Free access
Research Article
20 August 2020

Use of a Fluorescence-Based Assay To Measure Escherichia coli Membrane Potential Changes in High Throughput


Bacterial membrane potential is difficult to measure using classical electrophysiology techniques due to the small cell size and the presence of the peptidoglycan cell wall. Instead, chemical probes are often used to study membrane potential changes under conditions of interest. Many of these probes are fluorescent molecules that accumulate in a charge-dependent manner, and the resulting fluorescence change can be analyzed via flow cytometry or using a fluorescence microplate reader. Although this technique works well in many Gram-positive bacteria, it generates fairly low signal-to-noise ratios in Gram-negative bacteria due to dye exclusion by the outer membrane. We detail an optimized workflow that uses the membrane potential probe, 3,3′-diethyloxacarbocyanine iodide [DiOC2(3)], to measure Escherichia coli membrane potential changes in high throughput and describe the assay conditions that generate significant signal-to-noise ratios to detect membrane potential changes using a fluorescence microplate reader. A valinomycin calibration curve demonstrates this approach can robustly report membrane potentials over at least an ∼144-mV range with an accuracy of ∼12 mV. As a proof of concept, we used this approach to characterize the effects of some commercially available small molecules known to elicit membrane potential changes in other systems, increasing the repertoire of compounds known to perturb E. coli membrane energetics. One compound, the eukaryotic Ca2+ channel blocker amlodipine, was found to alter E. coli membrane potential and decrease the MIC of kanamycin, further supporting the value of this screening approach. This detailed methodology permits studying E. coli membrane potential changes quickly and reliably at the population level.


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 (15). 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 (610).
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, 1113). 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, 14, 15).
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 (18).
Fluorescent probes have been extensively used to measure relative membrane potentials in bacteria (1921). 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, 23). DiOC2(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 (2427). Alternatively, some investigators have utilized Gram-negative mutants with defective outer membranes with increased permeability to promote dye uptake (2830).
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 membrane energetics.
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.


Assay optimization and validation.

As an initial test of DiOC2(3) as a membrane potential sensor in E. coli, we examined the response of E. coli to 5 μM CCCP, a proton ionophore that should shift the membrane potential to ∼0 mV (the proton Nernst potential under these conditions). Although a small change was observed, the very low signal-to-noise ratio with DiOC2(3)-loaded cells that had been exposed to CCCP suggested that the assay needed to be further optimized. The Z′ value is often used in assay development to monitor this optimization process, since it captures the separation in signal between positive and negative controls (31). The positive control used in this calculation was cells exposed to CCCP, and the negative control was cells exposed to DMSO vehicle alone. Temperature, time of EDTA treatment, resuspension buffer components, DMSO concentration, and excitation and emission wavelengths were each optimized and are described below.
The concentration of DiOC2(3) and emission wavelength were optimized by obtaining emission spectra at different wavelengths in 15, 30, and 60 μM DiOC2(3). Based on Z′, 30 μM DiOC2(3) at a 670-nm emission wavelength generated the largest difference between positive and negative controls with the smallest overlap in error (see Fig. S1 in the supplemental material). We found performing the cell preparation steps and plate reader experiments at room temperature reduced variation between biological replicates.
DiOC2(3) and other membrane potential probes have reduced uptake in Gram-negative bacteria due to exclusion by porins. EDTA has been used to chelate divalent cations stabilizing the lipopolysaccharide molecules of the outer membrane, allowing for dye access to the inner membrane (32, 33). After testing 0, 5, 10, 15, and 20 min of EDTA treatment, we found that a 5-min EDTA treatment yielded the highest signal between controls with the least variation.
As expected, a 5-min EDTA treatment resulted in markedly increased uptake of DiOC2(3) at the population level, yielding a significant signal intensity between DMSO-treated cells and those depolarized with CCCP (Fig. 1A and B and Fig. S2). Fluorescence microscopy demonstrated that EDTA treatment resulted in a significant signal increase in single-cell measurements as well (Fig. 1C). To determine if EDTA affected the ability of molecules to gain access to the plasma membrane and if EDTA treatment significantly impacted the viability of E. coli, we monitored viable cell counts of cells treated with the potassium ionophore valinomycin. Despite generating an increase in dye signal, EDTA alone had no effect on viability (P = 0.4) (Fig. 2, first and fourth bars). Not surprisingly and consistent with literature demonstrating that EDTA enhances valinomycin uptake, EDTA treatment enhanced valinomycin-mediated lethality, decreasing the viable number of colony forming units (CFUs) more than 10-fold at 20 μM valinomycin (P = 0.007) (Fig. 2, third and sixth bars) (34, 35). More importantly, EDTA treatment was not required to see an effect of valinomycin on E. coli viability (P = 0.004) (Fig. 2, first and third bars). Valinomycin at both 5 and 20 μM significantly reduced the number of viable cells both with and without an initial EDTA treatment, demonstrating that large molecules, such as valinomycin (molecular weight, 1,111.32), can access the inner membrane in the absence of outer membrane destabilization (Fig. 2).
FIG 1 Effect of EDTA on DiOC2(3)-loaded E. coli. (A) The fluorescence spectra (excitation wavelength, 450 nm) of DiOC2(3)-loaded E. coli without (left) and with (right) EDTA treatment. The red dotted line indicates cells treated with 5 μM CCCP, whereas black is DMSO only. The arrow indicates the emission (Em.) wavelength at which subsequent experimental data were collected. AU, arbitrary units. (B) DiOC2(3) signal, normalized to the mean, at 670-nm emission from three biological replicates with and without 5 μM CCCP. n = 3. Error bars represent standard errors of the means (SEM). (C) Fluorescence microscopy images showing dye uptake in cells not subjected to (top) and subjected to (bottom) a 5-min EDTA treatment. The scale bar represents 3 μm.
FIG 2 Effects of EDTA and valinomycin on E. coli. Viable cell counts from E. coli without EDTA treatment or with a 5-min EDTA treatment plated on LB agar with 135 mM K+ in the presence of 0, 5, or 20 μM valinomycin. The % DMSO concentration remained constant under all three conditions. Error bars represent SEM. n = 4 to 7. Unpaired t tests, with Welch’s correction for unequal variances, were used for data analysis (**, P < 0.01).
Taken together, a final DMSO concentration of 1.5% [1.0% from compound, 0.5% due to DiOC2(3)] was chosen based on Z' optimization. A 6.1 (±0.9)-fold difference was observed using 670-nm emission, the wavelength identified as optimal by Z' analysis (Fig. 1B). The final optimized assay yields a Z' value of 0.8, suggesting that these assay conditions yield reproducible signal with a minimal number of false positives, making it ideal for use in high throughput (31).

Assay resolution and dynamic range.

To determine the resolution and dynamic range of the DiOC2(3) assay, experiments were carried out using valinomycin in different concentrations of extracellular KCl. Because valinomycin allows the diffusion of potassium ions (K+) down electrochemical gradient, it can be used to alter membrane potential by manipulating extracellular concentrations of K+. The approximate membrane potential change associated with such conditions can be calculated using the Nernst equation. For example, in the presence of valinomycin in high concentrations of extracellular K+, the Nernst potential should change such that K+ is driven into the cell, resulting in its depolarization. By normalizing the fluorescence exhibited by cells at each K+ concentration to the fluorescence in 0 mM KCl buffer, we were able to observe the effect of varying K+ when it can freely permeate the inner membrane.
As expected, in the absence of valinomycin, extracellular KCl had little effect on E. coli membrane potential. Furthermore, upon the addition of valinomycin, there was little change in membrane potential observed when extracellular K+ was approximately equal to K+ found in the 1× phosphate-buffered saline (PBS) in which cells were previously resuspended. However, when the extracellular K+ was increased to >10 mM, membrane depolarization was observed, an effect that increased with a rise in extracellular K+ (Fig. 3A and B).
FIG 3 Response of E. coli to valinomycin in various KCl concentrations. Fluorescence intensities of cells treated with valinomycin were normalized to fluorescence of valinomycin-treated cells in 0 KCl buffer (A) and to the corresponding buffers without valinomycin (1% DMSO only) (B). Error bars represent SEM. Three biological replicates were performed. (C) Assay dynamic range was determined by calculating the change in membrane potential upon valinomycin addition for each extracellular K+ concentration relative to 1 mM K+ (see Materials and Methods).
Increased extracellular K+ led to increased depolarization when cells were resuspended in buffers containing different concentrations of K+. The Nernst equation was used to calculate the relative change in membrane potentials in K+ concentrations ranging from 1 to 300 mM extracellular K+; membrane potential changes relative to 1 mM extracellular K+ are reported, since the concentration of intracellular K+ is not known but is the same under these assay conditions (Fig. 3A). Using this information, the dynamic range of the assay is at least 144 mV, and the resolution is approximately 12 mV (Fig. 3C).

Effects of compounds on E. coli membrane potential.

One prospective purpose of using DiOC2(3) in high throughput is to identify compounds that are capable of altering bacterial membrane potential; ultimately, these molecules could lead to effective antimicrobials that are employed alone or in synergy with currently available antibiotics. Therefore, it is critical that the assay is capable of characterizing compounds in a reliable way, including the generation of meaningful dose-response curves and 50% inhibitory concentration (IC50) values. As a control, we generated a dose-response curve of the proton ionophore CCCP (Fig. 4). As anticipated, DiOC2(3)-loaded E. coli exhibited increased depolarization in response to increasing CCCP concentrations. The data fit to an IC50 of 0.10 ± 0.03 μM, similar to values reported in other organisms (17, 36).
FIG 4 DiOC2(3)-loaded E. coli exhibits increased depolarization in response to increasing CCCP concentrations. Dose-response curves of the membrane potential response to CCCP indicate an IC50 of 0.10 ± 0.03 μM. Error bars represent SEM.
We then characterized the effects of several small-molecule inhibitors of interest, including amlodipine, antimycin, barium chloride, and sodium azide, at various concentrations (Fig. 5). Each of these molecules has reported effects on bioenergetics in other organisms, but it remains unclear if they would exert similar effects on E. coli. Amlodipine, a eukaryotic Ca2+ channel inhibitor used to treat angina, hyperpolarized the E. coli membrane, yielding an IC50 value of 100 μM. The maximum amlodipine concentration tested, 300 μM, led to a 3-fold increase in the fluorescence of DiOC2(3). Antimycin, an electron transport chain inhibitor, decreased fluorescence 2.7-fold at its maximum concentration tested. Barium chloride, a nonselective channel inhibitor, depolarized the cells, eliciting 50% of its response at 4.7 mM. Sodium azide generated similar results, with an IC50 of 1.2 mM. Antimycin, barium chloride, and sodium azide generated approximately 40% inhibition at the maximum concentrations tested; this is similar to the effect seen by resuspending cells in 300 mM KCl in the presence of 10 μM valinomycin (Fig. 3), suggesting that the membrane potential change is less than ∼144 mV based on E. coli's response to valinomycin.
FIG 5 Effects of amlodipine, antimycin, barium chloride, and sodium azide on E. coli membrane potential. Dose-response curves are shown for each compound tested; data were normalized to the fluorescence of cells treated with vehicle only (DMSO, water, or ethanol). The IC50 for each compound is shown. Not determined (ND) indicates that a dose-response curve was unable to be fit to the data. Three biological replicates were performed to determine membrane potential response to each compound; data points represent averages from these experiments. Error bars represent SEM.
We next wanted to verify that amlodipine was altering membrane potential using a DiOC2(3)-independent assay. To test this, we measured the MIC of kanamycin, an aminoglycoside antibiotic whose uptake is dependent on a strong negative membrane potential, in the presence of sublethal concentrations of amlodipine (100 μM) (5). Consistent with the hyperpolarizing abilities of amlodipine, there was a 2-fold decrease in the MIC of kanamycin (Fig. S3).


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.


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 Na2HPO4, 60 mM NaH2PO4, 10 mM glucose, 5 mM KCl, and 0.5 mM MgCl2 (36). NaOH was used to adjust the pH of the resuspension buffer to 7.0. 10× PBS consisted of 1.3 M NaCl, 70 mM Na2HPO4, and 30 mM NaH2PO4, and the pH was adjusted to 7.0. All buffers were sterilized using a 0.22-μm vacuum filter.

Culturing cells.

E. coli 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.

Fluorescence microscopy.

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+ or K+. 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 and B 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/zF ln([K+]out/[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).

Dose-response curves.

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.

Statistical analyses.

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 + 3σneg)/(|μpos – μneg|), 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.

Supplemental Material

File (aac.00910-20-s0001.pdf)
ASM does not own the copyrights to Supplemental Material that may be linked to, or accessed through, an article. The authors have granted ASM a non-exclusive, world-wide license to publish the Supplemental Material files. Please contact the corresponding author directly for reuse.


Mitchell P, Moyle J. 1967. Chemiosmotic hypothesis of oxidative phosphorylation. Nature 213:137–139.
Szmelcman S, Adler J. 1976. Change in membrane potential during bacterial chemotaxis. Proc Natl Acad Sci U S A 73:4387–4391.
Peterkofsky A, Gazdar C. 1979. Escherichia coli adenylate cyclase complex: regulation by the proton electrochemical gradient. Proc Natl Acad Sci U S A 76:1099–1103.
Mogi T, Anraku Y. 1984. Mechanism of proline transport in Escherichia coli K12. III. Inhibition of membrane potential-driven proline transport by syn-coupled ions and evidence for symmetrical transition states of the 2H+/proline symport carrier. J Biol Chem 259:7802–7806.
Taber HW, Mueller JP, Miller PF, Arrow AS. 1987. Bacterial uptake of aminoglycoside antibiotics. Microbiol Rev 51:439–457.
Strahl H, Hamoen LW. 2010. Membrane potential is important for bacterial cell division. Proc Natl Acad Sci U S A 107:12281–12286.
Stratford JP, Edwards CLA, Ghanshyam MJ, Malyshev D, Delise MA, Hayashi Y, Asally M. 2019. Electrically induced bacterial membrane-potential dynamics correspond to cellular proliferation capacity. Proc Natl Acad Sci U S A 116:9552–9557.
Prindle A, Liu J, Asally M, Ly S, Garcia-Ojalvo J, Süel GM. 2015. Ion channels enable electrical communication in bacterial communities. Nature 527:59–63.
Zhang H, Pan Y, Hu L, Hudson MA, Hofstetter KS, Xu Z, Rong M, Wang Z, Prasad BVV, Lockless SW, Chiu W, Zhou M. 2020. TrkA undergoes a tetramer-to-dimer conversion to open TrkH which enables changes in membrane potential. Nat Commun 11:547.
Yang CY, Bialecka-Fornal M, Weatherwax C, Larkin JW, Prindle A, Liu J, Garcia-Ojalvo J, Süel GM. 2020. Encoding membrane-potential-based memory within a microbial community. Cell Syst 10:417–423.
Damper PD, Epstein W. 1981. Role of the membrane potential in bacterial resistance to aminoglycoside antibiotics. Antimicrob Agents Chemother 20:803–808.
Peng B, Bin Su Y, Li H, Han Y, Guo C, Tian YM, Peng XX. 2015. Exogenous alanine and/or glucose plus kanamycin kills antibiotic-resistant bacteria. Cell Metab 21:249–262.
Silverman JA, Perlmutter NG, Shapiro HM. 2003. Correlation of daptomycin bactericidal activity and membrane depolarization in Staphylococcus aureus. Antimicrob Agents Chemother 47:2538–2544.
Park YK, Ko KS. 2015. Effect of carbonyl cyanide 3-chlorophenylhydrazone (CCCP) on killing Acinetobacter baumannii by colistin. J Microbiol 53:53–59.
Osei Sekyere J, Amoako DG. 2017. Carbonyl cyanide m-chlorophenylhydrazine (CCCP) reverses resistance to colistin, but not to carbapenems and tigecycline in multidrug-resistant Enterobacteriaceae. Front Microbiol 8:228.
Farha MA, Verschoor CP, Bowdish D, Brown ED. 2013. Collapsing the proton motive force to identify synergistic combinations against staphylococcus aureus. Chem Biol 20:1168–1178.
McAuley S, Huynh A, Czarny TL, Brown ED, Nodwell JR. 2018. Membrane activity profiling of small molecule: B. subtilis growth inhibitors utilizing novel duel-dye fluorescence assay. Medchemcomm 9:554–561.
CDC. 2013. Antibiotic resistance threats in the United States, 2013. Centers for Disease Control and Prevention, Atlanta, GA.
Novo D, Perlmutter NG, Hunt RH, Shapiro HM. 1999. Accurate flow cytometric membrane potential measurement in bacteria using diethyloxacarbocyanine and a ratiometric technique. Cytometry 35:55–63.
Shapiro HM. 2000. Membrane potential estimation by flow cytometry. Methods 21:271–279.
Te Winkel JD, Gray DA, Seistrup KH, Hamoen LW, Strahl H. 2016. Analysis of antimicrobial-triggered membrane depolarization using voltage sensitive dyes. Front Cell Dev Biol 4:29.
Tempelaars MH, Rodrigues S, Abee T. 2011. Comparative analysis of antimicrobial activities of valinomycin and cereulide, the Bacillus cereus emetic toxin. Appl Environ Microbiol 77:2755–2762.
Parsons JB, Yao J, Frank MW, Jackson P, Rock CO. 2012. Membrane disruption by antimicrobial fatty acids releases low-molecular-weight proteins from staphylococcus aureus. J Bacteriol 194:5294–5304.
Yang M, Jalloh AS, Wei W, Zhao J, Wu P, Chen PR. 2014. Biocompatible click chemistry enabled compartment-specific pH measurement inside E. coli. Nat Commun 5:4981.
Dumas E, Gao C, Suffern D, Bradforth SE, Dimitrijevic NM, Nadeau JL. 2010. Interfacial charge transfer between CdTe quantum dots and gram negative vs gram positive bacteria. Environ Sci Technol 44:1464–1470.
Kirchhoff C, Cypionka H. 2017. Boosted membrane potential as bioenergetic response to anoxia in Dinoroseobacter shibae. Front Microbiol 8:695.
Zhang L, Dhillon P, Yan H, Farmer S, Hancock R. 2000. Interactions of bacterial cationic peptide antibiotics with outer and cytoplasmic membranes of Pseudomonas aeruginosa. Antimicrob Agents Chemother 44:3317–3321.
Wu M, Maier E, Benz R, Hancock R. 1999. Mechanism of interaction of different classes of cationic antimicrobial peptides with planar bilayers and with the cytoplasmic membrane of Escherichia coli. Biochemistry 38:7235–7242.
Farha MA, Brown ED. 2010. Chemical probes of Escherichia coli uncovered through chemical-chemical interaction profiling with compounds of known biological activity. Chem Biol 17:852–862.
Ahmed S, Booth IR. 1983. The use of valinomycin, nigericin and trichlorocarbanilide in control of the protonmotive force in Escherichia coli cells. Biochem J 212:105–112.
Zhang JH, Chung TD, Oldenburg KR. 1999. A simple statistical parameter for use in evaluation and validation of high throughput screening assays. J Biomol Screen 4:67–73.
Mitchell WJ, Booth IR, Hamilton WA. 1979. Quantitative analysis of proton-linked transport systems. Glutamate transport in Staphylococcus aureus. Biochem J 184:441–449.
Leive L. 1968. Studies on the permeability change produced in coliform bacteria by ethylenediaminetetraacetate. J Biol Chem 243:2373–2380.
Pavlasova E, Harold FM. 1969. Energy coupling in the transport of beta-galactosides by Escherichia coli: effect of proton conductors. J Bacteriol 98:198–204.
West I, Mitchell P. 1972. Proton-coupled beta-galactoside translocation in non-metabolizing Escherichia coli. J Bioenerg 3:445–462.
Gentry DR, Wilding I, Johnson JM, Chen D, Remlinger K, Richards C, Neill S, Zalacain M, Rittenhouse SF, Gwynn MN. 2010. A rapid microtiter plate assay for measuring the effect of compounds on Staphylococcus aureus membrane potential. J Microbiol Methods 83:254–256.
Datsenko KA, Wanner BL. 2000. One-step inactivation of chromosomal genes in Escherichia coli K-12 using PCR products. Proc Natl Acad Sci U S A 97:6640–6645.

Information & Contributors


Published In

cover image Antimicrobial Agents and Chemotherapy
Antimicrobial Agents and Chemotherapy
Volume 64Number 920 August 2020
eLocator: 10.1128/aac.00910-20


Received: 20 May 2020
Returned for modification: 19 June 2020
Accepted: 30 June 2020
Published online: 20 August 2020


Request permissions for this article.


  1. CCCP
  2. DiOC2(3)
  3. E. coli
  4. Gram-negative
  5. amlodipine
  6. antimycin
  7. barium chloride
  8. high throughput
  9. membrane potential
  10. valinomycin



Department of Biology, Texas A&M University, College Station, Texas, USA
Department of Biology, Texas A&M University, College Station, Texas, USA
Department of Biology, Texas A&M University, College Station, Texas, USA


Address correspondence to Steve W. Lockless, [email protected].

Metrics & Citations


Note: There is a 3- to 4-day delay in article usage, so article usage will not appear immediately after publication.

Citation counts come from the Crossref Cited by service.


If you have the appropriate software installed, you can download article citation data to the citation manager of your choice. Simply select your manager software from the list below and click Download.

View Options

Figures and Media






Share the article link

Share with email

Email a colleague

Share on social media

American Society for Microbiology ("ASM") is committed to maintaining your confidence and trust with respect to the information we collect from you on websites owned and operated by ASM ("ASM Web Sites") and other sources. This Privacy Policy sets forth the information we collect about you, how we use this information and the choices you have about how we use such information.
FIND OUT MORE about the privacy policy