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Real-Time Evaluation of Bacterial Viability Using Gold Nanoparticles Takamasa Kinoshita, Kengo Ishiki, Dung Quang Nguyen, Hiroshi Shiigi, and Tsutomu Nagaoka Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.7b05439 • Publication Date (Web): 28 Feb 2018 Downloaded from http://pubs.acs.org on February 28, 2018

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Analytical Chemistry is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

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Analytical Chemistry

Takamasa Kinoshita, Kengo Ishiki, Dung Q. Nguyen, Hiroshi Shiigi＊, and Tsutomu Nagaoka Department of Applied Chemistry, Osaka Prefecture University, 1-2 Gakuen, Naka, Sakai, Osaka 599-8570, Japan ABSTRACT: Real-time evaluation of bacterial viability is important for various purposes such as hygiene management, development of antibacterial agents, and effective utilization of bacterial resources. Here, we demonstrate a simple procedure for evaluating bacterial viability using gold nanoparticles (Au NPs). The color of bacterial suspensions containing Au NPs strongly depended on the bacterial viability. We found that the dispersion state of Au NPs affected the color of the suspension, based on the interaction of Au NPs with substances secreted by the bacteria. This color change was easily recognized with the naked eye, and viability was accurately determined by measuring the absorbance at a specific wavelength. This method was applicable to various bacterial species, regardless of whether they were gram-positive or gramnegative.

Bacterial viability is of considerable interest in terms of hygiene management in various fields, such as the medical, pharmaceutical, agricultural, and food-related industries.1– 4 Infectious diseases and food poisoning caused by pathogenic bacteria not only cause widespread illness, but also erode confidence in the related industries. Therefore, it is necessary to develop a simple method to monitor bacterial sterilization that can be performed anywhere without acquired skills and large-scale equipment, in order to prevent or eliminate bacterial contamination. In addition, antimicrobial agents have been studied for efficient sterilization and establishment of a sustainable sterilized environment.5–7 To that end, it is important to evaluate the efficacy and sustainability of sterilization in real-time. Various attempts have been made to develop effective uses for bacteria and their products as green and renewable resources for humans. Probiotic bacteria have been utilized in fermented foods to provide physiological benefits for the human body.8 Microbial fertilizers are useful as substitutes for chemical fertilizers that raise concerns over toxicity to humans and soil contamination.9,10 Dissimilatory metal-reducing bacteria can be applied to remove toxic metal pollution and collect precious metals from the environment.11,12 Many researchers have attempted to develop microbial fuel cells to harness clean energy.13,14 For the efficient use of beneficial bacteria, it is necessary to generate an appropriate environment where bacteria are active and have high viability. However, conventional methods for evaluating bacterial viability are problematic due to the requirement for laborious and time-consuming procedures. Colony-counting methods usually require a 24–48 h cultivation and appropriate medium and temperature management, which depend upon the bacterial species involved. Although fluorescent methods can be used to identify living and dead cells by dye-staining, such

methods necessitate additional instruments, such as a fluorescent microscope, fluorometer, and centrifuge.15,16 Metal nanoparticles, which have unique optical properties based on localized surface plasmon resonance (LSPR), have attracted the attention of many researchers in various fields such as physics, chemistry, and biology.17–19 In particular, gold nanoparticles (Au NPs) have been successfully integrated into biosensing and imaging systems, owing to their excellent chemical stability and biocompatibility of metallic gold.20–25 Previously, we reported a simple method for detecting bacteria using metal nanoparticles as optical labels, focusing on the chemical structure of cell-surface molecules, regardless of whether the bacteria were alive or dead.26–29 It is also important to count living bacteria in real time because the hazardous and beneficial properties of bacteria both depend on the viability. Here, we demonstrated the utility of a simple colorimetric procedure in assessing bacterial viability with the naked eye, based on the dispersion states of Au NPs coated with 2-aminoethanethiol (AET) or citrate.

Chemicals and Materials. All chemicals were of reagent grade. Ultrapure water (>18 MΩ cm) sterilized by ultraviolet (UV) light was used for all experiments. Salmonella enterica (S. enterica) (NBRC13245), Escherichia coli (E. coli) (NBRC3972), Staphylococcus aureus (S. aureus) (NBRC102135), and Bacillus subtilis (B. subtilis) (NBRC3009) were purchased from the National Institute of Technology and Evaluation Biological Resource Center (NBRC). Chloroauric acid, 2-aminoethanethiol hydrochloride, sodium tetrahydroborate, trisodium citrate dehydrate, and 4-mercaptobenzoic acid (MBA) were purchased from Wako Pure Chemical Industries, Ltd. (Japan). SYTOTM 9

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Green Fluorescent Nucleic Acid Stain was purchased from ThermoFisher Scientific. Bacterial culture and purification. All bacterial cultures and experiments were performed in a biosafety level 2 laboratory, developed and managed in accordance with safety regulations. Strains of S. enterica, E. coli, and B. subtilis were cultured in agar growth medium (E-MC35, Eiken Chemical Co., Japan) at 303 K for 18 h. Strains of S. aureus were cultured in the same agar growth medium at 303 K for 48 h. A single colony was transferred to liquid growth medium (30 mL; E-MC35, Eiken Chemical Co., Japan) and incubated at 303 K for 18 h. A cultured suspension (10 mL) was centrifuged at 6,500 rpm for 15 min at 278 K, and the pellet was resuspended in ultrapure water (10 mL) by shaking for 1 min. This procedure was repeated 3 times. All resulting suspensions were diluted in ultrapure water to 1.5 × 109 cells mL−1. The suspensions were used within 15 min. It was confirmed that incubating bacteria in ultrapure water did not affect their viabilities (Table S1). Apparatus. Size distributions and zeta potentials of Au NPs were measured with a zeta-potential and particle-size analyzer (ELSZ-2Plus, Otsuka Electronics, Japan). UV– visible (vis) absorption spectra were measured with a UV– vis spectrometer (UV-3100PC, Shimadzu, Japan). Fieldemission scanning electron microscope (SEM) images were obtained with an S-4700 instrument (Hitachi, Japan) at an applied voltage of 10 kV. Prior to SEM observations, a mixture of an Au NP dispersion and a bacterial suspension (5.0 μL) was pipetted onto a conducting Si (111) wafer (p-type) and dried in atmospheric air for 1 h. Dark-field observations and measurements of lightscattering spectra. Dark-field microscopy was used to detect only the light scattered by a structure, while directly transmitted light was blocked using a dark-field condenser. Dark-field observations were made using an optical microscope (ECLIPSE Ni, Nikon, Japan) with a dark-field condenser, a 100 W halogen lamp, and a charge-coupled device camera (DS-Ri1, Nikon, Japan). Prior to observation, a mixture of an Au NP dispersion and a bacterial suspension (5.0 μL) was pipetted onto a glass slide and covered with a glass coverslip. Measurements of light-scattering spectra were taken using a miniature grating spectrometer (USB4000, Ocean Optics), which was connected to the microscope using an optical fiber. Light-scattering spectra in a circular area with a 10-μm diameter were measured using an optical fiber, a 400-μm core diameter, and a 40× objective lens. Synthesis and characterization of AET-Au NPs. Chloroauric acid (24 mM, 2.3 mL) and 2-aminoethanethiol (0.45 M, 0.19 mL) were added to 40 mL of ultrapure water, and the solution was stirred for 20 min at room temperature. After addition of sodium borohydride (10 mM, 10 μL), the solution was stirred vigorously for 10 min at room temperature in the dark. The resulting AET-Au NP dispersion (0.028 weight %) was stored in a glass bottle at 277 K. The mean diameter of the AET-Au NPs was 30.3 ± 4.7 nm, and the zeta potential was +34 mV. Synthesis and characterization of citrate-Au NPs. Chloroauric acid (24 mM, 0.75 mL) and sodium citrate (68

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mM, 0.56 mL) were added to 25 mL of ultrapure water, and the solution was stirred at 353 K for 20 min. The resulting citrate-Au NP dispersion (0.013 weight %) was stored in a glass bottle at 277 K. The mean diameter of the citrate-Au NPs was 32.7 ± 5.4 nm, and the zeta potential was –31 mV. Synthesis and characterization of MBA-Au NPs. Sodium hydrate (0.1 M, 0.15 mL) and MBA (0.0095 mmol) were added to an aqueous solution of sodium citrate solution (68 mM, 3.3 mL). This solution and chloroauric acid (24 mM, 0.78 mL) were added to 25 mL of ultrapure water, and the mixture was stirred at 298 K for 30 min. The resulting MBA-Au NP dispersion (0.013 weight %) was stored in a glass bottle at 277 K. The mean diameter of the MBA-Au NPs was 34.7 ± 9.7 nm, and the zeta potential was –36 mV. Fluorescent measurements. A suspension of S. enterica (1.5 × 109 cells mL−1) was heated in a water bath at 353 K for 30 min. The dead bacterial suspension was centrifuged (6,500 rpm, 15 min, 278 K) to obtain substances eluted from dead cells. The AET-Au NP dispersion (0.50 mL) was added into the supernatant, and the mixture was incubated for 15 min. The mixture was again centrifuged. SYTOTM 9 solution (3.0 μL) was added to the supernatant before and after addition of the Au NPs, and then the fluorescent spectra were measured with an FP-6300 spectrofluorometer (JASCO, Japan).

S. enterica is a bacterium that commonly causes food poisoning. Although S. enterica infects humans following ingestion of unheated foods such as eggs, meat, and milk, it is possible to sterilize these products by heat treatment at a temperature over 343 K. We treated a suspension of S. enterica in a water bath at 353 K for 30 min, and confirmed that the viability of bacteria was nearly 0% based on an evaluation by fluorescent staining (Figure S1). Bacterial suspensions with 0, 20, 40, 60, 80, and 100% viability were prepared by mixing heat-treated (0% viable) and cultivated (100% viable) bacterial suspensions. These suspensions were used for the experiments described below. A 0.50-mL dispersion of AET-conjugated Au NPs, which have a positive zeta potential (+34 mV), was added to a 100%-viable suspension of S. enterica (1.5 × 109 cells, 1.0 mL). No change in the color of the mixture was observed, even after a 12-h incubation. Although the cytotoxicity of Au NPs strongly depends on types of molecules that cover the NPs as a passivation layer, as well as the NP concentration and diameter,30-33 no influence on the viability was found after the incubation (Table S1). The color of suspensions containing increasing proportions of dead bacteria immediately changed within 15 min and shifted from red, through purple, to blue, in parallel with decreasing viabilities (Figure 1A). A substantial difference in the color of the mixture was observed between 40% and 60% viability. The absorption spectrum of a suspension containing living bacteria (100% viable) showed 1 peak at 530 nm that was attributable to Au NPs in a dispersed state (Figure 1B).19 The absorbance at 530 nm decreased along with a decrease in viability, and a broad absorption appeared in the longer-


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Analytical Chemistry wavelength region, indicating that the Au NPs were aggregated in suspension. To investigate the effectiveness of the viability evaluation based on these changes, we attempted sterilization using different methods, including incubation in an aqueous 70% ethanol solution for 24 h and drying under a vacuum at 298 K for 24 h. After adding the reddish Au NP dispersion into respective suspension, blueish suspensions were immediately obtained, and we could distinguish the color change with the naked eye (Figure S2). The changes in absorption spectra were similar with heattreated bacteria. After the mixture (5.0 μL) was dried on a conducting Si wafer, bacterial cells were observed using an SEM. In the case of living bacteria (100% viable), complete dispersion of the Au NPs was observed on the substrate (Figure 1C). In contrast, aggregation of the Au NPs was observed with the 100%-dead bacteria (Figure 1D). Aggregates were not observed with the bacterial cells. These results implied that Au NP aggregation was not induced by the bacteria themselves.

Figure 1. (A) Photograph and (B) absorption spectra of S. enterica suspensions with AET-Au NPs. The viabilities tested were 0, 20, 40, 60, 80, and 100%. SEM images of (C) 100%-viable and (D) 100%-dead S. enterica mixed with AET-Au NPs.

After centrifuging a 100%-dead bacterial suspension, we collected the supernatant and precipitate. Au NP dispersions (0.50 mL) were added to both the supernatant and precipitate, which were resuspended in ultrapure water (1.0 mL). The absorption spectrum of the resuspended precipitate showed a strong peak at 530 nm (Figure 2A), which was identical to that of the 100%-viable bacterial suspension (Figure 1B). In contrast, the supernatant spectrum demonstrated a broad absorption centering around 680 nm, indicating that the Au NPs had aggregated. These data implied that substances released from dead bacteria promoted Au NP aggregation, considering that Au NP aggregation was not induced by the bacteria themselves. The absorption spectrum of the supernatant without Au NPs showed a clear peak at 257 nm (Figure 2B). The peak intensity increased linearly with decreases in the bacterial viability. It is well known that nucleic acid has a maximum

absorption wavelength of approximately 260 nm (attributed to 4 component bases: adenine, cytosine, guanine, and thymine) and that proteins have an absorption maximum of ~280 nm (due to the presence of tryptophan, tyrosine, and phenylalanine residues).34 The ratio of intensities at 260 and 280 nm (Abs260 nm/Abs280 nm) is conventionally used to evaluate the purity of extracted nucleic acid. We found that nucleic acid dominated most of the substance released from the dead bacteria, as the Abs260 nm/Abs280 nm ratio of the supernatant was >2.0. Therefore, the AET-Au NPs appeared to interact with the negatively charged phosphate groups of the phosphodiester in the nucleic acid backbone and to aggregate in the 100%-dead bacterial suspension and supernatant. To investigate the interaction between the Au NPs and nucleic acids, we used a staining dye (SYTOTM 9), which expresses green fluorescence by binding with nucleic acids. The fluorescent spectra of the supernatant were immediately obtained (Ex: at 485 nm). The spectrum of the supernatant without Au NPs showed a clear peak intensity at 505 nm (Figure S3). However, no peak was observed in the spectrum of the supernatant obtained after adding Au NPs. These data indicated that nucleic acids interacted with Au NPs and were removed as precipitates by centrifugation. We conclude that Au NP aggregation was induced by electrostatic interactions between Au NPs and nucleic acids eluted from the dead bacteria.

Figure 2. (A) Absorption spectra of AET-Au NPs mixed with the supernatant and resuspended precipitate of a 100%-dead S. enterica sample. (B) Absorption spectra of supernatants from S. enterica suspensions. The viabilities were 0, 20, 40, 60, 80, 100%, as indicated. The inset shows the dependence of the absorbance at 257 nm on the bacterial viability.

Citrate-Au NPs, which have a negative zeta potential (– 31 mV), showed different behavior from AET-Au NPs in the bacterial suspensions. No change was observed in the color of the mixture, even 12 h after the addition of the citrateAu NP dispersion to a suspension of dead (0% viable) bacteria. The suspension showed typical absorption spectra for Au NP dispersions, with a peak at 530 nm. These findings indicated that the citrate-Au NPs were dispersed stably without aggregation, because the citrate-Au NPs repelled each other in the presence of nucleic acids. The color of the mixture gradually changed from red to purple with increasing bacterial viability (Figure 3A). The peak at 530 nm gradually decreased with increasingly broadened absorption in the longer-wavelength region (Figure 3B).



The absorbance at 750 nm correlated linearly with bacterial viability (Figure 3C). These data suggested that citrateAu NPs can facilitate accurate naked-eye determinations of bacterial viability. We examined the size effects of the citrate-Au NPs by focusing on the absorbance ratio at 750 nm in the spectra (Figure S4). The ABSLive/ABSDead ratios obtained from dead (ABSDead) and living (ABSLive) bacterial suspensions for each Au NP were 1.2 (5 nm, mean diameter), 1.9 (15 nm), and 2.0 (30 nm). When Au NPs with a mean diameter of 50 nm were used, the aggregates easily precipitated. Therefore, we concluded that Au NPs with a mean diameter of 30 nm were optimal for evaluating the viability.

sults, and that aggregation increases the peak light-scattering wavelength.21,36 Hence, it was found that the Au NPs first adsorbed to the surface of bacteria and aggregated during incorporation into the bacterial assembly.

Figure 4. (A) Dark-field images and (B) light-scattering spectra of a 100%-viable S. enterica suspension with citrate-Au NPs at 5 min, 1 h, and 2 h after Au NP addition. The acquisition time was 200 ms. (C) SEM image of 100%-viable S. enterica suspension mixed with citrate-Au NPs for 2 h.

Figure 3. (A) Photograph and (B) absorption spectra of S. enterica suspensions with citrate-Au NPs. The viabilities were 0, 20, 40, 60, 80, and 100%, as indicated. (C) Dependence of the absorbance at 750 nm on the bacterial viability.

It had been reported that citrate-covered Au NPs show excellent biocompatibility.30,32 We also confirmed that no change in viability occurred after the incubation (Table S1). Therefore, Au NP aggregation was induced by viable bacteria. The process of Au NP aggregation in the 100%-viable bacterial suspensions was tracked with dark-field microscopy. Immediately after adding the Au NPs, the bacterial cells were observed as rod-shaped, greenish light spots in a monodispersed state (Figure 4A).35 These bacteria gradually self-associated after colliding in suspension, forming 0.10-mm aggregates after 2 h. In addition, the color of light scattered from the assembled bacteria shifted from green to red. Light-scattering spectra were obtained for single and assembled bacteria. The spectrum of a single bacterium showed a peak intensity at 560 nm after the addition of Au NPs for 5 min (Figure 4B). However, 2 peaks (at 560 and 750 nm) were observed in the bacterial assembly after 1 h. The peak intensity at 750 nm increased substantially after a 2-h incubation. It is well-known that monodispersed Au NPs scatter light at 530 nm based on LSPR re-

In general, living bacteria adhere to solid surfaces and/or themselves by secreting adhesive extracellular polymeric substances (EPSs), resulting in the formation of a self-assembly known as a biofilm.37 EPSs are composed of various molecules, such as polysaccharides, polypeptides, and extracellular nucleotides.38 By SEM, we observed crystalline substances secreted by the bacteria 2 h after Au NP addition (Figure 4C). Au NP aggregates formed with the secreted substance rather than on the bacterial surface. Handy et al. reported that an undisturbed layer on a surface of an organism (owing to mucus secretion) promoted local nanoparticle densification.39 Accordingly, Au NP aggregation was induced as EPSs overcame the protective effect of citrate covering Au NPs. Although further investigation is necessary, it is expected that biofilm formation was promoted with the surrounding Au NPs serving as a scaffold, thereby inducing further aggregation of the Au NPs. Biofilm formation was not observed in an aqueous citrate solution after a 12-h incubation (Figure S5). Another type of Au NP (MBA-Au NP), in which the zeta potential was –36 mV, did not aggregate and induce biofilm formation after a 12-h incubation (Figure S6). It was confirmed that no difference in bacterial viability occurred before and after incubation, even in the MBA-Au NP dispersion. Therefore, the chemical species in solution and the zeta potential of the Au NPs were not directly related to biofilm formation. This indicates that biofilm formation depends on the protective effect of the molecule covering Au NPs.


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Analytical Chemistry The color change, based on the dispersion state of the Au NPs, made it possible to estimate the viability of S. enterica in a suspension. To confirm the utility of this method, we examined other types of gram-negative and gram-positive bacteria such as E. coli, S. aureus, and B. subtilis. Dead bacteria induced AET-Au NP aggregation within 15 min (Figure 5A). Although gram-positive bacteria slightly induced Au NP aggregation (even viable bacteria), color differences between suspensions of living and dead bacteria were clearly distinguishable with the naked eye. Therefore, Au NP aggregation depended strongly on nucleic acids released from dead bacterial cells, rather than differences in the surface structure of gram-positive and gram-negative bacteria. We also examined a mixture of these bacteria, in which the concentration of each cell in the suspension was 3.8 × 108 cells mL−1, because multiple species of bacteria formed microbial communities in natural biofilms.40 Changes occurring in the color and absorption spectra were similar to those of single species (Figure S7).

Figure 5. Photographs and absorption spectra of living and dead bacterial suspensions of (a) E. coli, (b) S. aureus, and (c) B. subtilis with (A) AET-Au NPs or (B) citrate-Au NPs.

Aggregation of citrate-Au NPs did not occur in the suspension of dead bacteria, but was induced by living bacteria (Figure 5B). Although a change in the color of a living S. enterica suspension took 90 min after adding the citrateAu NPs, the colors of the E. coli and S. aureus suspensions changed immediately, and no additional change in the absorption spectra observed after 30 min. With B. subtilis, the color change occurred gradually and was completed in >2 h. These findings indicated that the time required for Au NP aggregation depended on the bacterial species. It is well-known that some EPSs have different chemical structures depending on the bacterial species.41-44 Their secreting rates and proportions are also thought to depend on

the species. The differences may affect the adsorption and aggregation of Au NPs. For the bacterial mixture, color changes between living and dead bacteria were distinguishable with the naked eye (Figure S8). According to the dark-field images, the size of bacterial assemblies that underwent biofilm formation was smaller than those of single species even at same elapsed time after Au NP addition (Figures 4A and S9). This finding indicates that Au NP aggregation depends on biofilm formation because some bacterial species are symbiotic in a biofilm, whereas others exhibit antagonistic behaviors such as inhibiting the growth of biofilms.43,44 To apply this technique with authentic bacterial samples, it is important that the dispersion state of Au NPs is maintained in solution and that Au NP aggregation is induced only when bacteria are added. Therefore, the effects of the solution pH and ionic strength on the dispersion state of Au NPs were investigated. In AET-Au NP dispersions, wherein the pH was changed from 2.0 to 12, no differences in the color and absorption spectra were observed. This finding indicates that the dispersion state of AET-Au NPs is stable within this pH range. Similarly, the dispersion state of the citrate-Au NPs was maintained over a pH range of 4.0 to 9.0. The dispersion state of Au NPs was not affected when the ionic strength of the solutions was less than 0.2. Therefore, the Au NP dispersions showed excellent stability in suitable solutions containing bacterial species under our conditions. When using sample solutions with more ionic species or pH values outside of these ranges, it will be necessary to recover the bacteria by centrifugation and/or adjust the pH. Our data revealed that the Au NP concentration needed for evaluating the viability depended on the density of the bacterial suspension. The concentration of the Au NPs used here (0.013~0.028 weight %) was appropriate for samples containing approximately 108 cells mL−1. When a suspension contained ~106–107 cells mL−1, the Au NP dispersion (~1.0 × 10−4–3.0 × 10−3 weight %) was suitable for use. Based on the typical detection limit of spectrophotometers, our system can be applied to evaluate viabilities for samples containing bacteria at a density of 105 cells mL−1, with an Au NP dispersion of 1.0 × 10−5 weight %. On the other hand, a color change could be clearly distinguished with the naked eye with an Au NP dispersion of 1.0 × 10−3 weight %. Therefore, it is possible to evaluate the viability using suspensions containing bacteria at a density of 107 cells mL−1 with the naked eye.

We achieved real-time evaluation of bacterial viability based on the dispersion state of Au NPs in bacterial suspensions. The AET-Au NPs reflected the bacterial viability within 15 min based on a color change of the suspension, resulting from their aggregation with nucleic acids released from dead cells. In contrast, the citrate-Au NPs aggregated with living bacteria as they formed biofilms. The viability can be easily recognized with the naked eye and accurately determined by taking absorption spectrum measurements. This method enables rapid and simple



evaluation of bacterial viability, regardless of gram-positive or gram-negative status, compared to conventional methods. Moreover, naked-eye determinations of dead and alive bacteria were possible for mixtures of multiple bacteria. We believe that the methods presented here will contribute to various aspects of research related to hygiene management, antibacterial agents, and the effective use of beneficial bacteria.

Confirmation of bacterial viability, other supplementing spectral data, and microscopic images are presented. The Supporting Information is available free of charge on the ACS Publications website.

[email protected]

All authors contributed equally to this study. The authors declare no competing financial interest.

We acknowledge the financial support provided by the Ministry of Agriculture, Forestry, and Fisheries through a Science and Technology Research Promotion Program for the agriculture, forestry, fisheries, and food industries. We also acknowledge financial support from the Japan Society for the Promotion of Science (JSPS) through a Grant-in-Aid for Scientific Research (B) (KAKENHI 16H04137) and a Grant-in-Aid for Challenging Exploratory Research (KAKENHI 26620072). T.K. acknowledges financial support from the JSPS through a Grant-in-Aid for JSPS Research Fellowship (16J07230).

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for TOC only


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Analytical Chemistry

TOC 83x43mm (300 x 300 DPI)


Real-Time Evaluation of Bacterial Viability Using ... - ACS Publications

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