BioMed Research International

BioMed Research International / 2015 / Article

Review Article | Open Access

Volume 2015 |Article ID 679109 | 11 pages | https://doi.org/10.1155/2015/679109

Antibacterial Mechanisms of Polymyxin and Bacterial Resistance

Academic Editor: Roy Gross
Received29 Aug 2014
Accepted10 Nov 2014
Published15 Jan 2015

Abstract

Multidrug resistance in pathogens is an increasingly significant threat for human health. Indeed, some strains are resistant to almost all currently available antibiotics, leaving very limited choices for antimicrobial clinical therapy. In many such cases, polymyxins are the last option available, although their use increases the risk of developing resistant strains. This review mainly aims to discuss advances in unraveling the mechanisms of antibacterial activity of polymyxins and bacterial tolerance together with the description of polymyxin structure, synthesis, and structural modification. These are expected to help researchers not only develop a series of new polymyxin derivatives necessary for future medical care, but also optimize the clinical use of polymyxins with minimal resistance development.

1. Introduction

An enormous and growing threat that some bacteria are becoming resistant to almost all available antibiotics is proposed to the world [1]. So far, there is no breakthrough in developing new drugs to kill multidrug-resistance (MDR) microorganisms, and the use of β-lactam, quinolone, or aminoglycoside is ineffective. The class of polymyxin antibiotics is increasingly considered as the final option of antibiotic therapy for MDR bacteria that are resistant to almost all other currently available antibiotics [2, 3]. Polymyxins consist of polymyxins A~E, of which polymyxin B and polymyxin E (colistin) are currently used as clinical medicines. In general, they have a narrow antibacterial spectrum mainly against the Gram-negatives [4].

Polymyxin is an old class of nonribosomal cyclic lipopeptide antibiotics originally discovered in 1947 [5]. Since 1959, polymyxin E has been used for the treatment of Gram-negative bacterial infection. However, in the 1970s, clinical use of polymyxin E and polymyxin B was limited due to their serious nephrotoxicity and neurotoxicity after parenteral administration. Together with the emergence of less-toxic aminoglycosides and other antipseudomonal agents [6], its parenteral use was almost completely abandoned in the 1980s. The revival of polymyxin has been coming since the mid-1990s, due to the lack of novel antibiotics against prevalent MDR Gram-negative bacteria [7]. However, the concerns on their nephrotoxicity and neurotoxicity still remain. Accordingly, colistin methanesulfonate (CMS), a prodrug, is typically applied, from which the active compound is slowly released in the blood.

Bacteria are usually able to evolve different strategies to sense, respond, and adapt to bactericidal agents including polymyxin. Therefore, novel polymyxin derivatives with less toxicity and higher bactericidal activity are highly desirable. This communication mainly aims to summarize and discuss the current understanding of antibacterial mechanisms of polymyxin and the corresponding bacterial resistance. We hope that this will serve as an up-to-date reference for researchers to develop polymyxin analogues with better antibacterial activity and less adaptable bacterial tolerance.

2. Polymyxin Structure and Synthesis

2.1. Chemical Structure

The structure of polymyxin is usually described as shown in Figure 1(a) due to the most thorough investigation on polymyxin B and polymyxin E. Its basic structure is a cyclic heptapeptide with a tripeptide side chain acylated by a fatty acid at amino terminus [8]. Polymyxin B and polymyxin E (Table 1) share almost identical primary sequence with major difference present at position 6 where D-Phe (D-phenylalanine) in polymyxin B is replaced by D-Leu (D-leucine) in polymyxin E [9]. The intramolecular cyclic heptapeptide loop is linked between amino group of side chain on diaminobutyric acid (Dab) residue at position 4 and carboxyl group of C-terminal L-Thr (L-threonine) residue at position 10. Therefore, its decapeptide sequence includes three parts, namely, a heptapeptide loop, a tripeptide side chain, and a fatty acid chain [10]. Polymyxin also bears other remarkable structural features, including cationic (L-α-γ-Dab) residues, making it polycationic at pH 7.4, and two hydrophobic domains (N-terminal fatty acyl chain and D-Phe6-L-Leu7 segment on polymyxin B or D-Leu6-L-Leu7 segment on polymyxin E). The mixture of lipophilic and hydrophilic groups makes it amphipathic [9, 10]. In addition, three-dimensional NMR analysis has revealed that polymyxin molecule is folded to form two distinct faces for polar and hydrophobic domains, thereby conferring structural amphipathicity that is essential for its antibacterial activity [9].


PolymyxinFatty acidaR6b

Polymyxin B1MOAD-Phe
Polymyxin B2IOAD-Phe
Polymyxin E1 (colistin A)MOAD-Leu
Polymyxin E2 (colistin B)IOAD-Leu

MOA: 6-methyloctanoic acid; IOA: isooctanoic acid; Phe: phenylalanine; Leu: leucine.
bR6 means amino acid residue at position 6 on polymyxin.
2.2. Polymyxin Biosynthesis

Different from ribosomal peptides that are synthesized by translation of mRNA, polymyxin is produced by nonribosomal peptide synthetase system (NRPS), a multienzyme complex with modular structures [11]. The typical module of NRPS mainly consists of three core domains: adenylation (A) domain, thiolation (T) domain (phosphopantetheine attachment site or peptidyl carrier protein), and condensation (C) domain. The A-domain plays a role in specific recognition and activation of amino acid or hydroxy acid through the formation of an aminoacyl adenylate. Then, the activated amino acid will be covalently bonded to 4′-phosphopantetheinyl (4′PPant) cofactor on T-domain via thioester formation. The T-domain mainly functions as transportation of substrate and elongation of intermediate to catalytic centers. Subsequently, the C-domain will catalyze the elongation of peptidyl chain by attaching thioesterified amino acid on phosphopantetheinyl arm at the upstream of T-domain to amino acid at the downstream of T-domain [12]. It is worth noting that NRPS can also include additional modules, such as epimerization and termination domains.

The modules and domains can orderly get together to form gene cluster. The biosynthetic gene cluster of polymyxin is called pmx cluster, including five open reading frames, namely, pmxA, pmxB, pmxC, pmxD, and pmxE (Figures 1(b) and 1(c)). Accordingly, they encode three polymyxin synthetases, PmxA, PmxB, and PmxE, and two membrane transport proteins, PmxC and PmxD [13]. PmxA comprises four modules whose amino acid substrates are Leu on polymyxin E or Phe on polymyxin B, Thr, Dab and Dab, and a C-domain. PmxB, responsible for the termination of polymyxin synthesis, composes only one module with Thr as its amino acid substrate. PmxE has five modules whose amino acid substrates are Dab, Thr, Dab, Dab, and Dab, and a C-domain. Based on the polymyxin structure, the order of modules for amino acid assembly during polymyxin synthesis should be PmxE-PmxA-PmxB [14], consistent with the order of ten amino acid groups on polymyxin molecule.

3. Polymyxin Derivatives

As mentioned earlier, novel polymyxin derivatives with either higher antimicrobial activity or lower toxicity are highly promising. So far, researches on modification of polymyxin are mainly focused on the change of N-terminal fatty acyl chain length and hydrophobic domain of D-Phe6-L-Leu7 (polymyxin B) or D-Leu6-L-Leu7 (polymyxin E) and substitution of Dab side chains and amino acids [9].

The polymyxin toxicity is partly attributed to N-terminal fatty acyl segment [15]. The derivatives of polymyxin E with C9–C14 unbranched fatty acyl chains showed higher activity against polymyxin-resistant strains and Gram-positive bacteria with longer fatty acyl chain, whereas the derivatives with C10 and C12 fatty acyl chain were more effective against polymyxin-susceptible strains [16, 17]. The derivatives of polymyxin B with modified N-terminal fatty acyl chain have also been investigated to show that the analogue with intermediate length of N-terminal fatty acyl chain (octanoyl, C8) was optimal [18], while the ones with either longer (myristoyl, C14) [19] or shorter (acetyl, C2) [11, 20] chains displayed poorer antimicrobial activity. Moreover, the smaller acetyl nonapeptide analogues showed decreased antimicrobial activity against Escherichia coli and Salmonella enterica. Recently, it was revealed that, compared to polymyxin B with octanoyl (C8) fatty acyl chain [21], the analogues with N-terminal fatty acyl chains > C8 or 6-methyl moiety yielded decreased antimicrobial activity, due to the sterically hindered outer membrane (OM) insertion by fatty acyl moiety [22]. In addition, the substitution of N-terminus of polymyxin B with hydrophobic Fmoc group can significantly enhance antimicrobial activity and reduce toxicity [23].

The cationic Dab residue on polymyxin, particularly within the cyclic heptapeptide, plays a key role in polymyxin’s antimicrobial activity through electrostatic interaction with phosphates of lipid A on bacterial member. The Dab on polymyxin has three important features, including cationic character of side chain groups, two-methylene group of Dab side chain, and specific order of Dab residues within the primary sequence that gives the proper spatial distribution of positive charge [9]. Various synthetic or semisynthetic modifications have been applied to Dab in order to increase antimicrobial activity or minimize potential toxicity [24, 25]. The Nγ-benzyl derivatives of polymyxin B and polymyxin E were synthesized by substituting Dab sides with lipophilic groups. Because of the reduced cationic character, the Nγ-benzyl derivatives appeared to have higher activity against Gram-positive Staphylococcus aureus and lower activity against Gram-negative E. coli [24]. The polymyxin B derivatives with positively charged or polar side chain on modified Dab showed better antimicrobial activity than polymyxin B and broadened the antibacterial spectrum [25]. It has been found thatthe Dabs within the heptapeptide ring on polymyxin B were more critical than the ones in linear tripeptide segment for antimicrobial activity [26]. As a kind of aminoglycoside, polymyxin carries 5 positive charges. Its nephrotoxicity is due to the highly cationic nature of molecule. Recently, it was reported that the polymyxin analogue with substitution of Dabs at positions 1 and 3 with Thr, Ser, or aminobutyryl group reduced its nephrotoxicity [27].

The hydrophobic domain of D-Phe6-L-Leu7 (polymyxin B) or D-Leu6-L-Leu7 (polymyxin E) can also affect its antibacterial activity through insertion with bacterial OM [26]. The hydrophobic domain of polymyxin B was evaluated by replacing D-Phe6 with D-Trp or D-Tyr and substituting L-Leu7 with L-Phe or L-Ala. The substitution of D-Phe6 and L-Leu7 with D-Tyr and L-Ala, respectively, significantly reduced LPS affinity and OM permeabilizing activity of polymyxin. The substitution of D-Phe6 with D-Trp, despite the similar affinity to LPS, displayed marginally reduced OM permeabilizing activity. The substitution of D-Phe6 with L-Phe resulted in an almost complete loss of OM permeabilizing activity [28]. It was reported that the replacement of D-Phe6-L-Leu7 segment with dipeptide mimics caused the loss of activity against E. coli [29].

Besides the above modifications, the size of cyclic peptide ring [30], the length of N-terminal linear tripeptide segment [31], and the generation of mimetic compounds [32] are also involved in polymyxin modification. A series of polymyxin B nonapeptide analogs with a cyclic peptide ring in size from 20 to 26 atoms were synthesized [30]. It was found that, among them, the one with native 23 atoms displayed the best OM permeabilizing activity and provided the most ideal structural configuration for potent antimicrobial activity. The analogues with a tripeptide linear tail of Met-Leu-Phe at N-terminus exhibited 8 to 10 times less toxicity than parent molecules [31]. The analogs of polymyxin B were designed to form amphipathic structure when they bind to LPS through tandemly repeated sequences of alternating cationic (Lys) and nonpolar (Val or Phe) residues [32]. It was found that the new analogs had strong antimicrobial effects but lacked hemolytic activity, highlighting the importance of peptide amphipathicity.

4. Antibacterial Mechanism of Polymyxins

4.1. Membrane Lysis Death Pathway

In Gram-negative bacteria, OM acts as a permeability barrier. The initial target of polymyxin is LPS of OM. Polymyxin can selectively bind to LPS, coincident with its narrow spectrum of antibacterial activity against Gram-negative bacteria [9]. LPS is composed of three domains: innermost lipid A, central core oligosaccharide region, and outermost O-antigen chain [33]. Among them, the most important domain is lipid A which serves as a hydrophobic anchor with tight packing of fatty acyl chains to stabilize overall OM structure. Some divalent cations such as Ca2+ and Mg2+ usually serve as a bridge between the adjacent LPS molecules to stabilize monolayer [34, 35].

It is generally believed that the polymyxin kills bacteria through membrane lysis, as shown in Figure 2(a) (left). Firstly, the protonation of free γ-amines present on positively charged Dab residues provides a means of electrostatic attraction to negatively charged phosphate headgroups of lipid A, resulting in displacement of divalent cations (Ca2+ and Mg2+) [9, 10]. After this initial electrostatic interaction, the polymyxin molecule will insert its hydrophobic N-terminal fatty acyl chain and D-Phe6-L-Leu7 (polymyxin B) or D-Leu6-L-Leu7 (polymyxin E) segment into OM. This insertion will weaken the packing of adjacent lipid A, thus inducing the expansion of OM monolayer [10, 36]. Eventually, this facilitates the formation of destabilized areas through which polymyxin will cross OM [37, 38]. Finally, polymyxin will destroy the physical integrity of phospholipid bilayer of inner membrane (IM) through membrane thinning by straddling the interface of hydrophilic headgroups and fatty acyl chains [9], leading to IM lysis and cell death.

4.2. Vesicle-Vesicle Contact Pathway

An alternative mechanism, called vesicle-vesicle contact, has also been proposed [39, 40]. It is believed that polymyxin can mediate the contacts between periplasmic leaflets of IM and OM. The complex structure of OM consists of an inner phospholipid leaflet and an outer leaflet that predominantly contains LPS, proteins, and lipoproteins [10]. As shown in Figure 2(b) (right), polymyxin can bind to both anionic phospholipid vesicles, namely, inner phospholipid leaflets of OM and IM, and promote the exchange of phospholipids between vesicles. In brief, with the help of electrostatic interaction and two hydrophobic domains, the polymyxin molecule can enter into and cross OM. Then, polymyxin will induce the lipid exchange between leaflets of IM and OM, triggering the loss of specificity of phospholipid composition. This can potentially cause an osmotic imbalance, leading to cell lysis [39, 40]. It was reported that an analogue of polymyxin B with an intervening Dab residue in D-Phe6-L-Leu7 domain was much more effective in inducing lipid exchange through vesicle-vesicle contact and gave higher permeabilizing activity [41]. Another analogue of polymyxin B with substitution of D-Phe6 with D-Trp can bind to bacterial vesicles and induce the formation of vesicle-vesicle contact [42].

4.3. Hydroxyl Radical Death Pathway

A new report showed that polymyxin can possibly induce rapid cell death through the accumulation of hydroxyl radical (OH) (Figure 3). This hypothesis is based on the oxidative stress due to polymyxin-induced formation of reactive oxygen species (ROS), including superoxide (), hydrogen peroxide (H2O2), and OH in Gram-negative bacterial cells [43]. It has been hypothesized that will be induced when polymyxin molecules enter into and cross OM and IM [44, 45]. Then, will be converted to H2O2 by superoxide dismutases (SOD) present in cells. Subsequently, H2O2 will oxidize ferrous iron (Fe2+) to ferric iron (Fe3+), along with the formation of OH, which is called Fenton reaction [44, 45]. When the concentration of OH reaches an uncontrollable level, it will result in oxidative damage of DNA, lipids, and proteins and eventually cause cell death [44, 46]. In this process, the damage and resynthesis of Fe-S dependent proteins, especially Fe-S dependent dehydratase, such as dihydroxy-acid dehydratase (DHAD), are important. The exposed Fe-S cluster will be oxidized by to an unstable species with H2O2 formation and Fe2+ release. Similar to , H2O2 can also destroy the Fe-S cluster, leading to the loss of Fe3+ and inactivation of Fe-S dependent protein [43]. After damage by either or H2O2, the inactive Fe-S cluster can be repaired by protein YggX (a member of the SoxRS regulon) and a di-iron protein YtfE in the presence of Fe3+ [43], whose uptake will be strongly triggered by ferric uptake regulator. It has been demonstrated that the OH production will increase in polymyxin B- or polymyxin E-treated Acinetobacter baumannii, leading to rapid cell death [47]. Moreover, the killing of A. baumannii by polymyxins was delayed in the presence of inhibitors that can both directly and indirectly block the ROS production.

5. Mechanisms of Bacterial Resistance to Polymyxins

5.1. PhoP-PhoQ Two-Component System

It is becoming increasingly clear that polymyxin resistance in Gram-negative bacteria involves the multitier upregulation of a number of regulatory systems [48, 49]. The OM usually serves as a permeability barrier to protect Gram-negative bacteria from various antibiotics and chemicals [34]. The critical step of bactericidal activity of polymyxin is the electrostatic interaction between positively charged Dab residues on polymyxin and negatively charged phosphate groups on lipid A of LPS [9]. The bacterial cell is able to reduce the initial electrostatic attraction by reducing net negative charge of OM via lipid A modification, thereby increasing resistance to polymyxin. The most common polymyxin-resistance mechanism inbacteriais attributed to the shielding of phosphates on lipid A with positively charged groups, such as phosphoethanolamine (pEtN) and L-4-aminoarabinose (L-Ara4N) [5053], which is mediated by PhoP-PhoQ regulatory system encoded by phoP locus (Figure 4).

Activated by PhoP-PhoQ, the PmrA-PmrB encoded by pmrCAB operon is the major regulator to mediate the LPS modification in Gram-negative bacteria [54]. PmrA-dependent modification can occur on each of the three distinct LPS domains, namely, lipid A, core polysaccharide, and O-antigen chain. In the innermost lipid A, the interaction of either pEtN or L-Ara4N with lipid A will neutralize lipid A phosphates and confer resistance to polymyxin B [33, 55]. The ugd gene encoding UDP-glucose dehydrogenase and pbg gene encoding L-Ara4N transferase are both activated by PmrA. They are necessary for biosynthesis and incorporation of L-Ara4N [55]. On the other hand, an IM PmrC protein encoded by PmrA-activated pmrC gene is needed for pEtN incorporation into lipid A. In the central core polysaccharide region, the decoration of heptose (I) phosphate with pEtN can further increase the resistance to polymyxin B [56]. The PmrA-activated cptA gene encoding for pEtN phosphotransferase specific for the core is responsible for the modification of heptose (I) phosphate with pEtN. Moreover, the PmrA-activated PmrG protein which is normally introduced by RfaY protein is a phosphatase for removing the phosphate from heptose (II) phosphate [33]. In the outermost O-antigen chain, the increase of O-antigen length will result in the heightened resistance to polymyxin B, which can be boosted up by iron. The O-antigen synthesis of S. enterica is controlled by the products of wzzst and wzzfepE genes that are controlled by PmrA-PmrB regulatory system [57, 58]. The transcriptional induction of wzzst and wzzfepE is activated by PmrA through directly binding to their promoter, consequently increasing the amount of O-antigen in LPS and finally increasing resistance [57, 58].

The PhoP-PhoQ two-component system in S. enterica has been well characterized [54]. It acts as a master regulator of virulence and evasion of killing by polymyxin [59]. In response to sublethal concentrations of polymyxin, PhoQ, an IM sensor kinase, will phosphorylate the cytoplasmic regulator PhoP, leading to activation of PmrA-PmrB via PhoP-activated PmrD protein whose product affects the phosphorylation of PmrA [54, 6063]. Under extracytoplasmic Fe3+ or Al3+ and acidic pH [64, 65], the sensor PmrB promotes phosphorylation of its cognate regulator PmrA, resulting in the transcription of PmrA-activated genes [66] and repression of PmrA-repressed genes [67]. Consequently, the PmrA-PmrB system activates the expression of PmrC or Ugd/PbgP, necessary for the covalent modification of phosphate groups on lipid A [68]. In addition, the PmrA-PmrB system will use PmrR to inhibit the activity of LpxT, a constitutively synthesized IM enzyme that generates diphosphorylated lipid A at 1-position (1-PP) [69]. All these PmrA-regulated modifications will decrease the overall negative charge of LPS, thereby avoiding the interaction with positively charged Dab residues of polymyxin. Upon the removal of stress from polymyxin, the phosphorylated PmrA (PmrA-P) in cells will be downregulated to appropriate level through three ways. Firstly, the PmrA-P protein can be positively downregulated through transcription of pmrCAB operon [70]; secondly, the PhoP-PhoQ two-component system can control the expression of pmrD gene to repress PmrA-P protein [71]; thirdly, as an intrinsic feedback mechanism, PmrB will dephosphorylate PmrA-P [67].

5.2. Species-Specific Resistance Mechanisms

Besides the LPS-binding pathway regulated by PhoP-PhoQ system, there are other unique and often species-specific mechanisms in polymyxin resistance. Multidrug efflux pumps play an important role of polymyxin resistance in Gram-negative and Gram-positive pathogens. The MexAB-OprM efflux pump in Pseudomonas aeruginosa has been proposed to confer tolerance towards polymyxin E, due to the increase of mexAB-oprM expression in P. aeruginosa upon polymyxin E exposure [72, 73]. The AcrAB efflux pump encoded by acrAB operon can give Klebsiella pneumoniae resistance to polymyxin B [74]. Moreover, the AcrAB efflux pump is also associated with polymyxin resistance in E. coli [75]. A multidrug efflux pump NorM in Burkholderia vietnamien has been shown to contribute to polymyxin resistance [76]. All these efflux pumps are thought to transport and pump out polymyxins present in cells.

In addition, polymyxin resistance is also thought to be associated with the expression of OM proteins in bacteria. It has been believed that the OM protein OprH, a membrane stabilization protein, can promote resistance to polymyxin B in P. aeruginosa [77]. The OM protein OmpA in K. pneumoniae can help to clearinfections, conferring resistance to antimicrobial peptides [78]. It has been found that the absence of OmpA decreases the expression of capsule polysaccharide, thereby increasing susceptibility to polymyxin B [79]. The capsule polysaccharide could increase resistance of K. pneumoniae to polymyxins [80]. Since the capsule polysaccharides are anionic whereas polymyxins are cationic, the capsule polysaccharides can bind to polymyxin to reduce the amount of peptides reaching bacterial surface. This will neutralize the bactericidal activity of polymyxin, at last enhancing electrostatic interaction between capsule polysaccharide and polymyxin [81].

Recently, It was found that the complete loss of LPS could lead to high-level polymyxin E resistance in A. baumannii, clearly indicating that the interaction of polymyxin E with LPS is critical for bactericidal action against A. baumannii [82, 83]. It is believed that the complete loss of LPS will decrease the target ability of polymyxin to cell, thereby causing high-level polymyxin resistance.

6. Future Perspective

The usefulness of polymyxin B and polymyxin E has been clearly demonstrated by optimizing their clinical use and developing their derivatives with less nephrotoxicity than earlier believed and they have been used as bactericidal agents for around 5 decades. Though polymyxins are mainly applied to killing Gram-negative pathogens, there are increasing reports showing their anti-Gram-positive bacteria activity. This needs to be further investigated for better understanding, because much higher concentration of polymyxin is needed against Gram-positive bacteria than the one against Gram-negative bacteria.

Different from traditional membrane lysis mechanism in bacteria, the OH accumulation is a newly proposed mechanism for polymyxin-induced cell death. However, the pathway to induce OH generation in cells exposed to polymyxin is still unclear. Since Fenton reaction is considered as the possible pathway for OH formation, it is very desirable to carry out detailed characterization on the key components such as SOD, H2O2, and Fe-S cluster in this reaction to fully understand this new mechanism.

Different mechanisms of polymyxin-resistance have been found in bacteria. Resistance to the current polymyxins could become a big global health challenge, because this means that virtually no antibiotics will be available for treatment of serious infections caused by polymyxin-resistant “superbugs.” Therefore, development of a next generation of polymyxin is urgently required. In order to achieve this goal, deeper understanding of the mechanisms of polymyxin antibacterial activity and bacterial resistance is the first and most crucial step.

Conflict of Interests

The authors declare that there is no conflict of interests regarding the publication of this paper.

Authors’ Contribution

Zhiliang Yu and Juanping Qiu contributed equally to this work.

References

  1. G. H. Talbot, J. Bradley, J. E. Edwards Jr., D. Gilbert, M. Scheid, and J. G. Bartlett, “Bad bugs need drugs: an update on the development pipeline from the antimicrobial availability task force of the infectious diseases society of America,” Clinical Infectious Diseases, vol. 42, no. 5, pp. 657–668, 2006. View at: Publisher Site | Google Scholar
  2. L. B. Rice, “Challenges in identifying new antimicrobial agents effective for treating infections with Acinetobacter baumannii and Pseudomonas aeruginosa,” Clinical Infectious Diseases, vol. 43, no. 2, pp. S100–S105, 2006. View at: Publisher Site | Google Scholar
  3. J. Li and R. L. Nation, “Old polymyxins are back: is resistance close?” Clinical Infectious Diseases, vol. 43, no. 5, pp. 663–664, 2006. View at: Publisher Site | Google Scholar
  4. J. Li, R. L. Nation, R. J. Owen, S. Wong, O. Spelman, and C. Franklin, “Antibiograms of multidrug-resistant clinical Acinetobacter baumannii: promising therapeutic options for treatment of infection with colistin-resistant strains,” Clinical Infectious Diseases, vol. 45, no. 5, pp. 594–598, 2007. View at: Publisher Site | Google Scholar
  5. S. Biswas, J. M. Brunel, J. C. Dubus, M. Reynaud-Gaubert, and J. M. Rolain, “Colistin: an update on the antibiotic of the 21st century,” Expert Review of Anti-Infective Therapy, vol. 10, no. 8, pp. 917–934, 2012. View at: Publisher Site | Google Scholar
  6. A. Poudyal, B. P. Howden, J. M. Bell et al., “In vitro pharmacodynamics of colistin against multidrug-resistant Klebsiella pneumoniae,” Journal of Antimicrobial Chemotherapy, vol. 62, no. 6, pp. 1311–1318, 2008. View at: Publisher Site | Google Scholar
  7. A. P. Zavascki, L. Z. Goldani, J. Li, and R. L. Nation, “Polymyxin B for the treatment of multidrug-resistant pathogens: a critical review,” Journal of Antimicrobial Chemotherapy, vol. 60, no. 6, pp. 1206–1215, 2007. View at: Publisher Site | Google Scholar
  8. J. Li, R. L. Nation, R. W. Milne, J. D. Turnidge, and K. Coulthard, “Evaluation of colistin as an agent against multi-resistant Gram-negative bacteria,” International Journal of Antimicrobial Agents, vol. 25, no. 1, pp. 11–25, 2005. View at: Publisher Site | Google Scholar
  9. T. Velkov, P. E. Thompson, R. L. Nation, and J. Li, “Structure-activity relationships of polymyxin antibiotics,” Journal of Medicinal Chemistry, vol. 53, no. 5, pp. 1898–1916, 2010. View at: Publisher Site | Google Scholar
  10. T. Velkov, K. D. Roberts, R. L. Nation, P. E. Thompson, and J. Li, “Pharmacology of polymyxins: new insights into an “old” class of antibiotics,” Future Microbiology, vol. 8, no. 6, pp. 711–724, 2013. View at: Publisher Site | Google Scholar
  11. N. I. Martin, H. Hu, M. M. Moake et al., “Isolation, structural characterization, and properties of mattacin (polymyxin M), a cyclic peptide antibiotic produced by Paenibacillus kobensis M,” The Journal of Biological Chemistry, vol. 278, no. 15, pp. 13124–13132, 2003. View at: Publisher Site | Google Scholar
  12. C. Rausch, I. Hoof, T. Weber, W. Wohlleben, and D. H. Huson, “Phylogenetic analysis of condensation domains in NRPS sheds light on their functional evolution,” BMC Evolutionary Biology, vol. 7, article 78, 2007. View at: Publisher Site | Google Scholar
  13. M. Z. Ansari, G. Yadav, R. S. Gokhale, and D. Mohanty, “NRPS-PKS: a knowledge-based resource for analysis of NRPS-PKS megasynthases,” Nucleic Acids Research, vol. 32, pp. W405–W413, 2004. View at: Publisher Site | Google Scholar
  14. S.-K. Choi, S.-Y. Park, R. Kim et al., “Identification of a polymyxin synthetase gene cluster of Paenibacillus polymyxa and heterologous expression of the gene in Bacillus subtilis,” Journal of Bacteriology, vol. 191, no. 10, pp. 3350–3358, 2009. View at: Publisher Site | Google Scholar
  15. H. Tsubery, H. Yaakov, S. Cohen et al., “Neopeptide antibiotics that function as opsonins and membrane-permeabilizing agents for gram-negative bacteria,” Antimicrobial Agents and Chemotherapy, vol. 49, no. 8, pp. 3122–3128, 2005. View at: Publisher Site | Google Scholar
  16. S. Chihara, A. Ito, and M. Yahata, “Chemical synthesis and characterization of n fattyacyl monoaminoacyl derivatives of colistin nonapeptide,” Agricultural and Biological Chemistry, vol. 38, no. 10, pp. 1767–1777, 1974. View at: Publisher Site | Google Scholar
  17. S. Chihara, A. Ito, and M. Yahata, “Chemical synthesis, isolation and characterization of α-N-fattyacyl colistin nonapeptide with special reference to the correlation between antimicrobial activity and carbon number of fattyacyl moiety,” Agricultural and Biological Chemistry, vol. 38, no. 3, pp. 521–529, 1974. View at: Publisher Site | Google Scholar
  18. S.-Y. Liao, G.-T. Ong, K.-T. Wang, and S.-H. Wu, “Conformation of polymyxin B analogs in DMSO from NMR spectra and molecular modeling,” Biochimica et Biophysica Acta (BBA), vol. 1252, no. 2, pp. 312–320, 1995. View at: Publisher Site | Google Scholar
  19. S. Bhattacharjya, S. A. David, V. I. Mathan, and P. Balaram, “Polymyxin B nonapeptide: conformations in water and in the lipopolysaccharide-bound state determined by two-dimensional NMR and molecular dynamics,” Biopolymers, vol. 41, pp. 251–265, 1997. View at: Google Scholar
  20. J. Mares, S. Kumaran, M. Gobbo, and O. Zerbe, “Interactions of lipopolysaccharide and polymyxin studied by NMR spectroscopy,” Journal of Biological Chemistry, vol. 284, no. 17, pp. 11498–11506, 2009. View at: Publisher Site | Google Scholar
  21. J. A. Orwa, C. Govaerts, R. Busson, E. Roets, A. Van Schepdael, and J. Hoogmartens, “Isolation and structural characterization of colistin components,” The Journal of Antibiotics, vol. 54, no. 7, pp. 595–599, 2001. View at: Publisher Site | Google Scholar
  22. J. A. Orwa, C. Govaerts, R. Busson, E. Roets, A. Van Schepdael, and J. Hoogmartens, “Isolation and structural characterization of polymyxin B components,” Journal of Chromatography A, vol. 912, no. 2, pp. 369–373, 2001. View at: Publisher Site | Google Scholar
  23. K. Okimura, K. Ohki, Y. Sato, K. Ohnishi, and N. Sakura, “Semi-synthesis of polymyxin B (2–10) and colistin (2–10) analogs employing the trichloroethoxycarbonyl (Troc) group for side chain protection of α, γ-diaminobutyric acid residues,” Chemical and Pharmaceutical Bulletin, vol. 55, no. 12, pp. 1724–1730, 2007. View at: Publisher Site | Google Scholar
  24. N. M. Witzke and H. Heding, “Broad spectrum derivatives of polymyxin B and colistin,” The Journal of Antibiotics, vol. 29, no. 12, pp. 1349–1350, 1976. View at: Publisher Site | Google Scholar
  25. J. Weinstein, A. Afonso, M. Eugene Jr., and G. H. Miller, “Selective chemical modifications of polymyxin B,” Bioorganic & Medicinal Chemistry Letters, vol. 8, no. 23, pp. 3391–3396, 1998. View at: Publisher Site | Google Scholar
  26. K. Kanazawa, Y. Sato, K. Ohki et al., “Contribution of each amino acid residue in polymyxin b3 to antimicrobial and lipopolysaccharide binding activity,” Chemical and Pharmaceutical Bulletin, vol. 57, no. 3, pp. 240–244, 2009. View at: Publisher Site | Google Scholar
  27. M. Vaara, J. Fox, G. Loidl et al., “Novel polymyxin derivatives carrying only three positive charges are effective antibacterial agents,” Antimicrobial Agents and Chemotherapy, vol. 52, no. 9, pp. 3229–3236, 2008. View at: Publisher Site | Google Scholar
  28. H. Tsubery, I. Ofek, S. Cohen, M. Eisenstein, and M. Fridkin, “Modulation of the hydrophobic domain of polymyxin B nonapeptide: effect on outer-membrane permeabilization and lipopolysaccharide neutralization,” Molecular Pharmacology, vol. 62, no. 5, pp. 1036–1042, 2002. View at: Publisher Site | Google Scholar
  29. P. C. De Visser, N. M. A. J. Kriek, P. A. V. Van Hooft et al., “Solid-phase synthesis of polymyxin B1 and analogues via a safety-catch approach,” The Journal of Peptide Research, vol. 61, no. 6, pp. 298–306, 2003. View at: Publisher Site | Google Scholar
  30. H. Tsubery, I. Ofek, S. Cohen, and M. Fridkin, “Structure-function studies of polymyxin B nonapeptide: implications to sensitization of Gram-negative bacteria,” Journal of Medicinal Chemistry, vol. 43, no. 16, pp. 3085–3092, 2000. View at: Publisher Site | Google Scholar
  31. H. Tsubery, H. Yaakov, S. Cohen et al., “Neopeptide antibiotics that function as opsonins and membrane-permeabilizing agents for gram-negative bacteria,” Antimicrobial Agents and Chemotherapy, vol. 49, no. 8, pp. 3122–3128, 2005. View at: Publisher Site | Google Scholar
  32. V. Frecer, B. Ho, and J. L. Ding, “De novo design of potent antimicrobial peptides,” Antimicrobial Agents and Chemotherapy, vol. 48, no. 9, pp. 3349–3357, 2004. View at: Publisher Site | Google Scholar
  33. K. Nishino, F.-F. Hsu, J. Turk, M. J. Cromie, M. M. S. M. Wösten, and E. A. Groisman, “Identification of the lipopolysaccharide modifications controlled by the Salmonella PmrA/PmrB system mediating resistance to Fe(III) and Al(III),” Molecular Microbiology, vol. 61, no. 3, pp. 645–654, 2006. View at: Publisher Site | Google Scholar
  34. H. Nikaido, “Molecular basis of bacterial outer membrane permeability revisited,” Microbiology and Molecular Biology Reviews, vol. 67, no. 4, pp. 593–656, 2003. View at: Publisher Site | Google Scholar
  35. C. R. H. Raetz and C. Whitfield, “Lipopolysaccharide endotoxins,” Annual Review of Biochemistry, vol. 71, pp. 635–700, 2002. View at: Publisher Site | Google Scholar
  36. R. E. W. Hancock and D. S. Chapple, “Peptide antibiotics,” Antimicrobial Agents and Chemotherapy, vol. 43, no. 6, pp. 1317–1323, 1999. View at: Google Scholar
  37. J. P. Powers and R. E. Hancock, “The relationship between peptide structure and antibacterial activity,” Peptides, vol. 24, no. 11, pp. 1681–1691, 2003. View at: Publisher Site | Google Scholar
  38. P. Pristovšek and J. Kidrič, “The search for molecular determinants of LPS inhibition by proteins and peptides,” Current Topics in Medicinal Chemistry, vol. 4, no. 11, pp. 1185–1201, 2004. View at: Publisher Site | Google Scholar
  39. A. Clausell, M. Garcia-Subirats, M. Pujol, M. A. Busquets, F. Rabanal, and Y. Cajal, “Gram-negative outer and inner membrane models: insertion of cyclic cationic lipopeptides,” Journal of Physical Chemistry B, vol. 111, no. 3, pp. 551–563, 2007. View at: Publisher Site | Google Scholar
  40. Y. Cajal, J. Rogers, O. G. Berg, and M. K. Jain, “Intermembrane molecular contacts by polymyxin B mediate exchange of phospholipids,” Biochemistry, vol. 35, no. 1, pp. 299–308, 1996. View at: Publisher Site | Google Scholar
  41. A. Clausell, F. Rabanal, M. Garcia-Subirats, M. A. Alsina, and Y. Cajal, “Synthesis and membrane action of polymyxin B analogues,” Luminescence, vol. 20, no. 3, pp. 117–123, 2005. View at: Publisher Site | Google Scholar
  42. A. Clausell, F. Rabanal, M. Garcia-Subirats, M. A. Alsina, and Y. Cajal, “Membrane association and contact formation by a synthetic analogue of polymyxin B and its fluorescent derivatives,” The Journal of Physical Chemistry B, vol. 110, no. 9, pp. 4465–4471, 2006. View at: Publisher Site | Google Scholar
  43. J. A. Imlay, “The molecular mechanisms and physiological consequences of oxidative stress: lessons from a model bacterium,” Nature Reviews Microbiology, vol. 11, no. 7, pp. 443–454, 2013. View at: Publisher Site | Google Scholar
  44. M. A. Kohanski, D. J. Dwyer, B. Hayete, C. A. Lawrence, and J. J. Collins, “A common mechanism of cellular death induced by bactericidal antibiotics,” Cell, vol. 130, no. 5, pp. 797–810, 2007. View at: Publisher Site | Google Scholar
  45. J. Yeom, J. A. Imlay, and W. Park, “Iron homeostasis affects antibiotic-mediated cell death in Pseudomonas species,” The Journal of Biological Chemistry, vol. 285, no. 29, pp. 22689–22695, 2010. View at: Publisher Site | Google Scholar
  46. D. J. Dwyer, M. A. Kohanski, B. Hayete, and J. J. Collins, “Gyrase inhibitors induce an oxidative damage cellular death pathway in Escherichia coli,” Molecular Systems Biology, vol. 3, article 91, 2007. View at: Publisher Site | Google Scholar
  47. T. R. Sampson, X. Liu, M. R. Schroeder, C. S. Kraft, E. M. Burd, and D. S. Weiss, “Rapid killing of Acinetobacter baumannii by polymyxins is mediated by a hydroxyl radical death pathway,” Antimicrobial Agents and Chemotherapy, vol. 56, no. 11, pp. 5642–5649, 2012. View at: Publisher Site | Google Scholar
  48. L. Fernández, C. Álvarez-Ortega, I. Wiegand et al., “Characterization of the polymyxin B resistome of Pseudomonas aeruginosa,” Antimicrobial Agents and Chemotherapy, vol. 57, no. 1, pp. 110–119, 2013. View at: Publisher Site | Google Scholar
  49. S. A. Loutet, L. E. Mussen, R. S. Flannagan, and M. A. Valvano, “A two-tier model of polymyxin B resistance in Burkholderia cenocepacia,” Environmental Microbiology Reports, vol. 3, no. 2, pp. 278–285, 2011. View at: Publisher Site | Google Scholar
  50. L. Fernández, H. Jenssen, M. Bains, I. Wiegand, W. J. Gooderham, and R. E. W. Hancock, “The two-component system CprRS senses cationic peptides and triggers adaptive resistance in Pseudomonas aeruginosa independently of ParRS,” Antimicrobial Agents and Chemotherapy, vol. 56, no. 12, pp. 6212–6222, 2012. View at: Publisher Site | Google Scholar
  51. S. D. Breazeale, A. A. Ribeiro, A. L. McClerren, and C. R. H. Raetz, “A formyltransferase required for polymyxin resistance in Escherichia coli and the modification of lipid A with 4-amino-4-deoxy-L-arabinose: identification and function of UDP-4-deoxy-4-formamido-L-arabinose,” Journal of Biological Chemistry, vol. 280, no. 14, pp. 14154–14167, 2005. View at: Publisher Site | Google Scholar
  52. S. M. Moskowitz, R. K. Ernst, and S. I. Miller, “PmrAB , a two-component regulatory system of Pseudomonas aeruginosa that modulates resistance to cationic antimicrobial peptides and addition of aminoarabinose to lipid A,” Journal of Bacteriology, vol. 186, no. 2, pp. 575–579, 2004. View at: Publisher Site | Google Scholar
  53. Z. Zhou, A. A. Ribeiro, S. Lin, R. J. Cotter, S. I. Miller, and C. R. H. Raetz, “Lipid A modifications in polymyxin-resistant Salmonella typhimurium: PmrA-dependent 4-amino-4-deoxy-L-arabinose, and phosphoethanolamine incorporation,” The Journal of Biological Chemistry, vol. 276, no. 46, pp. 43111–43121, 2001. View at: Publisher Site | Google Scholar
  54. A. Kato, H. D. Chen, T. Latifi, and E. A. Groisman, “Reciprocal control between a bacterium’s regulatory system and the modification status of its lipopolysaccharide,” Molecular Cell, vol. 47, no. 6, pp. 897–908, 2012. View at: Publisher Site | Google Scholar
  55. H. Lee, F.-F. Hsu, J. Turk, and E. A. Groisman, “The PmrA-regulated pmrC gene mediates phosphoethanolamine modification of lipid A and polymyxin resistance in Salmonella enterica,” Journal of Bacteriology, vol. 186, no. 13, pp. 4124–4133, 2004. View at: Publisher Site | Google Scholar
  56. R. Tamayo, B. Choudhury, A. Septer, M. Merighi, R. Carlson, and J. S. Gunn, “Identification of cptA, a PmrA-regulated locus required for phosphoethanolamine modification of the Salmonella enterica serovar typhimurium lipopolysaccharide core,” Journal of Bacteriology, vol. 187, no. 10, pp. 3391–3399, 2005. View at: Publisher Site | Google Scholar
  57. M. A. Delgado, C. Mouslim, and E. A. Groisman, “The PmrA/PmrB and RcsC/YojN/RcsB systems control expression of the Salmonella O-antigen chain length determinant,” Molecular Microbiology, vol. 60, no. 1, pp. 39–50, 2006. View at: Publisher Site | Google Scholar
  58. M. M. Pescaretti, F. E. López, R. D. Morero, and 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, vol. 157, no. 9, pp. 2515–2521, 2011. View at: Publisher Site | Google Scholar
  59. J. Barchiesi, M. E. Castelli, G. Di Venanzio, M. I. Colombo, and E. G. Véscovi, “The Phop/PhoQ system and its role in Serratia marcescens pathogenesis,” Journal of Bacteriology, vol. 194, no. 11, pp. 2949–2961, 2012. View at: Publisher Site | Google Scholar
  60. F. C. Soncini and E. A. Groisman, “Two-component regulatory systems can interact to process multiple environmental signals,” Journal of Bacteriology, vol. 178, no. 23, pp. 6796–6801, 1996. View at: Google Scholar
  61. D. Shin and E. A. Groisman, “Signal-dependent binding of the response regulators PhoP and PmrA to their target promoters in vivo,” The Journal of Biological Chemistry, vol. 280, no. 6, pp. 4089–4094, 2005. View at: Publisher Site | Google Scholar
  62. E. García Véscovi, F. C. Soncini, and E. A. Groisman, “Mg2+ as an extracellular signal: environmental regulation of Salmonella virulence,” Cell, vol. 84, no. 1, pp. 165–174, 1996. View at: Publisher Site | Google Scholar
  63. A. Kato and E. A. Groisman, “Connecting two-component regulatory systems by a protein that protects a response regulator from dephosphorylation by its cognate sensor,” Genes and Development, vol. 18, no. 18, pp. 2302–2313, 2004. View at: Publisher Site | Google Scholar
  64. M. M. S. M. Wösten, L. F. F. Kox, S. Chamnongpol, F. C. Soncini, and E. A. Groisman, “A signal transduction system that responds to extracellular iron,” Cell, vol. 103, no. 1, pp. 113–125, 2000. View at: Publisher Site | Google Scholar
  65. J. C. Perez and E. A. Groisman, “Acid pH activation of the PmrA/PmrB two-component regulatory system of Salmonella enterica,” Molecular Microbiology, vol. 63, no. 1, pp. 283–293, 2007. View at: Publisher Site | Google Scholar
  66. D. Shin, E.-J. Lee, H. Huang, and E. A. Groisman, “A positive feedback loop promotes transcription surge that jump-starts Salmonella virulence circuit,” Science, vol. 314, no. 5805, pp. 1607–1609, 2006. View at: Publisher Site | Google Scholar
  67. W.-S. Yeo, I. Zwir, H. V. Huang, D. Shin, A. Kato, and E. A. Groisman, “Intrinsic negative feedback governs activation surge in two-component regulatory systems,” Molecular Cell, vol. 45, no. 3, pp. 409–421, 2012. View at: Publisher Site | Google Scholar
  68. L. A. Arroyo, C. M. Herrera, L. Fernandez, J. V. Hankins, M. S. Trent, and R. E. W. Hancock, “The pmrCAB operon mediates polymyxin resistance in Acinetobacter baumannii ATCC 17978 and clinical isolates through phosphoethanolamine modification of lipid A,” Antimicrobial Agents and Chemotherapy, vol. 55, no. 8, pp. 3743–3751, 2011. View at: Publisher Site | Google Scholar
  69. C. M. Herrera, J. V. Hankins, and M. S. Trent, “Activation of PmrA inhibits LpxT-dependent phosphorylation of lipid A promoting resistance to antimicrobial peptides,” Molecular Microbiology, vol. 76, no. 6, pp. 1444–1460, 2010. View at: Publisher Site | Google Scholar
  70. J. S. Gunn and S. I. Miller, “PhoP-PhoQ activates transcription of pmrAB, encoding a two-component regulatory system involved in Salmonella typhimurium antimicrobial peptide resistance,” Journal of Bacteriology, vol. 178, no. 23, pp. 6857–6864, 1996. View at: Google Scholar
  71. A. Kato, T. Latifi, and E. A. Groisman, “Closing the loop: the PmrA/PmrB two-component system negatively controls expression of its posttranscriptional activator PmrD,” Proceedings of the National Academy of Sciences of the United States of America, vol. 100, no. 8, pp. 4706–4711, 2003. View at: Publisher Site | Google Scholar
  72. H. P. Schweizer, “Efflux as a mechanism of resistance to antimicrobials in Pseudomonas aeruginosa and related bacteria: Unanswered questions,” Genetics and Molecular Research, vol. 2, no. 1, pp. 48–62, 2003. View at: Google Scholar
  73. S. J. Pamp, M. Gjermansen, H. K. Johansen, and T. Tolker-Nielsen, “Tolerance to the antimicrobial peptide colistin in Pseudomonas aeruginosa biofilms is linked to metabolically active cells, and depends on the pmr and mexAB-oprM genes,” Molecular Microbiology, vol. 68, no. 1, pp. 223–240, 2008. View at: Publisher Site | Google Scholar
  74. E. Padilla, E. Llobet, A. Doménech-Sánchez, L. Martínez-Martínez, J. A. Bengoechea, and S. Albertí, “Klebsiella pneumoniae AcrAB efflux pump contributes to antimicrobial resistance and virulence,” Antimicrobial Agents and Chemotherapy, vol. 54, no. 1, pp. 177–183, 2010. View at: Publisher Site | Google Scholar
  75. D. M. Warner and S. B. Levy, “Different effects of transcriptional regulators MarA, SoxS and Rob on susceptibility of Escherichia coli to cationic antimicrobial peptides (CAMPs): rob-dependent CAMP induction of the marRAB operon,” Microbiology, vol. 156, no. 2, pp. 570–578, 2010. View at: Publisher Site | Google Scholar
  76. C. C. Fehlner-Gardiner and M. A. Valvano, “Cloning and characterization of the Burkholderia vietnamiensis norM gene encoding a multi-drug efflux protein,” FEMS Microbiology Letters, vol. 215, no. 2, pp. 279–283, 2002. View at: Publisher Site | Google Scholar
  77. E. L. A. Macfarlane, A. Kwasnicka, M. M. Ochs, and R. E. W. Hancock, “PhoP-PhoQ homologues in Pseudomonas aeruginosa regulate expression of the outer-membrane protein OprH and polymyxin B resistance,” Molecular Microbiology, vol. 34, no. 2, pp. 305–316, 1999. View at: Publisher Site | Google Scholar
  78. P. Jeannin, G. Magistrelli, L. Goetsch et al., “Outer membrane protein A ( OmpA ): a new pathogen-associated molecular pattern that interacts with antigen presenting cells—impact on vaccine strategies,” Vaccine, vol. 20, no. 4, pp. A23–A27, 2002. View at: Publisher Site | Google Scholar
  79. E. Llobet, C. March, P. Giménez, and J. A. Bengoechea, “Klebsiella pneumoniae OmpA confers resistance to antimicrobial peptides,” Antimicrobial Agents and Chemotherapy, vol. 53, no. 1, pp. 298–302, 2009. View at: Publisher Site | Google Scholar
  80. M. A. Campos, M. A. Vargas, V. Regueiro, C. M. Llompart, S. Albertí, and J. A. Bengoechea, “Capsule polysaccharide mediates bacterial resistance to antimicrobial peptides,” Infection and Immunity, vol. 72, no. 12, pp. 7107–7114, 2004. View at: Publisher Site | Google Scholar
  81. E. Llobet, J. M. Tomás, and J. A. Bengoechea, “Capsule polysaccharide is a bacterial decoy for antimicrobial peptides,” Microbiology, vol. 154, no. 12, pp. 3877–3886, 2008. View at: Publisher Site | Google Scholar
  82. J. H. Moffatt, M. Harper, P. Harrison et al., “Colistin resistance in Acinetobacter baumannii is mediated by complete loss of lipopolysaccharide production,” Antimicrobial Agents and Chemotherapy, vol. 54, no. 12, pp. 4971–4977, 2010. View at: Publisher Site | Google Scholar
  83. J. H. Moffatt, M. Harper, B. Adler, R. L. Nation, J. Li, and J. D. Boyce, “Insertion sequence IS Aba11 is involved in colistin resistance and loss of lipopolysaccharide in Acinetobacter baumannii,” Antimicrobial Agents and Chemotherapy, vol. 55, no. 6, pp. 3022–3024, 2011. View at: Publisher Site | Google Scholar

Copyright © 2015 Zhiliang Yu et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

19860 Views | 3596 Downloads | 68 Citations
 PDF  Download Citation  Citation
 Download other formatsMore
 Order printed copiesOrder

We are committed to sharing findings related to COVID-19 as quickly and safely as possible. Any author submitting a COVID-19 paper should notify us at help@hindawi.com to ensure their research is fast-tracked and made available on a preprint server as soon as possible. We will be providing unlimited waivers of publication charges for accepted articles related to COVID-19.