lipase (PagL) that
NU
[94]
SC
intracellular infection, Infect Immun 69 (2001) 4276-4286.
catalyzes 3-O-deacylation of lipid A precursors in membranes of Salmonella
[95]
MA
typhimurium, J Biol Chem 276 (2001) 9083-9092. K. Kawasaki, R.K. Ernst, S.I. Miller, Inhibition of Salmonella enterica serovar
TE
D
Typhimurium lipopolysaccharide deacylation by aminoarabinose membrane modification, J Bacteriol 187 (2005) 2448-2457. K. Kawasaki, T. Manabe, Latency of the lipid A deacylase PagL is involved in producing
AC CE P
[96]
a robust permeation barrier in the outer membrane of Salmonella enterica, J Bacteriol 192 (2010) 5837-5840. [97]
K. Kawasaki, K. China, M. Nishijima, Release of the lipopolysaccharide deacylase PagL from latency compensates for a lack of lipopolysaccharide aminoarabinose modificationdependent resistance to the antimicrobial peptide polymyxin B in Salmonella enterica, J Bacteriol 189 (2007) 4911-4919.
[98]
J.V. Hankins, J.A. Madsen, D.K. Giles, J.S. Brodbelt, M.S. Trent, Amino acid addition to Vibrio cholerae LPS establishes a link between surface remodeling in gram-positive and gram-negative bacteria, Proc Natl Acad Sci U S A 109 (2012) 8722-8727.
39
ACCEPTED MANUSCRIPT [99]
T.W. Cullen, D.K. Giles, L.N. Wolf, C. Ecobichon, I.G. Boneca, M.S. Trent, Helicobacter pylori versus the host: remodeling of the bacterial outer membrane is
PT
required for survival in the gastric mucosa, PLoS Pathog 7 (2011) e1002454.
RI
[100] K. Nummila, I. Kilpelainen, U. Zahringer, M. Vaara, I.M. Helander, Lipopolysaccharides of polymyxin B-resistant mutants of Escherichia coli are extensively substituted by 2-
SC
aminoethyl pyrophosphate and contain aminoarabinose in lipid A, Mol Microbiol 16
NU
(1995) 271-278.
[101] H. Lee, F.F. Hsu, J. Turk, E.A. Groisman, The PmrA-regulated pmrC gene mediates
MA
phosphoethanolamine modification of lipid A and polymyxin resistance in Salmonella enterica, J Bacteriol 186 (2004) 4124-4133.
TE
D
[102] C.M. Reynolds, S.R. Kalb, R.J. Cotter, C.R. Raetz, A phosphoethanolamine transferase specific for the outer 3-deoxy-D-manno-octulosonic acid residue of Escherichia coli
AC CE P
lipopolysaccharide. Identification of the eptB gene and Ca2+ hypersensitivity of an eptB deletion mutant, J Biol Chem 280 (2005) 21202-21211. [103] N. Figueroa-Bossi, S. Lemire, D. Maloriol, R. Balbontin, J. Casadesus, L. Bossi, Loss of Hfq activates the sigmaE-dependent envelope stress response in Salmonella enterica, Mol Microbiol 62 (2006) 838-852. [104] A.D. Cox, J.C. Wright, J. Li, D.W. Hood, E.R. Moxon, J.C. Richards, Phosphorylation of the lipid A region of meningococcal lipopolysaccharide: identification of a family of transferases that add phosphoethanolamine to lipopolysaccharide, J Bacteriol 185 (2003) 3270-3277. [105] L.A. Lewis, B. Choudhury, J.T. Balthazar, L.E. Martin, S. Ram, P.A. Rice, D.S. Stephens, R. Carlson, W.M. Shafer, Phosphoethanolamine substitution of lipid A and
40
ACCEPTED MANUSCRIPT resistance of Neisseria gonorrhoeae to cationic antimicrobial peptides and complementmediated killing by normal human serum, Infect Immun 77 (2009) 1112-1120.
PT
[106] M.M. Hobbs, J.E. Anderson, J.T. Balthazar, J.L. Kandler, R.W. Carlson, J. Ganguly,
RI
A.A. Begum, J.A. Duncan, J.T. Lin, P.F. Sparling, A.E. Jerse, W.M. Shafer, Lipid A's
and men, MBio 4 (2013) e00892-00813.
SC
structure mediates Neisseria gonorrhoeae fitness during experimental infection of mice
NU
[107] T. Murata, W. Tseng, T. Guina, S.I. Miller, H. Nikaido, PhoPQ-mediated regulation produces a more robust permeability barrier in the outer membrane of Salmonella
MA
enterica serovar typhimurium, J Bacteriol 189 (2007) 7213-7222. [108] M.S. Trent, A.A. Ribeiro, S. Lin, R.J. Cotter, C.R. Raetz, An inner membrane enzyme in
TE
D
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
AC CE P
Chem 276 (2001) 43122-43131.
[109] M.A. Hamad, F. Di Lorenzo, A. Molinaro, M.A. Valvano, Aminoarabinose is essential for lipopolysaccharide export and intrinsic antimicrobial peptide resistance in Burkholderia cenocepacia(dagger), Mol Microbiol 85 (2012) 962-974. [110] N. Marr, A. Tirsoaga, D. Blanot, R. Fernandez, M. Caroff, Glucosamine found as a substituent of both phosphate groups in Bordetella lipid A backbones: role of a BvgASactivated ArnT ortholog, J Bacteriol 190 (2008) 4281-4290. [111] N.R. Shah, S. Albitar-Nehme, E. Kim, N. Marr, A. Novikov, M. Caroff, R.C. Fernandez, Minor modifications to the phosphate groups and the C3' acyl chain length of lipid A in two Bordetella pertussis strains, BP338 and 18-323, independently affect Toll-like receptor 4 protein activation, J Biol Chem 288 (2013) 11751-11760.
41
ACCEPTED MANUSCRIPT [112] N.R. Shah, R.E. Hancock, R.C. Fernandez, Bordetella pertussis lipid A glucosamine modification confers resistance to cationic antimicrobial peptides and increases resistance
PT
to outer membrane perturbation, Antimicrob Agents Chemother 58 (2014) 4931-4934.
RI
[113] X. Wang, A.A. Ribeiro, Z. Guan, S.C. McGrath, R.J. Cotter, C.R. Raetz, Structure and biosynthesis of free lipid A molecules that replace lipopolysaccharide in Francisella
SC
tularensis subsp. novicida, Biochemistry 45 (2006) 14427-14440.
NU
[114] N.J. Phillips, B. Schilling, M.K. McLendon, M.A. Apicella, B.W. Gibson, Novel modification of lipid A of Francisella tularensis, Infect Immun 72 (2004) 5340-5348.
MA
[115] X. Wang, A.A. Ribeiro, Z. Guan, C.R. Raetz, Identification of undecaprenyl phosphatebeta-D-galactosamine in Francisella novicida and its function in lipid A modification,
TE
D
Biochemistry 48 (2009) 1162-1172. [116] A.C. Llewellyn, J. Zhao, F. Song, J. Parvathareddy, Q. Xu, B.A. Napier, H. Laroui, D.
AC CE P
Merlin, J.E. Bina, P.A. Cotter, M.A. Miller, C.R. Raetz, D.S. Weiss, NaxD is a deacetylase required for lipid A modification and Francisella pathogenesis, Mol Microbiol 86 (2012) 611-627. [117] K. Ilg, K. Endt, B. Misselwitz, B. Stecher, M. Aebi, W.D. Hardt, O-antigen-negative Salmonella enterica serovar Typhimurium is attenuated in intestinal colonization but elicits colitis in streptomycin-treated mice, Infect Immun 77 (2009) 2568-2575. [118] M. Vonkavaara, S.T. Pavel, K. Holzl, R. Nordfelth, A. Sjostedt, S. Stoven, Francisella is sensitive to insect antimicrobial peptides, J Innate Immun 5 (2013) 50-59. [119] L. Pescaretti Mde, F.E. Lopez, R.D. Morero, M.A. Delgado, The PmrA/PmrB regulatory system controls the expression of the wzzfepE gene involved in the O-antigen synthesis of Salmonella enterica serovar Typhimurium, Microbiology 157 (2011) 2515-2521.
42
ACCEPTED MANUSCRIPT [120] R. Morona, C. Daniels, L. Van Den Bosch, Genetic modulation of Shigella flexneri 2a lipopolysaccharide O antigen modal chain length reveals that it has been optimized for
PT
virulence, Microbiology 149 (2003) 925-939.
RI
[121] D. Bravo, C. Silva, J.A. Carter, A. Hoare, S.A. Alvarez, C.J. Blondel, M. Zaldivar, M.A. Valvano, I. Contreras, Growth-phase regulation of lipopolysaccharide O-antigen chain
SC
length influences serum resistance in serovars of Salmonella, J Med Microbiol 57 (2008)
NU
938-946.
[122] R.W. Crawford, A.M. Keestra, S.E. Winter, M.N. Xavier, R.M. Tsolis, V. Tolstikov, A.J.
MA
Baumler, Very long O-antigen chains enhance fitness during Salmonella-induced colitis by increasing bile resistance, PLoS Pathog 8 (2012) e1002918.
TE
D
[123] Y. Lequette, E.P. Greenberg, Timing and localization of rhamnolipid synthesis gene expression in Pseudomonas aeruginosa biofilms, J Bacteriol 187 (2005) 37-44.
AC CE P
[124] M.E. Davey, N.C. Caiazza, G.A. O'Toole, Rhamnolipid surfactant production affects biofilm architecture in Pseudomonas aeruginosa PAO1, J Bacteriol 185 (2003) 10271036.
[125] L. Hall-Stoodley, J.W. Costerton, P. Stoodley, Bacterial biofilms: from the natural environment to infectious diseases, Nat Rev Microbiol 2 (2004) 95-108. [126] F. Mika, R. Hengge, Small Regulatory RNAs in the Control of Motility and Biofilm Formation in E. coli and Salmonella, Int J Mol Sci 14 (2013) 4560-4579. [127] C. de la Fuente-Nunez, F. Reffuveille, L. Fernandez, R.E. Hancock, Bacterial biofilm development as a multicellular adaptation: antibiotic resistance and new therapeutic strategies, Curr Opin Microbiol 16 (2013) 580-589.
43
ACCEPTED MANUSCRIPT [128] H. Altman, D. Steinberg, Y. Porat, A. Mor, D. Fridman, M. Friedman, G. Bachrach, In vitro assessment of antimicrobial peptides as potential agents against several oral
PT
bacteria, J Antimicrob Chemother 58 (2006) 198-201.
RI
[129] A. Folkesson, J.A. Haagensen, C. Zampaloni, C. Sternberg, S. Molin, Biofilm induced tolerance towards antimicrobial peptides, PLoS One 3 (2008) e1891.
SC
[130] N. Hoiby, T. Bjarnsholt, M. Givskov, S. Molin, O. Ciofu, Antibiotic resistance of
NU
bacterial biofilms, Int J Antimicrob Agents 35 (2010) 322-332. [131] J.A. Haagensen, M. Klausen, R.K. Ernst, S.I. Miller, A. Folkesson, T. Tolker-Nielsen, S.
MA
Molin, Differentiation and distribution of colistin- and sodium dodecyl sulfate-tolerant cells in Pseudomonas aeruginosa biofilms, J Bacteriol 189 (2007) 28-37.
TE
D
[132] S.J. Pamp, M. Gjermansen, H.K. Johansen, T. Tolker-Nielsen, Tolerance to the antimicrobial peptide colistin in Pseudomonas aeruginosa biofilms is linked to
AC CE P
metabolically active cells, and depends on the pmr and mexAB-oprM genes, Mol Microbiol 68 (2008) 223-240. [133] S. Molin, T. Tolker-Nielsen, Gene transfer occurs with enhanced efficiency in biofilms and induces enhanced stabilisation of the biofilm structure, Curr Opin Biotechnol 14 (2003) 255-261.
[134] L. Zhang, T.F. Mah, Involvement of a novel efflux system in biofilm-specific resistance to antibiotics, J Bacteriol 190 (2008) 4447-4452. [135] R.J. Gillis, K.G. White, K.H. Choi, V.E. Wagner, H.P. Schweizer, B.H. Iglewski, Molecular basis of azithromycin-resistant Pseudomonas aeruginosa biofilms, Antimicrob Agents Chemother 49 (2005) 3858-3867.
44
ACCEPTED MANUSCRIPT [136] A. Jolivet-Gougeon, M. Bonnaure-Mallet, Biofilms as a mechanism of bacterial resistance, Drug Discov Today Technol 11 (2014) 49-56.
PT
[137] S.M. Soto, Role of efflux pumps in the antibiotic resistance of bacteria embedded in a
RI
biofilm, Virulence 4 (2013) 223-229.
[138] D. de Beer, P. Stoodley, Z. Lewandowski, Measurement of local diffusion coefficients in
SC
biofilms by microinjection and confocal microscopy, Biotechnol Bioeng 53 (1997) 151-
NU
158.
[139] T.F. Mah, B. Pitts, B. Pellock, G.C. Walker, P.S. Stewart, G.A. O'Toole, A genetic basis
MA
for Pseudomonas aeruginosa biofilm antibiotic resistance, Nature 426 (2003) 306-310. [140] C. Vuotto, F. Longo, M.P. Balice, G. Donelli, P.E. Varaldo, Antibiotic Resistance
TE
D
Related to Biofilm Formation in Klebsiella pneumoniae, Pathogens 3 (2014) 743-758. [141] E. Drenkard, Antimicrobial resistance of Pseudomonas aeruginosa biofilms, Microbes
AC CE P
Infect 5 (2003) 1213-1219.
[142] A. Alvarez-Ordonez, A. Fernandez, A. Bernardo, M. Lopez, Comparison of acids on the induction of an Acid Tolerance Response in Salmonellatyphimurium, consequences for food safety, Meat Sci 81 (2009) 65-70. [143] S.I. Miller, A.M. Kukral, J.J. Mekalanos, A two-component regulatory system (phoP phoQ) controls Salmonella typhimurium virulence, Proc Natl Acad Sci U S A 86 (1989) 5054-5058. [144] S.I. Miller, J.J. Mekalanos, Constitutive expression of the phoP regulon attenuates Salmonella virulence and survival within macrophages, J Bacteriol 172 (1990) 24852490.
45
ACCEPTED MANUSCRIPT [145] E.A. Groisman, E. Chiao, C.J. Lipps, F. Heffron, Salmonella typhimurium phoP virulence gene is a transcriptional regulator, Proc Natl Acad Sci U S A 86 (1989) 7077-
PT
7081.
RI
[146] K.L. Roland, L.E. Martin, C.R. Esther, J.K. Spitznagel, Spontaneous pmrA mutants of Salmonella typhimurium LT2 define a new two-component regulatory system with a
SC
possible role in virulence, J Bacteriol 175 (1993) 4154-4164.
NU
[147] J.S. Gunn, S.I. Miller, PhoP-PhoQ activates transcription of pmrAB, encoding a twocomponent regulatory system involved in Salmonella typhimurium antimicrobial peptide
MA
resistance, J Bacteriol 178 (1996) 6857-6864.
[148] E.A. Groisman, J. Kayser, F.C. Soncini, Regulation of polymyxin resistance and
TE
D
adaptation to low-Mg2+ environments, J Bacteriol 179 (1997) 7040-7045. [149] E.G. Vescovi, Y.M. Ayala, E. Di Cera, E.A. Groisman, Characterization of the bacterial
AC CE P
sensor protein PhoQ. Evidence for distinct binding sites for Mg2+ and Ca2+, J Biol Chem 272 (1997) 1440-1443. [150] E.L. Macfarlane, A. Kwasnicka, R.E. Hancock, Role of Pseudomonas aeruginosa PhoPphoQ in resistance to antimicrobial cationic peptides and aminoglycosides, Microbiology 146 ( Pt 10) (2000) 2543-2554. [151] A. Kato, T. Latifi, E.A. Groisman, Closing the loop: the PmrA/PmrB two-component system negatively controls expression of its posttranscriptional activator PmrD, Proc Natl Acad Sci U S A 100 (2003) 4706-4711. [152] M.W. Bader, W.W. Navarre, W. Shiau, H. Nikaido, J.G. Frye, M. McClelland, F.C. Fang, S.I. Miller, Regulation of Salmonella typhimurium virulence gene expression by cationic antimicrobial peptides, Mol Microbiol 50 (2003) 219-230.
46
ACCEPTED MANUSCRIPT [153] M.W. Bader, S. Sanowar, M.E. Daley, A.R. Schneider, U. Cho, W. Xu, R.E. Klevit, H. Le Moual, S.I. Miller, Recognition of antimicrobial peptides by a bacterial sensor kinase,
PT
Cell 122 (2005) 461-472.
RI
[154] E. Choi, E.A. Groisman, D. Shin, Activated by different signals, the PhoP/PhoQ twocomponent system differentially regulates metal uptake, J Bacteriol 191 (2009) 7174-
SC
7181.
NU
[155] S.M. Richards, K.L. Strandberg, J.S. Gunn, Salmonella-regulated lipopolysaccharide modifications, Subcell Biochem 53 (2010) 101-122.
MA
[156] E. Garcia Vescovi, F.C. Soncini, E.A. Groisman, Mg2+ as an extracellular signal: environmental regulation of Salmonella virulence, Cell 84 (1996) 165-174.
149.
TE
D
[157] M. Otto, Bacterial sensing of antimicrobial peptides, Contrib Microbiol 16 (2009) 136-
AC CE P
[158] S.M. Richards, K.L. Strandberg, M. Conroy, J.S. Gunn, Cationic antimicrobial peptides serve as activation signals for the Salmonella Typhimurium PhoPQ and PmrAB regulons in vitro and in vivo, Front Cell Infect Microbiol 2 (2012) 102. [159] S.Y. Park, E.A. Groisman, Signal-specific temporal response by the Salmonella PhoP/PhoQ regulatory system, Mol Microbiol 91 (2014) 135-144. [160] E.J. Rubin, C.M. Herrera, A.A. Crofts, M.S. Trent, Making the connection: PmrD is required for modifications to Escherichia coli endotoxin that promote antimicrobial resistance, Antimicrob Agents Chemother (2015). [161] J.E. Moss, P.E. Fisher, B. Vick, E.A. Groisman, A. Zychlinsky, The regulatory protein PhoP controls susceptibility to the host inflammatory response in Shigella flexneri, Cell Microbiol 2 (2000) 443-452.
47
ACCEPTED MANUSCRIPT [162] A. Llama-Palacios, E. Lopez-Solanilla, C. Poza-Carrion, F. Garcia-Olmedo, P. Rodriguez-Palenzuela, The Erwinia chrysanthemi phoP-phoQ operon plays an important
PT
role in growth at low pH, virulence and bacterial survival in plant tissue, Mol Microbiol
RI
49 (2003) 347-357.
[163] M. Falord, U. Mader, A. Hiron, M. Debarbouille, T. Msadek, Investigation of the
SC
Staphylococcus aureus GraSR regulon reveals novel links to virulence, stress response
NU
and cell wall signal transduction pathways, PLoS One 6 (2011) e21323. [164] M. Li, Y. Lai, A.E. Villaruz, D.J. Cha, D.E. Sturdevant, M. Otto, Gram-positive three-
MA
component antimicrobial peptide-sensing system, Proc Natl Acad Sci U S A 104 (2007) 9469-9474.
TE
D
[165] M. Fridman, G.D. Williams, U. Muzamal, H. Hunter, K.W. Siu, D. Golemi-Kotra, Two unique phosphorylation-driven signaling pathways crosstalk in Staphylococcus aureus to
7986.
AC CE P
modulate the cell-wall charge: Stk1/Stp1 meets GraSR, Biochemistry 52 (2013) 7975-
[166] H. Sun, Y. Yang, T. Xue, B. Sun, Modulation of cell wall synthesis and susceptibility to vancomycin by the two-component system AirSR in Staphylococcus aureus NCTC8325, BMC Microbiol 13 (2013) 286. [167] M. Pietiainen, P. Francois, H.L. Hyyrylainen, M. Tangomo, V. Sass, H.G. Sahl, J. Schrenzel, V.P. Kontinen, Transcriptome analysis of the responses of Staphylococcus aureus to antimicrobial peptides and characterization of the roles of vraDE and vraSR in antimicrobial resistance, BMC Genomics 10 (2009) 429. [168] P. Monsieurs, S. De Keersmaecker, W.W. Navarre, M.W. Bader, F. De Smet, M. McClelland, F.C. Fang, B. De Moor, J. Vanderleyden, K. Marchal, Comparison of the
48
ACCEPTED MANUSCRIPT PhoPQ regulon in Escherichia coli and Salmonella typhimurium, J Mol Evol 60 (2005) 462-474.
PT
[169] P. Mandin, H. Fsihi, O. Dussurget, M. Vergassola, E. Milohanic, A. Toledo-Arana, I.
RI
Lasa, J. Johansson, P. Cossart, VirR, a response regulator critical for Listeria monocytogenes virulence, Mol Microbiol 57 (2005) 1367-1380.
SC
[170] J.L. Yu, L. Guo, Quantitative Proteomic Analysis of Salmonella enterica Serovar
NU
Typhimurium under PhoP/PhoQ Activation Conditions, J Proteome Res 10 (2011) 29923002.
Bacteriol 183 (2001) 1835-1842.
MA
[171] E.A. Groisman, The pleiotropic two-component regulatory system PhoP-PhoQ, J
TE
D
[172] R.K. Ernst, T. Guina, S.I. Miller, Salmonella typhimurium outer membrane remodeling: role in resistance to host innate immunity, Microbes Infect 3 (2001) 1327-1334.
AC CE P
[173] M.D. Snavely, C.G. Miller, M.E. Maguire, The mgtB Mg2+ transport locus of Salmonella typhimurium encodes a P-type ATPase, J Biol Chem 266 (1991) 815-823. [174] R. Tamayo, B. Choudhury, A. Septer, M. Merighi, R. Carlson, J.S. Gunn, Identification of cptA, a PmrA-regulated locus required for phosphoethanolamine modification of the Salmonella enterica serovar typhimurium lipopolysaccharide core, J Bacteriol 187 (2005) 3391-3399. [175] A.X. Tran, M.E. Lester, C.M. Stead, C.R. Raetz, D.J. Maskell, S.C. McGrath, R.J. Cotter, M.S. Trent, Resistance to the antimicrobial peptide polymyxin requires myristoylation of Escherichia coli and Salmonella typhimurium lipid A, J Biol Chem 280 (2005) 2818628194.
49
ACCEPTED MANUSCRIPT [176] K. Nishino, F.F. Hsu, J. Turk, M.J. Cromie, M.M. Wosten, E.A. Groisman, Identification of the lipopolysaccharide modifications controlled by the Salmonella PmrA/PmrB system
PT
mediating resistance to Fe(III) and Al(III), Mol Microbiol 61 (2006) 645-654.
RI
[177] S.M. Moskowitz, M.K. Brannon, N. Dasgupta, M. Pier, N. Sgambati, A.K. Miller, S.E. Selgrade, S.I. Miller, M. Denton, S.P. Conway, H.K. Johansen, N. Hoiby, PmrB
SC
mutations promote polymyxin resistance of Pseudomonas aeruginosa isolated from
NU
colistin-treated cystic fibrosis patients, Antimicrob Agents Chemother 56 (2012) 10191030.
MA
[178] L.A. Arroyo, C.M. Herrera, L. Fernandez, J.V. Hankins, M.S. Trent, R.E. Hancock, The pmrCAB operon mediates polymyxin resistance in Acinetobacter baumannii ATCC
TE
D
17978 and clinical isolates through phosphoethanolamine modification of lipid A, Antimicrob Agents Chemother 55 (2011) 3743-3751.
AC CE P
[179] M.D. Adams, G.C. Nickel, S. Bajaksouzian, H. Lavender, A.R. Murthy, M.R. Jacobs, R.A. Bonomo, Resistance to colistin in Acinetobacter baumannii associated with mutations in the PmrAB two-component system, Antimicrob Agents Chemother 53 (2009) 3628-3634.
[180] H.Y. Cheng, Y.F. Chen, H.L. Peng, Molecular characterization of the PhoPQ-PmrDPmrAB mediated pathway regulating polymyxin B resistance in Klebsiella pneumoniae CG43, J Biomed Sci 17 (2010) 60. [181] S. Nakka, M. Qi, Y. Zhao, The PmrAB system in Erwinia amylovora renders the pathogen more susceptible to polymyxin B and more resistant to excess iron, Res Microbiol 161 (2010) 153-157.
50
ACCEPTED MANUSCRIPT [182] A.K. Miller, M.K. Brannon, L. Stevens, H.K. Johansen, S.E. Selgrade, S.I. Miller, N. Hoiby, S.M. Moskowitz, PhoQ mutations promote lipid A modification and polymyxin
RI
Antimicrob Agents Chemother 55 (2011) 5761-5769.
PT
resistance of Pseudomonas aeruginosa found in colistin-treated cystic fibrosis patients,
[183] N.S. Ly, J. Yang, J.B. Bulitta, B.T. Tsuji, Impact of two-component regulatory systems
SC
PhoP-PhoQ and PmrA-PmrB on colistin pharmacodynamics in Pseudomonas aeruginosa,
NU
Antimicrob Agents Chemother 56 (2012) 3453-3456.
[184] R. Tamayo, A.M. Prouty, J.S. Gunn, Identification and functional analysis of Salmonella
MA
enterica serovar Typhimurium PmrA-regulated genes, FEMS Immunol Med Microbiol 43 (2005) 249-258.
TE
D
[185] M. Merighi, A. Carroll-Portillo, A.N. Septer, A. Bhatiya, J.S. Gunn, Role of Salmonella enterica serovar Typhimurium two-component system PreA/PreB in modulating PmrA-
AC CE P
regulated gene transcription, J Bacteriol 188 (2006) 141-149. [186] D. Kraus, S. Herbert, S.A. Kristian, A. Khosravi, V. Nizet, F. Gotz, A. Peschel, The GraRS regulatory system controls Staphylococcus aureus susceptibility to antimicrobial host defenses, BMC Microbiol 8 (2008) 85. [187] M. Meehl, S. Herbert, F. Gotz, A. Cheung, Interaction of the GraRS two-component system with the VraFG ABC transporter to support vancomycin-intermediate resistance in Staphylococcus aureus, Antimicrob Agents Chemother 51 (2007) 2679-2689. [188] S. Gruenheid, H. Le Moual, Resistance to antimicrobial peptides in Gram-negative bacteria, FEMS Microbiol Lett 330 (2012) 81-89. [189] M.G. Habets, M.A. Brockhurst, Therapeutic antimicrobial peptides may compromise natural immunity, Biol Lett 8 (2012) 416-418.
51
ACCEPTED MANUSCRIPT [190] O. Samuelsen, H.H. Haukland, H. Jenssen, M. Kramer, K. Sandvik, H. Ulvatne, L.H. Vorland, Induced resistance to the antimicrobial peptide lactoferricin B in
PT
Staphylococcus aureus, FEBS Lett 579 (2005) 3421-3426.
RI
[191] J. Mehla, S.K. Sood, Substantiation in Enterococcus faecalis of dose-dependent resistance and cross-resistance to pore-forming antimicrobial peptides by use of a
SC
polydiacetylene-based colorimetric assay, Appl Environ Microbiol 77 (2011) 786-793.
NU
[192] J.E. Pollard, J. Snarr, V. Chaudhary, J.D. Jennings, H. Shaw, B. Christiansen, J. Wright, W. Jia, R.E. Bishop, P.B. Savage, In vitro evaluation of the potential for resistance
MA
development to ceragenin CSA-13, J Antimicrob Chemother 67 (2012) 2665-2672. [193] C.R. Raetz, C.M. Reynolds, M.S. Trent, R.E. Bishop, Lipid A modification systems in
TE
D
gram-negative bacteria, Annu Rev Biochem 76 (2007) 295-329. [194] J.W. Handing, A.K. Criss, The Lipooligosaccharide Modifying Enzyme LptA Enhances
AC CE P
Gonococcal Defense Against Human Neutrophils, Cell Microbiol (2014). [195] J.V. Hankins, J.A. Madsen, D.K. Giles, B.M. Childers, K.E. Klose, J.S. Brodbelt, M.S. Trent, Elucidation of a novel Vibrio cholerae lipid A secondary hydroxy-acyltransferase and its role in innate immune recognition, Mol Microbiol 81 (2011) 1313-1329. [196] K. Kawasaki, R.K. Ernst, S.I. Miller, 3-O-deacylation of lipid A by PagL, a PhoP/PhoQregulated deacylase of Salmonella typhimurium, modulates signaling through Toll-like receptor 4, J Biol Chem 279 (2004) 20044-20048. [197] K.E. McAuley, P.K. Fyfe, J.P. Ridge, N.W. Isaacs, R.J. Cogdell, M.R. Jones, Structural details of an interaction between cardiolipin and an integral membrane protein, Proc Natl Acad Sci U S A 96 (1999) 14706-14711.
52
ACCEPTED MANUSCRIPT [198] O. Schneewind, D. Missiakas, Lipoteichoic acids, phosphate-containing polymers in the envelope of gram-positive bacteria, J Bacteriol 196 (2014) 1133-1142.
PT
[199] W. Fischer, Lipoteichoic acid and lipids in the membrane of Staphylococcus aureus, Med
RI
Microbiol Immunol 183 (1994) 61-76.
[200] K.S. Molnar, M. Bonomi, R. Pellarin, G.D. Clinthorne, G. Gonzalez, S.D. Goldberg, M.
SC
Goulian, A. Sali, W.F. DeGrado, Cys-scanning disulfide crosslinking and bayesian
NU
modeling probe the transmembrane signaling mechanism of the histidine kinase, PhoQ,
AC CE P
TE
D
MA
Structure 22 (2014) 1239-1251.
53
ACCEPTED MANUSCRIPT Table 1: Details of the modifications and the enzymes responsible for the LPS modifications that are detailed in Figure 2 and Figure3.
E.
coli,
RI
ArnT S. enterica, (orf5, aeruginosa PmrK, or ifbI)
Protein function Reference Adds glycine to the secondary acyl chain [29] of the hexa-acylated lipid A
PT
Organism V. cholerae
P. Adds L-Ara4N (4-amino-4-deoxy-L-arabinose) [108, 110] to one or both of the phosphate groups of lipid A
SC
Gene AlmG
F. Adds galactosamine (GalN) to the phosphate [110, 113, tularensis subspecies novicida substituent at the 1-position of lipid A 193]
ArnT ortholog
Bordetella pertussis
EptA (pmrC)
H. pylory, S. enterica, E. coli
EptB
S. enterica, E.coli
LptA
N. gonorrhoeae, meningitides
LpxF
H. pylory, F. tularensis
Dephosphorylates lipid A at position 4’ [99, 193] leaving unmodified hydroxyl group.
LpxM
E. coli
Adds myristate (C14:0) to the myristate on the [195] 3′-position of the glucosamine disaccharide base of the Lipid A
LpxN
V. cholerae
Adds of a 3-hydroxylaurate (3-OH C12:0) to [195] the myristate on the 3′-position of the glucosamine disaccharide base of the Lipid A
LpxO
S. enterica
Hydroxylzes the 2’ position of the myristoyl [72, 107] residue at the 3′ position of the glucosamine disaccharide base of the Lipid A
PagL
S. enterica
Removes 3-hydroxymyristate from position 3’
PagP
S. enterica
Adds palmitoyl group to PG on the outer [72, 196] membrane and Lipid A
NU
ArnT ortholog
MA
Adds of glucosamine GlcN to both phosphate [110] groups of lipid A
AC CE P
TE
D
Adds phosphoethanolamine (PEA) to the 4’ [99, 101] phosphate on lipid A
Adds phosphoethanolamine (PEA) to the 3- [102] deoxy-D-manno-octulosonic acid (Kdo) of the inner core
Neisseria Adds phosphoethanolamine (PEA) to the 4’ [104, 105, phosphate on lipid A 194]
54
[72, 196]
ACCEPTED MANUSCRIPT
MprF
S. aureus
VraFG
S. aureus
adds D-alanine to the Gro-P moiety of the [77] LTA responsible for the addition of lysine and [56, alanine to PG on the plasma membrane 62] AMP efflux pump
PT
S. aureus
AC CE P
TE
D
MA
NU
SC
RI
DltA
55
[169]
60,
ACCEPTED MANUSCRIPT Figures Legends Figure 1: Outer membrane of Gram-negative bacteria (S. enterica): Schematic
PT
representation of the inner and outer leaflets of the outer membrane. The outer leaflet consists of
RI
LPS and PG. The LPS moiety consists of lipid A- di-glucoseamine linked to fatty acids, and is the major component of the outer leaflet. This lipid A is linked to sugars of the core (colored
SC
hexagons), and the O-antigens (black-lined hexagons). The inner leaflet on the contrary, contains
NU
PG, DPG and LPG or APG [58, 62, 197].
MA
Figure 2: D-alanylation of LTA. Illustration of D-alanylation of LTA of S. aureus. D-alanine is added to the repeated unit of the LTA by the enzyme DltA, and contributes to bacterial resistance
TE
D
[198, 199] .
AC CE P
Figure 3: Lipid A and inner core modifications. The outer surface of Gram-negative bacteria is mostly LPS. This LPS consists of three components (i) lipid A; (ii) core (inner and outer core); (iii) O-antigen (from the inside to the outside). The cartoon summarizes the modifications that take place on the lipid A and the inner core, which were shown to be involved in AMP resistance and susceptibility. Each modification of the lipid A and inner core are colored in a matching color as the name of the enzyme responsible for it. Note * Not all modifications occur in all bacteria species, and they might be relevant to one or more bacterial species (a partial modification per species can be found in Table 1). ** The current illustration refers to a structure of E. coli and S. enterica, but other Gram-negative bacteria might have an altered structures of lipid A and inner core.
56
ACCEPTED MANUSCRIPT Figure 4: TCS activation contributes to AMP resistance. Several TCS are known to be involved in AMP resistance in Gram-positive and Gram-negative bacteria. In Gram-negative
PT
bacteria a sensor histidine kinase, PhoQ or PmrB, is located on the inner membrane (plasma
RI
membrane) and activated by several environmental signals (such as AMPs), some of which are indicated in the cartoon. Upon activation it undergoes phosphorylation and phosphorylates a
SC
response regulator, PhoP and PmrA respectively. The response regulator acts as transcription
NU
factor and activates many genes some of which were found to be related to AMP resistance. There is a crosstalk between these two systems via PmrD, which directly activates PmrA in a
MA
PmrB-independent manner. In Gram-positive bacteria GraRS (ApsSR) is another TCS that is known to be activated by AMPs and is involved in AMP resistance. [57, 107, 160, 177, 186, 187,
AC CE P
TE
D
200]
57
ACCEPTED MANUSCRIPT
AC CE P
TE
D
MA
NU
SC
RI
PT
Figure 1
58
ACCEPTED MANUSCRIPT
AC CE P
TE
D
MA
NU
SC
RI
PT
Figure 2
59
ACCEPTED MANUSCRIPT
AC CE P
TE
D
MA
NU
SC
RI
PT
Figure 3
60
ACCEPTED MANUSCRIPT
AC CE P
TE
D
MA
NU
SC
RI
PT
Figure 4
61
ACCEPTED MANUSCRIPT Declaration
AC CE P
TE
D
MA
NU
SC
RI
PT
The authors have no conflict of interest in this study
62
ACCEPTED MANUSCRIPT Highlights
AC CE P
TE
D
MA
NU
SC
RI
PT
1. An updated review on multidrug resistance bacteria against antimicrobial peptides. 2. Resistance towards AMPs can be developed, by modification of bacterial surface molecules, secretion of protective material and up-regulation or elimination of specific proteins. 3. Because of the general mechanisms of attachment and action of AMPs, bacterial resistance to AMPs often involves biophysical and biochemical changes to the bacterial cell wall. 4. The review focuses on the biophysical, surface and surrounding changes that bacteria undergo in acquiring resistance to AMPs.
63
S0005-2736(15)00179-0 doi: 10.1016/j.bbamem.2015.05.022 BBAMEM 81917
To appear in:
BBA - Biomembranes
Received date: Revised date: Accepted date:
10 February 2015 25 May 2015 26 May 2015
Please cite this article as: Reut Nuri, Tal Shprung, Yechiel Shai, Defensive Remodeling: How bacterial surface properties and biofilm formation promote resistance to antimicrobial peptides, BBA - Biomembranes (2015), doi: 10.1016/j.bbamem.2015.05.022
This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
PT
ACCEPTED MANUSCRIPT
SC
MA
Reut Nuri, Tal Shprung and Yechiel Shai*
NU
promote resistance to antimicrobial peptides
RI
Defensive Remodeling: How bacterial surface properties and biofilm formation
AC CE P
Rehovot, 76100 Israel.
TE
D
Department of Biological Chemistry, the Weizmann Institute of Science,
*To whom correspondence should be addressed, at the Department of Biological Chemistry, The Weizmann Institute of Science, Rehovot, 76100 Israel. Tel: 972-8-9342711; Fax: 972-89344112; E-mail: [email protected]
Key words Antimicrobial peptides, resistance, surface, biophysical properties, biochemical properties, biofilm, cross resistance.
1
ACCEPTED MANUSCRIPT Abstract .....................................................................................................................................................................................4
2.
Introduction ............................................................................................................................................................................4
3.
Bacterial resistance to AMPs as a result of changes in bacterial outer surface ..........................................6
PT
1.
Extracellular polysaccharides ................................................................................................................................7
3.2
Lipid modifications in bacterial membranes ...................................................................................................8
RI
3.1
Phosphatidylglycerol (PG) modifications ......................................................................................... 8
3.2.2
Changing the fatty acids composition ................................................................................................ 9
3.2.3
In-Vitro studies .......................................................................................................................................... 10
NU
SC
3.2.1
LTA D-alanylation of LTA in Gram-positive bacteria ................................................................................ 10
3.4
Surface modifications of Gram-negative bacteria....................................................................................... 11
MA
3.3
3.4.1
Acylation and deacylation of Lipid A ................................................................................................ 12
3.4.2
Addition of glycine to Lipid A .............................................................................................................. 13
D
3.4.3 De-phosphorylation, hydroxylation and addition of Phosphoethanolamine (PE) to Lipid A 14 Addition of amino-sugars to lipid A .................................................................................................. 15
3.4.5
Changing the length of O-antigen....................................................................................................... 16
TE
3.4.4
Modifications of the bacterial outer surface that influence peptide properties. ..................................... 17
5.
Mechanisms of bacterial AMP resistance in biofilm ............................................................................................ 17
6.
AC CE P
4.
5.1
Creation of sub-populations in the biofilm- .................................................................................................. 18
5.2
A higher mutation rate- ......................................................................................................................................... 19
5.3
Upregulation of efflux pumps - ........................................................................................................................... 19
5.4
Induction of TCSs within the biofilm- .............................................................................................................. 19
5.5
Diffusion in biofilm- ................................................................................................................................................ 20
Two component systems (TCSs) regulate AMP resistance .............................................................................. 20 6.1
TCS activation by AMPs ......................................................................................................................................... 21
6.2
Genes activated by TCS that are involved in AMP resistance ................................................................ 22
7.
Ways to overcome AMP resistance ............................................................................................................................ 23
8.
Summary ............................................................................................................................................................................... 25 2
ACCEPTED MANUSCRIPT
TE
D
MA
NU
SC
RI
PT
References............................................................................................................................................................................. 27
AC CE P
9.
3
ACCEPTED MANUSCRIPT
1. Abstract
PT
Multidrug resistance bacteria are a major concern worldwide. These pathogens cannot be treated
RI
with conventional antibiotics and thus alternative therapeutic agents are needed. Antimicrobial
SC
peptides (AMPs) are considered to be good candidates for this purpose. Most AMPs are short and positively charged amphipathic peptides, which are found in all forms of life. AMPs are
NU
known to kill bacteria by binding to the negatively charged bacterial surface, and in most cases cause membrane disruption. Resistance towards AMPs can be developed, by modification of
MA
bacterial surface molecules, secretion of protective material and up-regulation or elimination of specific proteins. Because of the general mechanisms of attachment and action of AMPs,
TE
D
bacterial resistance to AMPs often involves biophysical and biochemical changes such as surface rigidity, cell wall thickness, surface charge, as well as membrane and cell wall modification.
AC CE P
Here we focus on the biophysical, surface and surrounding changes that bacteria undergo in acquiring resistance to AMPs. In addition we discuss the question of whether bacterial resistance to administered AMPs might compromise our innate immunity to endogenous AMPs.
2. Introduction
The fight between bacteria and man, once thought to be at its end when antibiotics were discovered, has gained much attention in recent years. The enhanced and unregulated use of antimicrobials has increased the occurrence of bacteria resistant to multiple antibiotics [1, 2], thus urging the research community to develop new antibacterial agents. One of the most promising strategies explored is the use of antimicrobial peptides (AMPs) [3-6]. AMPs are hydrophobic and amphipathic short peptides, which in most cases have a net positive charge [4, 4
ACCEPTED MANUSCRIPT 7]. Initially AMPs were found in invertebrates [3] and later in all known forms of life including humans [8, 9], where they serve as part of our innate immune system [5, 10, 11]. AMPs rapidly
PT
neutralize a broad range of microbes, both Gram-positive and Gram-negative, mainly through
RI
binding and perturbing their cell envelope, which contains lipoteichoic acid (LTA) or lipopolysaccharide (LPS) respectively, as well as their cytoplasmic membranes [5, 6, 12]. This
SC
makes AMPs promising candidates in the quest to find new and better antimicrobial agents [13].
NU
It is believed that the chemical and physical characteristics of AMPs such as a positive net charge secondary structure and amphipathic nature are important for their antimicrobial
MA
activity [6, 14-16]. For example, in the case of Gram-negative bacteria, due to their net positive charge, AMPs have high affinity to the negatively charged LPS, and then can displace divalent
TE
D
cations that are naturally bound to it. However, it has been shown that some AMPs specifically bind oprI, a lipoprotein of Pseudomonas aeruginosa, located on the outer membrane and
AC CE P
suggested to serve as an AMP receptor. OprI is important for cell wall integrity and has homologs in other Gram-negative bacteria [17]. The binding of AMPs to the bacterial surface causes a local interference with the outer membrane and enables peptides to traverse it. When reaching the inner membrane, AMPs can function either without causing membrane disruption by penetrating the cell and affecting cytoplasmic components such as DNA and DNA-polymerase [18]. Alternatively/in addition, AMPs can function in membrane disruptive manner thus impairing cell integrity [18]. Various mechanisms were suggested for the membrane disruptive activity of AMPs [5, 6, 10, 19]. The main models include: (1) the barrel-stave model - peptides assemble to form pores with their hydrophobic surfaces facing the membrane core [20]; (2) the carpet model – peptides bind to the membrane in a carpet-like manner. When a threshold concentration has been reached, they act as
5
ACCEPTED MANUSCRIPT a detergent and disrupt the membrane [10, 21] and (3) the toroidal model - peptides form transient pores composed of peptides and lipids head groups [22, 23]. These pores were
PT
suggested to be as one of the stages in the carpet mechanism [5].
RI
Bacteria however, have evolved mechanisms to counteract AMPs. There are two main types of resistance, (i) induced resistance, caused by a transient induction of bacterial systems
SC
that help the bacteria cope with AMPs, and (ii) constitutive resistance as a result of genetic
NU
changes [24, 25]. Currently, there are several putative mechanisms known for bacterial resistance to AMPs [26-29]: remodeling of the bacterial surface by making it less negatively charged and
MA
less permeable [30-35], modulation of host immunity gene expression in order to suppress the release of native AMPs [36, 37], biofilm formation [38, 39], proteolytic degradation of AMPs
TE
D
by peptidases and proteases [40, 41], efflux pumps that expel the AMPs [27, 42, 43], and trapping molecules [44-46] and vesicles [47] that capture the peptides and prevent them from
AC CE P
reaching the bacteria.
In this review we focus on the mechanisms that affect the biophysical characteristics of the bacteria and bacterial surrounding such as capsid and biofilm, as well as biochemical changes of the outer surface, including the regulation of some of these processes by two component systems (TCSs).
3. Bacterial resistance to AMPs as a result of changes in bacterial outer surface The outer surface of Gram-positive and Gram-negative bacteria is negatively charged due to LTA and LPS, respectively. AMPs on the other hand, are hydrophobic, positively charged molecules, and therefore it is believed that AMPs are initially attracted to the bacterial surface 6
ACCEPTED MANUSCRIPT due to electrostatic interactions. In line with this, it is assumed that neutralization of the negative charge can reduce peptide binding to the bacterial surface, and will be further discussed. Note
Extracellular polysaccharides
RI
3.1
PT
that everal reviews were published on this subject [26, 48-51].
SC
Extracellular polysaccharides are polymers produced by some Gram-positive and Gram-negative
NU
bacteria. When covalently linked to the cell wall they are termed capsule [28]. The question of whether or not a capsule in this strain contributes to AMP resistance is still under debate [51].
MA
There are some reports indicating that capsules cause AMP-resistance [51-53]. For example, Klebsiella pneumoniae bacteria mutated in capsule production are more susceptible to AMPs and
D
bind better polymyxin B (PmB) compared to the wild-type (WT) [54]. Moreover, PmB was
TE
found to induce capsule production in these bacteria, and it was suggested that the capsule serves
AC CE P
as a shield towards AMPs, thus limiting their interaction with the bacterial surface [55]. In addition, it was shown that K. pneumonia, Streptococcus pneumonia and P. aeruginosa tend to naturally release capsule polysaccharides (CPS) fractions into the environment, which contributes to their resistance to AMPs. This secretion is enhanced in the presence of PmB. Supernatant containing CPS of these strains showed increased resistance in Escherichia coli strain [54]. Moreover, the authors demonstrated that anionic CPS but not cationic or neutral charged CPS of Streptococcus pneumonia, specifically bind dansyl-PmB [54]. In Group B Streptococcus it was demonstrated that bacteria mutated in capsule production did not demonstrate a higher susceptibility to several tested AMPs [35]. To conclude, some studies have demonstrated AMP resistance mediated by capsules and some show that capsules do not change the resistance. We suggest therefore that whether or not a capsule confers AMP resistance depends on the bacterial species, and perhaps on the AMP itself. 7
ACCEPTED MANUSCRIPT 3.2
Lipid modifications in bacterial membranes Phosphatidylglycerol (PG) modifications
PT
3.2.1
In Gram-positive bacteria, a gene encodes for a large membrane protein mprF responsible for the
RI
addition of lysine or alanine to phosphatidylglycerol (PG) to form lysyl-phosphatidylglycerol
SC
(LPG) and alanyl- phosphatidylglycerol (APG), respectively [56], as well as for the translocation of these products to the outer leaflet [56-58]. In several studies, mprF mutants of S. aureus were
NU
found to be more susceptible to AMPs, which might suggest that the addition of these amino
MA
acids to the membrane reduces susceptibility to AMPs [59] (A detailed review regarding AMP resistance mediated by mprF can be found in reference [60]). However, in another study it was
D
shown that there was no change in susceptibility of S. aureus mutants to mprF for the tested
TE
peptides [35].
AC CE P
The membrane of S. aureus consists mostly of unmodified PG (38-76% of the phospholipids), its dimer, diphosphatidylglycerol (DPG) also known as cardiolipin (5-30%) and about 14-38% of LPG, [58] (Figure 1). Some daptomycin (a lipopeptide) resistant S. aureus isolated strains, showed gain of function mutations in mprF and had a higher LPG content compared to the susceptible strains. These strains were found to be more resistant to thrombininduced platelet microbicidal proteins (tPMP), and showed a reduced net surface charge [61]. In line with that, less daptomycin was bound to their surface compared to the other strains, however the binding of classical AMPs was not examined in this study [61]. In the Gram-negative bacteria P. aeruginosa, a homolog for MprF was found, which is responsible for the addition of alanine to PG to form APG. This modification resulted in increased resistance to antimicrobial agents including the AMP protamine sulfate [62].
8
ACCEPTED MANUSCRIPT 3.2.2
Changing the fatty acids composition
It was suggested that membrane lipid composition can affect cell wall properties and change
PT
interactions with AMPs, and therefore the susceptibility to these agents [63]. S. aureus strains
RI
resistant to thrombin-induced platelet microbicidal protein 1 (tPMP-1), showed a higher
SC
proportion of longer and more unsaturated fatty acids in the membrane, as well as higher membrane fluidity compared to the control, but no differences in phospholipid composition [64].
NU
Daptomycin resistant S. aureus isolated strains also showed a higher membrane fluidity
MA
compared to the control [65-67]. Nevertheless, one study reports on a correlation between membrane fluidity and the level of resistance to AMPs [67]. Conversely, other AMP-resistant
D
bacteria showed a higher rigidity. Two S. aureus strains, one resistant to magainin 2 and the
TE
other to gramicidin D, had a higher proportion of LPG in their cell membrane compared to the control strain and both mutants showed higher rigidity and surface positive charge [68]. E.
AC CE P
faecalis pediocin-resistant strains showed a higher ratio of saturated and unsaturated fatty acids, as well as, reduction in branched fatty acids in these strains compared to the WT. These changes in fatty acid composition increase cell membrane rigidity, and are thought to reduce the insertion of pediocin (AMP) to the membrane [69]. In summary, membrane fluidity is not a direct indication for resistance to AMPs, and cell membrane can be more or less fluid in the context of the organism and the AMP. Comparison of nisin resistant and susceptible strains of Listeria monocytogenes revealed that the ratio of PG and DPG was higher than in the WT. An in-vitro nisin binding assay demonstrated that nisin binds better to a lipid-monolayer derived from the WT than the resistant strain [70]. Other studies reported that nisin resistant L. monocytogenes strains had a greater proportion of saturated fatty acids whereas the WT had more unsaturated ones [71]. In 9
ACCEPTED MANUSCRIPT Enterococcus faecalis, a Gram-positive bacterium, pediocin-resistant strains had more LPG and phosphoethanolamine (PEA) in their cell membrane compared to the WT [69]. AMP resistance
PT
in Salmonella enterica Typhimurium, a Gram-negative bacterium, was found to be mediated by
RI
increased levels of DPG and palmitoylated acyl-phosphatidylglyceroles in the inner leaflet of the outer membrane, which is mainly composed of glycerophospholipids. This palmitoylation is
In-Vitro studies
NU
3.2.3
SC
conducted by the protein pagP and regulated by the two-component system PhoPQ [72].
MA
An in- vitro study examined the effect of binding and pore formation ability of AMPs on large unilamellar vesicle (LUVs) composed of different proportion of lipids. LUVs were generated
D
containing 1-palmitoyl-2-oleoyl-phosphatidylglycerol (POPG) and different concentrations of
TE
either zwitterionic Gln-PE or positively charged Lys-PE. It was shown that while increasing the
AC CE P
concentration of the Gln-PE in the membrane does not change the level of bound cecropin A and mastoparan X (AMPs), the increase of the Lys-PE in the membrane reduced the binding. Pore formation assay revealed that increasing concentration of Lys-PE, starting with 10%, reduced the levels of pore formation by both cecropin A and mastoparan X, while 10% of Gln-PE increased pore formation. In both peptides however, higher concentrations resulted in reduced pore forming ability [73].
3.3
LTA D-alanylation of LTA in Gram-positive bacteria
The surface of Gram-positive bacteria consists of the cytoplasmic membrane, a thick peptidoglycan layer and lipoteichoic acid (LTA). It has been reported that D-alanylation of the LTA contributes to bacterial resistance to AMPs [74-76]. It was found that the dlt operon is responsible for the incorporation of D-alanine (D-ala) into teichoic acid, which results in an 10
ACCEPTED MANUSCRIPT increase in the positive charge of the bacterial surface. Mutants of dltA, the gene encoding a Dalanyl carrier protein ligase (that adds D-alanine to the Gro-P moiety of the LTA) showed
PT
increased susceptibility to AMPs, whereas overexpression of this gene confers increased
RI
resistance [77] (Figure 2, Table 1).
Reducing the net negative charge of the LTA is considered to contribute to the reduction of
SC
electrostatic interactions between the positively charged AMPs and the negatively charged
NU
bacterial surface, thus reducing susceptibility [34, 76, 78, 79]. However, recent experimental data on Group B Streptococus (GBS) indicate that there is no significant change in the amount of
MA
peptide bound to the dltA mutant bacteria compared to the WT. It was proposed that the higher resistance observed in the WT compared to the dltA mutant is due to an increase in cell wall
TE
D
density and reduced AMP penetration [35], i.e. the addition of D-alanine to Gro-P moiety of the LTA does not change the amount of bound peptide, but rather serves as a shield that prevents
AC CE P
peptides from entering the bacteria. No difference in cell wall thickness was found in this study [35] as well as in other studies [35, 61, 80]. However, several studies showed a thicker cell wall of different AMP- resistant bacteria [81, 82], which suggests that cell wall thickness might be, but is not necessarily, correlated with resistance. Importantly, the D-alanylation of the LTA contributes to the resistance of bacteria to only some AMPs [35] and so far, other known modifications of the LTA (e.g. the addition of phosphocholine and glycosylation) were not tested for their effect on resistance to antimicrobial peptides [83].
3.4
Surface modifications of Gram-negative bacteria
The surface of Gram-negative bacteria is complex and covered with several constituents such as LPS, proteinaceous curli fibers, cellulose and capsules [84-88]. LPS, which is thought to be the main surface element, consists of a conserved lipophilic component, lipid-A, linked with 11
ACCEPTED MANUSCRIPT different polysaccharides or oligosaccharides, giving it a net negative charge. The LPS composition is dynamic; it may be modified and is found to be diverse in length and composition
PT
amongst the different Gram-negative bacteria species [87]. This unique structure, besides helping
RI
to maintain cellular equilibrium, serves as a physical barrier and may provide protection against antibacterial agents [19, 89]. Some LPS modifications were shown to contribute to bacterial
SC
resistance to AMPs. The modifications of the LPS depend on bacterial species and
NU
environmental conditions, and it was shown that not all modifications are found in all bacteria species [71]. Here we will discuss some of the changes in the outer surface with respect to AMP
MA
resistance and susceptibility. Lipid A and inner core modifications that are known to affect AMP
Acylation and deacylation of Lipid A
TE
3.4.1
D
resistance are summarized in Figure 3 and Table 1.
AC CE P
Addition of an acyl chain to Lipid A was found to contribute to bacterial resistance to AMPs. PagP is a gene that encodes for an outer membrane palmitoyl transferase in S. enteric Typhimurium [72]. This enzyme adds additional palmitate (C16:0 acyl chain) to the lipid A moiety and this acylation is thought to increase the hydrophobic interactions between the acyl chains in the Lipid A, thus increasing outer membrane fluidity [90-92] (Figure 3). S. enterica Typhimurium mutants in pagP showed higher membrane permeability in response to AMPs compared to the control strain, as well as, Legionella pneumophila mutants in Rcp, a PagP homolog that showed a higher susceptibility to C18G and PmB [93] (Figure 3). To support the finding that lipid A acylation contributes to bacterial resistance to AMPs, deacylation was found to increase bacteria susceptibility to AMPs. PagL is an enzyme responsible for deacylation of lipid A. It is located on the outer membrane of several Gramnegative bacteria such as S. enterica Typhimurium, and removes R-3-hydroxymyristate from 12
ACCEPTED MANUSCRIPT position 3 of some lipid A precursors [94]. It was found that pagL is expressed upon PhoPQ activation but no deacylation of lipid A was detected under these conditions. Moreover, it was
PT
demonstrated that pagL is latent in the WT, and the regulation of the latency involves
RI
aminoarabinose modifications of the lipid A regulated by pmrA. Therefore, the pagL null mutation did not change cell permeability to ethidium compared to the WT [94]. However,
SC
permeability of pmrA-pagL double mutant was lower compared to pmrA mutant, that lacks
NU
aminoarabinose modifications on the lipid A, which indicates that deacylation by PagL increases the permeability of the membrane. In addition, it was shown that a WT strain containing a non-
MA
latent pagL vector possesses an increased susceptibility to various antimicrobial agents compared to the WT [95, 96]. However, investigating the effect of pmrA/pagL and pmrc/pagL double
TE
D
mutants on the susceptibility to PmB showed increased susceptibility to this AMP in the double mutants compared to pmrA and pmrC single mutants [97]. pagL mutant did not show increased
AC CE P
susceptibility compared to the WT, which might suggest that deacylation of lipid A is important to PmB resistance under certain conditions [97].
3.4.2
Addition of glycine to Lipid A
It was found that AlmEFG operon is involved in glycine and diglycine addition to the LPS of the Gram-negative bacteria Vibrio cholera. This modification confers polymyxin resistance in these species. The AlmEFG operon has some sequence homology with complexes involved in Dalanylation in Gram-positive bacteria [29, 98]. This modification process requires three proteins, AlmE activates glycine and transfers it to the carrier protein AlmF. The glycine is then transferred to AlmG which in turn transfers the glycine to the lipid A [29]. However, it was not yet reported how this modification confers Vibrio cholera resistance to polymyxin, and whether this system is present in other Gram-negative bacteria (Figure 3). 13
ACCEPTED MANUSCRIPT 3.4.3
De-phosphorylation, hydroxylation and addition of Phosphoethanolamine (PE) to Lipid A
PT
Studies have shown that de-phosphorylation of lipid A contributes to resistance against AMPs. It
RI
was shown that Helicobacter pylori possesses a protein called LpxE that is responsible for the
SC
de-phosphorylation of lipid A at position 1. This de-phosphorylation allows the addition of phosphoethanolamine (PEA) by EptA (PmrC), a PEA transferase. In addition, LpxF was found to
NU
dephosphorylate lipid A at position 4 leaving an unmodified hydroxyl group [99]. LpxE and
MA
LpxF mutants were shown to possess a higher susceptibility to PmB. In addition, a lipid A fully phosphorylated at positions 1 and 4 was shown to bind PmB more efficiently, rendering these
D
bacteria more susceptible to PmB [99]. It was initially found that the LPS of PmB resistant E.
TE
coli contain two to three fold higher amounts of PEA compared to the WT [100]. Addition of PEA to the LPS is also thought to change the net negative charge of the LPS and by that
AC CE P
contributes to the electrostatic repulsion of the positively charged AMPs [27]. PEA attachment can occur either on the phosphate group of the lipid A or of the inner core oligosaccharides (kdo). In E. coli and S. enterica Typhimurium, these modifications are conducted by two distinct enzymes. A pmrA-regulated PmrC (EptA) is responsible for the PEA incorporation into the lipid A [101], whilst the enzyme EptB couples the PEA moiety to the 3-deoxy-D-manno-octulosonic acid (Kdo) of the inner core polysaccharide [102]. However, EptB was not reported to be involved in antimicrobial resistance, and turned out to be regulated by other proteins rather than PmrA [103] (Figure 3). In other Gram-negative bacteria such as Neisseria gonorrhoeae and meningococcal bacteria, the gene lptA was found to encode for a protein with a similar activity as PmrC [104, 105]. Mutants in the pmrC of S. enterica Typhimurium and in lptA of N. gonorrhoeae were more 14
ACCEPTED MANUSCRIPT sensitive to PmB compared to the WT [105]. In addition, an in vivo study with N. gonorrhoeae showed that the addition of PEA to LPS confers a survival advantage compared to lptA mutant in
PT
human and rabbit [106]. It was found that pmrA mutants, that lack the aminoarabinose and PEA
RI
modifications on the lipid A, are more permeable to ethidium, which suggests that these modifications are involved in generating a robust permeation barrier [96]. However PmrA also
SC
regulates other genes that might affect membrane permeability. Hydroxylation of the myristate at
NU
position 2 on the lipid A in S. enterica Typhimurium by LpxO was shown to reduce permeability of the membrane and suggests that this hydroxylation can increase the hydrogen bond between
MA
neighboring molecules of LPS, which contributes to the outer membrane stabilization [107]
Addition of amino-sugars to lipid A
TE
3.4.4
D
(Figure 3).
AC CE P
There are several amino–sugars that are added to the lipid A moiety and are known to contribute to AMP resistance. These include (i) aminoarabinose, (ii) glucosamine, and (iii) galactoseamine (Figure 3).
(i) Addition of 4-amino-4-deoxy-L-arabinose (L-Ara4) to the phosphate group of the Lipid A correlated with acquired resistance to AMPs. L-Ara4 addition is done by ArnT, an enzyme regulated by PmrAB TCS [108]. The addition of L-Ara4 causes reduction of the net negative charge of the LPS, which is thought to reduce the attachment of positively charged AMPs, due to electrostatic repulsion [25, 94, 100]. In addition to its direct role in AMP resistance, the addition of L-ara4 to LPS has a regulatory effect on other processes related to LPS, such as the deacylation of lipid A by pagL (as described in section 3.4.1 above) [95], [96] and LPS export. It was found that L-ara4 synthesis is essential for bacteria viability since it is important for proper export of LPS molecules to the outer membrane of Burkholderia 15
ACCEPTED MANUSCRIPT cenocepacia [109]. It is possible that part of the AMP-resistance that is related to L-ara4 is due to its regulatory effects and not to direct effects of this modification.
PT
(ii) It has been reported that LPS undergoes a modification where glucosamine is added
RI
to the two phosphate groups of the lipid A in several species of Bordetella genus by ArnT homolog [110, 111]. This modification was found to confer resistance to AMPs, and reduce
SC
outer membrane perturbation by EDTA [112].
NU
(iii) Addition of galactosamine to lipid A was first discovered in Francisella tularensis Holarctica and Francisella novicida , and is done by ArnT homolog [113-115]. It was found later
MA
that in order for the galactosamine addition to occur, deacetylation of the lipid A needed to take place. This deacetylation is done by NaxD, and NaxD mutants were more susceptible to PmB
Changing the length of O-antigen
AC CE P
3.4.5
TE
D
compared to the parental strain [116 ].
To date, there is no conclusive evidence of whether the existence and length of the O-antigen moiety of the LPS affects AMP resistance. On the one hand, it was shown that a S. enterica Typhimurium mutant in O-antigen production is more sensitive to PmB compared to the WT strain. This mutant has a deletion in wbaP, a gene responsible for the addition of phosphogalactose to undecaprenyl-phosphate, a step necessary for O-antigen biosynthesis in this strain. However, this strain was still infectious and causes gut inflammation in mice models [117]. On the other hand, in F. tularensis it was shown that the absence of O-antigen did not cause a higher susceptibility to AMPs [118]. In addition, there is no conclusive answer of whether the length of the O-antigen in S. enterica Typhimurium affects resistance to AMPs. O-antigen length in these bacteria is defined by WZZst and fepE (WZZfepE) genes regulated by PmrAB TCS [119]. WZZst and fepE code 16
ACCEPTED MANUSCRIPT for a long O-antigen and a very long O-antigen, respectively [120, 121]. There is an ongoing debate on the question of whether the very long O-antigen is important for AMP resistance.
SC
RI
[119], while others showed no change in susceptibility [122] .
PT
Some claim that S. enterica Typhimurium mutants in FepE showed higher susceptibility to PmB
4. Modifications of the bacterial outer surface that influence peptide
NU
properties.
MA
Binding of AMPs to the LPS could change their chemical properties. It was shown [89] that a 12-mer peptide K5L7 has lower antibacterial activity compared to its diastereomer 4D-K5L7 in
D
which 4 L-amino acids were replaced with their enantiomers, D-amino acids. Nonetheless, when
TE
comparing the effect of these peptides on PG and phosphatidylethanolamine (PE) vesicles, an increase in the permeability of the corresponding vesicles was found for both. Furthermore, they
AC CE P
strongly bind LPS with a similar affinity; however each one of them was affected differently by the LPS. Both peptides have undefined structure in solution, but in LPS, K5L7 became aggregated whereas 4D-K5L7 did not. These results suggest that membrane binding is important for AMP activity, but it is not the only factor that affects the permeating activity of the peptides. It might also suggest that peptides can change their properties depending on the lipid they are bound to, and might explain the different activity of peptides on Gram-positive and negative bacteria. Later work also supports this hypothesis [63].
5. Mechanisms of bacterial AMP resistance in biofilms Bacterial biofilms are a colonization of bacteria on either a biotic or abiotic surface, where they form three dimensional multi-cell structures in which bacteria are embedded in an extracellular 17
ACCEPTED MANUSCRIPT matrix (ECM) composed of polysaccharides, DNA and proteins. Biofilms can be well structured and involve quorum sensing and formation of molecules that mediate migration of cells within
PT
the biofilm and formation of specific structures as seen in P. aeruginosa [123, 124]. Biofilms are
RI
important for bacterial survival in their natural environments [125], and protects the bacteria from the immune system and antibiotics including AMPs [126, 127]. Bacterial susceptibility to
SC
AMPs in biofilm was lower compared to the planktonic state [128, 129], however biofilm-
NU
resistance to AMPs was not extensively studied compared to other antimicrobial agents. Furthermore, no specific mechanisms for AMP-resistance in biofilm were discovered. It is
MA
assumed that mechanisms involved in the AMP-resistance in biofilms are similar to those known for other antimicrobial agents. Some of the putative mechanisms of resistance against
TE
D
antimicrobial agents in biofilms can theoretically be applied to AMPs and are therefore discussed in the following sections. A detailed review about resistance mechanisms to antibiotic in biofilm
5.1
AC CE P
has been published recently [130].
Creation of sub-populations in the biofilm-
P. aeruginosa biofilms that were grown in a flow chamber formed a mushroom shaped structure, and two well distinguished subpopulations were found in it [131]. When a biofilm was treated with colistin, there was a clear difference between the bacteria in the middle layer and those on the outer layer. The bacteria in the middle of the colony died while those on the outer layer were kept alive [131]. In addition, the bacteria on the outer surface and not in the middle were characterized with motility as was demonstrated after photo bleaching [131]. Resistant individuals were also found in bacteria with motility deficiencies. However these bacteria were not placed on the outer part of the biofilm, but were randomly incorporated in it. This might suggest that the motility is important for the arrangement of the different populations in the 18
ACCEPTED MANUSCRIPT biofilm, but not directly involved in AMP-resistance [131, 132]. Furthermore, pmr operon expression pattern as observed with GFP was correlated to the two bacterial subpopulations
PT
[131]. In addition, cells of the different subpopulations were characterized by different metabolic
RI
levels whereas cells in the outer surface were more metabolically active compared to the ones in
A higher mutation rate-
NU
5.2
SC
the middle. This suggests that metabolically active bacteria are more resistant to colistin [132].
It was found that bacteria in biofilm tend to have a higher mutation rate compared to the
MA
planktonic bacteria. The higher mutation rate can accelerate the natural occurring process of resistance to antimicrobial agents [130]. The accelerated mutation rate was found to involve a
D
reduced expression of DNA repair genes and mismatch repair system, as well as, elevation in
TE
oxidative stress [130]. In addition, several studies reported for a higher frequency of horizontal
5.3
AC CE P
gene transfer in biofilm. This is another aspect that contributes to a higher mutation rate [133].
Upregulation of efflux pumps -
Efflux pumps were shown to confer biofilm resistance in several bacterial strains in their planktonic state [27, 42, 43] and biofilm state [132, 134]. It was reported that efflux pumps undergo upregulation in biofilm compared to the planktonic state [135-137]. In addition, MexAB-OprM efflux pumps were found to be important for the development of tolerance to colistin in a sub-population of P. aeruginosa biofilm [132]. A detailed review about the role of efflux pumps in biofilm resistance to antibiotics can be found in reference [137].
5.4
Induction of TCSs within the biofilm-
19
ACCEPTED MANUSCRIPT P. aeruginosa resistant to colistin showed upregulation of pmr operon genes, which was suggested to contribute to bacterial biofilm tolerance due to LPS modifications [131, 132]. It was
PT
also suggested that the DNA that is found as part of the ECM reduces the level of divalent
Diffusion in biofilm-
SC
5.5
RI
cations in the environment thus activating PhoPQ TCS and conferring resistance to AMPs [38].
NU
It is not clear whether biofilm can reduce the diffusion rate of AMPs and in so doing help to resist them. It was shown that the diffusion rate of fluorescein, a small molecule that does not
MA
interact with the biofilm, is not affected by the presence of a biofilm. However, it was predicted that the diffusion of large molecules will be influenced by the biofilm [138]. Periplasmic glucan
D
was found to confer resistance to several antibiotics in P. aeruginosa, in a biofilm specific
TE
manner, where a mutant for ndvB, a gene necessary for periplasmic glucan, was shown to be
AC CE P
more susceptible to these agents [139]. In this work the authors suggested that glucans interact with the antibiotics, and therefore slow the diffusion of these molecules. However, resistance to AMPs was not checked in this respect. In addition, it was shown that P. aeruginosa and K. pneumonia biofilms reduce the diffusion rate of beta-lactam antibiotics only when betalactamase enzymes are present within the biofilms [130, 140]. For more details on diffusion of
antibiotics in P. aeruginosa biofilms the following review is recommended [141].
6. Two component systems (TCSs) regulate AMP resistance TCSs are commonly used in bacteria to sense and respond to environmental changes [142-145]. Many of the outer surface modifications that were discussed above are a result of activation of such systems via regulation of the expression of downstream genes as will be discussed in the
20
ACCEPTED MANUSCRIPT following section. TCSs consist of a membrane sensor that receive signals from the environment and transfers this information by activating a transcriptional response regulator through
PT
phosphorylation or de-phosphorylation [24, 143, 146-150]. In these systems, a receptor (usually
RI
histidine kinase) is situated in the cytoplasmic membrane, and is activated by an environmental signal. The sensor in turn, phosphorylates a cytoplasmic protein that acts as a transcription
SC
factor. The response involves activation of sets of genes such as virulence genes, ion transporters
NU
and membrane-remodeling genes that help the Gram-positive and Gram-negative bacteria to adjust better to the environment within hours. Some of these TCS are known to help bacteria
TCS activation by AMPs
D
6.1
MA
cope with AMPs and are activated by them.
TE
S. enterica PhoPQ TCS is known to be activated by low pH, low divalent cation concentration
AC CE P
and AMPs [151-153]. It has been shown recently that peptides with a higher positive charge were more potent in PhoPQ activation [15]. Studies on the mode of PhoQ activation by AMPs suggest that they displace divalent cations that bind to a highly acidic domain within the PhoQ [149, 153-158], which corroborates the finding that the more positively charged the AMP is, the more it tends to activate the PhoPQ [15]. Surprisingly, a histidine-rich AMP with a charge of only +1 under these conditions was highly potent in PhoPQ activation [15]. This supports a second mechanism for removing divalent cations and activation of PhoPQ by certain AMPs In that case, histidine, a known chelator for divalent cations, exerts this activity in the LPS environment, reducing their concentration in the environment leading to PhoPQ activation [15]. However, activation by different signals promotes different responses [159]. PmrAB TCS is known to be activated by high Fe+3 concentration [158]. In addition, there is a crosstalk between these two TCSs, and PhoPQ activated gene pmrD encodes for a protein activator that activates 21
ACCEPTED MANUSCRIPT pmrA, which in turn represses pmrD expression [151]. These systems have homologous systems in several Gram-negative bacteria, but the interaction between them in different species can be
PT
different. For example in E. coli another unknown bacterial system is involved in PmrAB
RI
activation in a PmrD dependent manner [160]. Both PhoPQ and PmrAB were found to be essential for the virulence of many bacteria species [143, 161, 162].
SC
GraRS (ApsRS) is a well-studied TCS with respect to AMPs resistance, that is found in
NU
several Gram-positive bacteria such as S. aureus and Staphylococcus epidermidis [163]. Similarly to PhoPQ activation in Gram-negative bacteria, it is assumed that AMPs bind a highly
MA
negative extracellular loop in the GraS (ApsS) sensor protein, and causes activation of this system [164]. GraRS was shown to crosstalk with another bacterial system Stk1/Stp1, where
TE
D
Stk1 phosphorylates the response regulator GraR, and thus enabling alanilation of the wall theichoic acid (WTA) by dlt operon, which is regulated by GraRS TCS. S. aureus mutants in the
AC CE P
phosphorylation site of Skt1on GraR showed a similar reduced level of alanine in the WTA [165]. However, Bacillus subtilis homolog for GraR, BceR, does not undergo Skp1 dependent phosphorylation [165]. Other TCS in S. aureus were also reported to play a role in vancomycin resistance. Such TCSs are WalKR, VraSR and AirSR (YhcSR) [166, 167].
6.2
Genes activated by TCS that are involved in AMP resistance
Upon activation, TCSs can activate tens and even hundreds of genes [168, 169]. Some of the genes activated by PhoPQ and PmrAB are able to change the composition of the LPS and other surface elements and so change their general surface thickness and charge, thus altering AMP’s effect on these bacteria [92, 100, 145, 170]. For example, PhoPQ induces pagP and pagL expression. pagP and pagL, as described before, encode for acyltransferases which catalyze 22
ACCEPTED MANUSCRIPT palmitoylation of lipid-A and glycerophospholipids of the inner leaflet of the outer membrane [72, 90]. On the other hand, pagL encodes a deacylase which removes a fatty acid from the
PT
lipid-A core [94]. This leads to increase or decrease of both the membrane thickness and the
RI
hydrophobicity of the LPS [90, 92]. Other PhoPQ and PmrAB activated genes are pmrE, pmrHJIFKLM operon that are involved in LPS charge modifications [3, 94, 100, 171-176].
SC
Because these systems induce genes that influence the hydrophobicity and charge of the LPS, it
NU
is highly logic to assume that they contribute to resistance toward AMPs. Indeed PhoPQ and PmrAB relevance to AMP-resistance was demonstrated by showing that phoP-KO or pmrA-KO
MA
bacteria displayed increased susceptibility to some AMPs (particularly PmB and colistin) compared to the WT [13, 177-183]. Despite extensive studies there are still genes induced by
TE
D
these systems that have an unknown function [184, 185]. In addition, most of the work done was limited to certain types of AMPs (particularly PmB, colistin and LL-37), which do not represent
AC CE P
the major classical AMPs (Figure 4).
GraRS TCS consists of GraS- a sensor protein, GraR (ApsR)- a response regulator and another accessory protein with an unknown function, GraX (ApsX). Mutations in each of these three genes increase susceptibility to AMPs [164, 186]. GraRS is known to regulate the expression of mprF, dlt operon and vraFG operon [57, 187], which are known to be involved in AMP resistance. Over expression of GraRS in S. aureus caused higher resistance to vancomycin [163]. As previously mentioned, MprF is responsible for the addition of lysine and alanine to PG on the plasma membrane, DltA is involved in the addition of D-alanine to the LTA, and VraFG is an AMP efflux pump. In Listeria monocytogenes a TCS named VirRS regulates dlt operon and mprF homolog, among other genes [169] (Figure 4).
7. Ways to overcome AMP resistance 23
ACCEPTED MANUSCRIPT Mammalian AMPs serve as part of the innate immune system, and consist of two main groups, cathelicidins and the cysteine rich defensins. Mutations in genes related to AMP production in
PT
humans and mice led to frequent bacterial infections, whilst transgenic mice expressing high
RI
amounts of human defensin 5 were protected from certain bacterial infections [188]. All known mechanisms of bacterial resistance to AMPs refer to general changes in bacteria properties that
SC
allegedly can be applied to all AMPs. Therefore, there is ongoing debate as to whether the use of
NU
AMPs in the clinic can lead to bacteria resistance to naturally occurring AMPs, and by that damage the activity of the innate immune system against various pathogens [13]. Indeed, a study
MA
showed that there is cross resistance between some AMPs, and thus pre-exposure to one peptide can lead to higher resistance to other peptides [189]. Lfcin B resistant S. aureus bacteria showed
TE
D
mild cross resistance with Indolicidin and a derivative of magainin (Ala3,13,18-magainin), while for the same strains no resistance was observed for LL-37 [190]. Induced S. enterica
AC CE P
Typhimurium resistant mutants to LL-37, CNY100HL and Wheat Germ Histones (WGH), showed cross resistance to the other peptides to a certain extent. Several lines of S. aureus that are resistant to daptomycin, a lipopeptide, showed a higher resistance to tPMPs and human neutrophil peptide-1 (hNP-1). However, resistance to PMB was not consistent in most of the lines [67].
Moreover, it has been also shown that there is no cross resistance between different AMPs. So far, it is suggested that peptides show cross resistance when they share common mechanisms of action [191], but this assumption should be further evaluated. In addition, in some cases the resistance to the AMPs was abolished after removing the AMP [129, 190, 192]. This kind of induced resistance is only transient and as a result of the activation of bacterial systems to environmental changes such as encountering an AMP. Therefore, it is indeed possible that the
24
ACCEPTED MANUSCRIPT use of AMPs for the treatment of bacterial infections will cause resistance to innate immune AMPs. However, a better understanding of the mechanisms that affect cross resistance between
PT
AMP, as well as the analysis of the nature of cross resistance, might allow this problem to be
RI
overcome, so that AMPs can be used as the first line treatment for infectious diseases. In addition, combined therapies using two different AMPs or a regular antibiotic and AMP might
NU
SC
also provide a solution to the cross resistance between AMPs.
8. Summary
MA
In summary, bacteria were found to acquire AMP resistance by two general mechanisms: (1) induced resistance, by induction of specific bacterial systems that contribute to AMP resistance.
TE
D
When the bacteria are no longer under these conditions the resistance will vanish. (2) Constitutive resistance which results from genetic changes. In this review we discussed briefly
AC CE P
these two general mechanisms which involve: (i) Extracellular polysaccharide secretion to form capsules that protect the bacteria from various antimicrobial agents including AMPs. (ii) Modifying bacteria surface molecules, such as LTA in Gram-positive bacteria with the addition of D-alanine. Modification of the LPS moieties: modification of the lipid A by the addition of glycine, amino sugars such as aminoarabinose, glucosamine and galactoseamine, dephosphorilation, hydroxylation , addition of phosphoethanolamine, addition and removal of acyl chains, and changing the length of O antigen moiety. (iii) Changes in bacterial surface which can cause alterations in peptide properties, and as such its effect on the bacteria.
25
ACCEPTED MANUSCRIPT (iv) Biofilm formation including the secretion of extra cellular matrix, which consists of polysaccharides, proteins and DNA, all of which form a barrier to AMPs.
PT
(v) Peptide activation of TCSs resulting in gene expression patterns, which in many cases
RI
regulate the above mechanisms, most of which induce biophysical changes to the bacteria, such as surface rigidity, cell wall thickness, surface charge and membrane fluidity.
SC
Importantly, despite the main concern that bacterial resistance to administered AMPs might
NU
cause resistance to natural AMPs and so sabotage the natural immune response, there are
MA
indications for only limited cross resistance between different AMPs.
D
Acknowledgements
TE
This investigation was supported in part by a grant from the Conseil Pasteur-Weizmann to PTC, TM and YS); the Israel Ministry of Health (IMOH), Grant No. 3-7291 and the European
AC CE P
Community's Seventh Framework Programme (FP7/2007-2013) under grant agreement n° 278998. Yechiel Shai is the incumbent of the Harold S. and Harriet B. Brady Professorial Chair in Cancer Research.
26
ACCEPTED MANUSCRIPT
B. Spellberg, R. Guidos, D. Gilbert, J. Bradley, H.W. Boucher, W.M. Scheld, J.G.
SC
[1]
RI
PT
9. References
Bartlett, J. Edwards, Jr., The epidemic of antibiotic-resistant infections: a call to action
NU
for the medical community from the Infectious Diseases Society of America, Clin Infect
[2]
MA
Dis 46 (2008) 155-164.
J.F. Acar, G. Moulin, Antimicrobial resistance: a complex issue, Rev Sci Tech 31 (2012)
H.G. Boman, Peptide antibiotics and their role in innate immunity, Annu Rev Immunol 13 (1995) 61-92.
T. Koprivnjak, A. Peschel, Bacterial resistance mechanisms against host defense
AC CE P
[4]
TE
[3]
D
23-31.
peptides, Cell Mol Life Sci 68 (2011) 2243-2254. [5]
M. Zasloff, Antimicrobial peptides of multicellular organisms, Nature 415 (2002) 389395.
[6]
Y. Shai, A. Makovitzky, D. Avrahami, Host defense peptides and lipopeptides: modes of action and potential candidates for the treatment of bacterial and fungal infections, Curr Protein Pept Sci 7 (2006) 479-486.
[7]
L.T. Nguyen, E.F. Haney, H.J. Vogel, The expanding scope of antimicrobial peptide structures and their modes of action, Trends Biotechnol 29 (2011) 464-472.
27
ACCEPTED MANUSCRIPT [8]
B. Agerberth, H. Gunne, J. Odeberg, P. Kogner, H.G. Boman, G.H. Gudmundsson, FALL-39, a putative human peptide antibiotic, is cysteine-free and expressed in bone
G.H. Gudmundsson, B. Agerberth, J. Odeberg, T. Bergman, B. Olsson, R. Salcedo, The
RI
[9]
PT
marrow and testis, Proc. Natl. Acad. Sci. U. S. A. 92 (1995) 195-199.
human gene FALL39 and processing of the cathelin precursor to the antibacterial peptide
Y. Shai, Mechanism of the binding, insertion and destabilization of phospholipid bilayer
NU
[10]
SC
LL-37 in granulocytes, Eur. J. Biochem. 238 (1996) 325-332.
membranes by alpha-helical antimicrobial and cell non-selective membrane-lytic
[11]
MA
peptides, Biochim Biophys Acta 1462 (1999) 55-70. U.H. Durr, U.S. Sudheendra, A. Ramamoorthy, LL-37, the only human member of the
TE
D
cathelicidin family of antimicrobial peptides, Biochim Biophys Acta 1758 (2006) 14081425.
H. Jenssen, P. Hamill, R.E. Hancock, Peptide antimicrobial agents, Clin Microbiol Rev
AC CE P
[12]
19 (2006) 491-511. [13]
A.K. Marr, W.J. Gooderham, R.E. Hancock, Antibacterial peptides for therapeutic use: obstacles and realistic outlook, Curr Opin Pharmacol 6 (2006) 468-472.
[14]
Z. Jiang, A.I. Vasil, J.D. Hale, R.E. Hancock, M.L. Vasil, R.S. Hodges, Effects of net charge and the number of positively charged residues on the biological activity of amphipathic alpha-helical cationic antimicrobial peptides, Biopolymers 90 (2008) 369383.
[15]
T. Shprung, A. Peleg, Y. Rosenfeld, P. Trieu-Cuot, Y. Shai, Effect of PhoP-PhoQ activation by broad repertoire of antimicrobial peptides on bacterial resistance, J Biol Chem 287 (2012) 4544-4551.
28
ACCEPTED MANUSCRIPT [16]
L.M. Yin, M.A. Edwards, J. Li, C.M. Yip, C.M. Deber, Roles of hydrophobicity and charge distribution of cationic antimicrobial peptides in peptide-membrane interactions, J
Y.M. Lin, S.J. Wu, T.W. Chang, C.F. Wang, C.S. Suen, M.J. Hwang, M.D. Chang, Y.T.
RI
[17]
PT
Biol Chem 287 (2012) 7738-7745.
Chen, Y.D. Liao, Outer membrane protein I of Pseudomonas aeruginosa is a target of
J.P. Powers, R.E. Hancock, The relationship between peptide structure and antibacterial activity, Peptides 24 (2003) 1681-1691.
R.E. Hancock, G. Diamond, The role of cationic antimicrobial peptides in innate host
MA
[19]
NU
[18]
SC
cationic antimicrobial peptide/protein, J Biol Chem 285 (2010) 8985-8994.
defences, Trends. Microbiol. 8 (2000) 402-410.
D
G.A. Woolley, B.A. Wallace, Model ion channels: gramicidin and alamethicin, J Membr
TE
[20]
Biol 129 (1992) 109-136.
Y. Shai, Mode of action of membrane active antimicrobial peptides, Biopolymers 66
AC CE P
[21]
(2002) 236-248. [22]
K. Matsuzaki, Why and how are peptide-lipid unteractions utilized for self defense? Magainins and tachyplesins as archyetypes, Biochim. Biophys. Acta. 1462 (1999) 1-10.
[23]
L. Yang, T.A. Harroun, T.M. Weiss, L. Ding, H.W. Huang, Barrel-stave model or toroidal model? A case study on melittin pores, Biophys J 81 (2001) 1475-1485.
[24]
E. Garcia Vescovi, F.C. Soncini, E.A. Groisman, The role of the PhoP/PhoQ regulon in Salmonella virulence, Res Microbiol 145 (1994) 473-480.
[25]
J.S. Gunn, K.B. Lim, J. Krueger, K. Kim, L. Guo, M. Hackett, S.I. Miller, PmrA-PmrBregulated genes necessary for 4-aminoarabinose lipid A modification and polymyxin resistance, Mol Microbiol 27 (1998) 1171-1182.
29
ACCEPTED MANUSCRIPT [26]
V. Nizet, Antimicrobial peptide resistance mechanisms of human bacterial pathogens, Curr Issues Mol Biol 8 (2006) 11-26. Y.L. Tzeng, K.D. Ambrose, S. Zughaier, X. Zhou, Y.K. Miller, W.M. Shafer, D.S.
PT
[27]
RI
Stephens, Cationic antimicrobial peptide resistance in Neisseria meningitidis, J Bacteriol 187 (2005) 5387-5396.
K.L. Nawrocki, E.K. Crispell, S.M. McBride, Antimicrobial Peptide Resistance
SC
[28]
[29]
NU
Mechanisms of Gram-Positive Bacteria, Antibiotics (Basel) 3 (2014) 461-492. J.C. Henderson, C.D. Fage, J.R. Cannon, J.S. Brodbelt, A.T. Keatinge-Clay, M.S. Trent,
MA
Antimicrobial peptide resistance of Vibrio cholerae results from an LPS modification pathway related to nonribosomal peptide synthetases, ACS Chem Biol 9 (2014) 2382-
TE
[30]
D
2392.
A.X. Tran, J.D. Whittimore, P.B. Wyrick, S.C. McGrath, R.J. Cotter, M.S. Trent, The
AC CE P
lipid A 1-phosphatase of Helicobacter pylori is required for resistance to the antimicrobial peptide polymyxin, J Bacteriol 188 (2006) 4531-4541. [31]
J.B. McPhee, M. Bains, G. Winsor, S. Lewenza, A. Kwasnicka, M.D. Brazas, F.S. Brinkman, R.E. Hancock, Contribution of the PhoP-PhoQ and PmrA-PmrB twocomponent regulatory systems to Mg2+-induced gene regulation in Pseudomonas aeruginosa, J Bacteriol 188 (2006) 3995-4006.
[32]
J.B. McPhee, S. Lewenza, R.E. Hancock, Cationic antimicrobial peptides activate a twocomponent regulatory system, PmrA-PmrB, that regulates resistance to polymyxin B and cationic antimicrobial peptides in Pseudomonas aeruginosa, Mol Microbiol 50 (2003) 205-217.
30
ACCEPTED MANUSCRIPT [33]
S.M. Moskowitz, R.K. Ernst, S.I. Miller, PmrAB, a two-component regulatory system of Pseudomonas aeruginosa that modulates resistance to cationic antimicrobial peptides and
A. Peschel, How do bacteria resist human antimicrobial peptides?, Trends Microbiol 10
RI
[34]
PT
addition of aminoarabinose to lipid A, J Bacteriol 186 (2004) 575-579.
(2002) 179-186.
R. Saar-Dover, A. Bitler, R. Nezer, L. Shmuel-Galia, A. Firon, E. Shimoni, P. Trieu-
SC
[35]
NU
Cuot, Y. Shai, D-alanylation of lipoteichoic acids confers resistance to cationic peptides in group B streptococcus by increasing the cell wall density, PLoS Pathog 8 (2012)
[36]
MA
e1002891.
G.H. Gudmundsson, P. Bergman, J. Andersson, R. Raqib, B. Agerberth, Battle and
TE
D
balance at mucosal surfaces--the story of Shigella and antimicrobial peptides, Biochem Biophys Res Commun 396 (2010) 116-119. B. Sperandio, B. Regnault, J. Guo, Z. Zhang, S.L. Stanley, Jr., P.J. Sansonetti, T. Pedron,
AC CE P
[37]
Virulent Shigella flexneri subverts the host innate immune response through manipulation of antimicrobial peptide gene expression, J Exp Med 205 (2008) 11211132. [38]
H. Mulcahy, L. Charron-Mazenod, S. Lewenza, Extracellular DNA chelates cations and induces antibiotic resistance in Pseudomonas aeruginosa biofilms, PLoS Pathog 4 (2008) e1000213.
[39]
C. Chan, L.L. Burrows, C.M. Deber, Helix induction in antimicrobial peptides by alginate in biofilms, J Biol Chem 279 (2004) 38749-38754.
[40]
M. Sieprawska-Lupa, P. Mydel, K. Krawczyk, K. Wojcik, M. Puklo, B. Lupa, P. Suder, J. Silberring, M. Reed, J. Pohl, W. Shafer, F. McAleese, T. Foster, J. Travis, J. Potempa,
31
ACCEPTED MANUSCRIPT Degradation of human antimicrobial peptide LL-37 by Staphylococcus aureus-derived proteinases, Antimicrob Agents Chemother 48 (2004) 4673-4679. A. Schmidtchen, I.M. Frick, E. Andersson, H. Tapper, L. Bjorck, Proteinases of common
PT
[41]
RI
pathogenic bacteria degrade and inactivate the antibacterial peptide LL-37, Mol Microbiol 46 (2002) 157-168.
D.M. Warner, W.M. Shafer, A.E. Jerse, Clinically relevant mutations that cause
SC
[42]
NU
derepression of the Neisseria gonorrhoeae MtrC-MtrD-MtrE Efflux pump system confer different levels of antimicrobial resistance and in vivo fitness, Mol Microbiol 70 (2008)
[43]
MA
462-478.
W.M. Shafer, X. Qu, A.J. Waring, R.I. Lehrer, Modulation of Neisseria gonorrhoeae
TE
D
susceptibility to vertebrate antibacterial peptides due to a member of the resistance/nodulation/division efflux pump family, Proc Natl Acad Sci U S A 95 (1998)
[44]
AC CE P
1829-1833.
H.C. Maisey, D. Quach, M.E. Hensler, G.Y. Liu, R.L. Gallo, V. Nizet, K.S. Doran, A group B streptococcal pilus protein promotes phagocyte resistance and systemic virulence, FASEB J 22 (2008) 1715-1724.
[45]
X. Lauth, M. von Kockritz-Blickwede, C.W. McNamara, S. Myskowski, A.S. Zinkernagel, B. Beall, P. Ghosh, R.L. Gallo, V. Nizet, M1 protein allows Group A streptococcal survival in phagocyte extracellular traps through cathelicidin inhibition, J Innate Immun 1 (2009) 202-214.
[46]
I.M. Frick, P. Akesson, M. Rasmussen, A. Schmidtchen, L. Bjorck, SIC, a secreted protein of Streptococcus pyogenes that inactivates antibacterial peptides, J Biol Chem 278 (2003) 16561-16566.
32
ACCEPTED MANUSCRIPT [47]
A.J. Manning, M.J. Kuehn, Contribution of bacterial outer membrane vesicles to innate bacterial defense, BMC Microbiol 11 (2011) 258. J.L. Anaya-Lopez, J.E. Lopez-Meza, A. Ochoa-Zarzosa, Bacterial resistance to cationic
PT
[48]
[49]
RI
antimicrobial peptides, Crit Rev Microbiol 39 (2013) 180-195.
J.S. Gunn, The Salmonella PmrAB regulon: lipopolysaccharide modifications,
M.R. Yeaman, N.Y. Yount, Mechanisms of antimicrobial peptide action and resistance,
NU
[50]
SC
antimicrobial peptide resistance and more, Trends Microbiol 16 (2008) 284-290.
Pharmacol Rev 55 (2003) 27-55.
A.O. Olaitan, S. Morand, J.M. Rolain, Mechanisms of polymyxin resistance: acquired
MA
[51]
and intrinsic resistance in bacteria, Front Microbiol 5 (2014) 643.
D
A. Jones, M. Georg, L. Maudsdotter, A.B. Jonsson, Endotoxin, capsule, and bacterial
TE
[52]
attachment contribute to Neisseria meningitidis resistance to the human antimicrobial
[53]
AC CE P
peptide LL-37, J Bacteriol 191 (2009) 3861-3868. H. Miajlovic, S.G. Smith, Bacterial self-defence: how Escherichia coli evades serum killing, FEMS Microbiol Lett 354 (2014) 1-9. [54]
E. Llobet, J.M. Tomas, J.A. Bengoechea, Capsule polysaccharide is a bacterial decoy for antimicrobial peptides, Microbiology 154 (2008) 3877-3886.
[55]
M.A. Campos, M.A. Vargas, V. Regueiro, C.M. Llompart, S. Alberti, J.A. Bengoechea, Capsule polysaccharide mediates bacterial resistance to antimicrobial peptides, Infect Immun 72 (2004) 7107-7114.
[56]
C.M. Ernst, S. Kuhn, C.J. Slavetinsky, B. Krismer, S. Heilbronner, C. Gekeler, D. Kraus, S. Wagner, A. Peschel, The lipid-modifying multiple peptide resistance factor is an oligomer consisting of distinct interacting synthase and flippase subunits, MBio 6 (2015).
33
ACCEPTED MANUSCRIPT [57]
C.M. Ernst, P. Staubitz, N.N. Mishra, S.J. Yang, G. Hornig, H. Kalbacher, A.S. Bayer, D. Kraus, A. Peschel, The bacterial defensin resistance protein MprF consists of separable
PT
domains for lipid lysinylation and antimicrobial peptide repulsion, PLoS Pathog 5 (2009)
[58]
RI
e1000660.
A. Peschel, R.W. Jack, M. Otto, L.V. Collins, P. Staubitz, G. Nicholson, H. Kalbacher,
SC
W.F. Nieuwenhuizen, G. Jung, A. Tarkowski, K.P. van Kessel, J.A. van Strijp,
NU
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-
[59]
MA
lysine, J Exp Med 193 (2001) 1067-1076.
S.J. Yang, N.N. Mishra, A. Rubio, A.S. Bayer, Causal role of single nucleotide
TE
D
polymorphisms within the mprF gene of Staphylococcus aureus in daptomycin resistance, Antimicrob Agents Chemother 57 (2013) 5658-5664. C.M. Ernst, A. Peschel, Broad-spectrum antimicrobial peptide resistance by MprF-
AC CE P
[60]
mediated aminoacylation and flipping of phospholipids, Mol Microbiol 80 (2011) 290299. [61]
A.S. Bayer, N.N. Mishra, G. Sakoulas, P. Nonejuie, C.C. Nast, J. Pogliano, K.T. Chen, S.N. Ellison, M.R. Yeaman, S.J. Yang, Heterogeneity of mprF sequences in methicillinresistant Staphylococcus aureus clinical isolates: role in cross-resistance between daptomycin and host defense antimicrobial peptides, Antimicrob Agents Chemother 58 (2014) 7462-7467.
[62]
S. Klein, C. Lorenzo, S. Hoffmann, J.M. Walther, S. Storbeck, T. Piekarski, B.J. Tindall, V. Wray, M. Nimtz, J. Moser, Adaptation of Pseudomonas aeruginosa to various
34
ACCEPTED MANUSCRIPT conditions includes tRNA-dependent formation of alanyl-phosphatidylglycerol, Mol Microbiol 71 (2009) 551-565. J.K. Muraih, J. Harris, S.D. Taylor, M. Palmer, Characterization of daptomycin
PT
[63]
RI
oligomerization with perylene excimer fluorescence: stoichiometric binding of phosphatidylglycerol triggers oligomer formation, Biochim Biophys Acta 1818 (2012)
A.S. Bayer, R. Prasad, J. Chandra, A. Koul, M. Smriti, A. Varma, R.A. Skurray, N. Firth,
NU
[64]
SC
673-678.
M.H. Brown, S.P. Koo, M.R. Yeaman, In vitro resistance of Staphylococcus aureus to
MA
thrombin-induced platelet microbicidal protein is associated with alterations in cytoplasmic membrane fluidity, Infect Immun 68 (2000) 3548-3553.
D
N.N. Mishra, A.S. Bayer, Correlation of cell membrane lipid profiles with daptomycin
TE
[65]
resistance in methicillin-resistant Staphylococcus aureus, Antimicrob Agents Chemother
[66]
AC CE P
57 (2013) 1082-1085.
T. Jones, M.R. Yeaman, G. Sakoulas, S.J. Yang, R.A. Proctor, H.G. Sahl, J. Schrenzel, Y.Q. Xiong, A.S. Bayer, 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 52 (2008) 269-278.
[67]
N.N. Mishra, J. McKinnell, M.R. Yeaman, A. Rubio, C.C. Nast, L. Chen, B.N. Kreiswirth, A.S. Bayer, In vitro cross-resistance to daptomycin and host defense cationic antimicrobial peptides in clinical methicillin-resistant Staphylococcus aureus isolates, Antimicrob Agents Chemother 55 (2011) 4012-4018.
35
ACCEPTED MANUSCRIPT [68]
T. Shireen, M. Singh, T. Das, K. Mukhopadhyay, Differential adaptive responses of Staphylococcus aureus to in vitro selection with different antimicrobial peptides,
R. Kumariya, S.K. Sood, Y.S. Rajput, N. Saini, A.K. Garsa, Increased membrane surface
RI
[69]
PT
Antimicrob Agents Chemother 57 (2013) 5134-5137.
positive charge and altered membrane fluidity leads to cationic antimicrobial peptide
A. Verheul, N.J. Russell, T.H.R. Van, F.M. Rombouts, T. Abee, Modifications of
NU
[70]
SC
resistance in Enterococcus faecalis, Biochim Biophys Acta 1848 (2015) 1367-1375.
membrane phospholipid composition in nisin-resistant Listeria monocytogenes Scott A,
[71]
MA
Appl Environ Microbiol 63 (1997) 3451-3457.
H. Zhou, J. Fang, Y. Tian, X.Y. Lu, Mechanisms of nisin resistance in Gram-positive
[72]
TE
D
bacteria, Annals of Microbiology 64 (2014) 413-420. Z.D. Dalebroux, S.I. Miller, Salmonellae PhoPQ regulation of the outer membrane to
[73]
AC CE P
resist innate immunity, Curr Opin Microbiol 17 (2014) 106-113. E. Cox, A. Michalak, S. Pagentine, P. Seaton, A. Pokorny, Lysylated phospholipids stabilize models of bacterial lipid bilayers and protect against antimicrobial peptides, Biochim Biophys Acta 1838 (2014) 2198-2204. [74]
S.M. McBride, A.L. Sonenshein, The dlt operon confers resistance to cationic antimicrobial peptides in Clostridium difficile, Microbiology 157 (2011) 1457-1465.
[75]
Z. Abi Khattar, A. Rejasse, D. Destoumieux-Garzon, J.M. Escoubas, V. Sanchis, D. Lereclus, A. Givaudan, M. Kallassy, C. Nielsen-Leroux, S. Gaudriault, The dlt operon of Bacillus cereus is required for resistance to cationic antimicrobial peptides and for virulence in insects, J Bacteriol 191 (2009) 7063-7073.
36
ACCEPTED MANUSCRIPT [76]
S.A. Kristian, V. Datta, C. Weidenmaier, R. Kansal, I. Fedtke, A. Peschel, R.L. Gallo, V. Nizet, D-alanylation of teichoic acids promotes group a streptococcus antimicrobial
PT
peptide resistance, neutrophil survival, and epithelial cell invasion, J Bacteriol 187 (2005)
[77]
RI
6719-6725.
F.C. Neuhaus, J. Baddiley, A continuum of anionic charge: structures and functions of D-
SC
alanyl-teichoic acids in gram-positive bacteria, Microbiol Mol Biol Rev 67 (2003) 686-
[78]
NU
723.
A. Peschel, H.G. Sahl, The co-evolution of host cationic antimicrobial peptides and
[79]
MA
microbial resistance, Nat Rev Microbiol 4 (2006) 529-536. A. Peschel, M. Otto, R.W. Jack, H. Kalbacher, G. Jung, F. Gotz, Inactivation of the dlt
TE
D
operon in Staphylococcus aureus confers sensitivity to defensins, protegrins, and other antimicrobial peptides, J Biol Chem 274 (1999) 8405-8410. S.J. Yang, C.C. Nast, N.N. Mishra, M.R. Yeaman, P.D. Fey, A.S. Bayer, Cell wall
AC CE P
[80]
thickening is not a universal accompaniment of the daptomycin nonsusceptibility phenotype in Staphylococcus aureus: evidence for multiple resistance mechanisms, Antimicrob Agents Chemother 54 (2010) 3079-3085. [81]
L. Cui, E. Tominaga, H.M. Neoh, K. Hiramatsu, Correlation between Reduced Daptomycin Susceptibility and Vancomycin Resistance in Vancomycin-Intermediate Staphylococcus aureus, Antimicrob Agents Chemother 50 (2006) 1079-1082.
[82]
S. Maisnier-Patin, J. Richard, Cell wall changes in nisin-resistant variants of Listeria innocua grown in the presence of high nisin concentrations, FEMS Microbiol Lett 140 (1996) 29-35.
37
ACCEPTED MANUSCRIPT [83]
M.G. Percy, A. Grundling, Lipoteichoic acid synthesis and function in gram-positive bacteria, Annu Rev Microbiol 68 (2014) 81-100. X. Zogaj, M. Nimtz, M. Rohde, W. Bokranz, U. Romling, The multicellular morphotypes
PT
[84]
RI
of Salmonella typhimurium and Escherichia coli produce cellulose as the second component of the extracellular matrix, Mol Microbiol 39 (2001) 1452-1463. A.P. White, D.L. Gibson, S.K. Collinson, P.A. Banser, W.W. Kay, Extracellular
SC
[85]
NU
polysaccharides associated with thin aggregative fimbriae of Salmonella enterica serovar enteritidis, J Bacteriol 185 (2003) 5398-5407.
C.E. de Rezende, Y. Anriany, L.E. Carr, S.W. Joseph, R.M. Weiner, Capsular
MA
[86]
polysaccharide surrounds smooth and rugose types of Salmonella enterica serovar
[87]
TE
D
Typhimurium DT104, Appl Environ Microbiol 71 (2005) 7345-7351. C. Alexander, E.T. Rietschel, Bacterial lipopolysaccharides and innate immunity, J
[88]
AC CE P
Endotoxin Res 7 (2001) 167-202. C.R. Raetz, C. Whitfield, Lipopolysaccharide endotoxins, Annu Rev Biochem 71 (2002) 635-700. [89]
N. Papo, Y. Shai, A molecular mechanism for lipopolysaccharide protection of Gramnegative bacteria from antimicrobial peptides, J. Biol. Chem. 280 (2005) 10378-10387.
[90]
R.E. Bishop, H.S. Gibbons, T. Guina, M.S. Trent, S.I. Miller, C.R. Raetz, Transfer of palmitate from phospholipids to lipid A in outer membranes of gram-negative bacteria, EMBO J 19 (2000) 5071-5080.
[91]
A.H. Delcour, Outer membrane permeability and antibiotic resistance, Biochim Biophys Acta 1794 (2009) 808-816.
38
ACCEPTED MANUSCRIPT [92]
L. Guo, K.B. Lim, C.M. Poduje, M. Daniel, J.S. Gunn, M. Hackett, S.I. Miller, Lipid A acylation and bacterial resistance against vertebrate antimicrobial peptides, Cell 95
M. Robey, W. O'Connell, N.P. Cianciotto, Identification of Legionella pneumophila rcp,
RI
[93]
PT
(1998) 189-198.
a pagP-like gene that confers resistance to cationic antimicrobial peptides and promotes
M.S. Trent, W. Pabich, C.R. Raetz, S.I. Miller, A PhoP/PhoQ-induced Lipase (PagL) that
NU
[94]
SC
intracellular infection, Infect Immun 69 (2001) 4276-4286.
catalyzes 3-O-deacylation of lipid A precursors in membranes of Salmonella
[95]
MA
typhimurium, J Biol Chem 276 (2001) 9083-9092. K. Kawasaki, R.K. Ernst, S.I. Miller, Inhibition of Salmonella enterica serovar
TE
D
Typhimurium lipopolysaccharide deacylation by aminoarabinose membrane modification, J Bacteriol 187 (2005) 2448-2457. K. Kawasaki, T. Manabe, Latency of the lipid A deacylase PagL is involved in producing
AC CE P
[96]
a robust permeation barrier in the outer membrane of Salmonella enterica, J Bacteriol 192 (2010) 5837-5840. [97]
K. Kawasaki, K. China, M. Nishijima, Release of the lipopolysaccharide deacylase PagL from latency compensates for a lack of lipopolysaccharide aminoarabinose modificationdependent resistance to the antimicrobial peptide polymyxin B in Salmonella enterica, J Bacteriol 189 (2007) 4911-4919.
[98]
J.V. Hankins, J.A. Madsen, D.K. Giles, J.S. Brodbelt, M.S. Trent, Amino acid addition to Vibrio cholerae LPS establishes a link between surface remodeling in gram-positive and gram-negative bacteria, Proc Natl Acad Sci U S A 109 (2012) 8722-8727.
39
ACCEPTED MANUSCRIPT [99]
T.W. Cullen, D.K. Giles, L.N. Wolf, C. Ecobichon, I.G. Boneca, M.S. Trent, Helicobacter pylori versus the host: remodeling of the bacterial outer membrane is
PT
required for survival in the gastric mucosa, PLoS Pathog 7 (2011) e1002454.
RI
[100] K. Nummila, I. Kilpelainen, U. Zahringer, M. Vaara, I.M. Helander, Lipopolysaccharides of polymyxin B-resistant mutants of Escherichia coli are extensively substituted by 2-
SC
aminoethyl pyrophosphate and contain aminoarabinose in lipid A, Mol Microbiol 16
NU
(1995) 271-278.
[101] H. Lee, F.F. Hsu, J. Turk, E.A. Groisman, The PmrA-regulated pmrC gene mediates
MA
phosphoethanolamine modification of lipid A and polymyxin resistance in Salmonella enterica, J Bacteriol 186 (2004) 4124-4133.
TE
D
[102] C.M. Reynolds, S.R. Kalb, R.J. Cotter, C.R. Raetz, A phosphoethanolamine transferase specific for the outer 3-deoxy-D-manno-octulosonic acid residue of Escherichia coli
AC CE P
lipopolysaccharide. Identification of the eptB gene and Ca2+ hypersensitivity of an eptB deletion mutant, J Biol Chem 280 (2005) 21202-21211. [103] N. Figueroa-Bossi, S. Lemire, D. Maloriol, R. Balbontin, J. Casadesus, L. Bossi, Loss of Hfq activates the sigmaE-dependent envelope stress response in Salmonella enterica, Mol Microbiol 62 (2006) 838-852. [104] A.D. Cox, J.C. Wright, J. Li, D.W. Hood, E.R. Moxon, J.C. Richards, Phosphorylation of the lipid A region of meningococcal lipopolysaccharide: identification of a family of transferases that add phosphoethanolamine to lipopolysaccharide, J Bacteriol 185 (2003) 3270-3277. [105] L.A. Lewis, B. Choudhury, J.T. Balthazar, L.E. Martin, S. Ram, P.A. Rice, D.S. Stephens, R. Carlson, W.M. Shafer, Phosphoethanolamine substitution of lipid A and
40
ACCEPTED MANUSCRIPT resistance of Neisseria gonorrhoeae to cationic antimicrobial peptides and complementmediated killing by normal human serum, Infect Immun 77 (2009) 1112-1120.
PT
[106] M.M. Hobbs, J.E. Anderson, J.T. Balthazar, J.L. Kandler, R.W. Carlson, J. Ganguly,
RI
A.A. Begum, J.A. Duncan, J.T. Lin, P.F. Sparling, A.E. Jerse, W.M. Shafer, Lipid A's
and men, MBio 4 (2013) e00892-00813.
SC
structure mediates Neisseria gonorrhoeae fitness during experimental infection of mice
NU
[107] T. Murata, W. Tseng, T. Guina, S.I. Miller, H. Nikaido, PhoPQ-mediated regulation produces a more robust permeability barrier in the outer membrane of Salmonella
MA
enterica serovar typhimurium, J Bacteriol 189 (2007) 7213-7222. [108] M.S. Trent, A.A. Ribeiro, S. Lin, R.J. Cotter, C.R. Raetz, An inner membrane enzyme in
TE
D
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
AC CE P
Chem 276 (2001) 43122-43131.
[109] M.A. Hamad, F. Di Lorenzo, A. Molinaro, M.A. Valvano, Aminoarabinose is essential for lipopolysaccharide export and intrinsic antimicrobial peptide resistance in Burkholderia cenocepacia(dagger), Mol Microbiol 85 (2012) 962-974. [110] N. Marr, A. Tirsoaga, D. Blanot, R. Fernandez, M. Caroff, Glucosamine found as a substituent of both phosphate groups in Bordetella lipid A backbones: role of a BvgASactivated ArnT ortholog, J Bacteriol 190 (2008) 4281-4290. [111] N.R. Shah, S. Albitar-Nehme, E. Kim, N. Marr, A. Novikov, M. Caroff, R.C. Fernandez, Minor modifications to the phosphate groups and the C3' acyl chain length of lipid A in two Bordetella pertussis strains, BP338 and 18-323, independently affect Toll-like receptor 4 protein activation, J Biol Chem 288 (2013) 11751-11760.
41
ACCEPTED MANUSCRIPT [112] N.R. Shah, R.E. Hancock, R.C. Fernandez, Bordetella pertussis lipid A glucosamine modification confers resistance to cationic antimicrobial peptides and increases resistance
PT
to outer membrane perturbation, Antimicrob Agents Chemother 58 (2014) 4931-4934.
RI
[113] X. Wang, A.A. Ribeiro, Z. Guan, S.C. McGrath, R.J. Cotter, C.R. Raetz, Structure and biosynthesis of free lipid A molecules that replace lipopolysaccharide in Francisella
SC
tularensis subsp. novicida, Biochemistry 45 (2006) 14427-14440.
NU
[114] N.J. Phillips, B. Schilling, M.K. McLendon, M.A. Apicella, B.W. Gibson, Novel modification of lipid A of Francisella tularensis, Infect Immun 72 (2004) 5340-5348.
MA
[115] X. Wang, A.A. Ribeiro, Z. Guan, C.R. Raetz, Identification of undecaprenyl phosphatebeta-D-galactosamine in Francisella novicida and its function in lipid A modification,
TE
D
Biochemistry 48 (2009) 1162-1172. [116] A.C. Llewellyn, J. Zhao, F. Song, J. Parvathareddy, Q. Xu, B.A. Napier, H. Laroui, D.
AC CE P
Merlin, J.E. Bina, P.A. Cotter, M.A. Miller, C.R. Raetz, D.S. Weiss, NaxD is a deacetylase required for lipid A modification and Francisella pathogenesis, Mol Microbiol 86 (2012) 611-627. [117] K. Ilg, K. Endt, B. Misselwitz, B. Stecher, M. Aebi, W.D. Hardt, O-antigen-negative Salmonella enterica serovar Typhimurium is attenuated in intestinal colonization but elicits colitis in streptomycin-treated mice, Infect Immun 77 (2009) 2568-2575. [118] M. Vonkavaara, S.T. Pavel, K. Holzl, R. Nordfelth, A. Sjostedt, S. Stoven, Francisella is sensitive to insect antimicrobial peptides, J Innate Immun 5 (2013) 50-59. [119] L. Pescaretti Mde, F.E. Lopez, R.D. Morero, M.A. Delgado, The PmrA/PmrB regulatory system controls the expression of the wzzfepE gene involved in the O-antigen synthesis of Salmonella enterica serovar Typhimurium, Microbiology 157 (2011) 2515-2521.
42
ACCEPTED MANUSCRIPT [120] R. Morona, C. Daniels, L. Van Den Bosch, Genetic modulation of Shigella flexneri 2a lipopolysaccharide O antigen modal chain length reveals that it has been optimized for
PT
virulence, Microbiology 149 (2003) 925-939.
RI
[121] D. Bravo, C. Silva, J.A. Carter, A. Hoare, S.A. Alvarez, C.J. Blondel, M. Zaldivar, M.A. Valvano, I. Contreras, Growth-phase regulation of lipopolysaccharide O-antigen chain
SC
length influences serum resistance in serovars of Salmonella, J Med Microbiol 57 (2008)
NU
938-946.
[122] R.W. Crawford, A.M. Keestra, S.E. Winter, M.N. Xavier, R.M. Tsolis, V. Tolstikov, A.J.
MA
Baumler, Very long O-antigen chains enhance fitness during Salmonella-induced colitis by increasing bile resistance, PLoS Pathog 8 (2012) e1002918.
TE
D
[123] Y. Lequette, E.P. Greenberg, Timing and localization of rhamnolipid synthesis gene expression in Pseudomonas aeruginosa biofilms, J Bacteriol 187 (2005) 37-44.
AC CE P
[124] M.E. Davey, N.C. Caiazza, G.A. O'Toole, Rhamnolipid surfactant production affects biofilm architecture in Pseudomonas aeruginosa PAO1, J Bacteriol 185 (2003) 10271036.
[125] L. Hall-Stoodley, J.W. Costerton, P. Stoodley, Bacterial biofilms: from the natural environment to infectious diseases, Nat Rev Microbiol 2 (2004) 95-108. [126] F. Mika, R. Hengge, Small Regulatory RNAs in the Control of Motility and Biofilm Formation in E. coli and Salmonella, Int J Mol Sci 14 (2013) 4560-4579. [127] C. de la Fuente-Nunez, F. Reffuveille, L. Fernandez, R.E. Hancock, Bacterial biofilm development as a multicellular adaptation: antibiotic resistance and new therapeutic strategies, Curr Opin Microbiol 16 (2013) 580-589.
43
ACCEPTED MANUSCRIPT [128] H. Altman, D. Steinberg, Y. Porat, A. Mor, D. Fridman, M. Friedman, G. Bachrach, In vitro assessment of antimicrobial peptides as potential agents against several oral
PT
bacteria, J Antimicrob Chemother 58 (2006) 198-201.
RI
[129] A. Folkesson, J.A. Haagensen, C. Zampaloni, C. Sternberg, S. Molin, Biofilm induced tolerance towards antimicrobial peptides, PLoS One 3 (2008) e1891.
SC
[130] N. Hoiby, T. Bjarnsholt, M. Givskov, S. Molin, O. Ciofu, Antibiotic resistance of
NU
bacterial biofilms, Int J Antimicrob Agents 35 (2010) 322-332. [131] J.A. Haagensen, M. Klausen, R.K. Ernst, S.I. Miller, A. Folkesson, T. Tolker-Nielsen, S.
MA
Molin, Differentiation and distribution of colistin- and sodium dodecyl sulfate-tolerant cells in Pseudomonas aeruginosa biofilms, J Bacteriol 189 (2007) 28-37.
TE
D
[132] S.J. Pamp, M. Gjermansen, H.K. Johansen, T. Tolker-Nielsen, Tolerance to the antimicrobial peptide colistin in Pseudomonas aeruginosa biofilms is linked to
AC CE P
metabolically active cells, and depends on the pmr and mexAB-oprM genes, Mol Microbiol 68 (2008) 223-240. [133] S. Molin, T. Tolker-Nielsen, Gene transfer occurs with enhanced efficiency in biofilms and induces enhanced stabilisation of the biofilm structure, Curr Opin Biotechnol 14 (2003) 255-261.
[134] L. Zhang, T.F. Mah, Involvement of a novel efflux system in biofilm-specific resistance to antibiotics, J Bacteriol 190 (2008) 4447-4452. [135] R.J. Gillis, K.G. White, K.H. Choi, V.E. Wagner, H.P. Schweizer, B.H. Iglewski, Molecular basis of azithromycin-resistant Pseudomonas aeruginosa biofilms, Antimicrob Agents Chemother 49 (2005) 3858-3867.
44
ACCEPTED MANUSCRIPT [136] A. Jolivet-Gougeon, M. Bonnaure-Mallet, Biofilms as a mechanism of bacterial resistance, Drug Discov Today Technol 11 (2014) 49-56.
PT
[137] S.M. Soto, Role of efflux pumps in the antibiotic resistance of bacteria embedded in a
RI
biofilm, Virulence 4 (2013) 223-229.
[138] D. de Beer, P. Stoodley, Z. Lewandowski, Measurement of local diffusion coefficients in
SC
biofilms by microinjection and confocal microscopy, Biotechnol Bioeng 53 (1997) 151-
NU
158.
[139] T.F. Mah, B. Pitts, B. Pellock, G.C. Walker, P.S. Stewart, G.A. O'Toole, A genetic basis
MA
for Pseudomonas aeruginosa biofilm antibiotic resistance, Nature 426 (2003) 306-310. [140] C. Vuotto, F. Longo, M.P. Balice, G. Donelli, P.E. Varaldo, Antibiotic Resistance
TE
D
Related to Biofilm Formation in Klebsiella pneumoniae, Pathogens 3 (2014) 743-758. [141] E. Drenkard, Antimicrobial resistance of Pseudomonas aeruginosa biofilms, Microbes
AC CE P
Infect 5 (2003) 1213-1219.
[142] A. Alvarez-Ordonez, A. Fernandez, A. Bernardo, M. Lopez, Comparison of acids on the induction of an Acid Tolerance Response in Salmonellatyphimurium, consequences for food safety, Meat Sci 81 (2009) 65-70. [143] S.I. Miller, A.M. Kukral, J.J. Mekalanos, A two-component regulatory system (phoP phoQ) controls Salmonella typhimurium virulence, Proc Natl Acad Sci U S A 86 (1989) 5054-5058. [144] S.I. Miller, J.J. Mekalanos, Constitutive expression of the phoP regulon attenuates Salmonella virulence and survival within macrophages, J Bacteriol 172 (1990) 24852490.
45
ACCEPTED MANUSCRIPT [145] E.A. Groisman, E. Chiao, C.J. Lipps, F. Heffron, Salmonella typhimurium phoP virulence gene is a transcriptional regulator, Proc Natl Acad Sci U S A 86 (1989) 7077-
PT
7081.
RI
[146] K.L. Roland, L.E. Martin, C.R. Esther, J.K. Spitznagel, Spontaneous pmrA mutants of Salmonella typhimurium LT2 define a new two-component regulatory system with a
SC
possible role in virulence, J Bacteriol 175 (1993) 4154-4164.
NU
[147] J.S. Gunn, S.I. Miller, PhoP-PhoQ activates transcription of pmrAB, encoding a twocomponent regulatory system involved in Salmonella typhimurium antimicrobial peptide
MA
resistance, J Bacteriol 178 (1996) 6857-6864.
[148] E.A. Groisman, J. Kayser, F.C. Soncini, Regulation of polymyxin resistance and
TE
D
adaptation to low-Mg2+ environments, J Bacteriol 179 (1997) 7040-7045. [149] E.G. Vescovi, Y.M. Ayala, E. Di Cera, E.A. Groisman, Characterization of the bacterial
AC CE P
sensor protein PhoQ. Evidence for distinct binding sites for Mg2+ and Ca2+, J Biol Chem 272 (1997) 1440-1443. [150] E.L. Macfarlane, A. Kwasnicka, R.E. Hancock, Role of Pseudomonas aeruginosa PhoPphoQ in resistance to antimicrobial cationic peptides and aminoglycosides, Microbiology 146 ( Pt 10) (2000) 2543-2554. [151] A. Kato, T. Latifi, E.A. Groisman, Closing the loop: the PmrA/PmrB two-component system negatively controls expression of its posttranscriptional activator PmrD, Proc Natl Acad Sci U S A 100 (2003) 4706-4711. [152] M.W. Bader, W.W. Navarre, W. Shiau, H. Nikaido, J.G. Frye, M. McClelland, F.C. Fang, S.I. Miller, Regulation of Salmonella typhimurium virulence gene expression by cationic antimicrobial peptides, Mol Microbiol 50 (2003) 219-230.
46
ACCEPTED MANUSCRIPT [153] M.W. Bader, S. Sanowar, M.E. Daley, A.R. Schneider, U. Cho, W. Xu, R.E. Klevit, H. Le Moual, S.I. Miller, Recognition of antimicrobial peptides by a bacterial sensor kinase,
PT
Cell 122 (2005) 461-472.
RI
[154] E. Choi, E.A. Groisman, D. Shin, Activated by different signals, the PhoP/PhoQ twocomponent system differentially regulates metal uptake, J Bacteriol 191 (2009) 7174-
SC
7181.
NU
[155] S.M. Richards, K.L. Strandberg, J.S. Gunn, Salmonella-regulated lipopolysaccharide modifications, Subcell Biochem 53 (2010) 101-122.
MA
[156] E. Garcia Vescovi, F.C. Soncini, E.A. Groisman, Mg2+ as an extracellular signal: environmental regulation of Salmonella virulence, Cell 84 (1996) 165-174.
149.
TE
D
[157] M. Otto, Bacterial sensing of antimicrobial peptides, Contrib Microbiol 16 (2009) 136-
AC CE P
[158] S.M. Richards, K.L. Strandberg, M. Conroy, J.S. Gunn, Cationic antimicrobial peptides serve as activation signals for the Salmonella Typhimurium PhoPQ and PmrAB regulons in vitro and in vivo, Front Cell Infect Microbiol 2 (2012) 102. [159] S.Y. Park, E.A. Groisman, Signal-specific temporal response by the Salmonella PhoP/PhoQ regulatory system, Mol Microbiol 91 (2014) 135-144. [160] E.J. Rubin, C.M. Herrera, A.A. Crofts, M.S. Trent, Making the connection: PmrD is required for modifications to Escherichia coli endotoxin that promote antimicrobial resistance, Antimicrob Agents Chemother (2015). [161] J.E. Moss, P.E. Fisher, B. Vick, E.A. Groisman, A. Zychlinsky, The regulatory protein PhoP controls susceptibility to the host inflammatory response in Shigella flexneri, Cell Microbiol 2 (2000) 443-452.
47
ACCEPTED MANUSCRIPT [162] A. Llama-Palacios, E. Lopez-Solanilla, C. Poza-Carrion, F. Garcia-Olmedo, P. Rodriguez-Palenzuela, The Erwinia chrysanthemi phoP-phoQ operon plays an important
PT
role in growth at low pH, virulence and bacterial survival in plant tissue, Mol Microbiol
RI
49 (2003) 347-357.
[163] M. Falord, U. Mader, A. Hiron, M. Debarbouille, T. Msadek, Investigation of the
SC
Staphylococcus aureus GraSR regulon reveals novel links to virulence, stress response
NU
and cell wall signal transduction pathways, PLoS One 6 (2011) e21323. [164] M. Li, Y. Lai, A.E. Villaruz, D.J. Cha, D.E. Sturdevant, M. Otto, Gram-positive three-
MA
component antimicrobial peptide-sensing system, Proc Natl Acad Sci U S A 104 (2007) 9469-9474.
TE
D
[165] M. Fridman, G.D. Williams, U. Muzamal, H. Hunter, K.W. Siu, D. Golemi-Kotra, Two unique phosphorylation-driven signaling pathways crosstalk in Staphylococcus aureus to
7986.
AC CE P
modulate the cell-wall charge: Stk1/Stp1 meets GraSR, Biochemistry 52 (2013) 7975-
[166] H. Sun, Y. Yang, T. Xue, B. Sun, Modulation of cell wall synthesis and susceptibility to vancomycin by the two-component system AirSR in Staphylococcus aureus NCTC8325, BMC Microbiol 13 (2013) 286. [167] M. Pietiainen, P. Francois, H.L. Hyyrylainen, M. Tangomo, V. Sass, H.G. Sahl, J. Schrenzel, V.P. Kontinen, Transcriptome analysis of the responses of Staphylococcus aureus to antimicrobial peptides and characterization of the roles of vraDE and vraSR in antimicrobial resistance, BMC Genomics 10 (2009) 429. [168] P. Monsieurs, S. De Keersmaecker, W.W. Navarre, M.W. Bader, F. De Smet, M. McClelland, F.C. Fang, B. De Moor, J. Vanderleyden, K. Marchal, Comparison of the
48
ACCEPTED MANUSCRIPT PhoPQ regulon in Escherichia coli and Salmonella typhimurium, J Mol Evol 60 (2005) 462-474.
PT
[169] P. Mandin, H. Fsihi, O. Dussurget, M. Vergassola, E. Milohanic, A. Toledo-Arana, I.
RI
Lasa, J. Johansson, P. Cossart, VirR, a response regulator critical for Listeria monocytogenes virulence, Mol Microbiol 57 (2005) 1367-1380.
SC
[170] J.L. Yu, L. Guo, Quantitative Proteomic Analysis of Salmonella enterica Serovar
NU
Typhimurium under PhoP/PhoQ Activation Conditions, J Proteome Res 10 (2011) 29923002.
Bacteriol 183 (2001) 1835-1842.
MA
[171] E.A. Groisman, The pleiotropic two-component regulatory system PhoP-PhoQ, J
TE
D
[172] R.K. Ernst, T. Guina, S.I. Miller, Salmonella typhimurium outer membrane remodeling: role in resistance to host innate immunity, Microbes Infect 3 (2001) 1327-1334.
AC CE P
[173] M.D. Snavely, C.G. Miller, M.E. Maguire, The mgtB Mg2+ transport locus of Salmonella typhimurium encodes a P-type ATPase, J Biol Chem 266 (1991) 815-823. [174] R. Tamayo, B. Choudhury, A. Septer, M. Merighi, R. Carlson, J.S. Gunn, Identification of cptA, a PmrA-regulated locus required for phosphoethanolamine modification of the Salmonella enterica serovar typhimurium lipopolysaccharide core, J Bacteriol 187 (2005) 3391-3399. [175] A.X. Tran, M.E. Lester, C.M. Stead, C.R. Raetz, D.J. Maskell, S.C. McGrath, R.J. Cotter, M.S. Trent, Resistance to the antimicrobial peptide polymyxin requires myristoylation of Escherichia coli and Salmonella typhimurium lipid A, J Biol Chem 280 (2005) 2818628194.
49
ACCEPTED MANUSCRIPT [176] K. Nishino, F.F. Hsu, J. Turk, M.J. Cromie, M.M. Wosten, E.A. Groisman, Identification of the lipopolysaccharide modifications controlled by the Salmonella PmrA/PmrB system
PT
mediating resistance to Fe(III) and Al(III), Mol Microbiol 61 (2006) 645-654.
RI
[177] S.M. Moskowitz, M.K. Brannon, N. Dasgupta, M. Pier, N. Sgambati, A.K. Miller, S.E. Selgrade, S.I. Miller, M. Denton, S.P. Conway, H.K. Johansen, N. Hoiby, PmrB
SC
mutations promote polymyxin resistance of Pseudomonas aeruginosa isolated from
NU
colistin-treated cystic fibrosis patients, Antimicrob Agents Chemother 56 (2012) 10191030.
MA
[178] L.A. Arroyo, C.M. Herrera, L. Fernandez, J.V. Hankins, M.S. Trent, R.E. Hancock, The pmrCAB operon mediates polymyxin resistance in Acinetobacter baumannii ATCC
TE
D
17978 and clinical isolates through phosphoethanolamine modification of lipid A, Antimicrob Agents Chemother 55 (2011) 3743-3751.
AC CE P
[179] M.D. Adams, G.C. Nickel, S. Bajaksouzian, H. Lavender, A.R. Murthy, M.R. Jacobs, R.A. Bonomo, Resistance to colistin in Acinetobacter baumannii associated with mutations in the PmrAB two-component system, Antimicrob Agents Chemother 53 (2009) 3628-3634.
[180] H.Y. Cheng, Y.F. Chen, H.L. Peng, Molecular characterization of the PhoPQ-PmrDPmrAB mediated pathway regulating polymyxin B resistance in Klebsiella pneumoniae CG43, J Biomed Sci 17 (2010) 60. [181] S. Nakka, M. Qi, Y. Zhao, The PmrAB system in Erwinia amylovora renders the pathogen more susceptible to polymyxin B and more resistant to excess iron, Res Microbiol 161 (2010) 153-157.
50
ACCEPTED MANUSCRIPT [182] A.K. Miller, M.K. Brannon, L. Stevens, H.K. Johansen, S.E. Selgrade, S.I. Miller, N. Hoiby, S.M. Moskowitz, PhoQ mutations promote lipid A modification and polymyxin
RI
Antimicrob Agents Chemother 55 (2011) 5761-5769.
PT
resistance of Pseudomonas aeruginosa found in colistin-treated cystic fibrosis patients,
[183] N.S. Ly, J. Yang, J.B. Bulitta, B.T. Tsuji, Impact of two-component regulatory systems
SC
PhoP-PhoQ and PmrA-PmrB on colistin pharmacodynamics in Pseudomonas aeruginosa,
NU
Antimicrob Agents Chemother 56 (2012) 3453-3456.
[184] R. Tamayo, A.M. Prouty, J.S. Gunn, Identification and functional analysis of Salmonella
MA
enterica serovar Typhimurium PmrA-regulated genes, FEMS Immunol Med Microbiol 43 (2005) 249-258.
TE
D
[185] M. Merighi, A. Carroll-Portillo, A.N. Septer, A. Bhatiya, J.S. Gunn, Role of Salmonella enterica serovar Typhimurium two-component system PreA/PreB in modulating PmrA-
AC CE P
regulated gene transcription, J Bacteriol 188 (2006) 141-149. [186] D. Kraus, S. Herbert, S.A. Kristian, A. Khosravi, V. Nizet, F. Gotz, A. Peschel, The GraRS regulatory system controls Staphylococcus aureus susceptibility to antimicrobial host defenses, BMC Microbiol 8 (2008) 85. [187] M. Meehl, S. Herbert, F. Gotz, A. Cheung, Interaction of the GraRS two-component system with the VraFG ABC transporter to support vancomycin-intermediate resistance in Staphylococcus aureus, Antimicrob Agents Chemother 51 (2007) 2679-2689. [188] S. Gruenheid, H. Le Moual, Resistance to antimicrobial peptides in Gram-negative bacteria, FEMS Microbiol Lett 330 (2012) 81-89. [189] M.G. Habets, M.A. Brockhurst, Therapeutic antimicrobial peptides may compromise natural immunity, Biol Lett 8 (2012) 416-418.
51
ACCEPTED MANUSCRIPT [190] O. Samuelsen, H.H. Haukland, H. Jenssen, M. Kramer, K. Sandvik, H. Ulvatne, L.H. Vorland, Induced resistance to the antimicrobial peptide lactoferricin B in
PT
Staphylococcus aureus, FEBS Lett 579 (2005) 3421-3426.
RI
[191] J. Mehla, S.K. Sood, Substantiation in Enterococcus faecalis of dose-dependent resistance and cross-resistance to pore-forming antimicrobial peptides by use of a
SC
polydiacetylene-based colorimetric assay, Appl Environ Microbiol 77 (2011) 786-793.
NU
[192] J.E. Pollard, J. Snarr, V. Chaudhary, J.D. Jennings, H. Shaw, B. Christiansen, J. Wright, W. Jia, R.E. Bishop, P.B. Savage, In vitro evaluation of the potential for resistance
MA
development to ceragenin CSA-13, J Antimicrob Chemother 67 (2012) 2665-2672. [193] C.R. Raetz, C.M. Reynolds, M.S. Trent, R.E. Bishop, Lipid A modification systems in
TE
D
gram-negative bacteria, Annu Rev Biochem 76 (2007) 295-329. [194] J.W. Handing, A.K. Criss, The Lipooligosaccharide Modifying Enzyme LptA Enhances
AC CE P
Gonococcal Defense Against Human Neutrophils, Cell Microbiol (2014). [195] J.V. Hankins, J.A. Madsen, D.K. Giles, B.M. Childers, K.E. Klose, J.S. Brodbelt, M.S. Trent, Elucidation of a novel Vibrio cholerae lipid A secondary hydroxy-acyltransferase and its role in innate immune recognition, Mol Microbiol 81 (2011) 1313-1329. [196] K. Kawasaki, R.K. Ernst, S.I. Miller, 3-O-deacylation of lipid A by PagL, a PhoP/PhoQregulated deacylase of Salmonella typhimurium, modulates signaling through Toll-like receptor 4, J Biol Chem 279 (2004) 20044-20048. [197] K.E. McAuley, P.K. Fyfe, J.P. Ridge, N.W. Isaacs, R.J. Cogdell, M.R. Jones, Structural details of an interaction between cardiolipin and an integral membrane protein, Proc Natl Acad Sci U S A 96 (1999) 14706-14711.
52
ACCEPTED MANUSCRIPT [198] O. Schneewind, D. Missiakas, Lipoteichoic acids, phosphate-containing polymers in the envelope of gram-positive bacteria, J Bacteriol 196 (2014) 1133-1142.
PT
[199] W. Fischer, Lipoteichoic acid and lipids in the membrane of Staphylococcus aureus, Med
RI
Microbiol Immunol 183 (1994) 61-76.
[200] K.S. Molnar, M. Bonomi, R. Pellarin, G.D. Clinthorne, G. Gonzalez, S.D. Goldberg, M.
SC
Goulian, A. Sali, W.F. DeGrado, Cys-scanning disulfide crosslinking and bayesian
NU
modeling probe the transmembrane signaling mechanism of the histidine kinase, PhoQ,
AC CE P
TE
D
MA
Structure 22 (2014) 1239-1251.
53
ACCEPTED MANUSCRIPT Table 1: Details of the modifications and the enzymes responsible for the LPS modifications that are detailed in Figure 2 and Figure3.
E.
coli,
RI
ArnT S. enterica, (orf5, aeruginosa PmrK, or ifbI)
Protein function Reference Adds glycine to the secondary acyl chain [29] of the hexa-acylated lipid A
PT
Organism V. cholerae
P. Adds L-Ara4N (4-amino-4-deoxy-L-arabinose) [108, 110] to one or both of the phosphate groups of lipid A
SC
Gene AlmG
F. Adds galactosamine (GalN) to the phosphate [110, 113, tularensis subspecies novicida substituent at the 1-position of lipid A 193]
ArnT ortholog
Bordetella pertussis
EptA (pmrC)
H. pylory, S. enterica, E. coli
EptB
S. enterica, E.coli
LptA
N. gonorrhoeae, meningitides
LpxF
H. pylory, F. tularensis
Dephosphorylates lipid A at position 4’ [99, 193] leaving unmodified hydroxyl group.
LpxM
E. coli
Adds myristate (C14:0) to the myristate on the [195] 3′-position of the glucosamine disaccharide base of the Lipid A
LpxN
V. cholerae
Adds of a 3-hydroxylaurate (3-OH C12:0) to [195] the myristate on the 3′-position of the glucosamine disaccharide base of the Lipid A
LpxO
S. enterica
Hydroxylzes the 2’ position of the myristoyl [72, 107] residue at the 3′ position of the glucosamine disaccharide base of the Lipid A
PagL
S. enterica
Removes 3-hydroxymyristate from position 3’
PagP
S. enterica
Adds palmitoyl group to PG on the outer [72, 196] membrane and Lipid A
NU
ArnT ortholog
MA
Adds of glucosamine GlcN to both phosphate [110] groups of lipid A
AC CE P
TE
D
Adds phosphoethanolamine (PEA) to the 4’ [99, 101] phosphate on lipid A
Adds phosphoethanolamine (PEA) to the 3- [102] deoxy-D-manno-octulosonic acid (Kdo) of the inner core
Neisseria Adds phosphoethanolamine (PEA) to the 4’ [104, 105, phosphate on lipid A 194]
54
[72, 196]
ACCEPTED MANUSCRIPT
MprF
S. aureus
VraFG
S. aureus
adds D-alanine to the Gro-P moiety of the [77] LTA responsible for the addition of lysine and [56, alanine to PG on the plasma membrane 62] AMP efflux pump
PT
S. aureus
AC CE P
TE
D
MA
NU
SC
RI
DltA
55
[169]
60,
ACCEPTED MANUSCRIPT Figures Legends Figure 1: Outer membrane of Gram-negative bacteria (S. enterica): Schematic
PT
representation of the inner and outer leaflets of the outer membrane. The outer leaflet consists of
RI
LPS and PG. The LPS moiety consists of lipid A- di-glucoseamine linked to fatty acids, and is the major component of the outer leaflet. This lipid A is linked to sugars of the core (colored
SC
hexagons), and the O-antigens (black-lined hexagons). The inner leaflet on the contrary, contains
NU
PG, DPG and LPG or APG [58, 62, 197].
MA
Figure 2: D-alanylation of LTA. Illustration of D-alanylation of LTA of S. aureus. D-alanine is added to the repeated unit of the LTA by the enzyme DltA, and contributes to bacterial resistance
TE
D
[198, 199] .
AC CE P
Figure 3: Lipid A and inner core modifications. The outer surface of Gram-negative bacteria is mostly LPS. This LPS consists of three components (i) lipid A; (ii) core (inner and outer core); (iii) O-antigen (from the inside to the outside). The cartoon summarizes the modifications that take place on the lipid A and the inner core, which were shown to be involved in AMP resistance and susceptibility. Each modification of the lipid A and inner core are colored in a matching color as the name of the enzyme responsible for it. Note * Not all modifications occur in all bacteria species, and they might be relevant to one or more bacterial species (a partial modification per species can be found in Table 1). ** The current illustration refers to a structure of E. coli and S. enterica, but other Gram-negative bacteria might have an altered structures of lipid A and inner core.
56
ACCEPTED MANUSCRIPT Figure 4: TCS activation contributes to AMP resistance. Several TCS are known to be involved in AMP resistance in Gram-positive and Gram-negative bacteria. In Gram-negative
PT
bacteria a sensor histidine kinase, PhoQ or PmrB, is located on the inner membrane (plasma
RI
membrane) and activated by several environmental signals (such as AMPs), some of which are indicated in the cartoon. Upon activation it undergoes phosphorylation and phosphorylates a
SC
response regulator, PhoP and PmrA respectively. The response regulator acts as transcription
NU
factor and activates many genes some of which were found to be related to AMP resistance. There is a crosstalk between these two systems via PmrD, which directly activates PmrA in a
MA
PmrB-independent manner. In Gram-positive bacteria GraRS (ApsSR) is another TCS that is known to be activated by AMPs and is involved in AMP resistance. [57, 107, 160, 177, 186, 187,
AC CE P
TE
D
200]
57
ACCEPTED MANUSCRIPT
AC CE P
TE
D
MA
NU
SC
RI
PT
Figure 1
58
ACCEPTED MANUSCRIPT
AC CE P
TE
D
MA
NU
SC
RI
PT
Figure 2
59
ACCEPTED MANUSCRIPT
AC CE P
TE
D
MA
NU
SC
RI
PT
Figure 3
60
ACCEPTED MANUSCRIPT
AC CE P
TE
D
MA
NU
SC
RI
PT
Figure 4
61
ACCEPTED MANUSCRIPT Declaration
AC CE P
TE
D
MA
NU
SC
RI
PT
The authors have no conflict of interest in this study
62
ACCEPTED MANUSCRIPT Highlights
AC CE P
TE
D
MA
NU
SC
RI
PT
1. An updated review on multidrug resistance bacteria against antimicrobial peptides. 2. Resistance towards AMPs can be developed, by modification of bacterial surface molecules, secretion of protective material and up-regulation or elimination of specific proteins. 3. Because of the general mechanisms of attachment and action of AMPs, bacterial resistance to AMPs often involves biophysical and biochemical changes to the bacterial cell wall. 4. The review focuses on the biophysical, surface and surrounding changes that bacteria undergo in acquiring resistance to AMPs.
63
