Membrane potential plays a crucial role in many important physiological processes in bacteria. It is a component of the proton-motive force and is used to power various membrane-embedded complexes, including ATP synthase, the flagellar motor, and various small-molecule transport systems (1–5
). Membrane potential has also been shown to be critical for bacterial cell division, proliferation, and signaling, and recent studies have begun to elucidate the mechanisms by which bacterial membrane potential is regulated (6–10
Bacterial membrane potential also plays a critical role in antibiotic susceptibility, highlighting the value in identifying membrane potential-altering compounds in the quest to combat multidrug-resistant pathogens (5
). For example, carbonyl cyanide 3-chlorophenylhydrazone (CCCP), a well-known proton ionophore, increases Enterobacteriaceae
susceptibility to polymyxins, while others have shown that hyperpolarizing conditions, such as those with the addition of alanine and glucose, are capable of reversing resistance to aminoglycosides (12
Recent publications have demonstrated the value in having high-throughput techniques for discovering novel membrane potential-altering compounds, which can be used alone or in synergy to increase the efficacy of commonly used antibiotics. Farha et al. (16
) and McAuley et al. (17
) have conducted membrane potential screens in Gram-positive Staphylococcus aureus
and Bacillus subtilis
, respectively, demonstrating that membrane potential-altering compounds could serve as novel antimicrobials. No such high-throughput membrane potential screen has been described in the Gram-negative E. coli
, a species of public health concern with the rise of carbapenem-resistant and extended-spectrum beta-lactamase-producing Enterobacteriaceae
Fluorescent probes have been extensively used to measure relative membrane potentials in bacteria (19–21
). For example, 3,3′-diethyloxacarbocyanine iodide [DiOC2
(3)], a cationic carbocyanine dye, has been used to measure the relative membrane potentials of bacteria upon treatment with antimicrobials (22
(3) is a positively charged fluorescent dye that accumulates within cells in a charge-dependent manner, shifting its fluorescence spectrum from green to red at high concentrations due to concentration-dependent dye stacking (19
). Therefore, an increase in red fluorescence is indicative of an increase in membrane potential (i.e., hyperpolarized relative to the previous state). However, many fluorescent dyes, including DiOC2
(3), have low signal-to-noise ratios when used in Gram-negative bacteria due to dye exclusion by the outer membrane, which limits their use in many assays. EDTA treatment has been employed to chelate the metal ions that stabilize the dye-excluding outer membrane, allowing the dye to more accurately report the voltage across the inner membrane (24–27
). Alternatively, some investigators have utilized Gram-negative mutants with defective outer membranes with increased permeability to promote dye uptake (28–30
Changes in DiOC2
(3) fluorescence are typically recorded using flow cytometry or, in some cases, fluorescence microscopy; therefore, most membrane potential measurements in bacteria are collected at relatively low throughput. As has been previously observed, performing DiOC2
(3) measurements in populations (i.e., on the plate reader) using available methods yielded extreme variation and low signal-to-noise ratios, which complicated its use for high-throughput experiments (25
). We describe a high-throughput approach that uses DiOC2
(3) to measure Gram-negative membrane potential changes using a fluorescence microplate reader. We optimized several assay parameters, including temperature, dimethyl sulfoxide (DMSO) concentration, growth stage, DiOC2
(3) concentration, time of EDTA treatment, and resuspension buffer components, to maximize dynamic range and minimize variation. We include the results from several assay validation experiments and a valinomycin curve to correlate membrane potential changes to relative fluorescence changes. The optimized assay has a high signal-to-noise ratio and reliably reports membrane potential changes in a high-throughput format. As proof of its utility, this assay was used to characterize the effects of four small molecules hypothesized to alter E. coli
In the future, this approach can be utilized to efficiently probe the effects of other small molecules, growth conditions, and mutant strains. Ultimately, this could lead to the identification of novel antimicrobials and targets.
Intentionally perturbing bacterial membrane potential is likely an effective antimicrobial target for several reasons. First, it is critical for several physiological processes in bacteria, including ATP synthesis, transport, motility, and cellular division; disruption of these key processes through alterations of membrane potential could readily impact the ability of bacteria to thrive in the host environment. Second, several proteins that are already known to influence bacterial membrane potential, such as electron transport chain components and ion channels, are located in the plasma membrane; therefore, compounds that target these likely do not require transport into the cytoplasm and are less likely to lead to resistance via efflux mechanisms. Lastly, even if not effective alone, membrane potential-altering compounds can increase susceptibility to other antimicrobial compounds and could reverse currently existing resistance mechanisms. To discover novel compounds that target membrane potential, it is imperative that we have access to high-throughput tools to study its modulation.
Here, we optimized the use of the fluorescent dye DiOC2(3) in high-throughput formats to robustly capture membrane potential changes in E. coli and presumably other Gram-negative organisms. A valinomycin calibration curve showed that these conditions have a dynamic range of at least 144 mV and can readily distinguish 12-mV changes; a similar calibration could be performed to approximate the membrane potential changes of any user’s experimental conditions. Experiments can be carried out in 384-well assay plates to utilize small volumes of costly reagents with little day-to-day variation. We validated this approach by measuring the effects of several control compounds on membrane potential, including CCCP, valinomycin, sodium azide, barium chloride, antimycin A, and amlodipine. Future work will involve screening additional molecules to determine their effects on membrane potential and characterizing the precise mechanisms of action of compelling compounds.
MATERIALS AND METHODS
Solutions and media.
All cultures were streaked from frozen glycerol stocks onto Difco Luria-Bertani agar (Miller) plates. From fresh plates (less than 1 week old), overnight cultures were inoculated in 5 ml Merck Luria-Bertani broth (Miller) in 15-ml round-bottom plastic tubes. Day cultures were grown in Luria-Bertani broth (Miller) by diluting 1:1,000 into no more than 25 ml in a 250-ml flask. The small-molecule inhibitors used in this study included the following: antimycin A (A8674; Sigma-Aldrich), amlodipine besylate (PHR1185; Sigma-Aldrich), barium chloride dihydrate (0974-0; J. T. Baker), and sodium azide (0639; VWR). Tissue culture-grade DMSO was used where indicated. Assay resuspension buffer was comprised of 130 mM NaCl, 60 mM Na2
, 60 mM NaH2
, 10 mM glucose, 5 mM KCl, and 0.5 mM MgCl2
). NaOH was used to adjust the pH of the resuspension buffer to 7.0. 10× PBS consisted of 1.3 M NaCl, 70 mM Na2
, and 30 mM NaH2
, and the pH was adjusted to 7.0. All buffers were sterilized using a 0.22-μm vacuum filter.
K-12 BW25113 cultures were streaked from frozen stocks onto Luria-Bertani agar (Miller) (37
). Overnight cultures were grown in 5-ml volumes from three colonies in plastic, round-bottom tubes. Day cultures were inoculated by diluting 1:1,000 into Luria-Bertani from overnight cultures. All day cultures were aerated by shaking at 250 rpm, 37°C, until they reached an optical density at 600 nm (OD600
) of 0.5 to 0.6. Kanamycin MIC experiments were performed by diluting overnight cultures to an OD600
of 0.001 in cation-adjusted BD BBL Mueller-Hinton II broth. Cells then were cultured in 96-well microplates for 20 h of shaking at 250 rpm, 37°C.
Valinomycin and EDTA viable cell counts.
E. coli K-12 BW25113 cells were grown to an OD600 of ∼0.5 in LB and centrifuged at 2,400 × g for 10 min to remove spent media. Cells were resuspended in 1× PBS and treated with either 10 mM EDTA or water control for 5 min. The EDTA-treated cultures were centrifuged and resuspended in 1× PBS. Tenfold serial dilutions were performed, and the cells were plated on LB agar containing 135 mM KCl in place of NaCl and supplemented with either 20 μM valinomycin or DMSO (vehicle control). Viable cell counts were performed following an 18-h incubation at 37°C.
High-throughput DiOC2(3) membrane potential measurements.
Mid-exponential phase (OD600 of 0.5 ± 0.1) E. coli was pelleted by centrifugation at 2,400 × g for 10 min to remove spent growth medium; all centrifugation steps were performed at room temperature. The cells were then resuspended in 1× PBS to an OD600 of 1.0. The cells were treated with 10 mM EDTA for 5 min and then recentrifuged at 2,400 × g for 10 min to remove EDTA. EDTA-treated E. coli cells were pelleted and resuspended to an OD600 of 1.0 in assay resuspension buffer (see “Solutions and media,” above, for the resuspension buffer recipe). A 6 mM DiOC2(3) stock in DMSO was added to cells for a final concentration of 30 μM. DiOC2(3) was obtained from ThermoFisher Scientific. DiOC2(3)-loaded cells were then added to a 96-well opaque microplate for a final volume of 200 μl. Similar results were observed in transparent 384-well plates for final volumes of 50 μl/well.
In cases where compound effects were tested, the compound was first added to the bottom of the well of the microplate prior to the addition of the DiOC2(3)-loaded cells, and the cells were mixed with the compound by pipetting up and down twice. A total reaction volume of 200 μl was used in each well. 5 min after incubation at room temperature, DiOC2(3) fluorescence was recorded using the BMG Labtech CLARIOstar microplate reader using 450-nm excitation. Red fluorescence intensity was recorded at 670-nm emission.
Cells were cultured as described above. Half of the cells were treated with 10 mM EDTA, and half were untreated controls. Cells were loaded with 30 μM DiOC2(3) and imaged on noncoated glass slides using the Zeiss LSM 780 NLO multiphoton microscope to capture red fluorescence (emission, 670 nm) upon excitation at 488 nm. The images were taken at a magnification of ×63 and at a zoom setting of 2× for a total magnification of 126×.
Valinomycin calibration curves.
The membrane potential assay was performed with resuspension buffer containing various concentrations of KCl. The total NaCl and KCl concentration was held to a constant 300 mM XCl concentration, where X is Na+
. Cells resuspended in the appropriate buffer were added to 96-well plates containing valinomycin for a final concentration of 10 μM valinomycin. The fluorescence was recorded after 20 min of incubation. The data shown in Fig. 3A
were normalized to either the 0 mM potassium buffer or to DMSO only, respectively. The membrane potential change is reported relative to 1 mM extracellular K+
based on the Nernst equation: change in ΔΨ = RT
/[1 mM K+
]), where R
is the ideal gas constant (8.314 J/mol·K), T
is temperature in Kelvin, z
is the valency of the ion, and F
is Faraday’s constant (95,484.56 C/mol).
DiOC2(3)-loaded cells were added to wells containing the indicated compound. The fluorescence was recorded after 20 min of incubation. Background fluorescence, obtained from reading control solutions that contained no cells, was subtracted at each compound concentration. The background-corrected fluorescence was then normalized to that of the vehicle-only sample (DMSO, ethanol, or water only). Amlodipine stocks were dissolved in 100% DMSO. Antimycin was dissolved in 95% ethanol. Sodium azide and BaCl2 were dissolved in filter-sterilized water.
Effect of amlodipine on kanamycin MIC.
Exponential-phase E. coli BW25113 was diluted in pH 6 Mueller-Hinton (100 mM morpholineethanesulfonic acid) to an OD600 of 0.001. Diluted culture was added to wells of a 96-well plate containing various concentrations of kanamycin and DMSO or amlodipine for a final concentration of 100 μM amlodipine. The plates were then aerated for 18 h at 250 rpm, 37°C, after which growth was observed. The MIC was the lowest concentration of kanamycin in which no visible growth occurred during this time period. Twofold dilutions of filter-sterilized kanamycin dissolved in water were used for the determination of the MIC.
Experiments were performed using three biological replicates with a minimum of three technical replicates, except where otherwise indicated. In all cases, error bars represent the standard errors of the means. Statistical differences were analyzed using t
tests in GraphPad Prism. Z' was calculated using the following equation: Z' = 1 – (3σpos
|), where pos is the positive control (CCCP) and neg is the negative control (DMSO) (31
We thank James Sacchettini for discussions on assay development and use of the ClarioSTAR microplate reader. We also thank the TAMU Image Analysis Laboratory, College of Veterinary Medicine & Biomedical Sciences, specifically Rola Barhoumi and Joseph A. Szule, for help with microscopy experiments. We would like to acknowledge Dwight Baker and Thomas Meek for providing their expertise on assay development, Sarah Beagle and Katrina Hofstetter for thoughtful discussions, and Pushkar Lele, Joseph Sorg, and Ming Zhou for comments on our manuscript.
We declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
This work was supported by grants from The Welch Foundation (A-1742), NIGMS (1R01GM132436), and TAMU STRP funds. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institute of General Medical Sciences or the National Institutes of Health.