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Online-monitoring of biofilm formation using nanostructured electrode surfaces
Materials Science & Engineering C 100 (2019) 178–185
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Online-monitoring of biofilm formation using nanostructured electrode surfaces Mohammed Sedkia,1, Rabeay Y.A. Hassana,b,
⁎,1
, Silvana Andreescuc, Ibrahim M. El-Sherbinya,
T ⁎
a
Nanomaterials Laboratory, Center for Materials Science, Zewail City of Science and Technology, 6th October City, 12588 Giza, Egypt Applied Organic Chemistry Department, National Research Centre (NRC), Dokki, 12622 Giza, Egypt c Department of Chemistry and Biomolecular Science, Clarkson University, Potsdam, NY 13699-5810, USA b
A R T I C LE I N FO
A B S T R A C T
Keywords: Nanostructured surfaces Microbial electrochemistry Monitoring biofilm formation Pseudomonas aeruginosa
The direct monitoring of biofilm formation enables valuable insights into the industrial processes, microbiology, and biomedical applications. Therefore, in the present study, nano-structured bioelectrochemical platforms were designed for sensing the formation of biofilm of P. aeruginosa along with monitoring its electrochemical/morphological changes under different stresses. Through the assay optimizations, the performances of different electrode modifiers such as reduced graphene oxide (rGO) nanosheets, hyperbranched chitosan nanoparticles (HBCs NPs), and rGO-HBCs nano-composite were tested to assess the influence of the electrode materials on biofilm progression. As a need for the anodic respiration, the bioelectrochemical responses of the adhered bacterial cells changed from a non-electrochemically active (planktonic state) to an electrochemically active (biofilm matrix) state. Our results demonstrated that electrode modifications with conductive nanostructured elements is highly sensitive and enable direct assay for the biofilm formation without any preachments. Consequently, the morphological changes in bacterial cell wall, upon switching from the planktonic state to the biofilm matrix were imaged using scanning electron microscopy (SEM), and the changes in cell wall chemical composition were monitored by the Energy Dispersive X-ray analysis (EDX). Thus, the designed microbial electrochemical system (MES) was successfully used to monitor changes in the biofilm matrix under different stresses through direct measurements of electron exchanges.
1. Introduction Biofilms are three-dimensional, heterogeneous community of microbial cells enclosed in an exopolysaccharide matrix that can irreversibly attach to solid surfaces or living tissues [1–4]. Biofilm formation is of high importance in many useful applications such as biocatalysis, degradation of chemical contaminants in wastewater, and microbial fuel cells [5]. However, these biofilms may have serious implications on public health and environment [6–8]. For instance, they cause biofouling, microbially-induced corrosion, contamination of implanted medical devices, and are responsible for the persistent infections in human body resulting in death of millions of people every year [9–12]. The early detection of pathogens, contaminants and virulence biomolecules plays a crucial role in the prevention and treatment of microbial infections [13,14]. Therefore, numerous methods for the detection of pathogens have been developed. However, the big problem is not to measure the microbe in its natural form, the challenge is to detect
the colonized microbes within the biofilms that are not neither accessible nor easy to detect. Microbial electrochemical systems (MESs) were shown to be the most effective techniques for studying the role of the conductive solid surfaces in the enhancements of the biofilm formation [15]. Monitoring the microbe-electrode interaction(s) is the main concept of the MESs. Degradable organic matters provide the necessary nutrition to living organisms. As a result of the microbial catalytic oxidation of the degradable organic compounds, free electrons are liberated through the electron transport chain (ETC). Under the anaerobic conditions, the liberated electrons are transferred to a molecule other than oxygen such as nitrate or sulfate [16]. In the MESs, microbes exploit their catalytic activity to transfer the liberated electrons to an electrode (as an electron acceptor), and consequently electric current is generated. The generated current density is directly proportional to the number of viable microbial cells (i.e. the metabolically active microbes) [14,17]. Three mechanisms have been proposed to explain the electron
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Corresponding authors at: Center for Materials Science, Zewail City of Science and Technology, 6th October City, 12588 Giza, Egypt. E-mail addresses: [email protected] (R.Y.A. Hassan), [email protected] (I.M. El-Sherbiny). 1 Both are sharing the first authorship. https://doi.org/10.1016/j.msec.2019.02.112 Received 28 September 2018; Received in revised form 4 February 2019; Accepted 28 February 2019 Available online 04 March 2019 0928-4931/ © 2019 Elsevier B.V. All rights reserved.
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Fig. 1. (a) XRD analysis of the thermally reduced graphene oxide (rGO) nanosheets, (b) FTIR spectra of rGO, HBCs and rGO-HBCs nanostructures, (c & d) SEM images of rGO-HBCs nanostructure.
used on both planktonic and biofilm cultures to monitor changes in cell wall chemical composition and characterize their physical connections.
transfer at microbe-electrode interface by exoelectrogens without artificial mediators. Firstly, certain exoelectrogenic bacteria can transfer electrons to an electrode surface through soluble redox active compounds secreted by microorganisms [18,19]. Other microbes were able to directly transfer electrons to anodes via outer cell conductive membrane localized proteins [20]. Lastly, electrons are delivered directly to the electrode through the naturally produced bacterial nanowires [21,22]. It was concluded that strongly adhered microbes produce the higher bioelectrochemical signals. Therefore, the formation of a biofilm on a conductive electrode surface could be detected [23,24]. However, the biofilm formation is strongly dependent on the electrode material and the morphological structures of the working electrode [25,26]. Hence, a better understanding of the microbial communication in relation to the morphological and physicochemical properties of the electrode material is needed. Herein, we report a microbial electrochemical sensor system for reliable and rapid monitoring and investigation of the Pseudomonas aeruginosa biofilm formation on modified electrodes. P. aeruginosa was selected as a target organism because of its high ability to form biofilms in wounds, burns, implanted devices, catheters, urinary tract, and in lungs of patients having cystic fibrosis [27,28]. We designed nanostructured bio-electrochemical systems to explore the P. aeruginosa–electrode interactions using cyclic voltammetry (CV) measurements and the scanning electron microscopy (SEM) were used to investigate the morphological characteristics of the biofilms. EDX spectroscopy was
2. Materials and methods All data generated or analyzed during this study are included in this published article and its Supplementary Information files. D (+) glucose was purchased from Fine-Chem Limited. LuriaBertani (LB) liquid media were used for cells cultivation. The electrode modifiers of reduced graphene oxide (rGO) nanosheets, hyperbranched chitosan nanoparticles (HBCs NPs), and rGO-HBCs nanostructure were synthesized in our lab. Paraffin oil and graphite powder used in carbon paste electrode (CPE) preparation were purchased from Fluka and Sigma Aldrich, respectively. Ciprofloxacin [1-cyclopropyl-6-fluoro-4oxo-7-(piperazin-1-yl)-1,4-dihydroquinoline-3-carboxylic acid] was bought from Sigma-Aldrich. All OD600 measurements were conducted by Thermo-Scientific Evolution 600 UV–Vis spectrophotometer. All electrochemical measurements were carried out using a computercontrolled Gamry Potentiostat/Galvanostat/ZRA G750 (Gamry, Pennsylvania, USA). The three-electrode electrochemical system used in these measurements comprised a modified-CPE with a surface area of 0.5 cm2 as the working electrode, a calomel electrode as a reference, and a Pt disc as auxiliary electrode.
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2.1. Synthesis and characterization of HBCs, rGO, rGO-HBCs nanostructures Hyperbranched chitosan nanoparticles (HBCs NPs) were prepared using our reported method [29] and then conjugated with rGO nanosheets via on/off sonication for 10 min. rGO nanosheets were produced by thermal reduction, at 200 °C for 2 h, of the GO synthesized using an improved Hummer's method [30]. The synthesized rGO nanosheets were characterized by X-ray Diffraction (XRD) (Shimadzu, Kyoto, Japan) in the 2θ range of 11° to 29°, with a step size of 0.01°, and a time per step of 0.5 s. In addition, the synthesized rGO, HBCs, and rGO-HBCs nanostructures were investigated using Fourier Transform Infrared (FTIR) spectroscopy (Thermo Scientific Nicolet iS10 FT-IR), and imaged by scanning electron microscopy (SEM) (FEI Quanta 250).
2.2. Preparation of the nanostructured electrodes The modification of the working electrodes was conducted by mixing 0.9 g synthetic graphite powder with 0.1 g of the modifiers (HBCs, rGO or rGO-HBCs nanomaterials) with 0.4 ml paraffin oil in a small mortar. The prepared pastes were filled and compressed into the electrode assembly producing HBCs-CPE, rGO-CPE, and rGO-HBCs-CPE electrodes with a surface area of 0.5 cm2. The electrode surface was polished before each measurement via scratching its tip against a smooth wet paper and electrochemically activated in electrolyte (0.01 M KCl) by 5 cycles in a potential range of −0.3 to 1.00 V. The measurement time was fixed at 15 S and the scan rate was set at 50 mV/ S.
2.3. Testing electrode materials' effect on biofilm formation P. aeruginosa's three cells suspensions of OD600 = 0.1 were incubated with the previously prepared and tested electrodes (Bare CPE, HBCs-CPE and rGO-HBCs-CPE) under anaerobic conditions for 20 h, and then the corresponding cyclic voltammetry (CV) measurements were conducted. The electrodes were washed thoroughly with PBS before each measurement. In addition, the electrochemical measurements were done in KCl 0.01 M, away from any secreted biomolecules. Moreover, bare-CPE and rGO-HBCs-CPE were incubated with P. aeruginosa at the same abovementioned conditions for five days and the CV measurements were carried out on the washed biofilm-electrode at different time intervals.
2.4. SEM and EDX analyses A carbon screen printed electrode was covered with rGO-HBCs and incubated with bacterial cells in LB media for five days under anaerobic and static conditions at 37 °C. It was then washed, dried, covered by a thin layer of sputtered gold to enhance the image quality and imaged by SEM (FEI Quanta 250). The elemental composition of different sections was analyzed by the Energy Dispersive X-ray (EDX) analysis. In addition, an EDX mapping analysis was conducted on the mature biofilm without gold sputtering to confirm the concentration of iron species at the grown biofilm matrices.
2.5. The effect of planktonic cells adherence on electrode activity
Fig. 2. (a) The CV of the biofilm formation at different electrode materials (Bare CPE, HBCs-CPE, rGO-HBCs-CPE) after 20 h of incubation time, (b) The CV measurements of the biofilm formation monitored at the bare CPE electrode at different time intervals from 1.5 days to 5.0 days, (c) The same as (b) using the rGO-HBCs-CPE electrode. All measurements were carried out in 0.1 KCl with a sensing time of 15 s and a scan rate of 50 mV/S.
The rGO-HBCs-CPE electrode was incubated with P. aeruginosa cells suspension with an OD600 = 0.1 in PBS (0.1 M, pH = 7.4) and glucose concentration of 8 g/L for 3 h at 37 °C, and the corresponding CVs were recorded at different time intervals. The experiment was repeated under the same conditions after removal of bacterial cells (supernatant) for each measurement. 180
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Fig. 3. SEM images of (a) P. aeruginosa planktonic cells at the carbon screen printed electrode surface, (b) the slimy biofilm matrix of P. aeruginosa at the rGO-HBCsCPE electrode after incubation for five days in LB media, (c) a closer view showing the connection between biofilm bacterial cells through flagella and pili.
confirmed by the disappearance of the hydroxyl stretching peaks at 3400 cm−1. The very small peaks of rGO appeared at 1723 and 1750 cm−1 can be attributed to the remaining ketone carbonyl and ester carbonyl of the partially rGO, respectively. The presence of a very small portion of partially rGO is intentionally made to enhance the binding ability between rGO and HBCs. On the other hand, the on/off sonication of rGO with HBCs resulted in successful formation of rGOHBCs nanocomposite as confirmed by the presence of characteristic FTIR peaks for both materials. Results of FTIR analysis are shown in Fig. 1b. The morphology of the prepared nanocomposite (rGO-HBCs) was studied using SEM imaging which showed semi-spherical HBCs with particle sizes around 400 nm interconnected within the rGO nanosheets network, Fig. 1c & d. Additional microscopic information about the prepared rGO and the rGO-HBCs after immersion in aqueous media for a week are provided in supplementary file (Fig. S1).
2.6. The effect of ciprofloxacin and acetone on P. aeruginosa biofilm After formation of a mature P. aeruginosa biofilm at the rGO-HBCsCPE electrode, the electrode was washed several times with PBS and then incubated with ciprofloxacin at a concentration twice as high as the MIC for two days and the corresponding CV measurements were collected. In addition, the mature electrode-biofilm was immersed in acetone for 10 min and then measured using CV to assess the degradation of the biofilm.
3. Results and discussion 3.1. Synthesis and characterization of rGO, HBCs and rGO-HBCs nanostructures Hyperbranched chitosan nanoparticles (HBCs NPs) were prepared using our reported method [31] and then conjugated with reduced graphene oxide (rGO) nanosheets with on/off sonication for 10 min. rGO nanosheets were produced via the thermal reduction of the improved Hummer's method synthesized GO [30] at 200 °C for 1 h. The Xray diffraction (XRD) analysis of the powder sample of rGO nanosheets was carried out in the 2theta range of 11° to 29°. The results, Fig. 1a, showed a strong peak at 2Ɵ of 23° for rGO [32] with a small shoulder at 12° for partially reduced GO with no peaks for graphite at 2Ɵ of 26°, confirming the successful reduction process. The FTIR analysis of the core-shell hyperbranched Cs matched previously reported data [31] with stretching peaks of its terminal amine groups noted at 3288–3388 cm−1, stretching peaks of ester carbonyl at 1723 cm−1 and the stretching peaks of CeH at 2800–2900 cm−1. In addition, the thermal reduction of GO at 200 °C in ambient atmosphere was
3.2. Influence of electrode materials on biofilm formation The working electrode material is the key player in developing MESs, whereas the microbial kinetics is directly linked to the reactions occurring at the electrode surface [33–37,38]. Previously, we have developed an efficient MES using rGO-HBCs nanostructure as an electrode modifier for detection of cell viability and metabolic activity of P. aeruginosa [39]. Here we extended this concept to study the effect of surface roughness, wettability and the electron transfer properties of the nanocomposite to identify the driving parameters that are responsible for the enhanced biofilm formation of P. aeruginosa. To understand the contribution of each material in the matrix, three different working electrodes were prepared: Bare-CPE, HBCs-CPE, and rGOHBCs-CPE. The electrodes were incubated in a fresh individual bacterial 181
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Fig. 4. A schematic illustration of the biofilm growth monitored by measuring the direct extracellular electron transfer ability of the biofilm matrix.
nanostructured electrode surface.
culture containing a suspension of P. aeruginosa cells (OD600 = 0.1) in anaerobic conditions and the corresponding CVs were recorded. Prior to each CV measurement, the electrode was washed thoroughly with phosphate buffer to remove extracellular secretions. The strongest bioelectrochemical responses were observed at the rGO-HBCs-CPE electrode, Fig. 2a. This high bioelectrochemical performance is attributed to the interconnected network and high conductivity of the rGO and HBCs nanocomposite network possessing fast electron transfer ability and high surface roughness in addition to the adhesion properties of HBCs polymeric nanoparticles. By comparison, bare-CPE and HBCs-CPE each alone, they did not show strong bioelectrochemical signals. Further, bare-CPE and rGO-HBCs-CPE were incubated with P. aeruginosa in the same conditions for five days and the CV measurements were carried out on the washed biofilm-electrode at different time intervals, Fig. 2b & c. The adhered cells to the CPE did not show obvious electrochemical signals indicating lower electrode conductivity, Fig. 2b. On the other hand, the peak currents of the biofilm(rGO-HBCs-CPE) electrode increased significantly from tens of micro amperes to around 3.0 mA, Fig. 2c. The current increased gradually in both electrodes until it reached a constant value after which no further increase was detected. This increase in electron transfer demonstrates formation of matured biofilms that enable direct electrical communication between the active P. aeruginosa biofilm and the
3.3. P. aeruginosa biofilms: how are extracellular electrons transferred? Physical connections through the bacterial appendages, microbial nanowires, cyt-c and ferric iron were identified as main contributors to the biofilm assisted-bioelectrochemical signals in few microorganisms such as Geobacter sulfurreducens, Shewanella oneidensis, and Thiobacillus denitrificans [40–42]. Cellular interactions and the contributing factors in the formation of P. aeruginosa biofilms (i.e. the P. aeruginosa-electrode interactions) were less explored. The direct continuous increase of electrical current that is generated from the connectivity of adhered bacterial cells to the nano-electrode surface is indicating formation of mature electrochemically active biofilms. To determine the microbial connectors, the morphology of the adhered bacterial cells at the screen-printed electrode modified surface in planktonic state and biofilm matrices were imaged using SEM. The images shown in Fig. 3a illustrated the typical bacillus (rod-like) morphology of planktonic cells without any appendages or physical connections between them. On the other hand, the SEM images of the P. aeruginosa biofilm, after incubation for five days with the electrode surface, demonstrated the coverage of bacterial cells in slimy secretions of polysaccharides as seen in Fig. 3b, and the direct physical 182
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Fig. 5. The SEM-EDX of P. aeruginosa biofilm illustrating the connection between biofilm cells and the percentage of iron at different sections; (a) at the cellular structure of planktonic cells, (b) at the cell wall of P. aeruginosa biofilm cells, and (c) at the flagella connecting biofilm cells to each other. The samples were covered with a thin gold sputtered film to enhance the images' resolution.
bacterial cells and the biofilm formed layer were analyzed by EDXspectroscopy. Results showed in Fig. 5 identified the elemental composition of planktonic bacterial cells, biofilm matrix and the conductive flagella. Surprisingly, the content of the biofilm outer-layer exhibited iron-complex concentrations ranging from 1.78% in biofilm matrix to 2.99% in flagella. Nevertheless, the planktonic cells contained negligible traces of iron (less than 0.31%). The EDX mapping results of the P. aeruginosa mature biofilm showed higher distribution of iron species at the thicker and more complex biofilm regions, confirming a direct relationship between the conductivity of the biofilm and the existence of iron species. This is illustrated in Fig. 6b as related to the captured images of the mapped EDX (Fig. 6a) on ETD detector. Typically, microorganisms in biofilms live in a home-made matrix of extracellular polymeric substances (EPS) [43]. The EPS support the mechanical stability of biofilms, mediate cell adhesion to the solid
connections between almost all cells with each other and with the substrate and biofilm matrix, Fig. 3c. The SEM images of these bacterial nanowires match the morphology and dimensions of flagella whose role in motility and/or the initial cellto-surface interactions has been demonstrated by O'Toole et al. They have shown that the defective flagella mutant (flgK) did not develop microcolonies on the solid substrate over the course of the experiments, suggesting the necessity of the flagella in microcolony formation [13]. Our results indicate that the signaling between the P. aeruginosa cells and the direct extracellular electron transfer observed may occur through purposely produced bacterial nanowires and their conductive elements. A schematic illustration of the biofilm growth at different stages of formation monitored by the corresponding conductivity of the biofilm matrix is shown in Fig. 4. Additional confirmatory SEM images are provided in the supplementary information (Fig. S3). The composition of the outer-layer of the dispersions of fresh 183
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(a)
(b)
Fig. 6. (a) The image of the mapped area using the ETD detector, and (b) EDX mapping showing distribution of iron of P. aeruginosa biofilm at the surface of rGOHBCs.
Fig. 7. (a) CVs of P. aeruginosa mature biofilm at the RGO-HBCs modified electrode in the presence of ciprofloxacin for two days, and (b) effect of the incubation of the formed biofilm-modified electrode in acetone for 10 min.
3.4. Adherence of the planktonic cells to the electrode surface
surfaces and form a cohesive, three-dimensional polymer network that interconnects biofilm cells. In most biofilms, living cells represent less than 10% of the total mass, whereas the rest of the matrix consists of water and secretions. The composition of biofilm matrices is dependent on the bacterial species and the environmental conditions, but in general they contain components such as polysaccharides, proteins and extracellular DNA [44,45]. From a mechanistic point of view, the extracellular DNA release in P. aeruginosa PAO1 biofilms is regulated by the iron availability [46]. On the other hand, the quorum-sensing systems are strongly involved in the formation of extracellular DNA in P. aeruginosa biofilms. Other reports showed that the level of iron is not only affecting the P. aeruginosa DNA release and biofilm formation but also the maturation of the biofilm and its structural development. Since iron was identified as a critical factor in the interaction between the pathogen and the host, in vitro studies proved that the sub-inhibitory concentrations of the iron chelator lactoferrin block the ability of P. aeruginosa biofilms formation. Thus, iron serves as a signaling element for biofilm development. Besides, P. aeruginosa produces siderophores with high-affinity for iron uptake that can not only chelate free iron but also ‘strip’ iron from host proteins such as lactoferrin [47–51].
The HBCs-rGO-CPE was incubated with cells suspension in a solution of PBS and glucose for 3 h, and the corresponding current of the secreted biomolecules was measured at different time intervals. Measurements were also performed on the supernatant after removal of bacterial cells in each time point in the same conditions. The electron transfer and the oxidation currents were inhibited in the presence of bacterial cells due to their adherence to electrode surface. The decrease in surface conductivity due to the attachment of cells reflects the poor conductivity of planktonic cells (Table S1). These findings support the assumption that the cell wall structure of P. aeruginosa changed from non-conductive planktonic cells to a conductive surface through the nanostructured elements of the electrode which promote biofilm formation. Conductivity has been gained by the production of natural nanowires rich in iron-containing materials (i.e. cytochromes or oxides). 3.5. Effect of stressors on mature biofilm formation rGO-HBCs modified nanostructured electrodes were incubated in bacterial cultures in absence and presence of ciprofloxacin, an 184
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antibiotic that is known to inactivate P. aeruginosa. The growth of electroactive biofilms and their electrochemical signals at the electrode surface was monitored to determine the effect of stress induced by ciprofloxacin. In these experiments, cells were exposed to the minimum inhibitory concentration (MIC), determined using the standard cell proliferation reagents WST-1 kit as shown in Fig. S4. The bioelectrochemical signals measured for the treated and untreated biofilms shown in Fig. 7a indicate that while the signals of the untreated biofilm were in a continuous increase, the activity of the treated biofilm was inhibited over the treatment time. The decrease in signals can be explained by the good penetration ability of ciprofloxacin through the biofilm layers, leading to partial destruction of the biofilm matrix [52,53]. Since P. aeruginosa biofilm is susceptible to ciprofloxacin, the proposed system can be used in to monitor biofilm formation under different microbiological conditions. In a second experiment, the effect of acetone, a strong organic solvent was investigated to assess its ability to destroy the electroactive biofilm matrix. The CV of the biofilm at the rGO-HBCs electrode immersed in acetone showed bioelectrochemical signal close to a bare electrode, which indicates the removal of the formed bio-electroactive microbial layer, likely by inducing its dissolution (Fig. 7b). Thus, monitoring the destruction of biofilm under different stressors is also possible.
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4. Conclusions P. aeruginosa is the most common model organism for studying biofilm development as it has long been associated with biofilm formation in chronic cystic fibrosis lung infections. Herein, the growth of electrochemically active biofilms of P. aeruginosa was investigated by using a nanostructured microbial electrochemical system. Sensing of the cellular changes was used to monitor the biofilm progression. Evidence from CVs, SEM and EDX experiments demonstrated the reason for the turnover from lower-electrochemical activities to higher electrochemical activities when the switch from planktonic life to the biofilm formation was reached. The proposed system was used to monitor biofilm destruction under different stressors which demonstrate the validity of this model in studying biofilm formation and measurements under a variety of microbiological conditions. Acknowledgments Authors would like to acknowledge Amr Hefnawy for his help in the synthesis of HBCs NPs. Also, all authors, particularly Dr. Rabeay Hassan are grateful for the group leader of Biological Systems Analysis (Prof. Dr Ursula Bilitewski, Helmholtz Centre for Infection Research, HZI, Braunschweig, Germany) for presenting the potentiostat (Gamry Potentiostat/Galvanostat/ZRA G750). Data availability The raw/processed data required to reproduce these findings cannot be shared at this time as the data also forms part of an ongoing study. Appendix A. Supplementary data Supplementary data to this article can be found online at https:// doi.org/10.1016/j.msec.2019.02.112. References [1] J. Azeredo, N.F. Azevedo, R. Briandet, N. Cerca, T. Coenye, A.R. Costa, M. Desvaux, G. Di Bonaventura, M. Hebraud, Z. Jaglic, M. Kacaniova, S. Knochel, A. Lourenco, F. Mergulhao, R.L. Meyer, G. Nychas, M. Simoes, O. Tresse, C. Sternberg, Crit. Rev. Microbiol. (2016) 1–39. [2] A.E. Barsoumian, K. Mende, C.J. Sanchez Jr., M.L. Beckius, J.C. Wenke, C.K. Murray, K.S. Akers, BMC Infect. Dis. 15 (2015) 223.
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Materials Science & Engineering C journal homepage: www.elsevier.com/locate/msec
Online-monitoring of biofilm formation using nanostructured electrode surfaces Mohammed Sedkia,1, Rabeay Y.A. Hassana,b,
⁎,1
, Silvana Andreescuc, Ibrahim M. El-Sherbinya,
T ⁎
a
Nanomaterials Laboratory, Center for Materials Science, Zewail City of Science and Technology, 6th October City, 12588 Giza, Egypt Applied Organic Chemistry Department, National Research Centre (NRC), Dokki, 12622 Giza, Egypt c Department of Chemistry and Biomolecular Science, Clarkson University, Potsdam, NY 13699-5810, USA b
A R T I C LE I N FO
A B S T R A C T
Keywords: Nanostructured surfaces Microbial electrochemistry Monitoring biofilm formation Pseudomonas aeruginosa
The direct monitoring of biofilm formation enables valuable insights into the industrial processes, microbiology, and biomedical applications. Therefore, in the present study, nano-structured bioelectrochemical platforms were designed for sensing the formation of biofilm of P. aeruginosa along with monitoring its electrochemical/morphological changes under different stresses. Through the assay optimizations, the performances of different electrode modifiers such as reduced graphene oxide (rGO) nanosheets, hyperbranched chitosan nanoparticles (HBCs NPs), and rGO-HBCs nano-composite were tested to assess the influence of the electrode materials on biofilm progression. As a need for the anodic respiration, the bioelectrochemical responses of the adhered bacterial cells changed from a non-electrochemically active (planktonic state) to an electrochemically active (biofilm matrix) state. Our results demonstrated that electrode modifications with conductive nanostructured elements is highly sensitive and enable direct assay for the biofilm formation without any preachments. Consequently, the morphological changes in bacterial cell wall, upon switching from the planktonic state to the biofilm matrix were imaged using scanning electron microscopy (SEM), and the changes in cell wall chemical composition were monitored by the Energy Dispersive X-ray analysis (EDX). Thus, the designed microbial electrochemical system (MES) was successfully used to monitor changes in the biofilm matrix under different stresses through direct measurements of electron exchanges.
1. Introduction Biofilms are three-dimensional, heterogeneous community of microbial cells enclosed in an exopolysaccharide matrix that can irreversibly attach to solid surfaces or living tissues [1–4]. Biofilm formation is of high importance in many useful applications such as biocatalysis, degradation of chemical contaminants in wastewater, and microbial fuel cells [5]. However, these biofilms may have serious implications on public health and environment [6–8]. For instance, they cause biofouling, microbially-induced corrosion, contamination of implanted medical devices, and are responsible for the persistent infections in human body resulting in death of millions of people every year [9–12]. The early detection of pathogens, contaminants and virulence biomolecules plays a crucial role in the prevention and treatment of microbial infections [13,14]. Therefore, numerous methods for the detection of pathogens have been developed. However, the big problem is not to measure the microbe in its natural form, the challenge is to detect
the colonized microbes within the biofilms that are not neither accessible nor easy to detect. Microbial electrochemical systems (MESs) were shown to be the most effective techniques for studying the role of the conductive solid surfaces in the enhancements of the biofilm formation [15]. Monitoring the microbe-electrode interaction(s) is the main concept of the MESs. Degradable organic matters provide the necessary nutrition to living organisms. As a result of the microbial catalytic oxidation of the degradable organic compounds, free electrons are liberated through the electron transport chain (ETC). Under the anaerobic conditions, the liberated electrons are transferred to a molecule other than oxygen such as nitrate or sulfate [16]. In the MESs, microbes exploit their catalytic activity to transfer the liberated electrons to an electrode (as an electron acceptor), and consequently electric current is generated. The generated current density is directly proportional to the number of viable microbial cells (i.e. the metabolically active microbes) [14,17]. Three mechanisms have been proposed to explain the electron
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Corresponding authors at: Center for Materials Science, Zewail City of Science and Technology, 6th October City, 12588 Giza, Egypt. E-mail addresses: [email protected] (R.Y.A. Hassan), [email protected] (I.M. El-Sherbiny). 1 Both are sharing the first authorship. https://doi.org/10.1016/j.msec.2019.02.112 Received 28 September 2018; Received in revised form 4 February 2019; Accepted 28 February 2019 Available online 04 March 2019 0928-4931/ © 2019 Elsevier B.V. All rights reserved.
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Fig. 1. (a) XRD analysis of the thermally reduced graphene oxide (rGO) nanosheets, (b) FTIR spectra of rGO, HBCs and rGO-HBCs nanostructures, (c & d) SEM images of rGO-HBCs nanostructure.
used on both planktonic and biofilm cultures to monitor changes in cell wall chemical composition and characterize their physical connections.
transfer at microbe-electrode interface by exoelectrogens without artificial mediators. Firstly, certain exoelectrogenic bacteria can transfer electrons to an electrode surface through soluble redox active compounds secreted by microorganisms [18,19]. Other microbes were able to directly transfer electrons to anodes via outer cell conductive membrane localized proteins [20]. Lastly, electrons are delivered directly to the electrode through the naturally produced bacterial nanowires [21,22]. It was concluded that strongly adhered microbes produce the higher bioelectrochemical signals. Therefore, the formation of a biofilm on a conductive electrode surface could be detected [23,24]. However, the biofilm formation is strongly dependent on the electrode material and the morphological structures of the working electrode [25,26]. Hence, a better understanding of the microbial communication in relation to the morphological and physicochemical properties of the electrode material is needed. Herein, we report a microbial electrochemical sensor system for reliable and rapid monitoring and investigation of the Pseudomonas aeruginosa biofilm formation on modified electrodes. P. aeruginosa was selected as a target organism because of its high ability to form biofilms in wounds, burns, implanted devices, catheters, urinary tract, and in lungs of patients having cystic fibrosis [27,28]. We designed nanostructured bio-electrochemical systems to explore the P. aeruginosa–electrode interactions using cyclic voltammetry (CV) measurements and the scanning electron microscopy (SEM) were used to investigate the morphological characteristics of the biofilms. EDX spectroscopy was
2. Materials and methods All data generated or analyzed during this study are included in this published article and its Supplementary Information files. D (+) glucose was purchased from Fine-Chem Limited. LuriaBertani (LB) liquid media were used for cells cultivation. The electrode modifiers of reduced graphene oxide (rGO) nanosheets, hyperbranched chitosan nanoparticles (HBCs NPs), and rGO-HBCs nanostructure were synthesized in our lab. Paraffin oil and graphite powder used in carbon paste electrode (CPE) preparation were purchased from Fluka and Sigma Aldrich, respectively. Ciprofloxacin [1-cyclopropyl-6-fluoro-4oxo-7-(piperazin-1-yl)-1,4-dihydroquinoline-3-carboxylic acid] was bought from Sigma-Aldrich. All OD600 measurements were conducted by Thermo-Scientific Evolution 600 UV–Vis spectrophotometer. All electrochemical measurements were carried out using a computercontrolled Gamry Potentiostat/Galvanostat/ZRA G750 (Gamry, Pennsylvania, USA). The three-electrode electrochemical system used in these measurements comprised a modified-CPE with a surface area of 0.5 cm2 as the working electrode, a calomel electrode as a reference, and a Pt disc as auxiliary electrode.
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2.1. Synthesis and characterization of HBCs, rGO, rGO-HBCs nanostructures Hyperbranched chitosan nanoparticles (HBCs NPs) were prepared using our reported method [29] and then conjugated with rGO nanosheets via on/off sonication for 10 min. rGO nanosheets were produced by thermal reduction, at 200 °C for 2 h, of the GO synthesized using an improved Hummer's method [30]. The synthesized rGO nanosheets were characterized by X-ray Diffraction (XRD) (Shimadzu, Kyoto, Japan) in the 2θ range of 11° to 29°, with a step size of 0.01°, and a time per step of 0.5 s. In addition, the synthesized rGO, HBCs, and rGO-HBCs nanostructures were investigated using Fourier Transform Infrared (FTIR) spectroscopy (Thermo Scientific Nicolet iS10 FT-IR), and imaged by scanning electron microscopy (SEM) (FEI Quanta 250).
2.2. Preparation of the nanostructured electrodes The modification of the working electrodes was conducted by mixing 0.9 g synthetic graphite powder with 0.1 g of the modifiers (HBCs, rGO or rGO-HBCs nanomaterials) with 0.4 ml paraffin oil in a small mortar. The prepared pastes were filled and compressed into the electrode assembly producing HBCs-CPE, rGO-CPE, and rGO-HBCs-CPE electrodes with a surface area of 0.5 cm2. The electrode surface was polished before each measurement via scratching its tip against a smooth wet paper and electrochemically activated in electrolyte (0.01 M KCl) by 5 cycles in a potential range of −0.3 to 1.00 V. The measurement time was fixed at 15 S and the scan rate was set at 50 mV/ S.
2.3. Testing electrode materials' effect on biofilm formation P. aeruginosa's three cells suspensions of OD600 = 0.1 were incubated with the previously prepared and tested electrodes (Bare CPE, HBCs-CPE and rGO-HBCs-CPE) under anaerobic conditions for 20 h, and then the corresponding cyclic voltammetry (CV) measurements were conducted. The electrodes were washed thoroughly with PBS before each measurement. In addition, the electrochemical measurements were done in KCl 0.01 M, away from any secreted biomolecules. Moreover, bare-CPE and rGO-HBCs-CPE were incubated with P. aeruginosa at the same abovementioned conditions for five days and the CV measurements were carried out on the washed biofilm-electrode at different time intervals.
2.4. SEM and EDX analyses A carbon screen printed electrode was covered with rGO-HBCs and incubated with bacterial cells in LB media for five days under anaerobic and static conditions at 37 °C. It was then washed, dried, covered by a thin layer of sputtered gold to enhance the image quality and imaged by SEM (FEI Quanta 250). The elemental composition of different sections was analyzed by the Energy Dispersive X-ray (EDX) analysis. In addition, an EDX mapping analysis was conducted on the mature biofilm without gold sputtering to confirm the concentration of iron species at the grown biofilm matrices.
2.5. The effect of planktonic cells adherence on electrode activity
Fig. 2. (a) The CV of the biofilm formation at different electrode materials (Bare CPE, HBCs-CPE, rGO-HBCs-CPE) after 20 h of incubation time, (b) The CV measurements of the biofilm formation monitored at the bare CPE electrode at different time intervals from 1.5 days to 5.0 days, (c) The same as (b) using the rGO-HBCs-CPE electrode. All measurements were carried out in 0.1 KCl with a sensing time of 15 s and a scan rate of 50 mV/S.
The rGO-HBCs-CPE electrode was incubated with P. aeruginosa cells suspension with an OD600 = 0.1 in PBS (0.1 M, pH = 7.4) and glucose concentration of 8 g/L for 3 h at 37 °C, and the corresponding CVs were recorded at different time intervals. The experiment was repeated under the same conditions after removal of bacterial cells (supernatant) for each measurement. 180
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Fig. 3. SEM images of (a) P. aeruginosa planktonic cells at the carbon screen printed electrode surface, (b) the slimy biofilm matrix of P. aeruginosa at the rGO-HBCsCPE electrode after incubation for five days in LB media, (c) a closer view showing the connection between biofilm bacterial cells through flagella and pili.
confirmed by the disappearance of the hydroxyl stretching peaks at 3400 cm−1. The very small peaks of rGO appeared at 1723 and 1750 cm−1 can be attributed to the remaining ketone carbonyl and ester carbonyl of the partially rGO, respectively. The presence of a very small portion of partially rGO is intentionally made to enhance the binding ability between rGO and HBCs. On the other hand, the on/off sonication of rGO with HBCs resulted in successful formation of rGOHBCs nanocomposite as confirmed by the presence of characteristic FTIR peaks for both materials. Results of FTIR analysis are shown in Fig. 1b. The morphology of the prepared nanocomposite (rGO-HBCs) was studied using SEM imaging which showed semi-spherical HBCs with particle sizes around 400 nm interconnected within the rGO nanosheets network, Fig. 1c & d. Additional microscopic information about the prepared rGO and the rGO-HBCs after immersion in aqueous media for a week are provided in supplementary file (Fig. S1).
2.6. The effect of ciprofloxacin and acetone on P. aeruginosa biofilm After formation of a mature P. aeruginosa biofilm at the rGO-HBCsCPE electrode, the electrode was washed several times with PBS and then incubated with ciprofloxacin at a concentration twice as high as the MIC for two days and the corresponding CV measurements were collected. In addition, the mature electrode-biofilm was immersed in acetone for 10 min and then measured using CV to assess the degradation of the biofilm.
3. Results and discussion 3.1. Synthesis and characterization of rGO, HBCs and rGO-HBCs nanostructures Hyperbranched chitosan nanoparticles (HBCs NPs) were prepared using our reported method [31] and then conjugated with reduced graphene oxide (rGO) nanosheets with on/off sonication for 10 min. rGO nanosheets were produced via the thermal reduction of the improved Hummer's method synthesized GO [30] at 200 °C for 1 h. The Xray diffraction (XRD) analysis of the powder sample of rGO nanosheets was carried out in the 2theta range of 11° to 29°. The results, Fig. 1a, showed a strong peak at 2Ɵ of 23° for rGO [32] with a small shoulder at 12° for partially reduced GO with no peaks for graphite at 2Ɵ of 26°, confirming the successful reduction process. The FTIR analysis of the core-shell hyperbranched Cs matched previously reported data [31] with stretching peaks of its terminal amine groups noted at 3288–3388 cm−1, stretching peaks of ester carbonyl at 1723 cm−1 and the stretching peaks of CeH at 2800–2900 cm−1. In addition, the thermal reduction of GO at 200 °C in ambient atmosphere was
3.2. Influence of electrode materials on biofilm formation The working electrode material is the key player in developing MESs, whereas the microbial kinetics is directly linked to the reactions occurring at the electrode surface [33–37,38]. Previously, we have developed an efficient MES using rGO-HBCs nanostructure as an electrode modifier for detection of cell viability and metabolic activity of P. aeruginosa [39]. Here we extended this concept to study the effect of surface roughness, wettability and the electron transfer properties of the nanocomposite to identify the driving parameters that are responsible for the enhanced biofilm formation of P. aeruginosa. To understand the contribution of each material in the matrix, three different working electrodes were prepared: Bare-CPE, HBCs-CPE, and rGOHBCs-CPE. The electrodes were incubated in a fresh individual bacterial 181
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Fig. 4. A schematic illustration of the biofilm growth monitored by measuring the direct extracellular electron transfer ability of the biofilm matrix.
nanostructured electrode surface.
culture containing a suspension of P. aeruginosa cells (OD600 = 0.1) in anaerobic conditions and the corresponding CVs were recorded. Prior to each CV measurement, the electrode was washed thoroughly with phosphate buffer to remove extracellular secretions. The strongest bioelectrochemical responses were observed at the rGO-HBCs-CPE electrode, Fig. 2a. This high bioelectrochemical performance is attributed to the interconnected network and high conductivity of the rGO and HBCs nanocomposite network possessing fast electron transfer ability and high surface roughness in addition to the adhesion properties of HBCs polymeric nanoparticles. By comparison, bare-CPE and HBCs-CPE each alone, they did not show strong bioelectrochemical signals. Further, bare-CPE and rGO-HBCs-CPE were incubated with P. aeruginosa in the same conditions for five days and the CV measurements were carried out on the washed biofilm-electrode at different time intervals, Fig. 2b & c. The adhered cells to the CPE did not show obvious electrochemical signals indicating lower electrode conductivity, Fig. 2b. On the other hand, the peak currents of the biofilm(rGO-HBCs-CPE) electrode increased significantly from tens of micro amperes to around 3.0 mA, Fig. 2c. The current increased gradually in both electrodes until it reached a constant value after which no further increase was detected. This increase in electron transfer demonstrates formation of matured biofilms that enable direct electrical communication between the active P. aeruginosa biofilm and the
3.3. P. aeruginosa biofilms: how are extracellular electrons transferred? Physical connections through the bacterial appendages, microbial nanowires, cyt-c and ferric iron were identified as main contributors to the biofilm assisted-bioelectrochemical signals in few microorganisms such as Geobacter sulfurreducens, Shewanella oneidensis, and Thiobacillus denitrificans [40–42]. Cellular interactions and the contributing factors in the formation of P. aeruginosa biofilms (i.e. the P. aeruginosa-electrode interactions) were less explored. The direct continuous increase of electrical current that is generated from the connectivity of adhered bacterial cells to the nano-electrode surface is indicating formation of mature electrochemically active biofilms. To determine the microbial connectors, the morphology of the adhered bacterial cells at the screen-printed electrode modified surface in planktonic state and biofilm matrices were imaged using SEM. The images shown in Fig. 3a illustrated the typical bacillus (rod-like) morphology of planktonic cells without any appendages or physical connections between them. On the other hand, the SEM images of the P. aeruginosa biofilm, after incubation for five days with the electrode surface, demonstrated the coverage of bacterial cells in slimy secretions of polysaccharides as seen in Fig. 3b, and the direct physical 182
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Fig. 5. The SEM-EDX of P. aeruginosa biofilm illustrating the connection between biofilm cells and the percentage of iron at different sections; (a) at the cellular structure of planktonic cells, (b) at the cell wall of P. aeruginosa biofilm cells, and (c) at the flagella connecting biofilm cells to each other. The samples were covered with a thin gold sputtered film to enhance the images' resolution.
bacterial cells and the biofilm formed layer were analyzed by EDXspectroscopy. Results showed in Fig. 5 identified the elemental composition of planktonic bacterial cells, biofilm matrix and the conductive flagella. Surprisingly, the content of the biofilm outer-layer exhibited iron-complex concentrations ranging from 1.78% in biofilm matrix to 2.99% in flagella. Nevertheless, the planktonic cells contained negligible traces of iron (less than 0.31%). The EDX mapping results of the P. aeruginosa mature biofilm showed higher distribution of iron species at the thicker and more complex biofilm regions, confirming a direct relationship between the conductivity of the biofilm and the existence of iron species. This is illustrated in Fig. 6b as related to the captured images of the mapped EDX (Fig. 6a) on ETD detector. Typically, microorganisms in biofilms live in a home-made matrix of extracellular polymeric substances (EPS) [43]. The EPS support the mechanical stability of biofilms, mediate cell adhesion to the solid
connections between almost all cells with each other and with the substrate and biofilm matrix, Fig. 3c. The SEM images of these bacterial nanowires match the morphology and dimensions of flagella whose role in motility and/or the initial cellto-surface interactions has been demonstrated by O'Toole et al. They have shown that the defective flagella mutant (flgK) did not develop microcolonies on the solid substrate over the course of the experiments, suggesting the necessity of the flagella in microcolony formation [13]. Our results indicate that the signaling between the P. aeruginosa cells and the direct extracellular electron transfer observed may occur through purposely produced bacterial nanowires and their conductive elements. A schematic illustration of the biofilm growth at different stages of formation monitored by the corresponding conductivity of the biofilm matrix is shown in Fig. 4. Additional confirmatory SEM images are provided in the supplementary information (Fig. S3). The composition of the outer-layer of the dispersions of fresh 183
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(a)
(b)
Fig. 6. (a) The image of the mapped area using the ETD detector, and (b) EDX mapping showing distribution of iron of P. aeruginosa biofilm at the surface of rGOHBCs.
Fig. 7. (a) CVs of P. aeruginosa mature biofilm at the RGO-HBCs modified electrode in the presence of ciprofloxacin for two days, and (b) effect of the incubation of the formed biofilm-modified electrode in acetone for 10 min.
3.4. Adherence of the planktonic cells to the electrode surface
surfaces and form a cohesive, three-dimensional polymer network that interconnects biofilm cells. In most biofilms, living cells represent less than 10% of the total mass, whereas the rest of the matrix consists of water and secretions. The composition of biofilm matrices is dependent on the bacterial species and the environmental conditions, but in general they contain components such as polysaccharides, proteins and extracellular DNA [44,45]. From a mechanistic point of view, the extracellular DNA release in P. aeruginosa PAO1 biofilms is regulated by the iron availability [46]. On the other hand, the quorum-sensing systems are strongly involved in the formation of extracellular DNA in P. aeruginosa biofilms. Other reports showed that the level of iron is not only affecting the P. aeruginosa DNA release and biofilm formation but also the maturation of the biofilm and its structural development. Since iron was identified as a critical factor in the interaction between the pathogen and the host, in vitro studies proved that the sub-inhibitory concentrations of the iron chelator lactoferrin block the ability of P. aeruginosa biofilms formation. Thus, iron serves as a signaling element for biofilm development. Besides, P. aeruginosa produces siderophores with high-affinity for iron uptake that can not only chelate free iron but also ‘strip’ iron from host proteins such as lactoferrin [47–51].
The HBCs-rGO-CPE was incubated with cells suspension in a solution of PBS and glucose for 3 h, and the corresponding current of the secreted biomolecules was measured at different time intervals. Measurements were also performed on the supernatant after removal of bacterial cells in each time point in the same conditions. The electron transfer and the oxidation currents were inhibited in the presence of bacterial cells due to their adherence to electrode surface. The decrease in surface conductivity due to the attachment of cells reflects the poor conductivity of planktonic cells (Table S1). These findings support the assumption that the cell wall structure of P. aeruginosa changed from non-conductive planktonic cells to a conductive surface through the nanostructured elements of the electrode which promote biofilm formation. Conductivity has been gained by the production of natural nanowires rich in iron-containing materials (i.e. cytochromes or oxides). 3.5. Effect of stressors on mature biofilm formation rGO-HBCs modified nanostructured electrodes were incubated in bacterial cultures in absence and presence of ciprofloxacin, an 184
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antibiotic that is known to inactivate P. aeruginosa. The growth of electroactive biofilms and their electrochemical signals at the electrode surface was monitored to determine the effect of stress induced by ciprofloxacin. In these experiments, cells were exposed to the minimum inhibitory concentration (MIC), determined using the standard cell proliferation reagents WST-1 kit as shown in Fig. S4. The bioelectrochemical signals measured for the treated and untreated biofilms shown in Fig. 7a indicate that while the signals of the untreated biofilm were in a continuous increase, the activity of the treated biofilm was inhibited over the treatment time. The decrease in signals can be explained by the good penetration ability of ciprofloxacin through the biofilm layers, leading to partial destruction of the biofilm matrix [52,53]. Since P. aeruginosa biofilm is susceptible to ciprofloxacin, the proposed system can be used in to monitor biofilm formation under different microbiological conditions. In a second experiment, the effect of acetone, a strong organic solvent was investigated to assess its ability to destroy the electroactive biofilm matrix. The CV of the biofilm at the rGO-HBCs electrode immersed in acetone showed bioelectrochemical signal close to a bare electrode, which indicates the removal of the formed bio-electroactive microbial layer, likely by inducing its dissolution (Fig. 7b). Thus, monitoring the destruction of biofilm under different stressors is also possible.
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4. Conclusions P. aeruginosa is the most common model organism for studying biofilm development as it has long been associated with biofilm formation in chronic cystic fibrosis lung infections. Herein, the growth of electrochemically active biofilms of P. aeruginosa was investigated by using a nanostructured microbial electrochemical system. Sensing of the cellular changes was used to monitor the biofilm progression. Evidence from CVs, SEM and EDX experiments demonstrated the reason for the turnover from lower-electrochemical activities to higher electrochemical activities when the switch from planktonic life to the biofilm formation was reached. The proposed system was used to monitor biofilm destruction under different stressors which demonstrate the validity of this model in studying biofilm formation and measurements under a variety of microbiological conditions. Acknowledgments Authors would like to acknowledge Amr Hefnawy for his help in the synthesis of HBCs NPs. Also, all authors, particularly Dr. Rabeay Hassan are grateful for the group leader of Biological Systems Analysis (Prof. Dr Ursula Bilitewski, Helmholtz Centre for Infection Research, HZI, Braunschweig, Germany) for presenting the potentiostat (Gamry Potentiostat/Galvanostat/ZRA G750). Data availability The raw/processed data required to reproduce these findings cannot be shared at this time as the data also forms part of an ongoing study. Appendix A. Supplementary data Supplementary data to this article can be found online at https:// doi.org/10.1016/j.msec.2019.02.112. References [1] J. Azeredo, N.F. Azevedo, R. Briandet, N. Cerca, T. Coenye, A.R. Costa, M. Desvaux, G. Di Bonaventura, M. Hebraud, Z. Jaglic, M. Kacaniova, S. Knochel, A. Lourenco, F. Mergulhao, R.L. Meyer, G. Nychas, M. Simoes, O. Tresse, C. Sternberg, Crit. Rev. Microbiol. (2016) 1–39. [2] A.E. Barsoumian, K. Mende, C.J. Sanchez Jr., M.L. Beckius, J.C. Wenke, C.K. Murray, K.S. Akers, BMC Infect. Dis. 15 (2015) 223.
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