A Microbial Platform Based on Conducting Polymers for Evaluating

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Cite This: Anal. Chem. XXXX, XXX, XXX−XXX

A Microbial Platform Based on Conducting Polymers for Evaluating Metabolic Activity Maki Saito, Kengo Ishiki, Dung Q. Nguyen, and Hiroshi Shiigi* Department of Applied Chemistry, Osaka Prefecture University, 1-2 Gakuen, Naka, Sakai, Osaka 599-8570, Japan

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ABSTRACT: Bacterial cells possessing a certain zeta potential are immobilized by electrochemical deposition within conducting polymers such as poly(3,4-ethylenedioxythiophene) (PEDOT) and polypyrrole (PPy). These conducting polymers serve as a biocompatible matrix for trapping bacteria on an indium−tin− oxide (ITO)-coated glass substrate. The biological functions of bacteria were not affected by the chemical structure and electrical conductivity of the matrix. The viability of the bacteria on the ITO glass was monitored by dark-field microscopy. The cell density of Escherichia coli increased logarithmically during incubation in nutrient broth medium, leading to definitive formation of a biofilm on PPy. The facultative E. coli anaerobe sustains metabolism under aerobic and anaerobic conditions, but proliferates more extensively in the presence of oxygen. The conducting PPy film also facilitates electrochemical evaluation of the respiratory activity of bacterial cells and establishes that facultative anaerobic and aerobic bacteria exhibit similar respiratory activities under aerobic conditions.

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conducting PPy-modified gold substrates in adhesion and proliferation of mouse stem cells has been demonstrated.16 PPy also indicated a good biocompatibility for various microorganisms such as fungi,17,18 yeast,19−21 and bacterial cells22 in the improvement of mechanical properties of cells and charge transfer efficiency from cells. We have previously used conducting polymers to immobilize bacterial cells on substrates.23−27 Bacterial cells are retained by the polymer matrix as anionic dopants due to the negative zeta potential generated by the phosphate groups on the lipopolysaccharides that comprise the outer membrane of the cell. Herein, we demonstrate the utility of conducting polymer films as a matrix for evaluating the biological properties by monitoring the growth and respiration of cells immobilized on conducting polymer-coated ITO substrates.

lthough pathogenic bacteria pose a threat to human life by causing food poisoning1 and infectious diseases,2,3 some strains find application in areas such as sewage purification,4,5 decomposition of toxic substances,6,7 and microbial fuel cell construction.8−10 Hence, a better understanding of the biological functions of bacteria is required to reduce the associated threat and increase the usefulness of these microorganisms, which, in turn, necessitates the quantitative evaluation of bacterial metabolic processes including growth and respiration. Bacterial growth and respiration have been assessed by cellcounting, staining, or gas chromatography.11−14 Conventional methods require one to individually prepare substrates and containers such as agar plates, flasks, and microplates for bacterial samples according to the purpose and method. One means of assessing bacterial metabolism is by immobilizing these entities on a glass slide or an electrode. Immobilization allows bacterial properties to be monitored by optical and/or electrochemical methods and can serve as a platform for carrying out biocatalytic metabolite production and constructing biofuel cells and biosensors.15 In many cases, living cells must be stabilized by confinement in a suitable matrix. The long-term viability and metabolic activity of confined bacteria are influenced by the chemical and mechanical properties of the matrix, including biocompatibility, structural porosity, and contraction. Immobilization of live bacteria is essential for reliable evaluation of their activity through microscopic observations and electrochemical measurements. From such a point of view, the usefulness of the conductive polymer as a matrix material has been clarified. The effectiveness of © XXXX American Chemical Society

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EXPERIMENTAL SECTION Chemicals and Materials. All chemical reagents were of analytical grade and were used as supplied without further purification, unless indicated otherwise. Ultrapure water (resistance >18 MΩ) was used in all experiments. Pyrrole was purchased from Wako Pure Chemical Industries (Japan), and 3,4-ethylenedioxythiophene (EDOT) was obtained from Sigma-Aldrich. Nutrient broth (NB) was obtained from Eiken Chemicals (E-MC35, Japan). Escherichia coli, Acetobacter

Received: May 21, 2019 Accepted: August 19, 2019 Published: August 19, 2019 A

DOI: 10.1021/acs.analchem.9b02350 Anal. Chem. XXXX, XXX, XXX−XXX

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Electrochemical Measurements. Voltammetric measurements were performed using a custom-made thin-layer electrolytic cell (Scheme 1). A piece of filter paper (No. 1, φ

xylinum, Pseudomonas aeruginosa, and Salmonella enterica were acquired from the Biological Resource Center, National Institute of Technology and Evaluation (NBRC, NITE, Japan). Bacterial Culturing. All experiments involving bacterial cultures were executed in a biosafety level 2 laboratory and designed and managed in accordance with safety regulations. A. xylinum was incubated in a medium agar plate (NBRC 350 medium) at 30 °C for 3 d. A single colony on the plate was suspended in 30 mL of liquid 350 medium. After cultivation, the suspension of A. xylinum (7.5 mL) was added to a glass flask containing 142.5 mL of the liquid 350 medium and incubated for 2 d. Other bacteria were incubated in an NB agar plate at 30 °C for 24 h. A single colony on the plate was suspended in 30 mL of liquid NB medium and cultured overnight upon shaking. The bacterial suspension then was centrifuged at 8000 rpm (7510g) for 5 min, the supernatant was removed, and the precipitate was resuspended in sterilized water. The procedure was repeated twice to obtain purified bacterial suspensions (1.0 × 109 CFU mL−1). PEDOT Film Fabrication. The surface of an ITO glass strip with dimensions of 26 × 77 mm2 (Kinoene Optics Inc., Japan) was covered with a UV-curing resin film using a Roland DG LEF12 inkjet printer, with the exception of a circular area (0.79 cm2) used as the working electrode.19−22 A platinum mesh and a Ag|AgCl electrode (filled with 3.0 M KCl) were used as counter and reference electrodes, respectively. The electrodes were placed in a glass cell that contained a solution (6.0 mL) of EDOT (10 mM) and purified A. xylinum in a phosphate buffer (pH 5.3). Electrochemical polymerization was carried out at +1.05 V (vs Ag|AgCl) for 900 s to obtain a cell-doped PEDOT film on the ITO electrode. The freshly prepared A. xylinum/PEDOT film was then rinsed with a copious amount of sterile water. PPy Film Fabrication. Pyrrole (100 mM) was added to the as-prepared bacterial suspension of E. coli, P. aeruginosa, or S. enterica in a phosphate buffer (pH 3.0). An ITO electrode covered with an insulating film to control the electrode area (0.13 cm2) was used as the working electrode.26−29 Electrochemical polymerization was carried out at +0.98 V (vs Ag| AgCl) for 100 s in the phosphate buffer containing bacterial cells to obtain a cell-doped PPy film. The PPy film-coated ITO electrode was rinsed with sterile water and then used as the working electrode. Spectroscopy. The A. xylinum/PEDOT-modified ITO glass strip was subjected to UV−vis spectrometry (V-730, Jasco, Japan) to confirm the formation of PEDOT. The glass strip was immersed into 30 mL of liquid 350 medium and cultured for 1 d. During incubation, the strip was removed from the medium, set in the sample chamber, and subjected to absorption spectrum measurement. Identical operations were carried out for a blank strip on which PEDOT was formed without A. xylinum. Microscopic Observations. The bacteria-doped conducting polymer film was observed using a dark-field microscope (ECLIPSE Ni, Nikon, Japan) equipped with a dark-field condenser, a 100-W halogen lamp, and a camera with a chargecoupled device (DS-Ri1, Nikon, Japan).30,31 Bacterial viability was evaluated by counting stained cells with a fluorescent microscope according to the manufacturer’s instructions for the BacLight bacterial viability kit (ThermoFisher Scientific). The above kit contained two fluorescent pigments, SYTO9, which strained both living and dead cells, and propidium iodide, which strained only dead cells.

Scheme 1. A Custom-Made Thin-Layer Electrolytic Cella

a

Inset shows a photograph of the PPy-modified ITO glass.

55 mm, Toyo Roshi Kaisha, Ltd., Japan) folded in half was placed on the PPy-modified ITO electrode. An appropriate amount of phosphate buffer (0.15 mL) was dropped on the filter paper, and another ITO glass slide was placed on the filter paper as a counter electrode. The assembly was fixed with an adhesive Teflon tape, and a Ag|AgCl reference electrode was inserted into the folded filter paper. Cyclic voltammograms (CVs) were recorded between −0.8 and +0.6 V at a scan rate of 10 mV s−1 under aerobic conditions at 310 K. The concentration of dissolved oxygen in the buffer was measured using an oxygen sensor (Firesting O2, PyroScience GmbH, Germany).

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RESULTS AND DISCUSSION The scanning electron microscopy (SEM) image in Figure 1Aa shows the A. xylinum cells incorporated in a PEDOT film. The difference in the contrast of the image arises from the difference in the electrical conductivity of the film components. The conductive PEDOT matrix appears light gray, whereas the insulating bacterial cells are dark gray. Most bacterial cells are rod-shaped, and their average width and length are 1.5 and 4.0 μm, respectively. The estimated population density is 8.8 × 105 cells cm−2. A dark-field image clearly demonstrates the presence of A. xylinum cells in the film (Figure 1Ba). The cells consist of 70% water and appear bright in rod-like form due to their greater incident-light scattering ability based on the difference in the refractive index between PEDOT (>1.5) and water (1.3) in the cell.32 The estimated viability of A. xylinum cells immobilized in PEDOT is greater than 90% (Figure S1). Although there is good correlation between the SEM and dark-field images of the film, the high vacuum condition results in a significant decrease in bacterial viability (≃0%). The ability to observe the bacteria at normal temperature and pressure using dark-field microscopy makes it possible to follow the cell growth without affecting its viability. Further, an A. xylinum/PEDOT film-coated ITO glass was immersed in liquid 350 medium and incubated aerobically at 303 K. After incubation, the film was rinsed with sterilized water and placed on the stage of a dark-field microscope. The cell density increased to 2.4 × 106 cells cm−2 after 12 h (Figure 1Bb). A. xylinum cells synthesize cellulose nanofibers from glucose.33 The production of bacterial cellulose known as a component of biofilm is very evident in the SEM image (Figure 1Ab). Although a cell density of 1.6 × 107 cells cm−2 B

DOI: 10.1021/acs.analchem.9b02350 Anal. Chem. XXXX, XXX, XXX−XXX

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cellulose layer. (C) Time dependence of the cell density of A. xylinum (n = 3). (D) UV−vis spectra of PEDOT (a) with and (b) without A. xylinum incubated in liquid 350 medium for 24 h under aerobic conditions.

was observed in the dark-field image after 24 h (Figure 1C), it became difficult to estimate the cell number based on SEM imaging due to the extensive production of cellulose nanofibers (Figure 1Ac) that ultimately covered the cells. After 42 h, the continued production of cellulose nanofibers leads to the formation of a white film (inset, Figure 1Ad), which prevents the observation of cells by dark-field microscopy (Figure 1Bd). Peeling the cellulose layer from the PEDOT film after 48-h incubation leads to a cell density of 2.7 × 106 cells cm−2 in the film adhered to the ITO glass. This value is comparable to the cell density in the original film (Figure 1Be). The SEM image of the cellulose layer reveals many A. xylinum cells (Figure 1Af). Cell growth occurs in the cellulose layer, but not in the liquid 350 medium. These observations are consistent with the formation of a biological film of cellulose fibers produced by A. xylinum cells. The UV−vis spectrum of an A. xylinum/PEDOT film obtained before incubation (0 h) exhibited a broad absorption at 700 nm well matched with that of the PEDOT film without A. xylinum (Figure 1D). Thus, bacteria did not affect the chemical structure of PEDOT. The absorption at 700 nm decreases in intensity and shifts toward 500 nm during 24h incubation. This behavior suggests that the bipolaron population decreases and the polaron population increases in the polymer backbone, thus initiating a change in PEDOT from a conducting to insulating state. Thus, bacterial growth and biofilm formation do not affect the chemical structure of PEDOT, as a PEDOT film without bacteria exhibits a similar spectral change. The SEM image of a conducting PPy film on ITO reveals the presence of 1.2 × 2.5 μm E. coli cells (Figure 2A) roughly half-embedded in the ∼800 nm-thick PPy film.34 The darkfield image shows the cells as light, rod-shaped bodies due to the difference in the refractive index between water and PPy (>1.5).35 The cells are well dispersed (Figure 2B) and have a density of 1.5 × 106 cells cm−2. Fluorescence observation indicates the cell viability to be greater than 99% (Figure S2). Therefore, PPy provides a favorable environment for bacteria, due to its greater biocompatibility than PEDOT.22 Growth of the facultative anaerobe, E. coli, was examined by dark-field microscopy. An E. coli/PPy film was immersed in liquid NB medium and incubated aerobically at 303 K. The cell density increased slightly to 1.6 × 106 cells cm−2 after 6-h incubation. Bacterial growth occurs in four stages comprising (i) lag, (ii) log, (iii) stationary, and (iv) death phases. No bacterial growth occurs during the lag phase, because essential cellular components including RNA and enzymes must first be synthesized.36 A significant increase in the cell density was observed after 9 h, indicating that E. coli had entered the log phase with cell division (Figure 2Bb). Biofilm formation was evident at places where bacteria had gathered. The estimated cell density in regions apart from the biofilm is 5.3 × 106 cells cm−2 (Figure 2C). The cell density ultimately reached 3.2 × 107 cells cm−2 excluding the biofilm region after 18 h with a marked increase in the amount of biofilm by this time. It is clear that it shows a more rapid growth curve, considering the number of bacteria in the biofilm during the period of 9−18 h.

Figure 1. (A) SEM and (B) dark-field images of an A. xylinum/ PEDOT film incubated in liquid 350 medium for (a) 0, (b) 12, (c) 24, (d) 42, and (e) 48 h under aerobic conditions. (f) SEM image of the cellulose layer peeled from the PEDOT film. Scale bars are 10 μm. The insets show photographs of the PPy-modified ITO glass and the C

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conditions (Figure S4), in line with the fact that these bacteria are facultative anaerobes that can sustain metabolism under both aerobic and anaerobic conditions. The estimated cell density of 8.0 × 106 cells cm−2 after 18-h incubation is ∼4 times less than that obtained under aerobic conditions (Figure 2Ca). Bacterial growth is strongly correlated with the level of adenosine triphosphate (ATP) as an energy source.39 In turn, ATP generation within a cell depends on the concentration of oxygen, which, as the final electron acceptor, drives the tricarboxylic acid cycle and accounts for greater ATP production under aerobic conditions (36 mol) than under anaerobic ones (2.0 mol) conditions.40,41 Therefore, E. coli cells grow more rapidly in the presence of oxygen. P. aeruginosa is an aerobic bacterium that exhibits metabolic activity in the presence of oxygen. Its cell density in a PPy film (Figure 3A) increases from an initially small value (1.5 × 106

Figure 3. (A) Dark-field images of P. aeruginosa in a PPy film. The PPy film incubated aerobically in liquid NB medium for (a) 0, (b) 9, and (c) 18 h at 303 K. Scale bars are 200 μm. (B) Time-dependence of the cell density of P. aeruginosa in (a) PPy film (left axis) and (b) liquid NB medium (right axis). Number of experiments was 3.

cells cm−2) to 2.6 × 107 cells cm−2 after 9-h incubation (Figure 3Ba). The cell density in the film decreases after 12 h, but the number of bacteria in the liquid medium increases significantly after this time (Figure 3Bb). This indicates that proliferated P. aeruginosa cells migrated from the PPy film to the liquid medium because P. aeruginosa growth is better sustained in the aerobic liquid medium than in the relatively anaerobic atmosphere of the PPy film. We evaluated the respiratory activity of bacterial cells in a PPy film using a thin-layer electrochemical cell (Scheme 1). We controlled the polymerization time for the formation of the PPy film on the ITO glass to regulate the cell density of E. coli. An increase in the thickness of the PPy film increased the apparent cell density on the ITO electrode (Figure 4A and B).26 We prepared the E. coli/PPy film with a polymerization time of 100 s (1.5 × 106 cells cm−2). In the initial CV recorded between +0.6 and −0.8 V vs Ag|AgCl, the Faradaic current that flows at potentials more negative than −0.3 V results from the reduction of dissolved oxygen in the electrolyte solution. There was no discernible difference in the CVs recorded with and without glucose at 0 min incubation time (Figure 4Ca). However, the oxygen reduction current decreased dramatically in the presence of glucose after 30 min (Figure 4Cb). This

Figure 2. (A) SEM images of an E. coli/PPy film: (a) top view and (b) 89°-angle view. (B) Dark-field images of a PPy film incubated aerobically in liquid NB medium for (a) 0, (b) 9, and (c) 18 h. (C) Time-dependence of the cell density of E. coli under aerobic and anaerobic conditions in (a) a PPy film excluding biofilm regions and (b) liquid NB medium (n = 3).

E. coli cell growth entered the stationary phase between 18 and 24 h (3.7 × 107 cells cm−2). We presume that the growth in the stationary phase is limited by the depletion of an essential nutrient.37,38 The cell viability gradually decreased thereafter, and almost all bacterial cells were dead after 48 h (Figure S3). This typical growth pattern suggests that E. coli cells maintain metabolic activity in the PPy matrix as well as in the liquid medium (Figure 2Cb). The result establishes PPy as a suitable environment for bacterial growth and confirms its combination with dark-field microscopy as a powerful tool for observing live cells under normal atmospheric conditions. E. coli cells in PPy films exhibit similar behavior under anaerobic D

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glucose (20 mM) is 1.2 × 10−11 mol min−1. Therefore, the net oxygen reduction rate for glucose utilization by E. coli is 8.0 × 10−12 mol min−1. The per cell rate of oxygen consumption by suspended E. coli is 1.9 × 10−17 mol min−1 per cell, which agrees with the electrochemical result. The same electrochemical experiments performed with S. enterica and P. aeruginosa demonstrate that facultative anaerobic and aerobic bacteria exhibit similar oxygen consumption (Figure S6). The oxygen reduction current decreased with time in the presence of glucose. They exhibit similar respiratory activities under aerobic conditions and high reproducibility with variations within 10% of viability (Table S1), and are in good agreement with the results obtained by optical sensor (Table S2). We also found that there was a large difference in the voltammograms between A. xylinum and the others. This is expected to be due to differences in glucose metabolism and should be investigated in detail by adding organics to replace glucose. These results establish PPy film as an appropriate platform for bacterial immobilization and activity monitoring. In conclusion, we successfully measured bacterial activities in conducting polymer films to show that this material provides a suitable environment for evaluating biological processes including bacterial growth and biofilm formation. Bacterial growth in the film equals that in a liquid medium. The conducting polymer matrix is a useful platform for evaluating the metabolic activity of facultative anaerobic bacteria. In addition, its biocompatibility and electrical conductivity facilitates quantitative evaluation of oxygen consumed by bacterial cells. Our findings are applicable to the analysis of living cells and to the development of electrode materials for biofuel cells and biosensors modified with exoelectrogenic bacteria such as Shewanella or Geobacter species.

Figure 4. (A) Dark-field images of a PPy film prepared by electrochemical polymerization for (a) 100 and (b) 300 s. Scale bars are 10 μm. The inset shows a cross-sectional SEM image of the film with marked thickness. (B) Dependence of E. coli cell density on the polymerization time used to obtain the PPy film (n = 5). (C) Aerobic CVs of a live E. coli/PPy film recorded in the thin-layer electrolysis cell containing 20 mM glucose (a) before incubation and (b,c) after 30 min incubation. Polymerization times of (a,b) 100 s and (c) 300 s were used.

change in current was not observed for the E. coli/PPy film in glucose-free electrolyte or for a PPy film without E. coli in a glucose-containing electrolyte. These results establish that the E. coli cells in the PPy film consume dissolved oxygen and utilize it to oxidize glucose according to the following equation: C6H12O6 + 6H 2O + 6O2 → 6CO2 + 12H 2O

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(1)

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.analchem.9b02350.

The decrease in the Faradaic current between −0.3 and −0.8 V is attributed to the consumption of dioxygen by E. coli based on eq 1. The accumulated charge of 20 μC estimated from the difference in the current responses obtained before and after incubation for 30 min equals the pink-colored area shown in Figure 4C. The quantities of electron and oxygen were calculated to be 0.21 and 0.11 nmol, respectively, based on eq 2. +

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O2 + 2H + 2e → H 2O2

ASSOCIATED CONTENT

S Supporting Information *

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Additional supporting microscopic and electrochemical data describing bacterial viability and respiratory activity (PDF)

AUTHOR INFORMATION

Corresponding Author

(2)

*E-mail: [email protected].

The estimated oxygen consumption by E. coli cells is 1.8 × 10−17 mol min−1 per cell. We found that the dissolved oxygen in the electrolyte solution decreased with an increase in the cell density of E. coli, which strongly affected the electrochemical response, and the oxygen was completely consumed after the incubation of a PPy film prepared by electrochemical polymerization for 300 s (2.6 × 106 cells cm−2). Oxygen consumption by suspended E. coli cells was measured to confirm the above result. An E. coli suspension (4.2 × 105 cells in 8 mL) was placed in a sealed container equipped with a fiber optic oxygen sensor. The concentration of the dissolved oxygen in the suspension decreased gradually in the absence of glucose due to its reaction with stored glycogen (Figure S5).42 Linear correlation between the concentration of dissolved oxygen and incubation time yields an oxygen reduction rate of 4.1 × 10−12 mol min−1. The oxygen reduction rate in the E. coli suspension including

ORCID

Hiroshi Shiigi: 0000-0002-1664-7816 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was supported by the Japan Society for the Promotion of Science through a Grant-in-Aid for Scientific Research (B) (KAKENHI 16H04137).

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