Abstract
The extracellular appendages of bacteria (flagella) that transfer electrons to electrodes are called bacterial nanowires. This study focuses on the isolation and separation of nanowires that are attached via Pseudomonas aeruginosa bacterial culture. The size and roughness of separated nanowires were measured using transmission electron microscopy (TEM) and atomic force microscopy (AFM), respectively. The obtained bacterial nanowires indicated a clear image of bacterial nanowires measuring 16 nm in diameter. The formation of bacterial nanowires was confirmed by microscopic studies (AFM and TEM) and the conductivity nature of bacterial nanowire was investigated by electrochemical techniques. Cyclic voltammetry (CV) and electrochemical impedance spectroscopy (EIS), which are nondestructive voltammetry techniques, suggest that bacterial nanowires could be the source of electrons—which may be used in various applications, for example, microbial fuel cells, biosensors, organic solar cells, and bioelectronic devices. Routine analysis of electron transfer between bacterial nanowires and the electrode was performed, providing insight into the extracellular electron transfer (EET) to the electrode. CV revealed the catalytic electron transferability of bacterial nanowires and electrodes and showed excellent redox activities. CV and EIS studies showed that bacterial nanowires can charge the surface by producing and storing sufficient electrons, behave as a capacitor, and have features consistent with EET. Finally, electrochemical studies confirmed the development of bacterial nanowires with EET. This study suggests that bacterial nanowires can be used to fabricate biomolecular sensors and nanoelectronic devices.
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1. Introduction
Flagella are electronically conductive. Nanowire architectures produced by microbes are microbial nanowires—also called bacterial nanowires. The definite biological role of flagella may vary among microorganisms [1], but they are present on the surfaces of all gram-negative (gram −ve) and some gram-positive (gram +ve) microorganisms [1–4]. In the present study, the gram −ve bacteria Pseudomonas aeroginosa (P. aeruginosa) was used for electric conductivity applications. Electron microscopy has been used for decades to image bacteria with flagella [5]. The images of bacterial nanowires under electron microscope show that the flagella of P. aeruginosa are numerous, relatively thin (8–12 nm), and emerge from one end of the bacterium [5–8]. The basic theory of organic energy generation is based on oxidation and reduction processes involving electron transfer within and/or outside the cells to terminal electron acceptors. In the extracellular respiration or extracellular electron transfer (electron transfer outside the cells), many mechanisms have been proposed, but most of them are not completely understood. Some of the mechanisms have shown potential, but there are still lacunae to be filled in each model and probably unknown mechanisms to be explored. Microbes can generate energy using various strategies.
The characteristics of bacterial nanowires are similar to those of polymer nanowires. Polymer nanowires have recently emerged as an alternative to metal and semiconducting nanowires because of their large conductivity, flexibility, and ease of synthesis [9, 10]. The mechanism of bacterial nanowire conductivity was observed, the oxidation of acetate was carried out at a restricted rate, and biofilms of limited thickness would grow on anodes. This supported the electron transfer rate between aligned cytochromes, as described by the super exchange (SEC) hypothesis. Based on spectroelectrochemical conductivity and cyclic voltammetry measurements, indications of reversible oxidation and reduction of biofilm-associated redox factors were observed, which strengthens the SEC hypothesis [11]. The electrochemical behavior and the redox states of bacterial nanowires are typically studied using cyclic voltammetry (CV) and electrochemical impedance spectroscopy (EIS) and have been documented in earlier studies [12, 13]. The electrochemical behavior of bacterial nanowire is dependent on many parameters, such as the applied potential, the choice of material and the surface area of the electrodes, composition of the electrolyte, and temperature.
The present study focuses on the isolation and separation of bacterial nanowires using a modified method [14] and the characterization of bacterial nanowires using atomic force microscopy (AFM) and transmission electron microscopy (TEM). Furthermore, the roughness of the bacterial nanowires was measured by atomic force microscopy and the conductivity of bacterial nanowires was measured using CV and EIS analyses. The study of the isolation and separation of bacterial nanowires in P. aeruginosa and the measurement of their conductivity is the first of its kind (schema
2. Experiment
2.1. Isolation of bacterial nanowires
Bacterial culture of P. aeruginosa (MTCC No. 2581) was procured from the Microbial Type Culture Collection and Gene Bank (MTCC), Chandigarh, India. The culture of P. aeruginosa was grown in luria betani (LB) medium from a −80 °C freezer stock. After several day's growth, 100 μl aliquots were taken and used to inoculate replicates of a 50 ml flask of LB medium. The isolation of bacterial nanowires was performed by centrifuging bacterial culture at 3000 rpm for 10 min followed by washing with double-distilled water 5 times. To the resulting pellet, 1 ml of distilled water was added. This bacterial suspension was imaged by AFM.
2.2. Separation of bacterial nanowires
Bacterial nanowires separation was done by means of a minor modification of the Chandlar–Kulasekharam method [15]. The whole bacterial culture grown in LB broth medium was deflagellated for 3 min. The aggregated cell and flagella were dispersed by vibromixing the culture with 0.4% NaCl and 0.02% sodium dodecyl sulphate (SDS) (1:10) for 3 min. After this, the cells were separated from the flagella using centrifugation at 16 000 g for 30 min. The supernatant was then filtered through a millipore (GSWP) 0.22 mm membrane and the flagella were resuspended into a solution with 0.4% of saline from the membrane surface by gently rubbing it with a curved glass rod.
2.3. Morphological studies
2.3.1. AFM
The flagella suspension was examined under AFM scanning probe microscope (Model: Shimadzu SPM 9500J2) to analyze the morphology and roughness of bacterial nanowires.
2.3.2. TEM
TEM analysis of bacterial nanowires of P. aeruginosa was carried out in an FEI 200 KeV, LaB6 filament, Tecnai T20 G2 TEM system. Filtered uranyl acetate (2% solution dissolved in distilled water) was used as the negative-staining reagent. The harvested samples were applied as droplet on a TEM grid, stained with 2% uranyl acetate, and dried in air after removing excess media on the grid using absorbent paper.
2.4. Electrochemical studies of bacterial nanowires
2.4.1. CV
Electrochemical experiments were carried out with a CHI 660B electrochemical workstation (CH Instruments Inc., USA) using a conventional one-compartment, three-electrode cell. A glassy carbon (GC) electrode was utilized as the working electrode (electrode area: 0.07 cm2), whereas a silver/silver chloride electrode (Ag/AgCl) was employed as the reference electrode, and a platinum coil was used as the counter electrode. The GC electrode was double-polished using alumina powder (0.05 μm) followed by sonication in double-distilled water for 3 min and used for the modification of the electrode. Experiments were carried out in deaerated 0.1 M H2SO4.
2.4.2. The bacterial nanowires film
The bacterial nanowires were coated under GC electrode by mixing a 0.5 mg ml−1 suspension of nanowire with 0.5% of Nafion polymer in double-distilled water under ultrasonication. Next, 10 μL of this suspension was cast on the cleaned GC electrode surface and allowed to dry at room temperature (20 ± 2 °C).
2.4.3. EIS
Electrochemical experiments were carried out with a CHI 660B electrochemical workstation (CH Instruments Inc., USA) using a conventional one-compartment, three-electrode cell. A GC electrode was utilized as the working electrode (electrode area: 0.07 cm2), whereas a silver/silver chloride electrode (Ag/AgCl) was employed as the reference electrode, and a platinum coil was used as the counter electrode. Ar was used to purge the solution to achieve an O2-free condition. All the electrochemical experiments were performed at room temperature.
EIS responses were obtained at GC electrodes modified with bacterial nanowires. Redox analyte is 1 mM of K3Fe(CN)6 in 0.1 M KCl. The electrode was polarized at 0.25 V and the frequency range was 1 Hz to 100 kHz. The [Fe(CN)6]3−/4− couple was used as the redox probe to study the electrical/conductive behavior of the bacterial nanowire. The Nyquist diagram of the complex impedance represents the imaginary versus the real part of the impedance. The Nyquist plot shows a semicircle at higher frequencies corresponding to the electron-transfer-limited process, and the linear portion at lower frequencies corresponding to the diffusion-limited process.
3. Results and discussion
3.1. Bacterial nanowires imaging
The AFM image of bacterial nanowires isolated from P. aeruginosa revealed extracellular nanowires from the cell bodies (figure 1). It is hard to observe the bacterial nanowire under scanning electron microscopy and so AFM was used as our assessment tool to ensure the presence of flagella. The presence or absence of bacterial nanowires depended on the methodology used to prepare the culture for imaging. The resulting image ensures the presence of bacterial nanowires to go for further separation studies. The extracellular electron transfer (EET) can be carried out from insoluble electron acceptors such as metal oxides and anodes in microbial fuel cells by P. aeruginosa bacterial nanowires. It is already reported that both Shewanella and Geobacter spp. can perform the electron flow from the inner to outer surface of cytochromes through EET or electron transfer, which is promoted by membrane contact with an insoluble electron acceptor [16, 17]. The isolation of nanowires from bacteria depends on growth conditions. Gorby et al [18] reported that chemostat cultures grown under O2-limited conditions (electron acceptor limitation) were observed under a scanning electron microscope, which showed images of Shewanella oneidensis MR-1 with attached nanowires.
Figure 1. AFM images obtained for the isolated bacterial nanowires from P. aeruginosa: (a) 2-D image and (b) 3-D image.
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Standard image High-resolution image3.2. Separation of bacterial nanowires
The separation of the bacterial nanowires was accomplished by the application of mechanical shearing force on bacterial cells. Since these bacteria are osmotically sensitive, mechanical shearing was carried out to the broth containing 2% (wt/vol) NaCl. Using differential centrifugation and filtration methods, crude bacterial nanowires separation was attained. The separated bacterial nanowires were of irregular lengths under AFM and a few visible contaminations were evident (figure 2). The TEM images presented in figure 3 show a negatively stained cell of P. aeruginosa separated bacterial nanowires. A single nanowire emanating from the bacterial cell was separated using a physical shearing method. The apparent lateral dimension of this nanowire is 16 nm. The effective separation of nanowires from the bacteria depends on the method used to prepare the culture for imaging. In a related work, when oxygen concentration exceeded 2% of air diffusion, the number of nanowires observed in chemostat cultures was few or none [18]. Similarly, Reguera et al [5] reported that pili production in Geobacter sulfurreducens was growth-regulated and it occurred only when the organisms were grown on Fe (III) oxide or fumarate at suboptimal temperatures below strictly anaerobic conditions.
Figure 2. AFM images of the separated bacterial nanowires from P. aeruginosa: (a) 2-D image and (b) 3-D image.
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Figure 3. TEM images of the separated bacterial nanowires from P. aeruginosa: (a) high-magnification image and (b) low-magnification image.
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Standard image High-resolution imageThe separation of P. aeruginosa bacterial nanowires by mechanical shearing of cells along with SDS resulted in nanowires of various lengths, with few broken or contaminated. The disintegration of bacterial nanowires using a sonication shearing method with SDS suggested that bacterial nanowires of P. aeruginosa are quite delicate. Meanwhile, these methods are highly reproducible and result in a higher yield of bacterial nanowires. Kalmokoff et al [19] reported that obtaining flagella of various lengths from the sheared cell preparation was not unexpected, but it was surprising because the nonionic surfactant (Triton X-100) preparation was a gentle method for recovering flagella. DePamphillis and Adler [20] found that for bacterial nanowires separation techniques in eubacteria, the use of nonionic surfactant does not appear to affect the morphology of bacterial nanowires. In this study, the nonionic surfactant used was SDS and the resulting image shows flagella of irregular lengths. This shows that the use of SDS achieves similar result to Triton X-100 and is also an effective method to separate bacterial nanowires [21].
The roughness of the bacterial nanowires was examined by AFM. In a log phase bacterial culture, we obserbed that the roughness peak attained a maximum height of 6.2 nm, average maximum roughness peak of 2.7 nm, maximum roughness valley depth of 5.1 nm, and average maximum roughness valley depth of 2.3 nm (figure 4). As the rough surface provides more space for the attachment, the roughness of nanowires is important because it plays a role in the efficient surface binding with other matrices.
Figure 4. Roughness measurement of the separated bacterial nanowires from P. aeruginosa.
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Standard image High-resolution image3.3. Electrochemical studies of bacterial nanowires
3.3.1. CV
CV is probably the most widely used analytical tool for examining the EET process in conductive bacterial nanowires [22–24]. CV was performed for the highly sensitive detection of EET. CV is used to measure the catalytic activity of bacterial nanowires, but the application of CV to bacterial nanowires requires the limiting potential range to prevent harmful oxidation or reduction conditions with the selection of informative scan rates [13, 25]. CV at low scan rates revealed stable catalytic features of bacterial nanowires [13]. For the CV measurements, a scan rate (20 mV s−1) at 20 ± 2 °C was chosen [13]. Under this condition, the rate of electron transfer from the bacterial nanowires to the electrode was enhanced. The CV technique can also be used to examine the rate limiting steps in current generation by bacterial nanowires, and the EET occurs in bacterial nanowires [24, 26, 27]. Figure 5(b) shows the CV of the bacterial nanowires electrode in 0.1 M H2SO4. A redox peak was obtained in the investigated potential range due to the presence of bacterial nanowires on the electrode surface. No redox peak was observed for the blank and is shown in (figure 5(a)), confirming that the redox peaks observed in the case of bacterial nanowires are due exclusively to the films. The reduction current appears to be high for the bacterial nanowire in comparison to oxidation current because of an increase in nanowire density, which enhances the number of electrons generated by oxidation, which in turn affects conductivity and capacity.
Figure 5. CVs of (a) the bare GC electrode and (b) bacterial-nanowire-modified GC electrode recorded in 0.1 M H2SO4, at a scan rate of 20 mV s−1.
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Standard image High-resolution imageThe appearance of an oxidation peak at 0.30 V and reduction peak at 0.27 V was caused by the presence of the nanowires on the electrode surface as shown in figure 5(b). The two clear peaks in the presence of a substrate, and the lack of a peak in the absence of substrate indicated that the electrons had been retrieved from oxidation as a result of bioelectricity. Compared with the blank, the redox peaks obtained for bacterial nanowire was well-defined and pronounced and showed both an oxidation and a reduction peak. A stable CV was obtained by repetitive measurements. Under these conditions, the rate of electron transfer increased rapidly at the bacterial nanowires electrode. This reaction, which is normally called a reversible catalytic wave, indicates the nonstop regeneration of electrons and EET from the film to the electrode [24]. In addition, considering the current responses of the redox peaks in the substrate, it is reasonable to assume that direct EET using nanowires was the primary mechanism. These results suggest that bacterial nanowires facilitated the EET between the bacterial nanowires and electrode.
3.3.2. EIS
To determine whether the resistivity values obtained from our two-contact devices include a significant contribution from contact resistance between electrodes and nanowires, EIS was used to measure the charge transfer resistance of bacterial nanowires as a function of their length. EIS is a suitable tool for measuring local electrical properties at the nanoscale materials, and has been increasingly used for the electrical characterization of biological molecules [28]. EIS is also newly employed to demonstrate transverse conduction through bacterial nanowires produced by P. aeruginosa. EIS is used to examine biofilm growth, biofilm conductivity, and electron transfer mechanisms [29, 30]. EIS is the most common alternating current method applied for the characterization of aqueous biointerfaces [24]. In most EIS methods, a small sinusoidal potential perturbation is applied to the sample. The frequency of this perturbation is changed in a range between a few mHz and 104 Hz. The resulting sinusoidal current is analyzed using fast Fourier transform techniques to calculate the impedance (Z) of the interface in the frequency domain to estimate the charge transfer resistance, diffusion at the surfaces covered by protein monolayers, charge transfer time constants, and mechanisms of EET [29, 30]. Applications of EIS in microbial systems, in which the sample is studied at the open circuit potential, are numerous. Therefore, EIS is performed typically at the open circuit potentials, which reveal the electron transfer characteristics of the active cells attached to electrodes [24].
Figure 6 shows comparative typical EIS Nyquist plots for blank and bacterial nanowire. EIS was performed at 0.0 V versus Ag/AgCl in potentiostatic mode. The impedance is expressed as a real part (plotted on the x-axis) and imaginary part (plotted on the y-axis). Each point on the plot represents the impedance at a certain frequency. The impedance at the high frequency limit is the ohmic resistance Rs, and the diameter of the semicircle Rp is the polarization resistance (charge transfer resistance or interfacial resistance) occurring at the surface of the electrode. The smaller arc radius suggests higher charge transfer efficiency [24, 29−31]. The presence of films containing bacterial nanowires reduced the charge transfer resistance. Figure 6 shows that the arc radius of the blank is smaller than that of the bacterial nanowires conductivity. This suggests that bacterial nanowires have the lowest resistance, which accelerates the EET. This trend shows an increase in electron transfer by bacterial nanowires. In a similar study, it was reported that the oxidation of bacterial nanowires generates an increasing number of electrons, which are transferred extracellularly to the electrode [29−31]. The voltage extensively moves the Fermi level away from this specific condition, resulting in lower conductance or even a reduction of current comparable to blank sample.
Figure 6. EIS recorded for the absence (blank) and presence of bacterial nanowires on to the electrode in 1 mM of K3Fe(CN)6 and 0.1 M KCl.
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Schema 1. Principle of using of bacterial nanowires as biomolecular sensor and nanoelectronic devices.
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The charge transfer resistance of the blank sample was about 7655 Ω cm2 and bacterial nanowires sample resistance was 3760 Ω cm2. This result shows that the bacterial nanowires resistance was decreased about two-times at the same time conductivity was increased. The presence of nanowires on the electrode surface showed a small charge transfer resistance in comparison to the blank sample. The decreased charge transfer resistance was attributed to the conductivity nature of the bacterial nanowires. Gorby et al [18] investigated the distance reliance of the current at various locations over bacterial nanowire appendages and excluded the possibility that the current was being carried by ionic conduction by the presence of water and dissolved ions. The same kind of work was done by El-Naggar et al [32] in S. oneidensis nanowires, which were found to be electrically conductive along micrometer-length scales with electron transport rates up to 109 s−1 at 100 mV of applied potential and a calculated resistivity on the order of 1 Ω cm. Leung et al [33] reported that the S. oneidensis nanowires show electronic behavior with a field effect mobility on the order of 10−1 cm2 V−1 s−1. The conductive flagella provide the opportunity to extend electron transfer capabilities well beyond the outer surface of the cells. These results show that the flagella of P. aeruginosa are highly conductive. The flagella are anchored in the periplasm and the outer membrane of cells, thus offering the possibility that flagella accept electrons from periplasmic and/or outer membrane electron transfer proteins.
4. Conclusion
The present study proves that, by using the method employed in this study, the isolation and separation of nanowires from P. aeruginosa is simple, cost-effective, and easily reproducible in any laboratory with commonly available instruments. Electrochemical methods (CV and EIS) were used to examine the conductivity of the bacterial nanowires to understand the EET phenomena. By choosing suitable techniques and situation, these nondestructive techniques permit the determination of electron transfer from bacterial nanowires. The measurements reported here suggest the need for further investigations on how to increase the bacterial nanowires conductivity. The bacterial nanowires give a green chemistry approach for efficient electron transfer and can be used as an electron source for various potential (bio) electronic applications such as microbial fuel cells, biosensors and organic solar cells.
Acknowledgments
The authors are grateful to the University Grants Commission (F.No.39-559/2010 (SR), dt.12.01.2011), Government of India, New Delhi and DST PURSE Scheme, Madurai Kamaraj University for their financial assistance.