Microbicides Alter the Expression and Function of RND-Type Efflux Pump AdeABC in Biofilm-Associated Cells of Acinetobacter baumannii Clinical Isolates

Abstract

Acinetobacter baumannii is a Gram-negative bacterium that causes nosocomial infections worldwide. This microbe's propensity to form biofilms allows it to persist and to survive on clinical abiotic surfaces for long periods. In fact, A. baumannii biofilm formation and its multidrug-resistant nature severely compromise our capacity to care for patients in hospital environments. In contrast, microbicides such as cetrimide (CT) and chlorhexidine (CHX) play important roles in the prevention and treatment of infections. We assessed the efficacy of CT and CHX, either alone or in combination, in eradicating A. baumannii biofilms formed by clinical isolates, by using stainless steel washers to mimic hard abiotic surfaces found in hospital settings. We demonstrated that increasing amounts of each microbicide, alone or in combination, were able to damage and to reduce the viability of A. baumannii biofilms efficaciously. Interestingly, the adeB gene of the resistance-nodulation-cell division (RND) family is predominantly associated with acquired resistance to antimicrobials in A. baumannii. We showed that CT and CHX adversely modified the expression and function of the RND-type efflux pump AdeABC in biofilm-associated A. baumannii cells. Furthermore, we established that these microbicides decreased the negative charges on A. baumannii cell membranes, causing dysregulation of the efflux pump and leading to cell death. Our findings suggest that CT and CHX, alone or in combination, can be used efficaciously for eradication of A. baumannii from hospital surfaces, in order to reduce infections caused by this nosocomial agent.

INTRODUCTION

Acinetobacter baumannii is a multidrug-resistant (MDR) Gram-negative bacterium capable of colonizing and causing disease in hospitalized patients, especially those with prolonged stays in the intensive care unit (ICU). The principal concern regarding A. baumannii is its ability to persist in the hospital environment, on various abiotic materials (1–3). This allows susceptible patients to come into contact with the microbe, which often results in outbreaks of ventilator-associated pneumonia, meningitis, septicemia, urinary tract infections, and wound infections (4). These infections are a challenge to treat, due to the emergence of multidrug-resistant (MDR) A. baumannii strains (5–7). It is thus clear why A. baumannii has become a potential emerging nosocomial threat throughout the world in the past 2 decades.

Microbial biofilms consist of bacteria enclosed in a polymeric matrix, which protects them against harsh environments (8). Biofilm-forming bacteria are more resistant to antimicrobials and disinfectants. A. baumannii biofilms can survive desiccation (1–3, 9), and microorganisms have been recovered from the patients' environments, including bed curtains, furniture, and hospital equipment, during outbreaks (10). Disinfection of patient rooms has been successful in reducing outbreaks of A. baumannii, which underscores the role of the hospital environment as a reservoir for A. baumannii (10). Disinfection and sterilization, as well as other aseptic techniques, are critical for the prevention and control of nosocomial infections. Therefore, careful assessment of the antimicrobial activity of disinfectants used in hospitals is necessary (11).

In Gram-negative bacteria, efflux pumps belonging to the resistance-nodulation-cell division (RND) family are considered some of the most important contributors to resistance to commonly used antimicrobial compounds (12). To date, three Acinetobacter drug efflux (Ade) RND systems, i.e., AdeABC (13), AdeFGH (14), and AdeIJK (15), have been characterized in A. baumannii. AdeABC and AdeFGH pumps are important in acquired resistance (16, 17), whereas AdeIJK contributes to intrinsic resistance (15). The AdeABC efflux pump is found in the majority of A. baumannii clinical isolates and primarily provides A. baumannii with antimicrobial resistance. This system consists of the AdeA, AdeB, and AdeC proteins, with AdeB being a member of the RND superfamily (13), responsible for aminoglycoside, β-lactam, chloramphenicol, erythromycin, and tetracycline resistance. Drug transport is driven by the transmembrane electrochemical gradient of protons. Due to the MDR and disinfection resistance of A. baumannii, it is necessary to investigate antimicrobial alternatives for eradicating A. baumannii and other intractable hospital-related microbes from clinical settings, in an effort to limit epidemics. One potential approach is the use of single or multiple microbicides to inhibit the function of efflux pumps in nosocomial agents, with the purpose of limiting disinfection resistance in clinical settings (18–21).

Microbicides are universally utilized to control microbial growth in household (e.g., hand washes and hard-surface disinfectants), medical (e.g., antiseptics, disinfectants, and additions to medical devices such as catheters and surgical dressings), and industrial (e.g., contamination control in industry and in food production) environments (22). The mechanisms of action of cationic agents, such as quaternary ammonium compounds (e.g., cetrimide [CT]) and biguanides (e.g., chlorhexidine [CHX]), involve direct interactions with the microbe's cell envelope, resulting in membrane disruption and leakage of cytoplasmic components (23). Despite the vast literature on the use of microbicides in hospital disinfection, limited studies have been performed to examine the efficacy of these compounds in eradicating microbial biofilms formed by clinical isolates.

In this study, we assessed the efficacy of CT and CHX, used alone or combination, in eradicating biofilms formed by A. baumannii clinical isolates grown on stainless steel washers (SSWs). We used SSWs as hard abiotic surfaces to grow the biofilms because this platform mimics the substrates found in hospital settings. This model was recently validated for study of the effects of commonly used disinfectants and environmental stressors on A. baumannii cells within biofilms (2). Furthermore, we investigated whether the expression and function of the AdeABC efflux pump in A. baumannii isolates was modified after treatment with microbicides, to identify the molecular mechanisms involved in microbial susceptibility to these antimicrobial agents.

MATERIALS AND METHODS

Acinetobacter baumannii.

A total of 8 A. baumannii clinical isolates (isolates 0057, 0248, 1422, 1611, 2098, 2231, 3659, and 7405) were included in the study and were selected because of their ability to form biofilms on SSWs (2). The isolates were derived from blood or wound cultures at the Walter Reed Medical Center (Washington, DC) or the Montefiore Medical Center (Bronx, NY). The antimicrobial susceptibility profile for each clinical isolate tested in this study was published previously (2). The strains were stored at −80°C in brain heart infusion (BHI) broth (BD, Franklin Lakes, NJ), with 40% glycerol, until use. Test organisms were grown in tryptic soy broth (TSB) (MP Biomedicals, Solon, OH) overnight at 37°C, using a rotary shaker set at 150 rpm. Growth was monitored by measuring the optical density at 600 nm (OD₆₀₀) using a microtiter plate reader (Bio-Tek, Winooski, VT).

Biofilm formation.

SSWs were used as a substrate to grow A. baumannii biofilms, as described previously (2). Briefly, for each strain, an inoculum of 4 × 10⁶ to 5 × 10⁶ A. baumannii cells was suspended in a 50-ml Erlenmeyer flask with 25 ml of TSB containing 6-mm-diameter SSWs. The flasks were then incubated for 24 h at 37°C on a rotary shaker set at 150 rpm. The SSWs containing A. baumannii biofilms were removed from the flask using flame-sterilized forceps. To remove nonadherent bacterial cells, the abiotic surface was washed three times with phosphate-buffered saline (PBS).

Susceptibilities of A. baumannii biofilms to microbicidal compounds.

To evaluate the susceptibilities of the A. baumannii cells within biofilms to antiseptic chemicals, 200 μl of PBS containing CT (0.1, 0.5, or 1%; Sigma, St. Louis, MO), CHX (0.1, 0.5, or 1%; Sigma), or a combination (0.1% CT plus CHX) was added to each well of microtiter plates containing A. baumannii biofilms on SSWs. Mature biofilms and microbicides were mixed for 0.5, 1, 5, or 10 min by use of a microtiter plate reader, to ensure uniform distribution, and were incubated at room temperature. After incubation, the viability of bacteria within the biofilms was quantified by CFU. The susceptibilities of the biofilm-associated cells to microbicides were determined by comparing the CFU of the biofilms coincubated with microbicides with the CFU of the biofilms grown in PBS.

Direct CFU determinations in biofilms.

SSWs were transferred from wells to microcentrifuge tubes, washed three times with 1 ml of PBS to remove the microbicide, and sonicated to detach the cells, as described previously (2). One hundred microliters of the dissociated cell suspension was transferred to another microcentrifuge tube, containing 900 μl of PBS. The suspension was then gently homogenized for 3 min. Two-fold serial dilutions of the suspensions were then performed, and 100-μl aliquots of each dilution were plated on tryptic soy agar (TSA) plates.

Crystal violet assay.

A. baumannii biofilm formation on SSWs was measured by crystal violet (CV) staining, modifying protocols described previously (24, 25). SSWs coated with A. baumannii biofilms were gently washed with PBS to remove nonadherent bacteria. Each SSW was added to a well of a 96-well plate, air dried, and stained with 0.1% CV solution for 15 min. SSWs were rinsed three times with distilled water (dH₂O), shaken vigorously with forceps to remove all excess dye, air dried, and transferred to a clean 96-well plate. A suspension of 30% acetic acid in dH₂O was added to the wells containing SSWs, to solubilize the CV, and the plate was incubated at room temperature for 15 min. Finally, the SSWs were removed from the wells, and solubilized CV was measured in a microtiter plate reader at 550 nm, using 30% acetic acid in dH₂O as a negative control.

Confocal microscopy.

The structural integrity of untreated biofilms and microbicide-treated biofilms was examined using the Live/Dead biofilm viability kit (Invitrogen, Carlsbad, CA) and confocal microscopy. Briefly, A. baumannii biofilms were grown for 24 h in 35-mm glass-bottom culture dishes (MatTek Corp., Ashland, MA), treated with microbicides alone or in combination for 1 min, rinsed three times with dH₂O, and incubated for 30 min at room temperature in 2 ml of dH₂O containing the fluorescent stains SYTO9 (6 μl) and propidium iodide (6 μl), with protection from light. The dishes were then rinsed three times with dH₂O to remove excess stain. SYTO9 (excitation wavelength, 500 nm; emission wavelength, 535 nm) labels live and dead bacteria, while propidium iodide (excitation wavelength, 600 nm; emission wavelength, 650 nm) penetrates only bacteria with damaged membranes. Propidium iodide binds to the cell's nucleic acids with greater affinity than SYTO9. Thus, with the appropriate mixture of the dyes, bacteria with intact cell membranes stain fluorescent green, whereas bacteria with damaged membranes stain fluorescent red. Microscopic examinations of biofilms formed in culture plates were performed with confocal microscopy, using an inverted Leica TCS SP5 confocal laser scanning microscope (Leica, Wetzlar, Germany).

Zeta potential measurements.

The zeta potential (ζ) was calculated in a zeta potential analyzer (ZetaPlus; Brookhaven Instruments Corp., Holtsville, NY). ζ is a measurement of charge (in millivolts), defined as the potential gradient that develops across the interface between a boundary liquid in contact with a solid and the mobile diffuse layer in the body of the liquid. It is derived from the equation ζ = (4πηm/D), where D is the dielectric constant of the medium, η is the viscosity, and m is the electrophoretic mobility of the particle.

RNA extraction and cDNA synthesis.

For RNA extraction, the experimental samples were processed as described for CFU determination. RNA extraction was performed using the RNeasy kit (Qiagen, Hilden, Germany), following the manufacturer's instructions. To remove any genomic DNA carryover, the samples were treated with DNase I (Qiagen) for 30 min at 37°C, followed by heat inactivation for 5 min at 65°C. Then, 1 μg of total RNA was used to synthesize cDNA with the Bio-Rad iScript reverse transcriptase kit (Bio-Rad, Berkeley, CA), following the manufacturer's instructions. The control reaction was set up using all components of the reaction mixture but without the reverse transcriptase enzyme (i.e., no reverse transcriptase [NRT]).

Reverse transcriptase quantitative PCR.

Analysis was carried out with the RND pump-encoding gene adeB. The primers used for reverse transcriptase quantitative PCR (qRT-PCR) analysis of adeB were as follows: forward, 5′-GGATTATGGCGACTGAAGGA-3′; reverse, 5′-AATACTGCCGCCAATACCAG-3′. The efficiency of each primer was tested by using a 10-fold serial dilution of the cDNA mixture, and only primers with efficiencies between 95% and 105% were used for the analysis. The expression of genes was determined by quantitative PCR using iQ SYBR Green Supermix (Bio-Rad). Two different control reactions were included in the analysis, i.e., a no-template control (NTC) and a NRT control. We used 16S rRNA as the reference gene (forward, 5′-CAGCTCGTGTCGTGAGATGT-3′; reverse, 5′-CGTAAGGGCCATGATGACTT-3′). Relative expression was determined using the cycle threshold (ΔΔC_T) method on a Mastercycler RealPlex2 system (Eppendorf, Hamburg, Germany). Reactions were set up using 300 nM primers and 5 μl of the cDNA template (diluted 1:10). The cycling conditions used were as follows: 55°C for 30 min and then 40 amplification cycles of 95°C for 15 s, 55°C for 30 s, and 72°C for 30 s. The samples were cooled to 55°C, and a melting curve for temperatures between 55°C and 95°C, with 0.5°C increments, was recorded. All reactions were carried out in triplicate. Target gene expression was measured using expression relative to that of the 16S reference gene, and A. baumannii strain 0057 was used as the control strain. Data analysis was carried out using Mastercycler ep realplex software.

Accumulation of ethidium bromide.

The kinetics of ethidium bromide (EtBr) accumulation in A. baumannii 0057 cells after treatment with single or combined microbicides were monitored with a fluorimetric assay, modifying protocols described previously (13). Cells were grown to an OD₆₀₀ of 0.5, pelleted carefully at room temperature, resuspended to an OD₆₀₀ of 0.2 in sodium phosphate buffer (pH 7.0), and returned to 37°C. EtBr was added at a final concentration of 2 μg/ml and, after 420 s of incubation, 0.5% CT or CHX, 0.1% CT plus CHX, or 256 μg/ml 1-(1-naphthylmethyl)-piperazine (NMP) was added. Stock solutions of microbicides and NMP were diluted in 0.1% dimethyl sulfoxide (DMSO) (Sigma). A solution of 0.1% DMSO was used as a control to rule out the possibility of cytotoxicity. The change in fluorescence intensity (excitation wavelength, 530 nm; emission wavelength, 600 nm), which is proportional to the quantity of intracellular dye, was recorded with a Synergy HT spectrofluorimeter (Bio-Tek) for 800 s following the addition of EtBr.

Statistical analysis.

All data were subjected to statistical analysis using GraphPad Prism 6.0 (GraphPad Software, La Jolla, CA). P values were calculated by analysis of variance (ANOVA) and adjusted by use of the Bonferroni correction. P values of <0.05 were considered significant.

RESULTS

Effects of single or combined microbicides on survival of A. baumannii biofilm-related cells.

To evaluate the susceptibilities of bacterial cells in mature biofilms of eight clinical isolates, A. baumannii biofilms on SSWs were incubated with CT, CHX, or the combination at different time points (0.5, 1, 5, and 10 min), and cell viability was determined by CFU analysis (Fig. 1A). Treatment of A. baumannii biofilms with 1% CT or 0.5 or 1% CHX alone or the combination of 0.1% CT plus CHX (P < 0.0001) significantly reduced approximately one-third of the bacterial population, compared to the control group, after 0.5 min of incubation (Fig. 1A). Notably, on average, A. baumannii cells within biofilms were susceptible to ≥0.5% CT or CHX alone, compared to the untreated control (P < 0.0001), after 1 min of incubation with microbicides (Fig. 1A). A. baumannii biofilms incubated with 0.1% CT plus CHX for similar times displayed considerably less CFU than did cells grown under most of the other conditions (P < 0.0001); the exception was cells within biofilms grown in the presence of 1% CT, which showed no significant statistical difference. Exposure of A. baumannii biofilms to microbicides alone or in combination for ≥5 min resulted in substantial reductions in the viability of biofilm-associated bacteria (Fig. 1A). Since we found that exposure to microbicidal compounds for 1 min resulted in reduction of 50% of the viable population for the majority of conditions, we used 1 min as the time of exposure to microbicides in all further experiments in this study.

FIG 1.

Effects of microbicidal compounds on A. baumannii biofilm-related bacteria formed on stainless steel washers (SSWs) for 24 h. Biofilm formation of A. baumannii strains was measured with CFU (A) and crystal violet (CV) staining (B) assays. Bacterial biofilms were exposed to cetrimide (CT) (0.1, 0.5, and 1%), chlorhexidine (CHX) (0.1, 0.5, and 1%), or a combination (0.1% each), and the viability and strength of the biofilms were compared with those of biofilms incubated in PBS. (A) Biofilms were treated with microbicides for 0.5, 1, and 5 min. Bars, averages of eight A. baumannii strains (n = 8) for each experimental condition; error bars, standard deviations (SDs). Significance (P < 0.05) was calculated by analysis of variance (ANOVA) and adjusted by use of the Bonferroni correction. *, #, ϕ, $, %, and &, significantly lower CFU than the untreated, 0.1% CT, 0.5% CT, 0.1% CHX, 0.5% CHX, and 1% CHX groups, respectively. (B) Top-down images of the CV solubilization assay with 30% acetic acid in dH₂O, with A. baumannii biofilms grown on SSWs after treatment with CT, CHX, or the combination for 1 min. All experiments in panels A and B were performed three times, with similar results being obtained each time.

Quantification of biofilms only in terms of cell viability provides a partial view of the actual biofilm quantities, as the extracellular matrix largely exceeds, in terms of mass, the cellular component. Thus, biofilm formation on SSWs was compared among the different A. baumannii strains by using the CV method, which stains both cellular and matrix component of biofilms (Fig. 1B). Similar to the CFU assay, A. baumannii cells within biofilms were susceptible to ≥0.5% CT and CHX alone (P < 0.0001). Bacterial biofilms treated with the combination of microbicidal compounds showed lower OD values than did those treated with ≤0.5% CT or CHX (P < 0.0001) (data not shown), but there was no statistical difference in comparison to biofilms exposed to 1% CT or CHX.

Additionally, we assessed the correlation between CFU and CV solubility assays in monitoring the biofilm cell viability of A. baumannii clinical isolates after incubation with microbicides alone or in combination (data not shown). A strong correlation was found between the results of the two assays (R² = 0.9954; P = 0.0001). CV solubility was found to have an exponential association with CFU counts. Therefore, cell viability was correlated with increased bacterial mass and extracellular matrix in the biofilms.

CT and CHX combined disrupt the architecture of A. baumannii biofilms.

Confocal microscopic examination was used to correlate the CFU and CV solubility assay results with the visible effects on biofilm structure (Fig. 2A and B). Regions of green fluorescence (SYTO9) represent viable cells; the red fluorescence (propidium iodide) indicates metabolically inactive or nonviable cells. Untreated A. baumannii strain 0057 biofilms grown in the presence of PBS alone showed a robust architecture, with a thickness of ∼70 μm and a majority of viable cells protected in the bottom of a structure consisting mainly of dead cells and exopolymeric matrix (Fig. 2A and B). Cell death is a normal component of multicellular development and an important mechanism for differentiation inside biofilms that facilitates dispersal of a subpopulation of surviving cells (26). Biofilms treated with 0.1% CT plus CHX exhibited decreased thickness (∼30 μm) of the exopolymeric matrix, structural disruption, and minimal numbers of viable cells (Fig. 2A and B).

FIG 2.

Confocal microscopy of A. baumannii strain 0057 biofilms after treatment with the combination of CT and CHX. (A) Representative images of biofilms showed viable (green [SYTO9]) and dead (red [propidium iodide]) cells. (B) The thickness and morphology of the bacterial biofilms can be observed in the z-stack reconstruction. For A and B, the photographs were taken at a magnification of ×63. Bars, 20 μm.

Microbicides alter the expression and function of the A. baumannii AdeABC efflux pump.

The A. baumannii gene adeB encodes the transmembrane protein of the AdeABC efflux pump, and disruption of this gene leads to the loss of MDR (13). All isolates in the present study were found to carry the adeB gene (data not shown). First, we investigated the impact of microbicides on the expression of the adeB gene by biofilm-associated cells by using qRT-PCR (Fig. 3A). A. baumannii biofilm-associated cells exhibited significantly reduced adeB expression after exposure to either 0.5% CT or CHX alone or in combination (P < 0.0001). Similarly, 256 μg/ml NMP, an efflux pump inhibitor (EPI) that has been shown to reverse MDR in A. baumannii (27), significantly decreased adeB gene expression, which suggests that single or combined microbicides may directly disrupt the functional activity of the AdeABC efflux pump (P < 0.0001).

FIG 3.

Role of the A. baumannii RND efflux pump after exposure to microbicidal compounds. (A) Changes in expression of the RND efflux pump-encoding gene adeB in A. baumannii clinical isolates after exposure to CT, CHX, or the combination. 1-(1-Naphthylmethyl)-piperazine (NMP) was used as the negative control, and 16S rRNA was used as the housekeeping gene control. Bars, averages (n = 8) for each experimental condition; error bars, SDs. Every isolate was tested in duplicate in two independent experiments. *, significance (P < 0.05) calculated by ANOVA and adjusted by use of the Bonferroni correction. (B) Ethidium bromide (EtBr) accumulation assay with A. baumannii strain 0057. Each reagent was added to the bacterial suspension at 420 s. This experiment was performed twice, with similar results being obtained each time. A.U., absorbance units.

EtBr, a toxic hydrophobic cation that is known to be a substrate of efflux systems, has been used to assess the contributions of efflux pumps and outer membrane permeability to cellular accumulation (13, 28). To confirm that the difference in microbicidal susceptibility shown by treated A. baumannii cells was due to inhibition of the efflux mechanism, we determined the accumulation of EtBr in A. baumannii strain 0057 (Fig. 3B). Cells under all of the conditions, except for heat-killed bacterial cells, displayed similar levels of EtBr uptake for 420 s. At 420 s, each culture was exposed to 0.5% CT, 0.5% CHX, 0.1% CT plus CHX, or NMP. Microbial cells treated with single or combined microbicides showed similar levels of EtBr accumulation. However, bacterial cells grown under these conditions showed much higher EtBr concentrations than untreated controls (P < 0.05). NMP-treated bacteria demonstrated greater and similar EtBr uptake, in comparison with microbicide-treated and heat-killed cells, respectively.

Microbicides reduce A. baumannii cellular charges.

CT and CHX are cationic compounds that interact with the negative charges of bacterial cell walls, resulting in destabilization of the cytoplasmic membranes where efflux pumps are located (23). We investigated whether microbicides are responsible for reductions in the negative charges on the surfaces of A. baumannii cells, affecting the expression and function of the A. baumannii AdeABC efflux pump. The surface charges of A. baumannii cells exposed to 0.5% CT or CHX or the combination of 0.1% CT and CHX were measured (data not shown). Exposure to single or combined microbicides significantly reduced the net negative surface charge of the A. baumannii cells (−24.32 ± 3.49 mV, −19.84 ± 4.29 mV, and −17.74 ± 3.67 mV with 0.5% CT, 0.5% CHX, and 0.1% CT plus CHX, respectively), compared to those of untreated (−42.69 ± 3.70 mV) and heat-killed (−31.53 ± 3.02 mV) bacterial cells (P < 0.05). Notably, A. baumannii cells treated with the combination of 0.1% CT and CHX showed considerably lower negativity than did microbial cells incubated with 0.5% CT.

DISCUSSION

Resistance to disinfection makes A. baumannii difficult to eradicate from hospital environments. More notably, biofilm formation by this microbe enhances its persistence and pathogenesis (9), as in cases of infections related to surgical implants, prosthetics, and catheters, all of which are common causes of nosocomial infections (10). Therefore, we exposed A. baumannii biofilm-associated cells to microbicides regularly used in household, health care, and industrial facilities. We found that CT, CHX, and a combination of the two microbicides effectively killed A. baumannii cells within biofilms. Our results suggest that exposing biofilms to ≥0.5% CT or CHX or 0.1% CT plus CHX for 1 min reduced by approximately 5-fold the numbers of viable bacteria on the SSWs. The percentages of microbicide solutions used in our experiments corresponded to the Centers for Disease Control and Prevention recommendations (29) (e.g., 0.1 to 2% CT and 0.05 to 1% CHX) for disinfection of hard surfaces such as SSWs. Similar percentages of microbicide solutions were recently used effectively against biofilms formed by periodontal pathogens (30–32) and single strains of relevant Gram-positive and Gram-negative pathogens, including A. baumannii (33). Interestingly, A. baumannii biofilm-associated cells have shown resistance to CHX, but effective concentrations used to kill these bacterial cells are comparable to those used in this study (33). Furthermore, CHX has been effectively combined with other antimicrobials to coat central venous catheters to prevent biofilm colonization by Gram-negative bacteria (34).

Several studies examining the efficacy of microbicides have used wide ranges of times of exposure to these agents, e.g., 30 s to 2 min (32), 5 to 20 min (30), or 24 h (33). We exposed A. baumannii biofilms grown on a hard surface to microbicides for 0.5, 1, 5, or 10 min. Our results revealed that a time of 1 min, which is practical to be used by maintenance employees, is appropriate for use in clinical settings to disinfect hard surfaces, especially surfaces that touch people's bare skin each day and any surfaces that could come into contact with uncovered infections. The fact that microbicides were mixed and uniformly distributed on the SSWs, effectively reducing the microbial population, suggests that the targeted approach of frequent thorough cleaning of touched surfaces and surfaces that have been exposed to infections might be an efficacious strategy for the eradication of biofilm-related infections in medical facilities.

We assessed the effects of CT plus CHX on mature A. baumannii biofilm architecture. Confocal microscopy showed that the microbicide combination considerably reduced the thickness of the biofilm architecture and increased the susceptibility of microbial cells. The modes of action of cationic microbicidal agents rely largely on interactions with the bacterial cell envelope, leading to membrane disruption and leakage of cytoplasmic organelles (23). In this regard, our zeta potential analysis demonstrated that microbicides had profound effects on the negative charges of A. baumannii cell membranes, which may translate into interference with surface colonization or adhesion and cell-cell interactions within the biofilms (35). In fact, heat-killed bacteria displayed more-negative cell surfaces than did microbicide-treated microorganisms, suggesting that the cationic nature of these antimicrobials significantly decreases cell surface negativity. For example, a net positive charge to the bacterial surface may disrupt the structural stability of the biofilm by dispersing the microbial cells into suspension, making them more susceptible to microbicidal damage as individual cells. A previous study showed that CHX-sensitive isolates of Pseudomonas stutzeri demonstrated peeling of the outer membrane, substantial loss of cytoplasmic electron-dense material, and extensive lysis (36). Moreover, colistin-susceptible A. baumannii strains exhibited reduced negative surface charges in the stationary phase (37), which is comparable to our results obtained with biofilm-related cells.

RND efflux systems present in A. baumannii have been shown to contribute to the organism's survival after exposure to disinfectants (21) or antimicrobial agents (13). Specifically, the AdeABC pump plays an important role in the acquired resistance of A. baumannii, being overexpressed in MDR strains. We examined the effects of microbicides, alone or in combination, on expression of the adeB gene by cells associated with mature biofilms. Interestingly, our findings indicate that single or combined microbicides similarly reduced the expression of adeB in biofilm-related cells. The EPI NMP, which interferes with A. baumannii multidrug resistance (27), was used as a control. A. baumannii isolates also exhibited 1-fold adeB expression, which agreed with the results obtained using microbicides alone or in combination. Inactivation of the adeB gene has been shown to increase A. baumannii susceptibility to microbicides (21). Recently, an adeB mutant displayed a defect in biofilm formation, behaving like Escherichia coli and Salmonella enterica, in which deletion or inhibition of the efflux pumps was associated with weakened biofilm formation (18–20).

We used fluorimetric analysis to investigate whether the microbicidal susceptibility seen in A. baumannii strains resulted from direct inhibition of the AdeABC pump. An efflux defect of the AdeABC pump was confirmed by comparing the levels of intracellular accumulation of EtBr by A. baumannii strain 0057 after treatment with single or combined microbicides. Our results indicate that inhibition of adeB by microbicidal substances increases EtBr accumulation, which supports the notion that the function of the AdeABC pump in microbial cells is altered upon treatment with CT or CHX or a combination of these microbicides. Moreover, the apparently faster penetration of EtBr in microbicide-treated A. baumannii cells, compared to untreated microbes, suggests greater permeability of the cell envelope. This is not surprising, given that CT and CHX are involved in the attraction and adsorption of cationic molecules to the cell surfaces of microorganisms, modifying cell membrane permeability and resulting in the loss of intracellular components and osmotic imbalance in the cells. Similarly, the AdeB protein contains 12 transmembrane segments that may be inactivated by alterations of the negative charges on cell membranes, resulting in impaired function and an inability to protect bacteria from antimicrobial activity.

Efforts to reduce the risk of nosocomial infections include appropriate programs for disinfecting surfaces, furniture, equipment, and physical areas, along with appropriate antisepsis of hands and use of gloves. There is also increasing interest in the incorporation of microbicides into medical devices such as catheters and surgical dressings, with the intention of inhibiting microbial colonization and biofilm formation (34, 38, 39). Despite the evident benefits of microbicides in the majority of these applications, concerns have been raised that their extensive use may select for bacteria with reduced susceptibility to them or cross-resistance to clinically relevant antibiotics (36, 40, 41). For example, triclosan can select for mutants of A. baumannii that display reduced susceptibilities to multiple antibiotics from chemically distinct classes, in addition to triclosan resistance (42). However, the stability of induced changes in microbicide susceptibility depends on the bacterium and the compound used (22). Although we did not observe acquired resistance for any of the clinical strains tested, further studies are warranted to examine the mechanisms of resistance of cells within biofilms to microbicides, including CHX and CT, in order to understand how bacteria avoid becoming eliminated from hospital environments. The lack of success in eliminating biofilms has become a major factor in outbreaks of infections and diseases related to contamination problems.

Studies assessing the susceptibilities of biofilm-related bacteria to antimicrobials are limited; therefore, we focused on identifying single microbicides or a combinations of microbicides, to establish a synergistic mechanism of biofilm inhibition in MDR A. baumannii. Future studies should monitor and address the mechanisms for the development of antimicrobial resistance by microbial biofilms, mainly in clinical environments, to minimize the negative effects on human health.

Funding Statement

L.R.M. is supported by NYIT College of Osteopathic Medicine start-up funds.