ABSTRACT
Plasma, a unique state of matter with properties similar to those of ionized gas, is an effective biological disinfectant. However, the mechanism through which nonthermal or “cold” plasma inactivates microbes on surfaces is poorly understood, due in part to challenges associated with processing and analyzing live cells on surfaces rather than in aqueous solution. Here, we employ membrane adsorption techniques to visualize the cellular effects of plasma on representative clinical isolates of drug-resistant microbes. Through direct fluorescent imaging, we demonstrate that plasma rapidly inactivates planktonic cultures, with >5 log10 kill in 30 s by damaging the cell surface in a time-dependent manner, resulting in a loss of membrane integrity, leakage of intracellular components (nucleic acid, protein, ATP), and ultimately focal dissolution of the cell surface with longer exposure time. This occurred with similar kinetic rates among methicillin-resistant Staphylococcus aureus (MRSA), Pseudomonas aeruginosa, and Candida albicans. We observed no correlative evidence that plasma induced widespread genomic damage or oxidative protein modification prior to the onset of membrane damage. Consistent with the notion that plasma is superficial, plasma-mediated sterilization was dramatically reduced when microbial cells were enveloped in aqueous buffer prior to treatment. These results support the use of nonthermal plasmas for disinfecting multidrug-resistant microbes in environmental settings and substantiate ongoing clinical applications for plasma devices.
INTRODUCTION
Plasmas are ionized gases that exhibit a plethora of applied temperature and physical properties. In biomedical applications, plasmas are supported by an electric field such that electrons receive external energy more rapidly than surrounding ions (5). Plasmas generate thermal energy (heat) when heavy particle temperatures equilibrate with electron temperature but are considered “nonthermal” when the cooling of ions and uncharged molecules is more effective than the energy transfer from electrons to gas (5). Nonthermal or “cold” plasmas produce a variety of short-lived and long-lived reactive components, including charged particles and UV radiation, without significantly raising temperature (25). Nonthermal plasmas (NTPs) are applied extensively in materials science to modify the properties of carbon-based materials but have also shown promise for applications in biology and medicine (14). Cells and tissues can be treated with NTPs via three general modes of transmission: (i) direct plasma, in which a biological target placed within the plasma stream functions as the ground electrode so that current passes through it; (ii) indirect plasma, in which plasma generated between two electrodes is blown onto a biological target via gas flow; and (iii) plasmas that use a grounded wire mesh electrode to direct current through the mesh rather than cells or tissue (14).
A variety of potential bioapplications have been described for nonthermal plasma, including disinfection of contaminated surfaces in food-processing and clinical settings (19). Indeed, studies have shown that NTPs efficiently inactivate viruses (27), microorganisms (26), and biomolecules such as DNA (22). However, the mechanism through which NTPs inactivate living cells is poorly understood. Several plausible hypotheses exist, including oxidative intracellular/DNA damage, disruption of cellular electrostatic potential, or chemical etching of the cellular surface (24). The lack of a standard protocol for preparing and treating cells with plasma, in addition to technical limitations associated with analyzing cells on surfaces rather than in solution, represent critical impediments for determining the exact mechanism through which NTPs inactivate microbes. Here we address these issues by adsorbing multidrug-resistant microbes onto growth-supportive membrane filters to directly image the physiological changes elicited by the full spectrum of plasma species (electrons, ions, reactive nitrogen and oxygen species, peroxide, hydroxyl radical, and UV) generated from a floating-electrode dielectric barrier discharge (DBD) plasma device (2, 6). This device produces a direct mode of plasma in normal atmospheric air when the applied voltage between the source electrode and the treated surface exceeds the breakdown voltage of surrounding air (6). We demonstrate that NTP rapidly inactivates planktonic cultures of methicillin-resistant Staphylococcus aureus (MRSA), Candida albicans, and Pseudomonas aeruginosa at similar kinetic rates by superficially modifying and permeabilizing the cell surface in a time-dependent manner.
MATERIALS AND METHODS
Strains and media.MRSA USA 300 (NRS384) was obtained from the Network on Antimicrobial Resistance in Staphylococcus aureus (17). Blood culture isolates of Pseudomonas aeruginosa (strain 09-010) and Candida albicans (strain 09-024) were obtained from the clinical repository of the Brooke Army Medical Center (BAMC) Molecular Biology Lab and the U.S. Army Institute of Surgical Research (USAISR). Pseudomonas aeruginosa 09-010 is resistant to ampicillin and tetracycline, while Candida albicans 09-024 is resistant to triazole antifungals (K. Mende, personal communication). Bacterial and yeast isolates were maintained on Trypticase soy agar (TSA) and yeast extract-peptone-dextrose (YPD) agar, respectively. Liquid suspensions were prepared in Luria broth (LB) or YPD broth. Using readings of optical density at 600 nm (OD600), we estimated cell density according to a 0.5 McFarland standard (A600 of 0.132 ≅ 1.5 × 108 CFU/ml, bacterial suspensions) or a common yeast convention (A600 of 1.0 ≅ 3 × 107 CFU/ml). These estimates were validated using CFU measurements of serially diluted cells.
Plasma treatment.Bacterial isolates were cultured in LB at 37°C for >6 h to mid- to late log phase and diluted to approximately 1 × 107 to 2 × 107 CFU/ml in isotonic buffer (0.85% NaCl). Candida albicans was cultured in YPD broth at 30°C to mid- to late log phase and diluted to approximately 1 × 106 to 2 × 106 CFU/ml. Approximately 1 ml of diluted culture was filtered through the matte side of 25-mm black Cyclopore membranes (0.2-μm pore size; GE Healthcare) using a Corning bottle top vacuum filter apparatus (0.22 μm cellulose acetate) and weak vacuum pressure. After filtration, Cyclopore membranes were transferred onto solid TSA (cell-side up) and exposed to plasma for the indicated times using a device described elsewhere (6). The plasma generator was set to 18.6 kV and 2,500 Hz (setting A) or 19.4 kV and 3,000 Hz (setting B), which provides ∼7 W (±1.5 W) total output. Plasma generation was visually and acoustically confirmed during treatment. An ∼24-mm round electrode was used to effectively treat the entire Cyclopore membrane surface. Unless otherwise stated, setting A was employed for all experiments.
To compare plasma efficacies between air and liquid interfaces, cells were captured on Cyclopore membranes as above and transferred onto solid TSA or YPD agar (cell-side up). Plasma was applied for 15 or 30 s over an air interface, after which 100 μl of 0.85% NaCl was added to each filter and cells were spread across the agar surface using glass beads. To test a liquid interface, 100 μl of 0.85% NaCl was applied to the center of each filter prior to NTP treatment, and glass beads were added after treatment to spread cells. Plates were incubated at 30°C or 37°C overnight and photographed the following day.
CFU analysis.To assess the ability of plasma-treated microbes to grow replicatively, clinical isolates were immediately replica plated or “stamped” onto fresh solid media by inverting and gently pressing Cyclopore membranes onto agar following plasma treatment. Cyclopore samples were removed prior to incubating the plates at 30°C (yeast) or 37°C (bacteria) overnight.
Fluorescence staining and analysis.Following NTP treatment, cells captured on Cyclopore membranes were stained with fluorescent dyes that are compatible with paraformaldehyde fixation. All staining and wash steps were performed on Cyclopore membranes using a Corning bottle top vacuum filter apparatus. Briefly, 0.5- to 1-ml volumes were carefully pipetted on top of the filters and washed away by applying weak vacuum pressure. To assess membrane integrity, cells were stained with 5 μg/ml ethidium homodimer-2 (Invitrogen) for 20 min, washed with isotonic buffer, fixed for 30 min with 4% paraformaldehyde in 0.85% NaCl solution, and then counterstained with Syto 9 (Invitrogen). To stain the cell wall of MRSA, cells captured on Cyclopore membranes were fixed in a similar manner and then stained with 100 μg/ml Texas Red-conjugated wheat germ agglutinin (Invitrogen) for 10 min. For cell wall analysis of Candida albicans, fixed cells were stained with 25 μM calcofluor white M2R (Invitrogen) for 30 min. To stain the outer lipid shell of Pseudomonas aeruginosa, cells were stained with 2.5 μg/ml FM1-43 FX (Invitrogen) for 2 min and then fixed. After several washes, Cyclopore membranes were transferred to glass slides and coverslipped with ∼10 μl of antifade reagent. Cells were imaged using a Zeiss AxioImager Z1 automated microscope equipped with 63× and 100× objectives. Images were prepared using AxioVision software (Zeiss, Thornwood, NY).
Quantitation of intracellular ATP.The BacTiter-Glo microbial viability assay (Promega) produces a luminescent signal that is proportional to the amount of ATP present within metabolically active cells. Following plasma treatment, Cyclopore membranes were immediately transferred to a 6-well plate and covered with 500 μl of LB broth. An equal volume of BacTiter-Glo reagent (Promega) was added to each well and incubated with mixing for 5 min. Aliquots were withdrawn in triplicate and measured in an opaque 96-well plate on a SpectraMax M2 plate reader (Molecular Devices) in luminescence mode. BacTiter-Glo was modified for use with Candida albicans by increasing the number of cells on Cyclopore membranes (∼3 × 107) and increasing reagent incubation time (10 min). For each experiment, the background luminescence of the cell medium alone was subtracted from each sample according to the manufacturer's instructions.
Protein and nucleic acid release.Following plasma treatment, Cyclopore membranes were immediately transferred to a 6-well plate and overlaid with 1 ml of sterile water. After brief incubation, eluates were collected from each well and centrifuged (5 min × 14,000 rpm) to remove any cells and/or debris that coeluted from the membrane filter. The optical density of the resulting supernatant was measured at 280 nm and 260 nm (in succession) in a quartz cuvette. To properly blank the spectrophotometer, a control sample (through which diluted rich medium was filtered and placed on solid agar) was processed in an identical manner.
In situ analysis of DNA strand breaks.After 7 s plasma treatment, Cyclopore membranes were fixed in 4% paraformaldehyde for 30 min and washed with isotonic buffer using a Corning bottle top vacuum filter apparatus. Cells on the filter surface were permeabilized with 0.1% Triton X-100 in 0.1% sodium citrate buffer and then transferred to a 6-well plate and incubated in terminal deoxynucleotidyltransferase-mediated dUTP-biotin nick end labeling (TUNEL) reagent using the fluoroscein in situ cell death detection kit (Roche) to label DNA strand breaks. Negative and positive controls were prepared according to the manufacturer's instructions. After TUNEL labeling, cells were counterstained with 5 μg/ml DAPI (4′,6-diamidino-2-phenylindole). Cyclopore membranes were mounted and coverslipped using ∼10 μl of antifade reagent, and then they were imaged as described above.
Oxyblot analysis.Approximately 3 × 109 CFU/ml were filtered onto Cyclopore membranes and exposed to plasma or air for 7 s in duplicate. To account for extracellular protein in the media, fresh medium was filtered through an additional set of Cyclopore membranes, which were then treated with plasma. All Cyclopore membranes were transferred to a 6-well plate and chilled for 10 min at −20°C prior to incubation with 300 μl B-PER reagent (Thermo Scientific). To prevent sample oxidation during extraction, dithiothreitol (DTT) was supplemented to 50 mM for one set of samples, and lysates were extracted for 15 min with rotation. DTT was withheld from the second set of samples to accurately measure the protein content of the extract using bicinchoninic acid (BCA) protein assay reagent (Thermo Scientific). DTT-treated lysates were transferred into microcentrifuge tubes and cleared by centrifugation (5 min × 13,000 rpm). A total of 15 μg of protein was derivatized using the Oxyblot protein oxidation detection kit (Millipore) following the manufacturer's instructions. Reagent volumes were scaled up accordingly. Derivatized lysates were separated by SDS-PAGE, transferred to polyvinylidene difluoride (PVDF), probed with anti-dinitrophenyl (DNP) antibody, and then detected by chemiluminescence following the manufacturer's instructions.
RESULTS
Plasma rapidly inactivates clinical microbial strains.Preparing and analyzing live cells on solid surfaces rather than in aqueous solution pose extra challenges to the study of how nonthermal plasma (NTP) inactivates microbes. Typically, cells are mounted on surfaces by spraying, smearing, or drying thin aqueous cell suspensions; for subsequent biological analysis (such as following NTP treatment), cells are generally recovered using a pipette or vortexer and then concentrated through centrifugation. Such intermediate steps may introduce additional experimental variables and risk segregating the original cell population into nonrepresentative “survivor” fractions after plasma treatment. To mitigate these concerns, we employed a membrane adsorption technique in which microbes were captured on the top surface of thin (7- to 20-μm thickness) Cyclopore polycarbonate membranes possessing 0.2-μm track-etched pores. These membrane filters are dyed with irgalan black to eliminate the natural autofluorescence of polycarbonate, thereby supporting fluorescence staining applications and epifluorescence microscopy. Using these filter membranes, microbial populations were imaged before and after plasma treatment with minimal physical or mechanical disruption. Importantly, Cyclopore membranes are biologically inert and support cell growth when placed on solid media (Fig. 1B).
Time course of plasma sterilization using a replica plating assay. (A) Approximately 107 total bacterial cells or ∼106 total yeast cells were filtered onto each Cyclopore membrane and exposed to air or plasma as indicated. After treatment, Cyclopore membranes were replica plated onto fresh media. Complete 7-log10 kills of Pseudomonas aeruginosa and MRSA USA 300 and a >5-log10 kill for Candida albicans were observed at 30 s of plasma treatment. Growth reduction is apparent following 1 s of plasma exposure. No significant differences are observed between plasma settings A (top rows in each panel) and B (bottom rows). Bar, 25 mm. (B) Representative microbial growth characteristics on the surface of Cyclopore membranes exposed to air or plasma. Approximately 107 cells (MRSA USA 300) were filtered onto Cyclopore membranes and grown overnight on TSA agar or exposed to air or plasma and then replica plated. No viable colonies were observed on the Cyclopore surface after 30 s of plasma treatment.
To investigate the mechanism through which NTP inactivates microbes, we tested three representative clinical isolates: MRSA USA 300, Pseudomonas aeruginosa, and Candida albicans. The latter were obtained from infected blood cultures and exhibit multidrug resistance (see Materials and Methods). To compare the disinfection rates of our experimental setup, approximately 106 to 107 cells of each strain were filtered onto the surface of black Cyclopore membranes, which were then placed on the surface of rich agar to maintain cell vitality and treated with plasma for various times. Immediately after treatment, microbes on the Cyclopore surface were replica plated onto fresh media to assess clonogenicity. As depicted in Fig. 1A, we observed that nonthermal plasma rapidly inactivated all three strains relative to control samples exposed to air, resulting in complete (7 log10) growth inhibition for MRSA and Pseudomonas aeruginosa and near-complete (>5 log10) growth reduction of Candida albicans after 30 s of NTP exposure. Partial cellular inactivation was apparent for all three strains even after 1 s of NTP treatment (Fig. 1A). After 30 s of plasma treatment, few if any viable cells were observed on the Cyclopore membrane surface as well (Fig. 1B). We observed no significant difference in disinfection rates between two similar NTP outputs. Overall, the observed kinetics of inactivation (<60 s) are consistent with the findings of other groups who have evaluated similar NTP devices on planktonic cultures (1, 10, 23, 27, 28).
Plasma disrupts the cell exterior.Since Cyclopore membranes are compatible with epifluorescence microscopy, we used a panel of fluorescent dyes with unique staining properties to investigate the structural integrity of microbes exposed to nonthermal plasma. Ethidium homodimer-2 (EthD-2), a red fluorescent nucleic acid dye that is normally excluded from intact (viable) cells, was employed to assess cell membrane integrity. After penetrating cells with damaged membranes, EthD-2 becomes fluorescent upon binding intracellular nucleic acids. As an added advantage, EthD-2 can be fixed in place using paraformaldehyde without altering its staining properties (16). Using membrane adsorption epifluorescence techniques, we observed that the incidence of EthD-2 increased strikingly in Candida albicans after plasma treatment compared to controls exposed to normal air (Fig. 2A). Unexpectedly, however, EthD-2 also accumulated at the cell periphery in plasma-treated Candida cells (Fig. 2A, white arrowheads). Since EthD-2 and related dyes are known to nonspecifically label the cell walls of certain cells (12), the unique staining pattern observed following NTP treatment might reflect structural modification of surface polysaccharides such that EthD-2 intercalates into these polymers in a manner analogous to DNA (12), although alternative explanations are possible (see Discussion).
Plasma modifies the staining properties of EthD-2 at the cellular surface and induces cell wall lesions in Candida albicans. (A) Time course of EthD-2 staining as a function of plasma exposure. In contrast to cells exposed to air, EthD-2 stains the cell surface in addition to the cell interior after brief plasma exposure (7 s, white arrowheads). EthD-2 fluorescence at the cell surface appears more punctate with longer plasma exposure (30 s, blue arrowheads). Intracellular nucleic acids are counterstained with Syto 9. (B) Calcofluor white M2R staining (pseudocolored) as a function of plasma exposure. Cell wall defects are visible with longer plasma treatment (blue arrowheads) and appear morphologically similar to cell surface lesions labeled by EthD-2 in panel A. Bar, 5 μm.
To better distinguish between the interior and exterior of Candida albicans, a green fluorescent nucleic acid dye (Syto 9) was used to counterstain intracellular nucleic acid pools. With longer NTP exposure (30 s), the staining pattern of EthD-2 along the cell periphery appeared more punctate (Fig. 2A, blue arrowheads) and seemed to indicate destruction of the cell wall. To verify these results, we stained Candida albicans with calcofluor white M2R, a fluorescent dye that exhibits selective cell wall binding. Indeed, following 30 s of NTP treatment, we observed large voids in calcofluor staining (Fig. 2B, blue arrowheads) that confirmed the localization pattern of EthD-2 (Fig. 2A, blue arrowheads), thus supporting the conclusion that NTP elicits acute cell wall damage over time.
Similar experiments were performed on MRSA and Pseudomonas aeruginosa to determine if plasma also altered the cellular surface of bacterial clinical isolates. Following short NTP exposure, the incidence of EthD-2 staining increased dramatically in both bacterial strains (Fig. 3A and C), indicating that NTP rapidly impairs the integrity of bacterial membranes. However, we were unable to clearly distinguish if EthD-2 labeled the bacterial cell wall in addition to the cell interior. To determine whether NTP affects the bacterial surface, we utilized two different fluorescent stains: for MRSA, a red fluorescent lectin (Texas Red-conjugated wheat germ agglutinin) that binds to the outer cell wall, and for Pseudomonas aeruginosa, an amphiphile (FM1-43 FX) that becomes fluorescent upon intercalating into the outer lipid membrane. As shown in Fig. 3B, visible cell wall damage was observed for MRSA following 30 s of NTP treatment (blue arrowheads) and mirrored the type of surface damage we observed with Candida albicans (Fig. 2B). Likewise, NTP elicited progressive alterations in the outer lipid membrane of Pseudomonas aeruginosa, concomitant with reduced FM1-43 FX fluorescence intensity and irregular cell morphology (Fig. 3D). When viewed in total, these patterns illustrate that NTP dramatically alters the microbial surface as a function of plasma duration, with increased membrane permeability and EthD-2 labeling preceding visible structural damage at the cell surface (Fig. 4).
Plasma elevates cellular EthD-2 staining and induces cell surface alterations in MRSA and Pseudomonas aeruginosa. (A) Time course of EthD-2 staining in MRSA as a function of plasma exposure. Cellular nucleic acids are counterstained with Syto 9 in overlay. Bar, 1 μm. (B) Staining pattern of Texas Red-conjugated wheat germ agglutinin (WGA) on MRSA as a function of plasma exposure. Cell wall lesions are visible following prolonged plasma treatment (30 s, arrowheads). Bar, 0.5 μm. (C) Time course of EthD-2 staining in Pseudomonas aeruginosa as a function of plasma exposure. Cellular nucleic acids are counterstained with Syto 9 in overlay. (D) FM1-43 FX staining in Pseudomonas aeruginosa as a function of plasma exposure. Overall cell morphology and FM1-43 FX staining intensity are progressively disrupted by plasma compared to control cells exposed to air. Bar, 2 μm.
EthD-2 staining precedes visible structural perturbation of the microbial surface during plasma treatment. The percentage of cells labeled with EthD-2 or depicting visible surface damage (based on calcofluor white M2R, Texas Red-conjugated WGA, or FM1-43 FX staining patterns) were scored from experiments depicted in Fig. 2 and 3. Data were pooled from different fields in which >100 bacterial cells or >50 yeast cells were scored per sample per treatment time and averaged.
Plasma rapidly permeabilizes cells.To confirm that NTP inactivates microbes by damaging the cellular surface, we measured intracellular ATP levels before and after plasma treatment using cells captured on Cyclopore membranes. All three microbial strains exhibited rapid declines in intracellular ATP levels (with similar kinetic rates) as a function of plasma duration (Fig. 5A). The steepest decline in ATP occurred within 7 s of plasma exposure, with levels dropping ∼50% after only 1 s of exposure (Fig. 5A). These results are consistent with microscopy images depicting rapid EthD-2 cellular uptake after short plasma treatment (Fig. 2A, 3A and C, and 4). Additionally, by measuring the optical density of the Cyclopore filter eluate, we observed a proportional increase in OD280 and OD260 readings after plasma treatment (Fig. 5B), thereby indicating leakage of nucleic acid and protein from microbial cells. However, we observed little to no evidence that NTP induced widespread genomic damage or oxidative stress during the first 7 s of treatment using an in situ TUNEL assay to detect DNA strand breaks and a separate assay to detect oxidized protein (Fig. 6). Nevertheless, given that cells are permeabilized by plasma over time and eventually release intracellular contents, secondary damage to DNA and protein may occur outside the cell or downstream of cell lysis with prolonged plasma exposures (9). Altogether, these data support our initial microscopy findings and confirm that plasma rapidly inactivates cells by permeabilizing the cell surface, resulting in loss of membrane integrity, leakage of intracellular components (ATP, nucleic acid, protein), and ultimately focal dissolution of the cell exterior as a function of treatment time.
Plasma rapidly permeabilizes microbial cells. (A) Rapid depletion of intracellular ATP as a function of plasma duration. ATP levels were quantified and normalized to that of untreated cells exposed to air. Intracellular ATP levels rapidly decline with similar kinetics for all tested microbes at two different plasma settings. (B) Leakage of protein and nucleic acids from plasma-treated cells. Cell-free eluates were prepared as described in Materials and Methods and analyzed for protein (OD280) and nucleic acid (OD260) content. Each bar represents the average OD value in arbitrary units (AU) from two independent experiments.
Plasma does not elicit widespread DNA strand breaks or irreversible oxidative modification of protein during the early phase of cellular inactivation. (A) In situ detection of DNA strand breaks using TUNEL. Pseudomonas aeruginosa was treated with air or plasma on Cyclopore membranes and stained with TUNEL reagent (green) and DAPI (violet pseudocolor). Only subtle TUNEL staining was observed after 7 s of plasma treatment. A TUNEL-positive control group was created by DNase I digestion. (B) Oxyblot detection of carbonylated protein before and after plasma treatment. Lysates of Pseudomonas aeruginosa were collected and analyzed as described in Materials and Methods. Control cells exposed to air exhibit several carbonylated bands after derivatization (+DNP), likely as a consequence of physiologic protein oxidation during exponential growth in rich media (4). Note that nonderivatized control lysate (−DNP) emits no signal, indicating that the α-DNP antibody is specific. Only subtle increases in protein carbonyls are observed after plasma treatment (see arrows).
The cellular environment modulates plasma sterilization.To further test our hypothesis that NTP targets the cellular surface, we assayed whether microbial sterilization could be affected by introducing an aqueous layer between the cell surface and the plasma stream. To this end, microbes were exposed to plasma with or without prior addition of 100 μl of isotonic buffer onto the Cyclopore membrane filter surface. This small liquid interface dramatically reduced the rate at which plasma inactivated cell growth for all three clinical isolates compared to an air interface (Fig. 7). These results demonstrate that plasma species are superficial and may be quenched by the cellular environment, including perhaps cells positioned closer to the electrode in 3-dimensional (3-D) space within the cell suspension. Indeed, similar studies have hinted that the effects of plasma may be modulated by overlaying cell biomass in addition to soluble components within aqueous media (3, 11). By analogy, our findings likely substantiate why log10 magnitudes of NTP sterilization are less robust for microbes enclosed in biofilm slime or wound effluent compared to planktonic cells in isolation (8, 11).
Cellular inactivation is diminished by the presence of a liquid interface between microbes and the plasma stream. (A) Comparison of MRSA inactivation across an air or liquid interface. Cells were filtered onto Cyclopore membranes and treated as described in Materials and Methods. The presence of a liquid partition between the cells and the plasma stream greatly diminishes sterilization efficacy relative to cells treated across air. (B) Comparison of Candida albicans and Pseudomonas aeruginosa inactivation for plasma applied across air or liquid interfaces for 15 s. The presence of a liquid partition supports the growth of microbial lawns that appear yellow (Candida) or green (Pseudomonas). In contrast, cells exposed to plasma across an air interface are completely inactivated or produce only sparse colonies.
DISCUSSION
Materials scientists routinely use NTPs to modify the surface characteristics of carbon-based polymers. Studies show that NTPs generated from DBD devices etch surfaces through more-or-less random ion bombardment, resulting in a time-dependent physical deposition of oxygen and nitrogen species from normal atmospheric air (24). Here, we report that analogous physical damage occurs on the surface of multidrug-resistant microbes, ultimately leading to partial cell lysis and the release of intracellular components with longer treatment times. Using membrane adsorption techniques to mount and stain cells on a supportive surface for epifluorescence microscopy, we observe that brief exposure to plasma (∼7 s) elicits a rapid loss of cell membrane integrity (poration), while longer exposures (∼30 s) result in visible lesions on the microbial surface (rupture of cell wall or outer lipid membrane). These results agree with data obtained through more indirect forms of imaging (e.g., scanning electron microscopy, atomic force microscopy, and Gram stain) by groups testing similar plasma devices (2, 8, 13, 15, 20, 26, 28) and are consistent with recent evidence that nonthermal plasma induces membrane lipid peroxidation (9). Analogous superficial effects are reported following the treatment of cultured human and mammalian cells with NTP (25). Concomitant with cell poration and lysis, we demonstrate that intracellular ATP is rapidly depleted and that cellular protein/nucleic acids gradually leak from the cell as a function of plasma duration.
Interestingly, we observed that the high-affinity nucleic acid dye EthD-2 localizes specifically to the cell wall of Candida albicans after plasma treatment. The reason behind this unexpected observation is unclear and warrants further investigation. This localization pattern may point to NTP-induced structural changes in the polysaccharide cell wall that perhaps mimic nucleic acid structure (12), as EthD-2 and related dyes normally emit fluorescence when bound and intercalated into DNA. However, since EthD-2 carries a net positive charge, we cannot rule out the possibility that EthD-2 may be recruited to the cell surface via electrostatic interaction with negatively charged species deposited by plasma (15). Alternatively, EthD-2 may stain nucleic acids leaking from the periphery of NTP-treated cells, as we and others have shown that plasma rapidly permeabilizes cell membranes and extracts intracellular nucleic acids (18). Prior to the onset of visible membrane damage, we observed only minor increases in DNA damage and protein oxidation, indicating that genetic damage and oxidative stress do not play primary roles in NTP-mediated microbial sterilization, consistent with the findings of other groups (7, 26, 27). However, significant DNA damage has been reported in bacteria with longer plasma exposure times (9). In light of the data presented here, it is possible that the complete breakdown of the microbial surface may promote secondary damage to intracellular components as a function of time.
It is tempting to speculate that NTP-induced damage to the cell wall may also explain certain “viable but nonculturable” (VBNC) phenotypes reported under some plasma conditions (2, 8). Indeed, cell wall-deficient microorganisms such as “L-form” bacteria and yeast spheroplasts (created in the lab by inhibiting cell wall synthesis or enzymatically damaging the cell wall) are nonculturable in standard nonisotonic laboratory media (21). The risk of producing VBNC phenotypes is generally reduced with longer plasma exposure (2), which, as demonstrated in this study, continues to damage the cell surface and lead to partial or complete cell lysis. However, because plasma is superficial, we also demonstrate that the physical environment of the cell is a critical modulator of NTP sterilization efficacy. This property bears important implications for further biomedical development of nonthermal plasmas.
ACKNOWLEDGMENTS
We thank Gary Friedman, Gregory Fridman, and Alexander Fridman at Drexel University for building the plasma device and facilitating helpful discussions and Stephen Davis and Lisa Plano at the University of Miami for providing MRSA USA 300 and for valuable feedback on clinically relevant plasma settings. We also thank members of BAMC and USAISR, including Katrin Mende, Clinton Murray, and Charles Guymon, for providing microbial strains.
This work was supported by the Defense Advanced Research Projects Agency (government contract number NBCW911NF0910063).
The views, opinions, and/or findings contained in this article are those of the authors and should not be interpreted as representing the official views or policies, either expressed or implied, of the Defense Advanced Research Projects Agency or the Department of Defense.
The methods outlined in this study utilized track-etch membranes manufactured by Whatman, a GE Healthcare company.
FOOTNOTES
- Received 20 September 2011.
- Returned for modification 13 October 2011.
- Accepted 29 December 2011.
- Accepted manuscript posted online 9 January 2012.
- Copyright © 2012, American Society for Microbiology. All Rights Reserved.
