Electrochemical Control with High Spatiotemporal Resolutions for Extracellular pH Microenvironment

Abstract

The local pH values and hence the pH microenvironment surrounding an organism critically affect its metabolism and physiology in both prokaryotes and eukaryotes. Despite the ubiquitous spatiotemporal heterogeneity of local pH conditions in biology, a biocompatible method that spatiotemporally programs the pH microenvironment remains elusive. Here we report an electrochemical approach that spatiotemporally controls pH microenvironments for potential microbial applications. A quinone-based redox couple undergoes electrochemical proton-coupled electron transfer (PCET) that releases or consumes protons and hence changes the local pH. We designed a biocompatible hydroquinone/quinone redox couple that has minimal interference with bacterial aerobic respiration thanks to its hydrophilic nature and the specially targeted redox potential. Deploying such a uniquely designed redox couple with interdigitated electrodes in a microfluidic device leads to electrochemical control of the pH microenvironment with fast temporal modulation (~10¹ s) and high spatial resolution (~10⁰ μm) suitable for biological applications. The concept of deploying biocompatible electrochemistry offers a new venue toward perturbing biological microenvironments at the cellular if not subcellular level.

Graphical Abstract

INTRODUCTION

The extracellular pH environment surrounding an organism that measures the local availability of proton is essential for every aspect of biological physiology and metabolism.^1–3 Oxidative phosphorylation, the most important route of ATP generation in aerobic respiration for most of organisms on this planet,^4,5 relies on a suitable proton motive force and local proton availability for transmembrane proton translocation.⁶ Yet the biochemical metabolism itself also affects the local extracellular pH and hence the pH microenvironments in the vicinity of the organisms. The Warburg effect in cancerous cells leads to locally acidic conditions in tumor tissues.⁷ Fermentative respiration commonly secretes organic acid that also changes the local pH, whose values typically ranges between 5 and 7 (Figure 1a),^8–10 especially in the extracellular space of biofilms.^11–13 From the perspective of bioenergetics, the transfers of protons and electrons, hence energy, are so ubiquitously coupled that there are hardly any biological phenomena completely independent of local pH conditions. The close interplay between organisms and their microenvironments leads to ubiquitous spatiotemporal heterogeneities of pH microenvironments in nature. Tools that characterize or perturb such biological pH microenvironments are critical for our advanced understanding of various prokaryotic and eukaryotic systems.

Figure 1.

The design of an electrochemical PCET reaction for extracellular pH modulation. (a) The spatial pH profiles observable in bacterial systems, as an example of the ubiquitous pH heterogeneity in biology. (b) The concept of generating pH gradients via electrochemical proton-couple electron transfer (PCET) reactions. (c) Schematic of electrochemically generated pH gradient via a PCET-based redox couple of quinone (Q) and hydroquinone (H₂Q), facilitated by interdigitated electrodes (IDEs) potentially in a microfluidic device. E_ox and E_red, the electrochemical potentials on the oxidizing and reducing electrodes, respectively. (d) Latimer diagram at pH = 7.0 for the design of hydroquinone/quinone derivatives given the redox potentials (E°) in the O₂ chemistry and the individual components of the membrane-bound electron transport chain (ETC). This work tested two candidate hydroquinone derivatives, 2,5-dihydroxybenzenesulfonate potassium (DHBS) and 4,5-dihydroxybenzene-1,3-disulfonic acid (BQDS). At pH = 7.0, E_RHE (V) vs. RHE = E_SHE (V) s. SHE – 0.413 V. RHE, reversible hydrogen electrode; SHE, standard hydrogen electrode.

The capability of spatiotemporally perturbing pH microenvironments for biological applications remains limited. Extensive efforts have been devoted towards the characterization of biological pH microenvironments particularly via fluorescent microscopy.^11,14–17 Yet there are only limited means for researchers to realize spatiotemporal perturbation toward extracellular pH at the micrometer-scale level, particularly under experimental conditions commensurate with optical microscopy. Microfluidic systems create spatial pH gradients by mixing laminar fluidic flows of different pHs,^18–21 but the spatial resolution is limited to about 10² μm^18–21 and takes at least several minutes (~10² s) for a temporal perturbation to be stably achieved.²⁰ Such a spatiotemporal resolution is unsatisfying, especially given the fledging interests in physiology and metabolism under single-cell if not subcellular levels.^22–24 The potential fast modulation of local pH within seconds if not at the subsecond level also opens up new lines of inquiry for the study of proton translocation across cellular membrane and oxidative phosphorylation that were not possible otherwise.²⁵ As the cellular membrane potential is closely related to the local pH in the proximity of organisms, fast dynamic change of local pH offers a valuable perturbation tool to study the biomachinery related to proton translocation without the potential interference from any transcriptional and post-transcriptional responses. It is necessary to develop tools that perturb pH extracellular microenvironments faster with a smaller spatial resolution.

We hypothesize that the biocompatible electrochemistry of proton-coupled electron transfer (PCET) in the extracellular space offers a new venue of programmable pH microenvironments for both prokaryotic and eukaryotic organisms. The electrochemical PCET reaction^26,27 consumes or produces protons in an electrode’s vicinity, which will lead to a diffusion-based microscopic pH gradients in weakly buffered aqueous solutions near neutral pH (Figure 1b). As we recently demonstrated for the microenvironments of O₂ and reactive oxygen species,^28,29 the spatial profile of chemical microenvironments can be programmed with high spatial resolution (10⁰ μm), potentially via inverse design based on machine learning,²⁹ by fine-tuning electrodes’ morphology and spatial arrangement. Meanwhile, not only does the magnitude of applied electrochemical potential E_appl provide one more degree of freedom for concentration’s spatial profile. The temporal sequence of E_appl controls the microenvironments in a time-dependent manner with a time resolution of seconds if not tens of seconds (10⁰–10¹ s).^29–31 Therefore, in a microfluidic device equipped with a predesigned electrochemical configuration such as interdigitated electrodes (IDEs), a programmable electrochemical PCET process is proposed to reinforce our limited capability of controlling extracellular pH microenvironments with spatiotemporal resolutions at single-cell if not subcellular levels (Figure 1c). Although electrochemically driven pH gradients have been reported,^32,33 the deployment of the PCET reaction to facilitate higher spatial resolutions has yet been tested experimentally despite its attractiveness and the apparent paucity of tools for extracellular pH perturbation.

One major challenge in fulfilling such a tantalizing concept is the biocompatibility and orthogonality of the pH-controlling electrochemical PCET process. As we aim to electrochemically perturb only the extracellular pH microenvironments, the electrochemical PECT reaction ought to be not only compatible with the biological moieties but also induce no undesired cellular responses, particularly under aerobic/microaerobic conditions when the cellular bioenergetic is sensitive to the extracellular pH and hence the proton motive force.^5,6 Our strategy to address this challenge is to design an electrochemical PCET reaction with two unique properties: First, we will deploy highly hydrophilic molecules of the PCET reaction that have minimal physical interactions with cellular structures and do not penetrate through the cellular membrane; second, the redox species are chosen so that the corresponding redox potential does not interfere much if not any either with the dissolved O₂ or with the cellular electron transport chain (ETC)³⁴ (Figure 1d), in order to minimize the potential formation of reactive oxygen species (ROS) and the potential disruption of membrane-bound oxidative phosphorylation. We surmise such a design strategy will minimize the interaction between the redox molecule and organisms and hence reduce the chances of potential alternation of cellular metabolism and physiology. In this context, this work reports our progress toward a biocompatible PCET reaction for the extracellular modulation of extracellular pH microenvironments.

Here we report the design of a hydroquinone/quinone PCET redox couple, proposed to be compatible and orthogonal with biological moieties, that serves to electrochemically program pH microenvironments spatiotemporally. Thanks to its hydrophilic nature and suitable redox potential, 2,5-dihydroxybenzenesulfonate potassium (DHBS) as the hydroquinone component is found to have minimal interference with the growth of the opportunistic pathogen Pseudomonas aeruginosa^35,36 and allows for electrochemical modulation of local pH via the two-electron PCET process. Tested in an IDE-equipped microfluidic channel, we demonstrate biocompatible electrochemical programming of spatiotemporal pH profiles in solution with spatial and temporal resolutions (about 10⁰ μm and 10¹ s), unachieved in other pH-controlling microfluidic devices. Such results showcase the feasibility of controlling pH concentration profiles with biocompatible PCET reactions, creating opportunities of future research to perturb extracellular pH microenvironments at single-cell if not subcellular levels.

RESULTS AND DISCUSSION

Design Considerations for a Biocompatible PCET Reaction Based on Quinone Derivatives.

Three criteria were implemented when designing the PCET process for electrochemical pH perturbation in extracellular space. First, because an electrochemical PCET process relies on the stoichiometric consumption or generation of protons to change local pH, the PCET redox couples ought to have high solubilities that at least are similar to the buffer concentration in the biological medium (usually 10⁰–10¹ mM). This led us to consider the redox couples based on synthetic hydroquinone/quinone chemistry, which undergo bidirectional PCET processes in water^37,38 and are known to have high aqueous solubilities thanks to the surging interests in aqueous redox flow batteries.^39–41 Second, as we propose to entail minimal interference between the redox couples and organisms, hydroquinone/quinone derivatives with highly negatively charged functional groups (e.g., sulfonate group) were favored. Not only will those negatively charged hydroquinone/quinone derivatives entail electrostatic repulsive interactions from the negatively charged cellular surface,^42–44 these hydrophilic molecules are also considered incapable of permeating through the cellular membranes for any alternation of metabolisms in the cytosol.

The third criterion for the PCET redox couple is a suitable redox potential that minimizes unwarranted side reactions with dissolved O₂ and the membrane-bound ETC in aerobic respiration (Figure 1d). Hydroquinone, the two-electron, two-proton reduction product from quinone, is known to reduce O₂ into H₂O₂,⁴⁵ a potent ROS species, when thermodynamically its redox potential E° is more negative than the redox potential of O₂ reduction H₂O₂ ($E{°}_{{\text{O}}_2/{\text{H}}_2{\text{O}}_2}$ in Figure 1d).⁴⁶ Therefore, the E° of hydroquinone/quinone redox that suits our design should be more positive than $E{°}_{{\text{O}}_2/{\text{H}}_2{\text{O}}_2}$. Meanwhile, E° should be more negative than the redox potential of the water oxidation reaction ($E{°}_{{\text{O}}_2/{\text{H}}_2\text{O}}$ in Figure 1d), in order to avoid water oxidation from the oxidized quinone species. Last, the redox’s interaction with the membrane-bound ETC should be mitigated by a judicious selection of hydroquinone/quinone’s E°. We conjecture that there will be minimal interaction if the E° is more positive than that redox species in Complex IV of ETC, the cytochrome a and a₃ (cyt a and cyt a₃, respectively, in Figure 1d),³⁴ when the electron and hence energy flux in the ETC will not be altered had the hydroquinone/quinone redox underwent electron transfers with the membrane bound ETC. Overall, although the concept of electrochemical pH modulation sounds straightforward with seemingly a large amount of candidate hydroquinone/quinone derivatives, the multiple criteria from various considerations indeed render a limited range of options for the PECT redox couple.

Under the aforementioned criteria, two hydroquinone derivatives, 5-dihydroxybenzenesulfonate potassium (DHBS) and 4,5-dihydroxybenzene-1,3-disulfonic acid (BQDS) (Figure 1d), both negatively charged at almost all pH ranges in aqueous solution, were chosen as the hydroquinone components for the candidate redox couple. Both molecules are commercially available and have been developed for the application of aqueous redox flow batteries thanks to their stability and high aqueous solubility (up to 0.7 and 0.3 M).⁴¹ The negative charge and electron-withdrawing nature of the sulfonic group are proposed to yield minimized cellular interactions and suitable redox potentials of 0.9 and 1.1 V vs. a reversible hydrogen electrode (RHE) at pH = 7.0 for DHBS and BQDS, respectively (Figure 1d).^39,41 As will be shown below, both DHBS and BQDS have been tested, while DHBS was found to be more appropriate for electrochemically controlled pH microenvironments.

Electrochemical Characterizations for Two Hydroquinone/Quinone Redox Couples.

Experiments of cyclic voltammetry on glassy carbon, platinum (Pt), and gold (Au) electrodes (see Methods) were conducted in 1× phosphate-buffered saline (PBS) solution (pH = 7.2, medium 1.1 in Table S1) to examine the electrochemical PCET processes for 5 mM of BQDS (Figure 2a) and DHBS (Figure 2b). Bidirectional PCET-based reduction and oxidation were observed for both quinone derivatives on all electrode materials, while kinetically faster for DHBS based on the smaller potential differences between the better-defined cathodic and anodic peaks. Estimated based on mid-point potentials, the measured E° of BQDS and DHBS are 0.9 and 1.1 V vs. RHE, respectively, consistent with literature values of 0.88 and 1.10 V vs. RHE reported in 0.5 M phosphate buffer.^39,41

Figure 2.

Electrochemical characterizations of BQDS and DHBS. (a) and (b) Cyclic voltammograms for BQDS (a) and DHBS (b) at 1× phosphate-buffered saline solution (PBS, pH = 7.2, medium 1.1 in Table S1) with glassy carbon, platinum (Pt), and gold (Au) working electrodes. (c) and (d) the current densities of cyclic voltammograms for the anodic and cathodic peaks (i_p) against the square root of scan rate v (v^1/2) for BQDS (c) and DHBS (d). (e) and (f) the inverse of oxidative current (1/I) on Au rotating disk electrode (RDE) against the inverse of square root of the angular rotation rate (ω^–1/2) for BQDS (e, at 1.5 V vs. RHE) and DHBS (f, at 1.45 V vs. RHE) in 10 mM phosphate buffer (medium 1.2 in Table S1) The electron transfer number n determined from the Koutecký–Levich equation is 1.06 and 2.10 for BQDS and DHBS, respectively. All data were iR corrected in N₂ with a AgCl/Ag (1 M KCl) reference and Au mesh counter.

Plotting the current densities i_p of anodic and cathodic peaks when available against the square roots of scan rates (v^1/2) in voltammetry leads to linear relationships for all electrode materials for both BQDS (Figure 2c) and DHBS (Figure 2d). This indicates diffusion-based redox behavior and minimal adsorption, thanks to the highly negatively charged nature of those hydroquinone/quinone derivatives. The observation that the hydrophobic glassy carbon, analogous to the lipid bilayer of cellular membranes, does not induce any observable surface adsorption in Figure 2c and d is supportive of our design that a hydrophilic, negatively charged quinone will incur muted interaction with biological moieties in solution.

Experiments of a rotating disk electrode (RDE) in a Au electrode (Figure 2e,f and Supplementary Figure S1; see Methods, medium 1.2 in Supplementary Table S1) were conducted for both BQDS and DHBS in order to extract the number of electrons transferred during the redox phenomena in Figure 2a and b. At 1.5 V vs. RHE, the inverse of the oxidative current (1/I) for the reduced BQDS is plotted against the inverse of the square root of the angular rotation rate (1/ω^1/2) (Figure 2e) for the analysis based on the Koutecký–Levich equation.^47,48 Based on a diffusion coefficient of 2.7 × 10⁻⁶ cm²/s,^48,49 the number of electron transfer (n) is calculated as 1.06. In contrast, the same analysis for DHBS at 1.45 V vs. RHE (Figure 2f) yields n = 2.10 based on a diffusion coefficient of 4.28 × 10⁻⁶ cm²/s.⁵⁰ These results show that BQDS at neutral pH undergoes a one-electron, one-proton electrochemical process with a semiquinone radical as the oxidized species, while DHBS at the same condition undergoes a targeted two-electron, two-proton one between the hydroquinone and quinone redox species.³⁷ Such results are consistent with the faster PCET kinetics for DHBS observed by cyclic voltammetry (Figures 1a and 2b). Compared to BQDS’s semiquinone radical as the oxidized counterpart in the PCET process, the electrochemical hydroquinone/quinone interconversion of DHBS is considered less prone to side reactions.

Biocompatibility Assay for Two Hydroquinone/Quinone Redox Couples.

Biocompatibility tests were conducted by observing the effects of either BQDS or DHBS towards the aerobic growth of Pseudomonas aeruginosa (see Methods), a Gram-negative, opportunistic bacterial pathogen³⁵ demonstrating extensive pH heterogeneities (pH ∈ [5.5, 7.2]) within its biofilms.^16,51 Two different strains of P. aeruginosa, moderately virulent PAO1^36,52 and hyper-virulent PA14,³⁶ were both tested with biological triplicates (n = 3). In Figure 3a, the optical density at 600 nm (OD₆₀₀), a surrogate of cell density, is plotted as a function of time for P. aeruginosa PAO1 in phosphate-buffered growth medium (pH = 7.2, medium 2.1 in Table S2) in the absence of any organic substrate (yellow), 20 mM BQDS (dark blue), 20 mM DHBS (light blue), or 30 mM succinate (red). Since appreciable bacterial growth was only observed with the addition of succinate, neither BQDS nor DHBS serves as organic substrate to support cellular growth. There is no significant impact on bacterial growth when the phosphate buffer concentration of the medium is within the range of 10 to 43 mM (Figure 3b), suggesting that the buffer capability of the bacterial culture solution may not be critical within a certain concentration range.

Figure 3.

Aerobic growth of Pseudomonas aeruginosa in the presence of BQDS and DHBS. (a) Aerobic growth curves for P. aeruginosa PAO1 in phosphate-buffered growth medium (medium 2.1 in Table S2) with the addition of succinate (red), BQDS (dark blue), DHBS (light blue), and no addition of any organic sources (yellow). OD₆₀₀, optical density at 600 nm. (b) Aerobic growth curves for P. aeruginosa PAO1 under different concentration of phosphate buffer (C_buffer) adapted from the growth medium (medium 2.1 in Table S2). (c) and (d) Aerobic growth curves for P. aeruginosa PAO1 under different concentrations of BQDS (c) and DHBS (d) along with the abiotic controls (medium 2.2 in Table S2). (e) and (f) Aerobic growth curves and the corresponding doubling times in two different pH conditions (pH = 5.0 and 7.0) for P. aeruginosa strains PAO1 (e) and PA14 (f) in growth mediums buffered by phosphate and 3-(N-morpholino) propanesulfonic acid (MOPS) (Medium 2.2 in Table S2 and medium 3.1 in Table S3), respectively. Biological triplicates (n = 3). Error bars represent standard deviations. p values calculated via a two-sample unequal variance t-test.

The addition of BQDS or DHBS up to 15 mM did not significantly hinder the growth of P. aeruginosa PAO1 (Figure 3, panels c and d, respectively), at 40-mM succinate growth medium with 10 mM phosphate buffer (pH = 7.2, medium 2.2 in Supplementary Table S2). Indeed, in the case of DHBS, the minimum inhibition concentration, defined as the lowest concentration of a chemical that will inhibit the visible growth of an organism,⁵³ was estimated to be between 128 to 256 mM based on bacterial growth in a 96-well plate (Supplementary Figure S2, see Methods, medium 2.2 in Supplementary Table S2); the half-maximal inhibitory concentration (IC₅₀), defined as the concentration of inhibitor necessary to halve the reaction rate observed under specified assay condition,⁵⁴ was estimated to be about 100 mM based on spot assay on the agar plate (Supplementary Figure S3, see Methods, medium 2.2 in Supplementary Table S2). These results suggest that bacterial growth and hence cellular metabolism is not significantly perturbed by the added hydroquinones, particularly DHBS, until the hydroquinone’s concentration reaches about 100 mM.

As the hydroquinone/quinone redox is designed to alter the extracellular pH microenvironments, the biocompatibilities of BQDS and DHBS were further tested at different pH conditions by evaluating the doubling times based on the growth curves of PAO1 and PA14 strains (see Methods, medium 2.2 in Supplementary Table S2 and medium 3.1 in Supplementary Table S3, respectively). As the extracellular pH for biofilms of P. aeruginosa typically ranges roughly between 5 and 7,^10,51 the bacterial doubling times at pH = 5.0 and 7.0 were both investigated. For the PAO1 strain, the growth curves and doubling times with the addition of 15 mM BQDS or 15 mM DHBS were compared with the controls of bare succinate growth medium (Figure. 3e; medium 2.2 in Supplementary Table S2). There are no statistically significant differences (p value > 0.05, two-sample unequal variance t-test) in terms of doubling time with the addition of BQDS or DHBS at both conditions of pH = 5.0 and 7.0, although in general P. aeruginosa grows more slowly at the suboptimal condition of pH = 5.0. Similar experiments were conducted for the PA14 strain in the growth medium of 10 mM 3-(N-morpholino)-propanesulfonic acid (MOPS) buffer (see Methods, medium 3.1 in Supplementary Table S3) (Figure 3f). For the PA14 strain, there are no statistically significant differences (p value > 0.05, two-sample unequal variance t-test) in terms of doubling time with the addition of BQDS or DHBS at both conditions of pH = 5.0 and 7.0, notwithstanding the condition of pH = 5.0 with the addition of BQDS (p value = 0.001). This indicates that in acidic medium, bacterial growth was inhibited by BQDS, potentially owing to its one-electron PCET process that yields reactive semiquinone radicals. Nonetheless, overall there is minimal interference from the hydroquinone/quinone species, particularly for DHBS, towards the aerobic growth of P. aeruginosa at different pH conditions, reinforcing our hypothesis that a highly negatively charged hydroquinone/quinone redox couple with suitable electrode potential will be orthogonal towards cellular metabolism.

The Experimental Design for Electrochemically Controlled pH Microenvironments.

We chose to deploy a DHBS-based redox couple for the experimental demonstration of electrochemical pH perturbation. Our electrochemical characterizations indicate that at neutral pH, DHBS undergoes the presumed two-electron, two-proton PCET process between the hydroquinone and quinone forms with relatively fast reaction rates, despite the challenging kinetics for stepwise PCET at neutral pH based on the kinetic model developed by E. Laviron.⁵⁵ Moreover, while BQDS and DHBS are both appreciably noninterfering toward the aerobic growth of P. aeruginosa, our evaluation of doubling times suggests that DHBS does not hinder bacterial growth even at suboptimal pH = 5.0, a condition when BQDS may be less innocent. The BQDS’s noninnocence toward bacterial growth is putatively due to the generated semiquinone from an O₂-driven one-electron, one-proton oxidation, whose radical nature poses BQDS a less benign redox couple for biological systems.

A microfluidic device of Au IDEs under a constant flow rate was constructed under a fluorescent confocal microscope to demonstrate the proposed electrochemical control of pH microenvironments (Figure 4a,b and Supporting Information, respectively; see Supporting Information, Note S1 for the rationale of choosing Au as electrode materials). Two pairs of Au finger electrodes, each of 20-μm width with a spacing of either 10 or 50 μm, were designated as two working electrodes in a bipotentiostat configuration with a Ag pseudo-reference electrode and indium–tin oxide counter electrode (Figure 4b, see Methods). As schematically depicted in Figure 1c, the anodic working electrode with potential E_ox oxidizes hydroquinone (H₂Q) into quinone (Q) with the release of two protons, while the cathodic working electrode with potential E_red reduces quinone to hydroquinone with the consumption of two protons. The diffusion of chemical species between the electrode pairs is expected to yield a lower pH near the oxidizing electrode and a higher pH near the reducing one, hence, a one-dimensional pH gradient normal to the individual finger electrodes.

Figure 4.

Steady-state pH gradient generated by DHBS-based electrochemistry in biological mediums. (a) and (b) pH-sensing ratiometric fluorescent probe seminaphthorhodafluor-4F 5-(and-6) carboxylic acid (C-SNARF-4) (a) was deployed for the quantification of a pH gradient in a microfluidic channel equipped with Au IDEs (b), whose two working electrodes (W.E. 1 and W.E. 2) electrochemically oxidize and reduce the DHBS redox couple with potential E_ox and E_red, respectively. Indium–tin oxide (ITO), counter electrode (C.E.); Ag-coated ITO, pseudo-reference electrode (R.E.). λ_ex, excitation wavelength; λ_em, emission wavelength. (c) The two-dimensional (2D) mapping about the ratio of fluorescent emissions (653 vs. 592 nm), 5 μm above the electrode surface, overlayed with the locations of W.E. 1 and W.E. 2. d, the distance between W.E. 1 and W.E. 2. The directions normal and parallel to the finger electrodes are defined as the x and y axes, respectively. (d) and (e) one-dimensional (1D) pH gradients against the x coordinate in growth mediums buffered by phosphate (d, medium 2.2 in Supplementary Table S2) and MOPS (e, medium 3.1 in Table S3) under different combinations of E_ox, E_red, and d values. The dots and shaded areas represent the average and standard deviation of the pH values across all y coordinates at the same x coordinates. The red and blue bars represent the positions of the oxidizing and reducing electrodes with potentials E_ox and E_red vs. RHE, respectively. Bulk solution pH = 7.2; fluidic flow rate: 0.1 mL/min. Additional experimental conditions can be found in Methods.

The hypothesized pH gradient, presumably controllable by the E_ox and E_red values, was spatially mapped within the IDE-bearing microfluidic channel by confocal microscope at a spatial resolution of about 650 nm with pH-sensitive ratiometric fluorescent probe seminaphthorhodafluor-4F 5-(and-6) carboxylic acid (C-SNARF-4),^9,11,51 whose ratio of emission intensities at 592 and 653 nm (Figure 4a) reports the local pH after the establishment of a calibration curve (Supplementary Figure S5). Control experiments with C-SNARF-4 showed that within our detection limit the pH probe does not inhibit the PCET processes (Supplementary Figure S6), and the DHBS-based electrochemical process does not degrade the probe’s optical property (Supplementary Figure S7). In order to observe the maximal pH gradients exerted by electrochemistry without significant optical interference from the metallic Au electrodes, the microscope focal plane was fixed at 5 μm above the electrode (see Methods, Note S2). Because the local pH microenvironment near the electrode differs more profoundly from the bulk as the one at 5 μm above the electrode, our experimentally reported pH gradient is an underestimate of the actual pH gradient that many biological systems will indeed experience. While a confocal image and hence the pH spatial profile is two-dimensional (Figure 4c), the aforementioned one-dimensional nature of the presumed pH gradient allows us to present the generated pH spatial profile as a one-dimensional plot by averaging the pH values at the same x coordinate along the direction normal to the finger electrodes.

Spatiotemporal Programming of pH Microenvironments by DHBS-Based Electrochemistry.

Steady-state pH gradients programmed by E_appl values were demonstrated in two of the most commonly used biological buffer solutions, phosphate-buffered and MOPS-buffered growth medium (Figure 4d,e; medium 2.2 and 3.1 in Supplementary Tables S2 and S3, respectively). The averaged pH values under different combinations of time-invariant E_appl values (E_ox and E_red), along with the measurement standard deviations, were plotted along the x coordinate normal to the finger electrodes where the location of oxidizing and reducing electrodes was marked as red and blue bars, respectively (Figure 4d,e, see Methods). In both mediums at a flow rate of 0.1 mL/min, pH spatial gradients between the anodic and cathodic electrodes were observed and, as expected, with a higher pH near the anode and a lower pH near the cathodic. An increase of E_ox under a constant E_red will lead to a much larger pH difference and hence a steeper pH gradient; ditto with a decrease of E_red under a constant E_ox. Our observed pH gradients are within the range for the biofilms of P. aeruginosa.⁵¹ The spatially repeated pattens of pH gradients are a testament of the reliability of the generated pH spatial gradients.

Moreover, reducing the distance d between oxidizing and reducing finger electrodes from 50 μm (left half of Figure 4d and e) to 10 μm (right half of Figure 4d and e) leads to more pronounced spatial pH gradients. The largest slope of pH gradient amounts to 0.02 pH/μm (E_ox = 1.47 V, E_red = 0.27 V, d = 10 μm in MOPS medium). Such a large slope of pH gradient can hardly be achieved in conventional methods based on passive diffusion or convection in lieu of active electrochemical modulation in microfluidic channels (Supplementary Table S4).^{18–21,56,57} Our results indicate that the electrochemical PCET process is capable of creating a 12-mV difference in the proton motive force across a distance of 1 μm or 0.12 V across 10 μm. Given the sensitivity of oxidative phosphorylation toward the proton motive force, our tool can pose an appreciable perturbation toward an organism’s aerobic respiration and overall metabolism at the single-cell level for prokaryotes and subcellular level for eukaryotic systems.

The temporal resolution of electrochemically driven pH perturbation was shown to be at 10⁰–10¹ s with good repeatability. One proposed benefit of electrochemically controlled pH microenvironments is the high temporal resolution, as electrochemical processes can be quickly turned on or off with the application or withdrawal of electrochemical potentials. In MOPS medium (medium 3.1 in Supplementary Table S3), we examined the temporal pH response with the provision of E_appl on and off under different periodicities ranging from 2 to 30 min. Given the high-dimensionality of the time-dependent pH gradients (Supplementary Video S1), the local pH values 5 μm above the centers of the reducing (pH_red, top row in Figure 5) and oxidizing electrodes (pH_ox, bottom row in Figure 5) were plotted every 5 s (see Methods). We have observed good repeatability of the established pH microenvironments across all periodicity values (Supplementary Figure S8). Moreover, stable pH values were achieved within about 10 s after the voltage switches, even in cases when the periodicity is only 2 min (Figure 5). Such a fast temporal resolution of pH modulation is at least one order-of-magnitude faster than the ~10² s ones in conventional methods based on passive diffusion or convection (Supplementary Table S4).^18,20,56,57 Thanks to the fast-switching nature of electrochemistry, it is feasible now to perturb the extracellular pH microenvironments faster than the responses of cellular translation and transcription^58–60 at single-cell if not subcellular levels.

Figure 5.

Temporal responses of electrochemically controlled pH microenvironments. The local pH values 5 μm above the centers of the reducing (top, pH_red, “Cathode”) and oxidizing electrodes (bottom, pH_ox, “Anode”) were plotted every 5 s with the provision of E_appl on and off under different periodicities ranging from 2 min to 30 min. When E_appl values were applied, E_red = 0.27 V and E_ox = 1.47 V vs. RHE. 10-mM MOPS growth medium (bulk pH = 7.2; medium 3.1 in Table S3). An exemplary evolution of time-dependent pH gradient when periodicity = 1 min is shown in Supplementary Video S1.

A Simulation-Driven Inquiry into the General Applicability of Electrochemical pH Control.

Additional numerical simulations based on finite-element methods were conducted to inquire into the parameter range compatible with electrochemically controlled pH microenvironments, as we envision the presented tool will be generally applicable toward different biological systems. A two-dimensional model built on COMSOL Multiphysics (version 5.3) was constructed to reflect not only the cross-section of electrochemical microfluidic setup (d = 50 μm) under a constant lamellar flow but also the electrochemical properties of Au IDEs based on our experimental results (left in Figure 6a, see Methods). Same as our experimental evaluation about the temporal resolution of electrochemical pH perturbation (Figure 5), the pH profiles including pH_ox and pH_red were numerically calculated under different conditions (right in Figure 6a, Supplementary Figure S9). As a surrogate of the pH range achievable by electrochemistry, pH difference ΔpH = pH_red – pH_ox was calculated as a function of E_ox, E_red, phosphate buffer concentration (C_buffer), and DHBS concentration (C_DHBS). When E_red = 0.67 V vs RHE (Figure 6b), there exists a wide range of parameter combinations when ΔpH > 0.7, the common pH differences across bacterial biofilms^9,61 and the cancerous tissues.^62–64 This indicates that the presented method of electrochemical pH modulation is amenable to many different extracellular mediums.

Figure 6.

Simulation inquiry into the general applicability of electrochemical pH modulation. (a) Numerical simulation based on a finite-element method was implemented to model the cross-sectional pH profiles near the IDEs in the microfluidic channel. pH_ox and pH_ed, pH values 5 μm above the centers of the oxidizing and reducing electrodes under E_ox and E_red (W.E. 1 and W.E. 2), respectively. (b) ΔpH, the difference between pH_red and pH_ox, is calculated in PBS solution as a function of C_buffer, E_ox, and C_DHBS. d = 50 μm; E_red = 0.67 V vs RHE; flow rate, 0.1 mL/min.

Generally, larger value of ΔpH is calculated under a relatively smaller C_buffer, larger C_DHBS, and larger E_ox; meanwhile, E_red poses a much smaller impact towards ΔpH (Supplementary Figure S10), and a smaller d value tends to reduce the magnitude of ΔpH (Supplementary Figure S11), although boosting the slope of pH gradient (Figure 4d and e). A smaller ΔpH is expected with a smaller d value because it is harder to maintain a pH difference between IDEs of a closer distance. When the solution buffering capability decreases (smaller C_buffer), or there are more protons released/consumed from hydroquinone/quinone redox (larger C_DHBS), the electrochemical PCET process will pose a much larger perturbation towards the extracellular pH (Supplementary Figure S12). Because ambiently the predominant species for DHBS is the reduced hydroquinone form owing to its positive redox potential (0.9 V vs RHE from Figure 2b), a change of E_ox value will lead to a significant change of local proton releases from hydroquinone oxidation, while a relatively mild change of proton consumption will incur from the change of the E_red value. As we know now that not every parameter is equally important toward maximizing ΔpH, such insights will benefit future designs of electrochemical pH microenvironments for biological appliations.

CONCLUSION

While pH microenvironments are ubiquitous in biological systems, spatiotemporal perturbation of local pH conditions for biological application has been rare and challenging to achieve. The rarity of reported spatiotemporal pH modulation resides in the lack of effective methods that change local pH environments with high biocompatibility and good spatial/temporal resolution. Based on the equally ubiquitous PCET nature of electrochemical processes, this work reports the feasibility of using electrochemical PCET reactions from biocompatible quinone/hydroquinone redox couple to generate biologically relevant pH gradients in bacterial growth medium with a steeper spatial gradient and faster temporal response. As the extent of the PCET process can be controlled by the applied electrochemical potential, the generated spatiotemporal pH profiles are indeed digitally programmed by electric signals. Deploying biocompatible electrochemistry with judicious device design offers a venue to transduce digital signals into spatiotemporal profiles of local pH and hence a larger degree of freedom to perturb and investigate microbial systems under heterogeneous microenvironments in the future.

METHODS

More detailed experimental procedures are documented in the Supplementary Information. In general, electrochemical characterizations were conducted in a H-cell with a three-electrode setup with the electrolyte of 1× PBS solution (Tables S1 and S2) and 5 or 20 mM of BQDS or DHBS. An Au mesh was used as the counter electrode, and an Ag/AgCl (1M KCl) electrode was used as the reference electrode. The redox potential of the Ag/AgCl (1M KCl) electrode was calibrated by a Zobell’s solution⁶⁵ with a redox potential of 227 mV vs. RHE. The PAO1 and PA14 strain of P. aeruginosa was obtained from Dianne Newman’s lab at Caltech. The bacteria were cultured at 30 °C in glassy tubes for all of the growth curve measurements if not mentioned specially. Two buffer media were used: phosphate-buffered and MOPS succinate mediums. The phosphate-buffered succinate medium (Table S2) includes 4.15 mM KH₂PO₄, 38.85 mM K₂HPO₄, 42.8 mM NaCl, 9.3 mM NH₄Cl, 40 mM sodium succinate, 1 mM MgSO₄, and 1 mL/L 1× SL-10 trace element solution. The MOPS succinate medium (Table S3) includes 100 mM MOPS, 50 mM NH₄Cl, 3.7 mM KH₂PO₄, 1 mM MgSO₄, 40 mM sodium succinate, and 7.2 μM FeSO₄. The pH was adjusted to 7.2 using HCl or NaOH. Absorbance at 600 nm (OD₆₀₀) was measured using an Agilent Cary 60 UV-Vis spectrophotometer. Each condition was tested in triplicates. Biocompatibility was determined via monitoring culture OD₆₀₀ values under different medium compositions every 2–8 h, depending on the stages of bacterial growth, for 2 days to collect the growth curve. In the determination of the minimum inhibition concentration and IC₅₀, serial dilution of bacterial culture at ratios of 1/10, 1/10², 1/10², 1/10³, 1/10⁴, and 1/10⁵ were conducted, and bacterial growth curves were obtained by monitoring the OD₆₀₀ values every 20 min for 24 h.

A three-electrode electrochemical system was established in a microfluidic flow device. An ITO glass (SPI supplies, 30–60, 2240 mm, 1.5) substrate was etched in 6 M HCl to create two distinct conductive regions. One region served as a counter electrode, while the other was coated with the Ag paste to function as a pseudo reference electrode. Gold interdigitated electrodes with varying gap widths of 10 and 50 μm (NanoSPR LLC, BA1810 & BA1850) were used as working electrodes. Each interdigitated electrode consisted of two pairs of Au finger electrodes, each with a width of 20 μm width and a variable in-between spacing of either 10 or 50 μm. A cap designed in AutoCAD was fabricated using a 3D printer (Dremel Digilab 3D45) and assembled with the electrodes and tubing. The epoxy (Loctite 1C 1C) was used to seal the device and prevent leakage. The confocal microscopy measurements were conducted using an inverted laser confocal microscope (LEICA SP8 SMD) with an HC PL FLUOTAR 10×/0.30 dry lens. The excitation wavelength was set to 488 nm, and signals were collected from two channels: 592 ± 30 nm and 653 ± 30 nm. The pinhole was set as 0.28 A.U. (20 μm). The x-y resolution was approximately 6.5×10² nm, and the z resolution was approximately 4.8×10³ nm. A syringe pump was used to flow 10 mM phosphate-buffered or MOPS succinate medium (Table S2 and Table S3) containing 15 mM hydroquinone and 10 μM C-SNARF-4. A series of flow rates (0.01, 0.05, 0.1, 0.2, 0.4 mL/min) were tested, and a typical flow rate of 0.1 mL/min was used for all the main figures. Succinate mediums of varying pH values (5.0, 5.5, 6.0, 6.5, 7.0, 7.5) were used to calibrate the pH before electrochemical measurements.

Supplementary Material

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acselectrochem.5c00054.

Detailed experimental methods, additional discussions, supplementary tables and figures (PDF)

Video of the spatiotemporal profiles of pH microenvironments (PDF)