Electrochemical and spectroelectrochemical characterization of bacteria and bacterial systems

Abstract

Microbes, such as bacteria, can be described, at one level, as small, self-sustaining chemical factories. Based on the species, strain, and even the environment, bacteria can be useful, neutral or pathogenic to human life, so it is increasingly important that we be able to characterize them at the molecular level with chemical specificity and spatial and temporal resolution in order to understand their behavior. Bacterial metabolism involves a large number of internal and external electron transfer processes, so it is logical that electrochemical techniques have been employed to investigate these bacterial metabolites. In this mini-review, we focus on electrochemical and spectroelectrochemical methods that have been developed and used specifically to chemically characterize bacteria and their behavior. First, we discuss the latest mechanistic insights and current understanding of microbial electron transfer, including both direct and mediated electron transfer. Second, we summarize progress on approaches to spatiotemporal characterization of secreted factors, including both metabolites and signaling molecules, which can be used to discern how natural or external factors can alter metabolic states of bacterial cells and change either their individual or collective behavior. Finally, we address in situ methods of single-cell characterization, which can uncover how heterogeneity in cell behavior is reflected in the behavior and properties of collections of bacteria, e.g. bacterial communities. Recent advances in (spectro)electrochemical characterization of bacteria have yielded important new insights both at the ensemble and the single-entity levels, which are furthering our understanding of bacterial behavior. These insights, in turn, promise to benefit applications ranging from biosensors to the use of bacteria in bacteria-based bioenergy generation and storage.

Figure 1.

Schematic diagram describing direct (left) and mediated (right) electron transfer in the microbial system. Adapted with permission from reference Copyright 2020 Progress in Chemistry.

Figure 2.

Cyclic voltammograms of a G. sulfurreducens biofilm on an indium-tin-oxide (ITO) electrode in (A) anodic (acetate to CO₂) and (B) cathodic (fumarate to succinate) modes. (C) UV-visible spectra of biofilms after anodic and cathodic scans. (D) Schematic illustrating plausible EET mechanism in cathodic mode. (E) Schematic and energy diagram showing EET from G. sulfurreducens directly to TiO₂ under visible illumination. OM = outer membrane. (F) Current density obtained from G. sulfurreducens under dark and illuminated conditions. Panels A-D are adapted with permission from ref. Copyright 2020 American Chemical Society. Panel E and F are adapted with permission from ref. Copyright 2020 Elsevier B.V.

Figure 3.

(A) Current densities obtained from E. coli immobilized on carbon paper electrode with nine different redox mediators at six different concentrations. NR: neutral red, PYO: pyocyanin, BAPD: benzo(A)phenazine-7,12-dioxide, MPMS: 1-methoxy-5-methylphenazinium methyl sulfate (MPMS), PMS: phenazine methosulfate, PES: phenazine ethosulfate, PHZ: phenazine, OHPHZ: 1-hydroxyphenazine, and PCX: phenazine-1-carboxamide. (B) Current densities as a function of time for S. oneidensis with five different redox mediators. AQS: 9,10-anthraquinone-2-sulfonic acid, AQDS: 9,10-anthraquinone-2,6-disulfonic acid, FMN: flavin mononucleotide, 2HNQ: 2-hydroxy-1,4-napthoquinone, and RF: riboflavin. (C) Scanning electron micrographs showing S. oneidensis biofilm formation as a function of redox mediator and time. (D) Current density obtained from S. oneidensis with and without outer membrane vesicles (OMVs). Panel A is adapted with permission from ref. Copyright 2021 The Electrochemical Society. Panels B and C are adapted with permission from ref. Copyright 2020 American Chemical Society. Panel D is adapted with permission from ref. Copyright 2019 American Chemical Society.

Figure 4.

(A) Schematic diagram showing SECM in combination with 3D printed microtrap system for mapping PYO spatial distributions. (B) Electrochemical current maps of PYO secreted ny P. aeruginosa using the SECM approach of panel A. Scale bar = 10 μm. (C) Left: schematic of dual ring nanopore electrode array (NEA) in contact with P. aeruginosa. Right: schematic showing redox cycling of phenazine in NEA. (D) Redox cycling induced cyclic voltammetry of P. aeruginosa as a function of oprical density (OD). Panels A and B are adapted with permission from ref. Copyright 2014 National Academy of Sciences. Panel C and D are adapted with permission from ref. Copyright 2021 The Royal Society of Chemistry.

Figure 5.

Surface-enhanced Raman spectra of PYO as a function of (A) pH, and (B) electrochemical potential. Adapted with permission from ref. Copyright 2019 American Chemical Society.

Figure 6.

(A) Different ways of detecting bacteria-electrode collisions along with their current-time responses. Top panel: bacterial cell blocking the UME electroactive area; Middle panel: bacterial cell catalyzing reduction; Bottom panel: regeneration of redox species by bacterial cell. (B) Time-lapse fluorescence images showing E. coli attaching to the UME surface. (C) Current-time trace corresponding to panel B. Panel A is adapted with permission from ref. Copyright 2018 American Chemical Society. Panels B and C are adapted with permission from ref. Copyright 2018 Elsevier Ltd.

Figure 7.

(A) Fluorescence images of S. oneidensis with thioflavin T obtained as a function of electrochemical potential. (B) Electrochemical potential- and fluorescence intensity-time traces for three individual cells marked in panel A. Adapted with permission from ref.. 2020 National Academy of Sciences.