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Journal of Antimicrobial Chemotherapy logoLink to Journal of Antimicrobial Chemotherapy
. 2012 Aug 16;67(11):2665–2672. doi: 10.1093/jac/dks276

In vitro evaluation of the potential for resistance development to ceragenin CSA-13

Jake E Pollard 1, Jason Snarr 1, Vinod Chaudhary 1, Jacob D Jennings 1, Hannah Shaw 1, Bobbie Christiansen 1, Jonathan Wright 1, Wenyi Jia 2,†, Russell E Bishop 2, Paul B Savage 1,*
PMCID: PMC3468081  PMID: 22899801

Abstract

Objectives

Though most bacteria remain susceptible to endogenous antimicrobial peptides, specific resistance mechanisms are known. As mimics of antimicrobial peptides, ceragenins were expected to retain antibacterial activity against Gram-positive and -negative bacteria, even after prolonged exposure. Serial passaging of bacteria to a lead ceragenin, CSA-13, was performed with representative pathogenic bacteria. Ciprofloxacin, vancomycin and colistin were used as comparators. The mechanisms of resistance in Gram-negative bacteria were elucidated.

Methods

Susceptible strains of Staphylococcus aureus, Pseudomonas aeruginosa and Acinetobacter baumannii were serially exposed to CSA-13 and comparators for 30 passages. MIC values were monitored. Alterations in the Gram-negative bacterial membrane composition were characterized via mass spectrometry and the susceptibility of antimicrobial-peptide-resistant mutants to CSA-13 was evaluated.

Results

S. aureus became highly resistant to ciprofloxacin after <20 passages. After 30 passages, the MIC values of vancomycin and CSA-13 for S. aureus increased 9- and 3-fold, respectively. The Gram-negative organisms became highly resistant to ciprofloxacin after <20 passages. MIC values of colistin for P. aeruginosa and A. baumannii increased to ≥100 mg/L after 20 passages. MIC values of CSA-13 increased to ∼20–30 mg/L and plateaued over the course of the experiment. Bacteria resistant to CSA-13 displayed lipid A modifications that are found in organisms resistant to antimicrobial peptides.

Conclusions

CSA-13 retained potent antibacterial activity against S. aureus over the course of 30 serial passages. Resistance generated in Gram-negative bacteria correlates with modifications to the outer membranes of these organisms and was not stable outside of the presence of the antimicrobial.

Keywords: bacterial resistance, antimicrobial peptide mimics, serial passaging, lipid A modifications

Introduction

The continued emergence of antibiotic- and antimicrobial-resistant bacteria is an impetus for the development of novel means of controlling bacterial growth. The actions of antimicrobial peptides (AMPs) constitute an ancient and ubiquitous means of controlling bacterial growth, and may provide an approach for controlling bacterial growth without readily engendering resistance.1–3 It is apparent that AMPs have played a key role in the innate immune defences of most higher organisms for eons. Nevertheless, bacteria generally remain susceptible to AMPs. Over 1500 examples of AMPs have been isolated from organisms ranging from mammals to insects, and consideration of the variation in the primary sequences of these AMPs suggests that they may have evolved independently dozens of times. The sustained susceptibility of bacteria to AMPs has been a factor in motivating the research into the development of these antimicrobials for clinical use. Challenges encountered in the clinical use of AMPs include the relatively high costs of production of peptide therapeutics and issues of stability in the presence of proteases.

Because bacteria have been exposed to AMPs for such long periods of time, it is not surprising that they have generated means of lessening the impact of AMPs. These means include the release of proteases that degrade AMPs4 and modifications to their membrane structures, which are the primary targets of most AMPs.2,5–8 In Gram-negative bacteria, two-component signal transduction systems, including PhoP-PhoQ and PmrA-PmrB, lead to modifications to the lipid A portion of lipopolysaccharide (LPS).5,6,8–10 Production of PagP (AMP resistance and lipid A acylation protein) is controlled by these systems and this protein transfers additional palmitate to lipid A, which decreases susceptibility to some AMPs.11 A key step in the activity of cationic AMPs is the association with negatively charged bacterial membrane components. To decrease charge recognition, these signal transduction systems lead to modification of the phosphate groups on lipid A with ethanolamine and 4-aminoarabinose.2,5,6,8,9 The positive charges of these compounds offset the negative charges of the phosphate esters and decrease susceptibility to AMPs. A similar tactic is employed by Gram-positive bacteria, e.g. lysine is added to phosphatidyl glycerol to decrease the net negative charge of the bacterial membrane.10 These methods of evading AMPs are due to the action of existing gene products; consequently, they are generally considered adaptational, though mutations leading to membrane modifications have been observed. Though adaptational resistance mechanisms decrease the antibacterial activity of AMPs, resistant bacteria revert to susceptible forms when gene expression regulated by the two-component systems ceases to cause modification of the bacterial membrane components.

In an effort to understand the structural requirements for the antibacterial activities of AMPs and to overcome some of the disadvantages of the use of peptide therapeutics, we have developed non-peptide mimics of AMPs, termed ceragenins.12 These compounds are based on a bile acid scaffold designed to mimic the facially amphiphilic morphology of AMPs. Ceragenins are broad-spectrum antimicrobial agents containing a lipase-resistant ether functionality and, because they are not peptide based, they are insensitive to deactivation by proteases. In addition, they are relatively simple to prepare on a large scale. CSA-13 (Figure 1) is a lead ceragenin and has been the subject of multiple studies (for a recent review see Lai et al.12).

Figure 1.

Figure 1.

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Structure of CSA-13, including a perspective drawing showing its conformation.

As mimics of AMPs, we anticipated that the ceragenins would not readily engender bacterial resistance and that if less susceptible forms of bacteria were identified, these would display the same adaptational modifications to membrane structure seen in response to AMPs. The propensity of bacteria to generate resistance can be evaluated using serial exposure of organisms to antimicrobial agents.13–15 In these experiments, bacteria are exposed to incrementally varied concentrations of the test antibacterial agent, and organisms that grow at the highest concentration of the antimicrobial are cultured and re-exposed to the antibacterial agent. This procedure is repeated and the MIC of the antimicrobial is monitored.

To test the hypothesis that ceragenins are unlikely to engender resistance, we exposed standard strains of pathogenic Gram-negative and -positive bacteria to serial passages of CSA-13 and monitored alterations in the MIC values. We performed 30 serial passages, and used ciprofloxacin, vancomycin and colistin as comparators. To understand modifications to the membrane components in response to serial exposure to these antimicrobials, the structures of lipid A from Gram-negative bacteria were monitored.

Materials and methods

Bacterial strains and reagents

Ceragenin CSA-13 was prepared as described previously.16 Ciprofloxacin, colistin and vancomycin were purchased from Sigma–Aldrich (St Louis, MO, USA). Stock solutions of CSA-13, vancomycin and colistin were prepared in deionized water. Ciprofloxacin stock solutions were prepared in 0.1 N HCl. The following bacterial strains were purchased from ATCC (Manassas, VA, USA): Staphylococcus aureus ATCC 25923, Pseudomonas aeruginosa ATCC 27853 and Acinetobacter baumannii ATCC 19606.

MIC/MBC values

The MIC/MBC values of each compound were determined in tryptic soy broth (TSB) using the microbroth dilution method described by the CLSI. To observe small changes in the MIC values, smaller increments in antimicrobial concentrations were made than typically used for standard MIC measurements. For MIC assays, bacteria were incubated at 37°C for 18–24 h with gentle shaking (∼50 rpm). The MIC values were determined versus bacterial concentrations of ∼5 × 105 cfu/mL. Bacterial quantification was performed by plating each sample on tryptic soy agar (TSA) and counting colonies. MBCs were defined as >5 log reductions in bacterial counts (i.e. below the 1 log detection limit).

Serial passage experiments

A modified serial passage protocol was developed based on previously described methods.13–15 Briefly, MIC values for wild-type strains were first determined and recorded. After incubation, serial passaging was initiated by harvesting bacterial cells growing at the highest concentration of the antimicrobial (just below the MIC) and inoculating into fresh TSB. This inoculum was subjected to another MIC assay. After an 18–24 h incubation period, cells growing in the highest concentration of the antimicrobial from the previous passage were once again harvested and assayed for the MIC. The process was repeated for 30 passages. Antimicrobial concentrations were adjusted during the process to compensate for rising MIC values. Resistant strains were maintained with mild selection on TSA containing 15 mg/L CSA-13, 5 mg/L vancomycin, 25 mg/L ciprofloxacin or 15 mg/L colistin.

Lipid A isolation and characterization

An established method was used to isolate LPS and truncate it to lipid A.17 Briefly, lyophilized bacterial cells (20 mg) were suspended in TRI reagent (500 μL) (Sigma–Aldrich). Cells were vortexed and incubated at 37°C for 20 min. Chloroform (200 μL) was added and the resulting mixture was vigorously vortexed. Cells were centrifuged at 5000 g for 10 min to separate the aqueous and organic phases. The aqueous phase was transferred to a new tube and the organic phase was re-extracted with water (3 × 300 μL) to completely remove LPS from the organic phase. All the aqueous extracts were pooled and lyophilized. LPS was further purified by adding cold, aqueous MgCl2 (1 mL of a 0.375 M solution). The resulting mixture was centrifuged at 10 000 g for 10 min to obtain semi-pure LPS as a white precipitate. To remove phospholipids, the semi-pure LPS was washed with a mixture of chloroform and methanol [200 μL of a cold 2 : 1 (v/v) mixture] and centrifuged at 10 000 g for 5 min. The precipitate was dissolved in an aqueous buffer containing SDS [1 mL of a 1% solution in 10 mM sodium acetate buffer (pH 4.5)]. The resulting mixture was heated at 100°C for 1 h. This mixture was lyophilized, washed twice with cold HCl in ethanol (0.02 N) to remove SDS and then with 95% cold ethanol, followed by drying under vacuum. The resulting material was dissolved in dichloromethane/methanol (1 : 1, v/v) and analysed via mass spectrometry (electrospray, time-of-flight analysis).

Resistance stability testing

Serial passages of resistant strains were performed without CSA-13 selection, based on methods previously described.18 Briefly, a parent culture of the resistant organism was grown up in TSB containing 10 mg/L CSA-13 for mild selection. After overnight growth under selection, the culture was diluted 1 : 1000 into fresh TSB and was grown again without selection. After each subsequent passage without selection, MIC values were determined and recorded as described previously. Passages without selection continued for 17 passages.

Observation of the impact of PagP on CSA-13 susceptibility in Escherichia coli

MIC values were determined for E. coli MC1061 (wild-type) in lysogeny broth containing 100 mg/L streptomycin and for E. coli WJ0124 (MC1061 pagP::amp) in lysogeny broth containing 100 mg/L ampicillin.19

Strain identification using PCR, 16S DNA sequencing and BLAST analysis

PCR amplification of 16S rDNA was performed using the following primers: E338for, ACT CCT ACG GGA GGC AGC AGT; and E1177rev, CGT CAT CCC CAC CTT CC.20 The PCR was performed in a Bio-Rad thermal cycler for 35 cycles using a Platinum Pfx Polymerase Kit (Invitrogen, Carlsbad, CA, USA) under the following conditions: denaturing, 95°C for 45 s; annealing, 55°C for 45 s; elongation, 68°C for 45 s; and final elongation at 68°C for 60 s. All PCR products were then confirmed by electrophoresis on a 1% agarose gel containing ethidium bromide. Reaction products were then purified using a Qiagen PCR Purification Kit (Qiagen, Valencia, CA, USA). The samples were sequenced, and the sequences were aligned and compared in a BLAST search to confirm the identity of each bacterial strain.

Results

The initial MIC values of ciprofloxacin, vancomycin and CSA-13 for S. aureus were 0.3, 1.0 and 0.3 mg/L, respectively (Table 1). After 10 serial passages the MIC of ciprofloxacin increased to >10 mg/L, and by 18 passages the MIC increased to >100 mg/L (Figure 2). The MIC of vancomycin increased slightly after a few passages, and continued to rise modestly over 30 passages to reach 9 mg/L. Slight variations in the MIC of CSA-13 were observed over 30 passages, but the MIC did not rise above 1 mg/mL. To verify that the test organism had not been contaminated by another, less susceptible organism, at the conclusion of the study the strains were verified as S. aureus using 16S rDNA PCR followed by DNA sequencing. To demonstrate that cross-resistance was not generated, the MIC values of CSA-13 were determined with the bacteria that were resistant to ciprofloxacin and vancomycin. These MIC values were largely unchanged from the initial strain (MIC values of 0.1 and 0.4 mg/L, respectively).

Table 1.

MICs for the indicated strains before [wild-type(wt)] and after 30 serial exposures to the indicated antimicrobial agents (vancR, vancomycin; csaR, CSA-13; cipR, ciprofloxacin; colR, colistin)

MIC (mg/L)
CSA-13 ciprofloxacin colistin vancomycin
S. aureuswt 0.3 0.3 1.0
S. aureusvancR 0.4 0.5 9.0
S. aureuscsaR 0.9 0.3 1.0
S. aureuscipR 0.1 >200 1.0
P. aeruginosawt 2.0 0.2 0.5 —
P. aeruginosacolR 2.0 0.2 >200 —
P. aeruginosacsaR 20 0.1 7.0 —
P. aeruginosacipR 2.0 >200 0.5 —
A. baumanniiwt 3.0 0.6 0.4 —
A. baumanniicolR 4.0 0.4 >200 —
A. baumanniicsaR 32 0.5 0.3 —
A. baumanniicipR 3.0 >250 0.4 —
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Figure 2.

Figure 2.

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MIC values for S. aureus of the designated antimicrobials after the number of serial passages indicated.

Standard strains of P. aeruginosa and A. baumannii were used in serial passaging experiments as representative of Gram-negative bacteria. Ciprofloxacin and colistin were used as comparators of CSA-13, and all three antibacterial agents were active against the parent strains (Table 1). With P. aeruginosa, resistance to ciprofloxacin was observed after <10 passages and the MIC value reached 200 mg/L after 12 passages (Figure 3). Resistance to colistin required additional passages and significant resistance was not seen until 15 passages had been completed. At that point, resistance increased significantly and reached 200 mg/L after 21 passages. A modest and steady increase in the MIC value of CSA-13 was observed over ∼20 passages for P. aeruginosa. Further changes in the susceptibility to CSA-13 were relatively small and after 30 passages the MIC value reached 20 mg/L, with an MBC of 28 mg/L. With A. baumannii, the MIC of colistin reached 200 mg/L after 19 passages (Figure 4). Though P. aeruginosa generated resistance to ciprofloxacin after relatively few passages, the generation of significant resistance by A. baumannii required >15 passages. Nevertheless, after 21 passages, the MIC of ciprofloxacin for A. baumannii reached 250 mg/L. For A. baumannii, the MIC of CSA-13 increased slightly at each passage and ultimately reached a value of just above 30 mg/L. As described for the experiments with S. aureus, after the serial passaging was completed, the strain identities were confirmed by characterizing 16S rDNA. To confirm that cross-resistance did not occur, the MIC values of CSA-13 for ciprofloxacin- and colistin-resistant strains of P. aeruginosa and A. baumannii were determined. The MICs were unchanged relative to the parent strain. In addition, the organisms that were resistant to CSA-13 were fully susceptible to ciprofloxacin.

Figure 3.

Figure 3.

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MIC values for P. aeruginosa of the designated antimicrobials after the number of serial passages indicated.

Figure 4.

Figure 4.

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MIC values for A. baumannii of the designated antimicrobials after the number of serial passages indicated.

As a mimic of AMPs, it was expected that the generation of resistance to CSA-13 among Gram-negative bacteria would involve modifications to the lipid A portion of LPS. To avoid pleiotropic effects associated with PhoPQ knockouts,11 we initially focused on AMP resistance due to lipid A palmitoylation. The production of PagP is under the control of the PhoPQ system and the additional palmitate chain incorporated into lipid A by PagP confers a degree of resistance.21,22 To determine whether the activity of PagP would impact the susceptibility of Gram-negative bacteria to CSA-13, we measured MIC values for well-characterized strains of E. coli, one with an inducible gene for PagP.19 With these strains, PagP did not impact the MIC of CSA-13; it was 1.6 mg/L for both the wild-type and PagP deletion strains.

To further understand the impact of membrane modification on the susceptibility to CSA-13, we isolated and characterized lipid A from the CSA-13-resistant strains of P. aeruginosa and A. baumannii using established methods.17 We also isolated lipid A from the parent strains of both bacteria. Mass spectrometry was used to identify modifications to lipid A. Mass signatures for hexa-acylated lipid A were found with the parent strains (data from experiments with P. aeruginosa are shown in Figure 5). In the resistant strains, 4-aminoarabinose-modified lipid A was prevalent, while it was absent (not detected) in the parent strains. These results are consistent with those observed with resistance generation by Gram-negative bacteria to AMPs.

Figure 5.

Figure 5.

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Mass spectra of lipid A from P. aeruginosa. LPS was isolated by extraction and partially hydrolysed to give lipid A. (a) Lipid A from CSA-13-resistant P. aeruginosa: *diphosphoryl lipid A, **4-aminoarabinose-appended lipid A. (b) Lipid A from CSA-13-susceptible P. aeruginosa: ***diphosphoryl lipid A.

The expectation was that resistance to CSA-13 was under adaptational control; consequently, it was anticipated that outside of the selection pressure of CSA-13, the Gram-negative organisms would revert back to susceptible forms. To test this hypothesis, serial passages outside of the presence of the antimicrobial were conducted and MIC values were measured at each reculturing of the organisms (every 24 h). After a short induction period, MIC values began to fall and within 15 passages began to approach the values measured with the fully susceptible organisms (Figure 6).

Figure 6.

Figure 6.

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Reversion of CSA-13-resistant A. baumannii and P. aeruginosa to susceptible forms after serial passages outside of the presence of CSA-13; 3 day running averages are shown.

Discussion

S. aureus, P. aeruginosa and A. baumannii were selected for these studies because they are important human pathogens. In addition, drug resistance among these organisms is prevalent and substantially limits treatment options. The emergence of resistant strains has led to studies of resistance mechanisms and these are relatively well characterized.23,24 Fluoroquinolone resistance in S. aureus, including resistance to ciprofloxacin, occurs through mutations in genes coding for gyrases25 and these mutations can occur relatively rapidly, leading to large increases in MIC values. Intermediate resistance to vancomycin in S. aureus can originate from cell wall thickening.26 Mechanisms of resistance to AMPs in Gram-positive organisms include the export of proteases and modifications to lipid head groups of membrane phospholipids.4,10 Because CSA-13 is not peptide based, it is not degraded by proteases and, apparently, if any modifications to the membrane lipids occurred, they had minimal impact on the antibacterial activities of CSA-13.

Mutations in gyrase genes also lead to fluoroquinolone resistance in Gram-negative bacteria,27 and few passages led to ciprofloxacin resistance in both S. aureus and P. aeruginosa; however, resistance generation was delayed with A. baumannii. Colistin resistance in Gram-negative bacteria is mediated by a variety of mechanisms, including modifications to the membrane structure and membrane permeability.25,28–30 Membrane modifications are controlled by the PmrA-PmrB two-component system, which leads to lipid modifications that are associated with the generation of resistance to AMPs in general. It was surprising, therefore, that the generation of resistance to colistin and CSA-13 followed very different paths. Nevertheless, highly colistin-resistant (MICs ≥256 mg/L) A. baumannii strains have been shown to be susceptible to AMPs,31 and, similarly, colistin-resistant A. baumannii and P. aeruginosa strains were susceptible to CSA-13. The MIC of colistin for the CSA-13-resistant strain was somewhat elevated as compared with the parent strain for P. aeruginosa (Table 1). These results argue that different mechanisms of resistance were generated in response to CSA-13 and colistin, with CSA-13 resistance impacting colistin activity, but not the inverse. The polymyxins, including colistin, are only weakly active against Gram-positive bacteria, in contrast to AMPs, which in general are active against Gram-negative and -positive organisms. A mechanism of action of the polymyxins has been proposed that is distinct from that of AMPs in general.7 Consequently, the differences in the mechanisms of action of colistin and CSA-13 may have led to the generation of the different mechanisms of resistance.

In earlier work, similarities in the antibacterial activities of AMPs and ceragenins, including CSA-13, have been demonstrated.32–34 These similarities include affinity for LPS, the ability to depolarize the cytoplasmic membranes of Gram-negative and -positive organisms, and blebbing of the outer membranes of Gram-negative bacteria in the presence of sublethal concentrations of the antimicrobials. The addition of 4-aminoarabinose to lipid A in response to exposure to CSA-13 suggests that bacteria respond to ceragenins through the same adaptational mechanisms employed when in the presence of AMPs. However, the conditions necessary to permit these adaptational mechanisms to deploy have to be carefully controlled. As with AMPs, the MIC values of CSA-13 are similar to the corresponding MBC values. This is particularly true with Gram-negative bacteria, e.g. the MIC values of CSA-13 for the parent strains of P. aeruginosa and A. baumannii are 2 and 3 mg/L, while the MBC values are 4 and 4 mg/L, respectively. Consequently, to achieve the pressure to activate the adaptational modification to lipid A, concentrations of CSA-13 had to be carefully controlled: if the concentrations were too low, insufficient pressure was placed on the organisms; and if the concentrations were too high, the organisms were eradicated. In potential therapeutic applications of ceragenins, it is unlikely that this level of control would be achieved.

The generation of bacterial resistance can be associated with metabolic costs. For example, colistin resistance in A. baumannii leads to differences in the expression of 35 different proteins and a slowing of growth.35 The metabolic costs of generating 4-aminoarabinose and appending it on lipid A probably contribute to the reversion of the resistant bacteria to susceptible forms outside of the pressure of CSA-13.

Conclusions

The ubiquity of AMPs argues that they are capable of controlling bacterial growth in many environments without readily engendering resistance. Nevertheless, bacteria are not defenceless; adaptive responses can decrease susceptibility to AMPs. Although CSA-13 appears to mimic the antibacterial activity of AMPs, because it is not peptide based, it is not degraded by proteases released by bacteria and its MIC value for S. aureus remains below 1 mg/L after 30 serial passages. Lipid A modifications impact the susceptibility of Gram-negative bacteria to AMPs and CSA-13. However, resistance generated by P. aeruginosa and A. baumannii to CSA-13 was modest in comparison with that generated to ciprofloxacin and colistin by these organisms. These results demonstrate similarities in the responses of bacteria to AMPs and CSA-13. Considering the interest in the use of AMPs clinically to treat a variety of infections, CSA-13 appears to be a possible alternative for use in augmenting and replacing the antibacterial activities of endogenous AMPs.

Funding

This work was supported by the National Institute of Allergy and Infectious Diseases [U01 AI082209-01 (P. B. S.)] and the Canadian Institutes of Health Research [Operating Grant MOP-84329 (R. E. B.)].

Transparency declarations

P. B. S. is a consultant for N8 Biomedical; other authors have no conflicts to declare.

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