Biological Cost of Different Mechanisms of Colistin Resistance and Their Impact on Virulence in Acinetobacter baumannii Alejandro Beceiro,a Antonio Moreno,a,b Nathalie Fernández,a Juán A. Vallejo,a Jesús Aranda,a,b,c Ben Adler,d,e Marina Harper,d,e John D. Boyce,d Germán Boua ‹Servicio de Microbiología-INIBIC, Complejo Hospitalario Universitario A Coruña (CHUAC), A Coruña, Spaina; Servicio de Microbiología, Complejo Hospitalario Pontevedra, Pontevedra, Spainb; Departament de Genètica i Microbiologia, Facultat de Biociències, Universitat Autònoma de Barcelona (UAB), Cerdanyola del Vallès, Barcelona, Spainc; Department of Microbiology, Monash University, Melbourne, Victoria, Australiad; Australian Research Council Centre of Excellence in Structural and Functional Microbial Genomics, Monash University, Melbourne, Victoria, Australiae

A

cinetobacter baumannii is a Gram-negative bacterium that has emerged in recent decades as an important hospital pathogen. A. baumannii infections are especially problematic in intensive care, trauma, and burn units and result in a range of infectious syndromes (1). A. baumannii strains are intrinsically resistant to desiccation and disinfectants and have shown a great capacity to acquire or develop antibiotic resistance (2, 3). Many strains are now resistant to almost all antibiotics classes and are designated multidrug resistant (MDR). One of the last-line treatment options with activity against MDR strains is the polymyxins, especially colistin (polymyxin E) (3, 4). Previously used in the 1960s and 1970s, the polymyxins were largely abandoned due to their perceived toxicity; however, they have recently been reintroduced for human therapy with new and revised dosing regimens (5, 6). However, pandrug-resistant isolates, including those resistant to colistin, have already been observed (4, 7, 8). Infections with these strains are potentially untreatable. The recent emergence of pandrug resistance underscores the urgency of understanding the implications that multi- and pandrug resistance have on the fitness and virulence of this species. Polymyxins are polycationic antimicrobial peptides with activity primarily against Gram-negative bacteria. The polyanionic bacterial lipopolysaccharide (LPS) is the initial target, and the amphiphilic nature of polymyxin is critical for its interaction with the hydrophobic lipid A component of LPS (9). Two different colistin resistance mechanisms have been characterized in A. baumannii (10). One involves total inactivation of the lipid A biosynthetic pathway and complete loss of surface LPS. Different molecular events, such as deletions, point mutations, or insertions, can inactivate any of the first three genes (lpxA, lpxC, and lpxD) in the

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lipid A biosynthetic pathway (11–13). The total loss of LPS in strains with this resistance mechanism prevents the essential interaction between it and colistin, giving rise to very high colistin MICs. The second mechanism, first suggested by Adams et al. (14) and then studied in depth by Beceiro et al. (15) and Arroyo et al. (16), is mediated by the PmrAB two-component system. It has been shown that mutations in pmrA or pmrB and/or increased expression of these genes leads to the addition of phosphoethanolamine (PEtn) to the hepta-acylated form of lipid A, due to the activity of the PEtn transferase pmrC gene located upstream of pmrAB. These lipid A modifications decrease the negative charge of LPS and occur in both laboratory mutants and clinical isolates of A. baumannii (15, 16). Similarly, colistin resistance due to LPS inactivation has also been identified in laboratory mutants and recent clinical isolates (11, 12). Little is known about the effects that acquisition of these different resistance mechanisms has on the fitness and virulence of A. baumannii. Given that colistin is one of the most important antimicrobial options against MDR A. baumannii, and, more significantly, considering that tigecycline has shown no better efficacy

Antimicrobial Agents and Chemotherapy

Received 23 July 2013 Returned for modification 16 August 2013 Accepted 29 October 2013 Published ahead of print 4 November 2013 Address correspondence to Alejandro Beceiro, [email protected], or Germán Bou, [email protected]. Copyright © 2014, American Society for Microbiology. All Rights Reserved. doi:10.1128/AAC.01597-13

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Two mechanisms of resistance to colistin have been described in Acinetobacter baumannii. One involves complete loss of lipopolysaccharide (LPS), resulting from mutations in lpxA, lpxC, or lpxD, and the second is associated with phosphoethanolamine addition to LPS, mediated through mutations in pmrAB. In order to assess the clinical impacts of both resistance mechanisms, A. baumannii ATCC 19606 and its isogenic derivatives, AL1851 ⌬lpxA, AL1852 ⌬lpxD, AL1842 ⌬lpxC, and ATCC 19606 pmrB, were analyzed for in vitro growth rate, in vitro and in vivo competitive growth, infection of A549 respiratory alveolar epithelial cells, virulence in the Caenorhabditis elegans model, and virulence in a systemic mouse infection model. The in vitro growth rate of the lpx mutants was clearly diminished; furthermore, in vitro and in vivo competitive-growth experiments revealed a reduction in fitness for both mutant types. Infection of A549 cells with ATCC 19606 or the pmrB mutant resulted in greater loss of viability than with lpx mutants. Finally, the lpx mutants were highly attenuated in both the C. elegans and mouse infection models, while the pmrB mutant was attenuated only in the C. elegans model. In summary, while colistin resistance in A. baumannii confers a clear selective advantage in the presence of colistin treatment, it causes a noticeable cost in terms of overall fitness and virulence, with a more striking reduction associated with LPS loss than with phosphoethanolamine addition. Therefore, we hypothesize that colistin resistance mediated by changes in pmrAB will be more likely to arise in clinical settings in patients treated with colistin.

Biological Cost of A. baumannii Colistin Resistance

TABLE 1 Bacterial strains used in this study Strain

Description

Colistin MIC (␮g/ml)

ATCC 19606

A. baumannii type strain; parent of AL1842 ⌬lpxC, AL1851 ⌬lpxA, and AL1852 ⌬lpxD

1

AL1851 ⌬lpxA AL1852 ⌬lpxD

Derivate of ATCC 19606; colistin resistant; 445-bp deletion within lpxA Derivate of ATCC 19606; colistin resistant; single base deletion at nucleotide [nt] 364 of lpxD and frame shift after K317 Derivate of ATCC 19606; colistin resistant; 84-bp deletion within lpxC Derivate of ATCC 19606; colistin resistant; single amino acid substitution (Ala227Val) in PmrB. Ala227Val is adjacent to the conserved histidine at the site of phosphorylation (His228). A. baumannii clinical strain carrying the carbapenemase OXA-24, isolated in a Spanish nosocomial outbreak in 1997 Derivate of A. baumannii ABRIM; colistin resistant; single amino acid substitution (Asn353Tyr) in PmrB. This substitution is inside the ATP binding site and may have an effect on the phosphorylation of His228 and thereby on the phosphorylation levels of PmrA. E. coli strain used for maintenance of C. elegans

64 ⬎128

American Type Culture Collection 12 12

128 64

12 15

1

21

32

15

NAa

24

AL1842 ⌬lpxC ATCC 19606 pmrB

ABRIM pmrB

OP50 a

NA, not applicable.

than commonly used antibiotics against MDR Gram-negative bacteria in general (17), it is critical that we understand the effect of resistance on fitness. López-Rojas et al. (18) have studied the effects of pmrAB mutations and PEtn addition to lipid A and have shown that these genetic and structural changes carry a biological cost. Subsequently, the clinical significance of colistin resistance in different isolates that developed in vivo has been discussed by López-Rojas et al. (19) and Rolain et al. (20), with some strains showing differences in their abilities to cause infection and showing changes on the two-component system pmrAB. To our knowledge, there are no studies comparing the impacts on fitness of the two different colistin resistance mechanisms that have been described in A. baumannii. Here, we analyze, by means of in vitro and in vivo models, the biological cost of colistin resistance developing via each of the two mechanisms. We have determined how these changes affect both overall fitness and virulence in order to assess the importance that these resistance mechanisms may have in the clinic in the near future. Importantly, we show that there is significant impact on virulence and fitness after acquisition of colistin resistance, and this biological cost is significantly more pronounced when the pathogen lacks LPS rather than when it produces PEtn-modified LPS. MATERIALS AND METHODS Bacterial strains, media, and susceptibility testing. The A. baumannii strains used in this study are listed in Table 1. The colistin-resistant (Col-R) A. baumannii lpx mutants derived from ATCC 19606 (AL1851 ⌬lpxA, AL1852 ⌬lpxD, and AL1842 ⌬lpxC) were originally isolated by Moffatt et al. (12), while the ATCC 19606 pmrB mutant was isolated by Beceiro et al. (15). In this study, we compared the differences in fitness and virulence between the wild-type (WT) strain ATCC 19606 and each of the lpx or pmrB mutants. Thus, although the strain background of all the mutants is ATCC 19606, two parental strains of ATCC 19606 were used to obtain the colistin-resistant mutants in two different laboratories. In order to rule out any possible differences in fitness and virulence of the two WT strains with different origins, assays were performed with both ATCC 19606 WT strains, and all results were similar (data not shown). Also, a multiresistant A. baumannii clinical strain (ABRIM) involved in a hospital outbreak (21) and its colistin-resistant counterpart with point modifica-

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tions in the pmrB gene (15) were included in virulence studies using Caenorhabditis elegans. The strains were cultured at 37°C in Luria-Bertani (LB) broth or on LB medium containing 1.5% agar. When necessary, the medium was supplemented with colistin at 10 mg/liter. The bacterial strains were frozen in LB-glycerol (20%) and were maintained at ⫺80°C. Colistin susceptibility testing was performed with microdilution and agar dilution, following the CLSI and BSAC criteria, respectively (22, 23). The Escherichia coli strain OP50 was used in the C. elegans assays for maintenance of the worms and was also used as a nonvirulent control strain (24). In the C. elegans assays, the worms were placed on freshly prepared nematode growth (NG) medium plates on which E. coli OP50 had been spread as a source of food. Growth curve assay. In vitro fitness was assessed under noncompetitive conditions by measuring the growth rates of all the strains and mutants described in Table 1 independently. Briefly, 50 ml of LB medium in 250-ml Erlenmeyer flasks was inoculated with approximately 0.5 ⫻ 108 CFU of each strain in exponential growth phase and incubated at 37°C for 20 h with constant shaking at 180 rpm. The bacterial concentration (expressed in log10 CFU per milliliter) was determined at 0, 1, 2, 3, 5, 7, and 20 h by spreading serial 10-fold dilutions onto LB agar plates. Four independent experiments were performed for each strain, and the results were compared using Student’s t test. The growth rate constant (␮) was calculated on the basis of the exponential segment of the growth curve and defined as ln2/g, where g is the doubling time or mean generation time (25). The Graphpad Prism 6.01 (La Jolla, CA, USA) statistical package was used to statistically analyze the results of all experiments. In vitro competition experiments. In order to measure the relative fitness of each colistin-resistant mutant compared with the parental strain, an in vitro competitive index (CI) was determined. Exponentially growing cells of each mutant and the wild-type strain were mixed in a 1:1 proportion (2.5 ⫻ 107 CFU of each strain) and grown in 50 ml of LB broth at 37°C at 180 rpm for 20 h. The number of cells corresponding to each strain of the pair was determined at 5 and 20 h by spreading serial 10-fold dilutions onto LB agar plates with or without 10 mg/liter of colistin. The number of cells of each strain in the input sample was also determined by direct plating onto medium with and without colistin. The CI was defined as the ratio of the numbers of CFU of the resistant mutant and the parental strain. Four independent competition experiments were performed to calculate the median values of each CI. Statistical analyses were performed

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ABRIM

Source or reference

Beceiro et al.

with Student’s t test, and differences were considered statistically significant at a P value of ⬍0.05. LIVE/DEAD fluorescence microscopy on infected A549 monolayers. A549 human alveolar epithelial cells were cultured in Dulbecco modified Eagle medium (DMEM) supplemented with 10% fetal bovine serum, 100 mg/liter of penicillin, and 100 mg/liter streptomycin at 37°C in the presence of 10% CO2. For cell viability assays, monolayers of A549 cells were cultured in 24-well plates to a density of 1 ⫻ 105 cells per well. Later, the cells were infected with 2 ⫻ 107 CFU/well of A. baumannii in new DMEM without antibiotics and grown for 20 h at 37°C (28, 29). The number of inoculated bacteria was determined by direct plating. A LIVE/ DEAD fluorescence microscopy kit (Cellstain double-staining kit; Fluka, Buchs, Switzerland) was used according to the manufacturer’s instructions to measure cell viability postinfection. Briefly, at 20 h postinfection, the A549 cells were incubated for 15 min at 37°C with phosphate-buffered saline (PBS) containing a mixture of the two fluorescent molecules to obtain simultaneous fluorescent staining; calcein-AM, is able to stain viable cells (green), while propidium iodide, a nucleus-staining dye that is unable to penetrate the cell membranes of viable cells, can stain only dead cells (red). Microscopic images of the stained cells (alive and dead) were obtained using an inverted fluorescence microscope (Nikon Eclipse Ti) and analyzed with the NIS Elements Br software package. The excitation absorbance of calcein-AM was 490 nm, and emission was at 515 nm, while the excitation range of propidium iodide was 535 nm and emission was 617 nm. At least six replicates of each assay were analyzed, and the statistical significance was determined using Student’s t test. Fitness in the systemic murine model. Female BALB/c mice 5 to 8 weeks of age and weighing approximately 20 g were housed under specificpathogen-free conditions. Antibiotic-free pelleted food and autoclaved water were provided ad libitum. All experiments involving mice were approved by the Animal Ethics Committee of the University Hospital Complex A Coruña. Competitive-growth assays were performed to measure the in vivo fitness of each mutant in a murine systemic-infection model. Each of the mutant strains was mixed with the parental strain, ATCC 19606, at a ratio of 1:1, and approximately 3.0 ⫻ 108 CFU in 250 ␮l of PBS was inoculated intraperitoneally into groups of 5 mice (30, 31). As previously described for the in vitro competitive-growth assays, the concentration of each strain in the input culture was determined by direct plate counting. The mice were euthanized with an overdose of thiopental sodium 20 h after inoculation. Livers were aseptically extracted, weighed, and homogenized in 1.5 ml of ice-cold LB medium in a Mixer Mill dismembrator (Retsch, Germany). The number of CFU of each strain and the CI were calculated as described for in vitro competitive-growth experiments. Statistical analysis was also performed as described for the in vitro experiments. Virulence assays in the mouse systemic-infection model. The ATCC 19606 lpxC and pmrB mutant strains were analyzed for virulence using the previously described mouse systemic-infection model (26), in

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terms of mortality and lethal dose, as well as by measuring the survival time of the infected mice. Groups of BALB/c mice were inoculated with the ATCC 19606 WT strain or the AL1842 ⌬lpxC or pmrB mutant strain and monitored carefully for signs of disease for 7 days. To calculate 50% lethal doses (LD50), groups of mice were infected by intraperitoneal injection with 250 ␮l of the different bacterial suspensions of ATCC 19606 WT (1 ⫻107, 4 ⫻107, 12 ⫻107, and 40 ⫻107 CFU/ml), the ATCC 19606 pmrB mutant (1 ⫻107, 3 ⫻107, 12 ⫻107, and 40 ⫻ 107 CFU/ml), and AL1842 ⌬lpxC (3 ⫻107, 12 ⫻107, 25 ⫻107, 40 ⫻107, and 80 ⫻ 107 CFU/ml). Indeed, a second assay was performed to measure the survival time; groups of 10 BALB/c mice were inoculated with bacterial suspensions containing 3 ⫻ 108 CFU/ml of the 19606 ATCC WT strain or the AL1842 ⌬lpxC or pmrB mutant. Any animals that survived to 7 days were euthanized with thiopental. Survival data were compared using the log rank test, with a P value of ⬍0.05 considered statistically significant (26, 27). Virulence studies in a C. elegans model. Previous work had shown that for Pseudomonas aeruginosa and A. baumannii, the maturity and quantity of C. elegans broods is inversely related to the virulence of the infecting strain (32–34). Therefore, we used C. elegans as a second infection model. Briefly, the E. coli strain OP50 was grown on 5.5-cm NG medium plates at 37°C for 24 h, and these plates were used for growth and maintenance of the worms (35). Each of the A. baumannii strains, ATCC 19606, ABRIM, and their colistin-resistant mutants, was seeded from overnight cultures in NG medium and grown at 37°C for 24 h. The eggs of C. elegans N2 Bristol were hatched in M9 medium, and worms in the first larval stage (L1) were arrested at 24 h by transfer to 20°C. Later, the L1 worms were added to the NG medium plates together with each A. baumannii strain to study. E. coli OP50 was also used as a nonvirulent control strain. Later, one C. elegans worm in the last larval stage (L4) was placed over each A. baumannii strain and incubated for 24 h at 25°C on a new NG medium plate. Daily, the worms were placed in a new plate seeded with the same strain. The worm progeny and eggs were counted every day for 5 days to determine their viability. A Nikon SMZ-745 dissecting microscope was used for the visual examination. Six independent replicates were performed with each strain (32, 34).

RESULTS

Effects of different colistin resistance mutations on bacterial growth and fitness. To determine whether the different LPS changes associated with each colistin resistance mechanism affect the bacterial growth rate, growth curves were performed for all the A. baumannii strains studied (Table 1 and Fig. 1). The rate of increase in bacterial cells is equal to the growth rate constant (␮) times the number of cells; therefore, determination of ␮ gives a measure of fitness or replication capacity (36). The growth rates of the AL1851 ⌬lpxA, AL1852 ⌬lpxD, and AL1842 ⌬lpxC mutants

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FIG 1 In vitro growth of ATCC 19606 and the colistin-resistant laboratory derivates. The numbers of CFU per milliliter were determined at 1, 2, 3, 5, 7, and 20 h. The means of four independent replicates (for each growth curve) are shown. The error bars represent the standard deviation (SD).

Biological Cost of A. baumannii Colistin Resistance

TABLE 2 A. baumannii doubling times and growth rates measured in the exponential phase of growth over the first 300 mina Strain

Doubling time (g) (min)

Growth rate (␮) (h⫺1)

ATCC 19606 AL1851 ⌬lpxA AL1842 ⌬lpxC AL1852 ⌬lpxD ATCC 19606 pmrB

27 ⫾ 3 49 ⫾ 6 40 ⫾ 3 83 ⫾ 8 32 ⫾ 1

1.56 ⫾ 0.27 0.85 ⫾ 0.09 1.03 ⫾ 0.09 0.49 ⫾ 0.03 1.32 ⫾ 0.03

a

Data were extracted from Fig. 1.

FIG 2 Relative in vitro competition indexes of colistin-resistant mutants. The CI values were calculated as the number of Col-R mutants divided by the number of bacteria of the Col-S parental strain of A. baumannii, and the median CI values are shown in parentheses. Each circle represents the CI obtained in each in vitro replicate. Experiments were performed at 5 and 20 h. In all cases, the differences in bacterial counts were statistically significant (P ⬍ 0.01). The error bars represent the standard deviation.

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FIG 3 Relative in vivo competition indexes determined in a mouse systemicinfection model over 20 h. The CI values were calculated as the number of Col-R mutants divided by the number of bacteria of the Col-S parental strain of A. baumannii, and the median CI values are shown in parentheses. Circles represent the CI obtained in each in vivo replicate. The differences in bacterial counts were statistically significant (P ⬍ 0.01). The error bars represent the standard deviation.

respectively), but the relative growth rate compared with the wildtype strain was clearly higher than those of the LPS-deficient strains. Competitive growth rates were also determined in vivo using the systemic BALB/c mouse model. Bacteria recovered from the livers of infected mice were used to calculate the CI of the colistin-resistant mutants compared to the wild-type control. The in vivo growth rates were similar to those measured in vitro at 20 h (Fig. 3). All the Col-R mutants showed a significant reduction in fitness, with a more marked effect observed for the ⌬lpxA, ⌬lpxD, and ⌬lpxC mutants (CI ⬍ 0.01, CI ⫽ 0.02, and CI ⫽ 0.09) than for the pmrB mutant (CI ⫽ 0.35). Effect on virulence of both colistin resistance mechanisms. (i) Infection of A549 human alveolar cells. The capacity of each of the A. baumannii strains to reduce the viability of A549 respiratory alveolar cells was assessed using LIVE/DEAD staining and fluorescence microscopy. Following incubation of each of the A. baumannii strains with the A549 cells, the numbers of live (green) and dead (red) A549 respiratory cells were determined (Fig. 4A). Coincubation of the ATCC 19606 WT strain with the A549 cells resulted in a 65% decrease in A549 cell viability. Coincubation of the A549 cells with the A. baumannii pmrB mutant resulted in a similar 57% reduction in viability, but incubation with each of the lpx mutants gave only a slight reduction in A549 cell viability (Fig. 4B). Thus, the lpx mutants showed highly reduced ability to kill A549 respiratory cells compared to both the wild-type and pmrB mutant strains (P ⬍ 0.01). Once again, the AL1852 ⌬lpxD mutant presented a higher cost, showing the least killing of the A549 alveolar cells (4%), approximately one-third that of both the ⌬lpxC and ⌬lpxA mutants (12 and 14%, respectively) (Fig. 4B). (ii) Virulence in a mouse model of systemic infection. The virulence of the wild-type strain and the AL1842 ⌬lpxC and pmrB colistin-resistant mutants was assessed in a mouse systemic-infection model. Mortality with the ATCC 19606 WT strain at inocula of 1 ⫻ 107, 4 ⫻ 107, 12 ⫻ 107, and 40 ⫻ 107 CFU/ml was 0, 0, 80, and 100%, respectively; with the ATCC 19606 pmrB mutant, the mortality at inocula of 1 ⫻ 107, 3 ⫻ 107, 12 ⫻ 107, and 40 ⫻ 107 CFU/ml was 0, 0, 80, and 100%, respectively; and finally, with the

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measured during the exponential growth phase (␮ ⫽ 0.85 ⫾ 0.09, 1.03 ⫾ 0.09, and 0.49 ⫾ 0.03, respectively) were significantly lower than that of the ATCC 19606 WT (␮ ⫽ 1.56 ⫾ 0.27) (Table 2 and Fig. 1). Indeed, after 20 h, the number of cells in the LPS-deficient mutant cultures was approximately 1 log unit lower than for the parental strain. Similarly, the pmrB mutant also failed to reach the same cell numbers as the WT strain; however, the difference was approximately 0.5 log unit. However, the growth rate of the pmrB mutant strain in exponential growth phase (␮ ⫽ 1.32 ⫾ 0.03) was close to that of the parent strain and significantly higher than for each of the lpx mutant strains. In vitro competition experiments were also performed to determine the relative growth rates of each of the strains, and data are presented as the CI (number of Col-R mutants/number of Col-S parental strain). In all cases, colistin resistance was associated with a significant fitness cost in vitro (Fig. 2). Measurements were performed at 5 and 20 h, and the results were similar at each time point. After 5 h, the median CIs of AL1851 ⌬lpxA, AL1852 ⌬lpxD, and AL1842 ⌬lpxC relative to the WT strain were 0.14, 0.01, and 0.7, respectively, and they were even lower after 20 h growth (CI ⫽ 0.02, CI ⬍ 0.01, and CI ⬍ 0.01, respectively). As was observed with the growth rate experiments, the AL1852 ⌬lpxD mutant showed the highest fitness cost. The pmrB mutant also displayed a significantly reduced CI (0.22 and 0.26 at 5 and 20 h,

Beceiro et al.

infected with each of the A. baumannii strains and stained with the LIVE/DEAD Cellstain double-staining kit. Healthy cells with intact membranes are stained green, and dead cells with permeabilized membranes are stained red. A549 cells were incubated with the A. baumannii strains ATCC 19606 WT, AL1851 ⌬lpxA, Al1852 ⌬lpxD, AL1842 ⌬lpxC, and ATCC 19606 pmrB for 20 h or left uninfected. (B) Quantification of A549 cell death caused by A. baumannii ATCC 19606 WT and AL1851 ⌬lpxA, AL1852 ⌬lpxD, and AL1842 ⌬lpxC mutants and the pmrB mutant. The results of 6 independent experiments are shown as means and SD. *, P ⬍ 0.01 between the parental strain and each of the designated mutants; **, not statistically different.

AL1842 ⌬lpxC mutant, the mortality at inocula of 3 ⫻ 107, 12 ⫻ 107, 25 ⫻ 107, 40 ⫻ 107, and 80 ⫻ 107 CFU/ml was 0, 0, 0, 20, and 60%, respectively. With these results, the LD50 were 10 ⫻ 107 CFU/ml for the ATCC 19606 strain, 8.8 ⫻ 107 CFU/ml for the ATCC 19606 pmrB mutant, and 63.5 ⫻ 107 CFU/ml for the AL1842 ⌬lpxC mutant, with the last showing a remarkably higher LD50. A second assay was performed to measure survival rates (Fig. 5). Seven of 10 mice infected with 3 ⫻ 108 CFU/ml of the wild-type strain died within the first 48 h, and 8 of 10 mice infected with the pmrB mutant strain also died in the first 48 h. Therefore, the pmrB mutation does not affect virulence in the mouse systemic model. Conversely, only one mouse infected with the AL1842 ⌬lpxC mutant died during the 7-day course of the experiment, indicating that the strain is highly attenuated for virulence in this model (P ⫽ 0.02). The AL1842 ⌬lpxC mutant was selected for

FIG 5 Survival of BALB/c (n ⫽ 10 per group) mice following intraperitoneal

infection with 3 ⫻ 108 CFU of the ATCC 19606, AL1842 ⌬lpxC, or ATCC 19606 pmrB strain. No difference in survival was found between the mice infected with the pmrB mutant or the parental strain, but survival was significantly higher in mice infected with the AL1842 ⌬lpxC mutant (P ⬍ 0.01).

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testing, as the previous growth experiments had shown that the mutant was similar in fitness to the AL1851 ⌬lpxA mutant and had a less marked fitness defect than the AL1852 ⌬lpxD mutant. These data clearly show that the production of LPS is critical for full virulence, at least in this mouse infection model, whereas the addition of phosphoethanolamine to lipid A is not as relevant to mouse mortality. (iii) Inhibition of reproduction in C. elegans. The invertebrate C. elegans model (34, 37) was used as a second measure of bacterial virulence, with modifications. Instead of measuring C. elegans death, we analyzed inhibition of proliferation in infected worms, similar to the method previously described for P. aeruginosa (37). Worms infected with the wild-type ATCC 19606 A. baumannii strain showed decreased fertility of 20.5% compared with worms cultured only in the presence of E. coli OP50 (Fig. 6A and Table 3); however, all the ⌬lpx mutants showed decreased virulence and were even less virulent than the E. coli OP50 control. The progeny production in the presence of the ⌬lpxA, ⌬lpxD, and ⌬lpxC mutants was 50.2, 58.9, and 36.8% higher than that with the wild-type A. baumannii strain. The ATCC 19606 pmrB mutant strain reduced C. elegans brood size slightly more than the WT strain, with 5.8% decreased fertility, although the difference was not statistically significant (P ⬍ 0.05). All ⌬lpx mutants showed significant differences compared with the WT strain (P ⬍ 0.01). The other isogenic colistin-susceptible (Col-S)/colistin-resistant pair of A. baumannii strains included in the assay, ABRIM and ABRIM pmrB, showed no difference in fertility rates of C. elegans, with only 1% higher progeny production in the presence of the ABRIM pmrB strain (Fig. 6B and Table 3). In neither assay was a decrease in virulence with C. elegans worms observed when A. baumannii strains developed colistin resistance due to changes in pmrB.

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FIG 4 Abilities of different A. baumannii strains to cause cell death of A549 alveolar cells. (A) Fluorescence microscopy images of human alveolar A549 cells

Biological Cost of A. baumannii Colistin Resistance

DISCUSSION

To date, two mechanisms of colistin resistance have been characterized in A. baumannii, one involving complete LPS loss and the other involving modification of LPS by addition of PEtn. While

TABLE 3 C. elegans fertility assay Quantification (per day) on daya: Strain

1

2

3

4

Total no. of progenya

Difference from controlb (%)

A. baumannii ATCC 19606 AL1851 ⌬lpxA AL1852 ⌬lpxD AL1842 ⌬lpxC ATCC 19606 pmrB ABRIM ABRIM pmrB

45 70.2 73.7 62.5 46.33 37.3 38.8

99.3 158.7 202.7 147.2 92 83.7 75

47.7 87.3 61.3 72.3 37.2 76.5 75.3

2.33 0.7 0.7 2.3 9.7 2.8 13.5

194.2 ⫾ 6.9 316.9 ⫾ 12 338.3 ⫾ 26 284.3 ⫾ 50 180.2 ⫾ 22 200.3 ⫾ 13 202.7 ⫾ 30.3

79.5 129.7 138.4 116.3 73.7 81.9 82.9

E. coli OP50

38.6

122.4

73.2

10.2

244.4 ⫾ 28

100

a

Mean quantification and total no. of progeny by worm. b Percentage of progeny compared to E. coli OP50.

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FIG 6 C. elegans fertility assays. The number of progeny from L4 stage worms was monitored for 5 days in the presence of the nonvirulent control E. coli OP50, the A. baumannii ATCC 19606 WT strain and its colistin-resistant mutants (A), or A. baumannii ABRIM and ABRIM pmrB (B). The error bars represent the standard deviation.

Moffat et al. (12) characterized the development of colistin resistance in A. baumannii as due to LPS loss in mutants derived from the ATCC 19606 type strain and recent clinical isolates, analysis of the current literature suggests that colistin resistance resulting from pmrAB mutations is more common, based on literature searches. The complete loss of LPS production can result from mutations in any of the genes lpxA, lpxC, and lpxD. Indeed, point mutations and deletions in these genes have been observed, and insertion sequence (IS) elements, such as ISX03 and ISAba11, either intrinsic to the A. baumannii genome or from other species, can insert into the lpx genes and inactivate them (11). However, to date, few studies have described colistin resistance by LPS loss in clinical isolates of A. baumannii. One such study, published by Park et al., described a colistin-resistant clinical isolate from South Korea that was shown to be resistant via inhibition of biosynthesis of lipid A (12, 38). The second mechanism of colistin resistance, resulting from the addition of phosphoethanolamine to lipid A, has been described a number of times in clinical isolates. Rolain et al. (7) analyzed the molecular basis of antimicrobial resistance in isogenic isolates of A. baumannii and characterized colistin-resistant isolates that developed during failed therapy with colistin and rifampin; the PmrAB system was implicated in the in vivo development of the resistance. Similarly, in the same study that characterized colistin-resistant pmrB mutants in ATCC 19606, Beceiro et al. also identified clinical isolates with pmrB mutations (15). In another study, Yoon et al. (39) analyzed 14 clinical isolates that developed colistin resistance during failed treatment, and all were observed to have mutations in pmrA or pmrB. These mutants showed fitness decreases of between 10 and 18%, indicating that the fitness or virulence cost due to changes in the PmrAB twocomponent system is likely variable and dependent on the specific location, type, and number of mutations (39, 40). Recently, bacterial whole-genome sequencing of clinical isolates recovered from patients following colistin treatment again showed the involvement of the pmr locus. Other studies have also identified clinical isolates carrying this mechanism of resistance (16). In this study, we have shown that the growth rate of the wildtype strain ATCC 19606 was slightly higher than that of the colistin-resistant pmrB mutant but clearly higher with the inactivation of lpx genes. Both types of colistin-resistant mutants showed reduced relative growth rates in vitro and in vivo, with the lpx mutants showing a more marked reduction. Incubation of each of the

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virulences of the strains in the mouse model, with differences in the LD50 and LD100 of approximately 1 log unit. The Col-R isolate studied, despite the genetic changes suffered, was able to develop meningitis in a patient. In our study, while the colistin-resistant pmrB mutant used showed reduced fitness with a decreased relative growth rate both in vitro and in vivo, we observed no difference in the virulence of the strain in a mouse systemic-infection model or in the C. elegans fertility model. Also, no differences in virulence were observed with the clinical isolate ABRIM and its colistin-resistant counterpart in the C. elegans model. In conclusion, the in vitro and in vivo assays performed show that the changes implicated in development of colistin resistance by pmrAB mutations produce a fitness loss greater than the virulence loss. However, this biological cost seems not to be enough to prevent the emergence of in vivo colistin-resistant clinical isolates by this mechanism. The mechanisms underlying selection of the two different types of colistin resistance mutations have not been analyzed in detail, but it is possible that the different selection regimes used to obtain the mutants play a key role in the specific mutations selected. The lpx mutants have generally been obtained by selection in one single passage on Mueller-Hinton agar plates with 10 mg/ liter of colistin sulfate, and the mutants obtained have very high colistin MICs (ⱖ128 mg/liter) (11, 12). Conversely, colistin-resistant strains with pmrAB mutations have generally been obtained by multiple passages in increasing concentrations of colistin and give rise to strains with lower colistin MICs. For example, strains selected by passage in LB broth with 2 to 8 mg/liter of colistin give resistant mutants with MICs of 16 to 64 mg/liter (15), and strains selected on colistin gradient LB agar plates containing 4 to 50 mg/liter colistin give resistant mutants with MICs of 2 to 8 mg/ liter (16). Another important difference between these two resistance mechanisms is that resistant strains resulting from pmrAB mutation and PEtn modification of lipid A do not show altered susceptibility to other antimicrobials (44), while the LPS-deficient mutants are highly susceptible to other antibiotics, such as azithromycin, cefepime, and teicoplanin (12). This is likely due to the altered membrane permeability of these strains and is consistent with the high fitness cost of the resistance. Therefore, we propose that acquisition of high-level colistin resistance in a single step requires more drastic genetic and structural changes with greater effects on fitness and virulence. This may explain the higher number of colistin-resistant pmrAB mutants in clinical isolates that have been reported in the literature. Antibiotic selection pressure is a constant feature of the hospital environment, and this leads to the gradual development of antibiotic resistance. It is likely that colistin resistance is also readily selectable in vivo during monotherapy, since 90% of the A. baumannii isolates prevent the development of colistin resistance at 128 mg/liter, the mutant prevention concentration (MPC), which is much higher than the concentration of the antimicrobial in plasma (46). Therefore, it seems certain that the increased use of colistin will result in greater numbers of colistin-resistant isolates in the near future. In this work, we have shown that there is a greater fitness burden associated with development of colistin resistance via LPS loss than via pmrB mutation, suggesting that there will be a higher frequency of colistin-resistant clinical isolates modulated by the PmrAB system. Resistance due to lipid A modifications has a relatively low cost, especially with respect to A. baumannii virulence, and it is readily selectable under laboratory

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mutants with A549 alveolar cells showed that each of the colistinresistant strains demonstrated reduced ability, compared to the wild-type strain, to decrease the viability of the A549 cells. Importantly, all the ⌬lpx mutants showed reduced ability to impair C. elegans fertility in NG medium, and the lpxC mutant, but not the pmrB mutant, showed reduced virulence in the mouse systemicinfection model. Also, using a different bacterial strain in another genetic background, such as the ABRIM strain, the resistance due to mutation in pmrB caused a decrease in virulence in the worm model. Taken together, these results show that there is a clear fitness and virulence loss associated with colistin resistance but that this genetic burden is higher in mutants that lack LPS than in those that have phosphoethanolamine attached to lipid A. Furthermore, it appears that the different lpx mutants have different fitness costs, with the lpxD mutant showing the highest fitness cost and the least ability to reduce the viability of A549 alveolar respiratory cells. Also, this mutant showed higher colistin MICs, so there may be a correlation between the inactive lpx gene, colistin MICs, and the cost of this genetic load. A transcriptomic analysis of the colistin-resistant lpxA A. baumannii mutant indicated that many genes implicated in cell envelope and membrane integrity were differentially expressed in the mutant (13). Indeed, the mutant clearly needed to alter the expression of many different systems involved in membrane biogenesis, lipoprotein transport, exopolysaccharide production, and drug efflux to survive the loss of LPS. While these LPS-deficient colistin-resistant strains still elaborate an outer membrane, there are substantial differences in the integrity and permeability of this outer membrane. Ultimately, the strains that develop colistin resistance due to lpx mutations need to modulate the composition and structure of the bacterial surface in response to the lack of LPS (13, 41). A proteomic study of a colistin-resistant ATCC 19606 strain with changes in pmrB showed differences in the expression of some proteins, with most showing reduced production in the colistin-resistant mutant. This analysis also showed a biological cost associated with the acquisition of colistin resistance, as we have also shown here (42). However, these protein expression changes do not appear to be nearly as dramatic as the gene expression changes observed following inactivation of lipid A biosynthesis. Likewise, the results presented here agree with those of Lin et al. (43), who showed that treatment of A. baumannii-infected mice with a chemical LpxC inhibitor reduced the production of LPS and completely protected the mice from lethal infection, highlighting the importance of surface LPS for A. baumannii virulence. In a previously published study by López-Rojas et al. (18), the authors showed impaired fitness and virulence of a colistin-resistant A. baumannii strain with mutations in pmrB. Also, Lesho et al. have described a collection of colistin-resistant isolates recovered after treatment with colistin, where the operon pmrCAB seems to be implicated in all cases (44). Although virulence was not tested by the authors, the colistin-resistant strains showed fitness costs, similar to the results obtained in the in vitro assays in our study. Also, recently, López-Rojas et al. analyzed a colistin-resistant clinical isolate of A. baumannii due to changes in the pmrA gene and observed significant differences in growth both in vitro and in vivo when the resistant strain and the colistin-susceptible parental isolate were grown together, with results similar to those obtained in our study comparing the ATCC 19606 strain and the pmrB mutant (45). However, in that study, there were differences in the

Biological Cost of A. baumannii Colistin Resistance

conditions (15, 16). Therefore, we propose that to avoid the emergence of resistance to colistin, especially when monotherapy is used, it is desirable to combine colistin with rifampin, carbapenems, or tigecycline (10) to reduce the development of colistin resistance in A. baumannii by either mechanism.

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This study was funded by grants from the European Community, FP 7 (ID, 278232) (MagicBullet); the Ministerio de Ciencia e Innovación; Instituto de Salud Carlos III, cofinanced by the Spanish Network for Research in Infectious Diseases (REIPI RD12/0015); and Fondo de Investigación Sanitaria (grant PI12/00552). We acknowledge Jennifer Moffat for kindly supplying the A. baumannii lpx mutants.

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