Interfacial engineering of graphenic carbon electrodes by antimicrobial polyhexamethylene guanidine hydrochloride for ultrasensitive bacterial detection

Carbon 159 (2020) 185e194

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Interfacial engineering of graphenic carbon electrodes by antimicrobial polyhexamethylene guanidine hydrochloride for ultrasensitive bacterial detection Lingli Zhu a, Liang Wang a, **, Xinqi Zhang a, Ting Li a, Yilei Wang a, Muhammad Adil Riaz b, Xiao Sui b, Ziwen Yuan b, Yuan Chen b, * a

Tianjin Key Laboratory of Organic Solar Cells and Photochemical Conversion, School of Chemistry and Chemical Engineering, Tianjin University of Technology, Tianjin, 300384, China School of Chemical and Biomolecular Engineering, The University of Sydney, NSW, 2006, Australia

b

a r t i c l e i n f o

a b s t r a c t

Article history: Received 31 October 2019 Received in revised form 13 December 2019 Accepted 14 December 2019 Available online 16 December 2019

Pathogenic bacteria are severe threats to public health. Existing bacterial detection methods are often time-consuming and costly. Impedance-based electrochemical sensors using carbon electrodes have been explored for bacterial detection. However, the pristine carbon surface is inefficient in attracting bacterial cells. Here, we demonstrate an interfacial engineering method for graphenic carbon (GC) electrodes by using a common antibacterial material to achieve ultrasensitive bacterial detection. Polyhexamethylene guanidine hydrochloride (PHMG) is a broad-spectrum antimicrobial material. We first conjugated perylene bisimide (PBI) with PHMG to form a new PBI-PHMG compound. PBI-PHMG with an optimal PBI content can retain PHMG’s intrinsic antibacterial activity while PBI anchors PBI-PHMG on the GC surface via strong pi-pi interactions. The resulting PBI-PHMG modified GC electrodes have positively charged surfaces, which effectively attract and inactivate bacterial cells. Cytoplasm materials released from damaged cells change the impedance of GC electrodes significantly, leading to considerably enhanced bacterial detection sensitivity down to 2 CFU mL
1 for both E. coli and S. aureus within 30 min of incubation time. The facile preparation, short detection time and high sensitivity of the impedance sensors based on the modified GC electrodes are promising for practical applications such as portable devices for point-of-use bacterial detection. © 2019 Elsevier Ltd. All rights reserved.

Keywords: Graphenic carbon Impedance sensor Bacterial detection Polyhexamethylene guanidine hydrochloride Antibacterial material

1. Introduction The bacterial infection is a significant threat to human life. Notably, the potential spread of drug-resistant bacteria may lead to devastating death tolls and high healthcare costs [1e3]. Rapid detection of pathogenic bacteria is essential for point-of-care diagnostic tools [4,5], which can help prevent bacterial infections and increase patient survival rates. Traditional methods of detecting pathogenic bacteria include plate colony counting, polymerase chain reaction (PCR), and enzyme-linked immunosorbent assay (ELISA) [6]. The colony counting method requires separation,

* Corresponding author. ** Corresponding author. E-mail addresses: [email protected] (L. Wang), [email protected] (Y. Chen). https://doi.org/10.1016/j.carbon.2019.12.035 0008-6223/© 2019 Elsevier Ltd. All rights reserved.

identification, culturing and counting of bacterial cells over a few days, which is labor-intensive and time-consuming. Although PCR and ELISA are much faster, they require expensive instruments and reagents [7e9]. Further, cell lysis and DNA extraction required by sample pretreatment for PCR and ELISA may bring additional complications [10]. Using electrochemical sensors for bacterial detection offers the advantages of high specificity, selectivity, and sensitivity, low cost, and high speed [11e14]. Impedance-based electrochemical sensors monitor impedance changes of electrodes upon bacterial attachment [15]. They do not require cell labeling, are easy to operate, provide reproducible results, and are also compatible with complex solutions, such as food, beverages, and bio-samples [16e21]. The current research of impedancebased bacterial sensors focuses on improving their performance by using microelectrode arrays to enhance their signal intensity [22,23] or immobilizing bacteria-specific biomolecules on electrodes (e.g., peptide, DNA, or antibodies) to enhance bacterial

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attachment [24e28]. However, the complex fabrication of microelectrode arrays and the instability of biomolecules limit their practical applications. It is desirable to develop simpler, low cost, and more sensitive electrodes for impedance-based bacterial sensors. Graphenic carbon (GC) materials consist of mostly hydrophobic sp2 carbon, including 2D graphene materials, as well as 3D carbonaceous solids containing graphene layers. They can be used as electrode materials in impedance-based bacterial sensors. However, sensors based on common GC materials have two problems. First, the hydrophobic surface of the pristine carbon surface is inefficient for bacterial cell attachment [29,30]. Although O-containing functional groups can be introduced on a carbon surface to increase its hydrophilicity, these functional groups would significantly compromise the electrical conductivity of carbon materials. Second, although GC materials, especially graphene and graphene oxide, have some antibacterial activity [31,32], their antibacterial activity is usually not high, often requiring several hours to inactivate pathogenic bacteria completely [33,34]. We propose a strategy to modify the interface between carbon electrodes and bacterial cells by assembling other antibacterial materials on the surface of carbon electrodes. The modified carbon electrodes may help to capture, inactivate, and damage bacterial cells more efficiently. Cytoplasm materials released from damaged bacterial cells trapped on carbon electrodes are expected to significantly change their electrical conductivity, resulting in much more sensitive bacterial detection sensors. Polyhexamethylene guanidine hydrochloride (PHMG) is an antimicrobial material with a broad-spectrum activity against Gram-positive and Gram-negative bacteria, fungi, yeasts, and viruses [35,36]. At a concentration lower than 1 wt%, PHMG is less toxic or harmless to humans and animals than many other commonly used disinfectants [35]. Thus, PHMG has been widely used as an antiseptic in the food industry and medical field, such as mouthwash solutions [37,38] and solid surface disinfectants [39]. PHMG possesses multiple positive charges, which disturb negative charges presented on cell walls of microorganisms. It may diffuse through cell membranes and bind to phospholipid molecules in lipid bilayers, which destabilizes the osmotic equilibrium and eventually destroys cytoplasmic membranes, causing the leakage of cell contents [40]. Thus, we propose to assemble the antimicrobial PHMG on carbon electrodes, which may lead to much more sensitive impedance-based bacterial detection sensors. Here, we first designed and synthesized a new perylene bisimide (PBI)-containing PHMG (PBI-PHMG) compound. PBI was introduced to anchor PHMG quickly on GC materials via strong pipi interactions between PBI and graphenic basial planes. Next, we compared the antimicrobial activity of PBI-PHMG with PHMG to ensure that PBI-PHMG still retains the antimicrobial activity of PHMG. Besides, GC electrodes were prepared by growing GC materials on porous Ni foams. Afterward, PBI-PHMG was assembled on GC electrodes to construct PBI-PHMG conjugated GC electrodes (denoted as PPG electrodes). The morphological and physicochemical properties of PPG electrodes were characterized by scanning electron microscopy (SEM) and X-ray photoelectron spectroscopy (XPS). The bacterial detection performance of GC and PPG electrodes was compared. Our results show that conjugated PBI-PHMG changes the interface between carbon surface and bacterial cells, which significantly increases the binding affinity of bacteria on GC electrodes. The resulting PPG electrodes can efficiently detect both E. coli and S. aureus at different concentrations with short detection time.

2. Experimental section 2.1. Chemicals and cell culture media Perylene-3,4,9,10-tetracarboxylic dianhydride (PDA), PHMG, and shellac biopolymer were from Energy Chemical Co., Ltd, China. Porous Ni foams were from Shanxi Lizhiyuan Battery Material Co., Ltd, China. Isopropanol and ethanol were from Tianjin Guangfu Fine Chemical Research Institute, China. All chemicals were analytical grade and used as received. The preparation of Luria-Bertani (LB) broth and LB agar followed the method described in the Supplementary data (SD). 2.2. Synthesis of PBI-PHMG Five PBI-PHMB samples were synthesized by tuning the feeding amount of PDA in the synthesis from 1.25, 2.50, 5.00, 10.0 to 15.0 wt %, and they are denoted as PBI1.25-PHMB, PBI2.50-PHMB, PBI5.00PHMB, PBI10.0-PHMB, and PBI15.0-PHMB, respectively. Taking the synthesis of PBI5.00-PHMG as an example, PHMG (1.9 g) was mixed with PDA (100 mg) in 8 mL of n-butanol. The red mixture was refluxed at 120  C for 16 h in an N2 atmosphere. The resulting upper solvent was decanted, while the remaining dark-red solid (i.e., crude PBI5.00-PHMG) was dispersed in a small amount of dilute hydrochloric acid to recover guanidine hydrochloride groups, which decompose during the reflux. 100 mL of ethanol was then used to dissolve the resulting solid, and the solution was centrifuged at 5000 rpm for 10 min to eliminate unreacted solid PDA particles. Afterward, the supernatants were evaporated using a rotary evaporator to yield a solid material, which was washed by 100 mL of isopropanol for three times, and then dried under vacuum to yield a dark red powder (i.e., PBI5.00-PHMG). 2.3. Preparation of GC and PPG electrodes Porous Ni foams (6.0  50  1.0 mm) were first cleaned by sonication in acetone for 10 min to remove organic contamination. Cleaned Ni foams were dipped in a shellac biopolymer solution for 10 min, and then naturally dried for 24 h. The yellow color shellac solution was prepared by dissolving pure shellac solids (400 mg) in ethanol (20 mL) at room temperature. The shellac-coated Ni foams were then annealed at 600  C in N2 for 30 min, leading to the formation of a GC coating on the Ni surface, which is a porous carbon material with multiple graphenic layers. PPG electrodes were prepared by incubating GC electrodes in 10 mL of PBI-PHMG aqueous solution (1.5 mg mL
1) for 24 h and then washed by DI water. 2.4. Physicochemical property characterization 1

H NMR spectra of PBI-PHMG were obtained using a Fouriertransform nuclear magnetic resonance (FT-NMR) spectrometer (AVIII 400, Bruker, Switzerland). Ultraviolet (UV) and fluorescence (FL) spectra of aqueous solutions of the five PBI-PHMG samples (0.25 mg mL
1) were tested using a UVevisible (UVeVis) spectroscopy (U-3310 spectrophotometer, Hitachi, Japan) and an FL spectroscopy (F-4500, Hitachi, Japan), respectively. The physical structures of Ni, GC, and PPG electrodes were examined by an SEM (Merlin Compact, Carl Zeiss, Germany) at 10.00 kV. Raman spectra of PBI-PHMG, GC, and PPG electrodes were recorded using a highresolution laser confocal micro-Raman spectrometer (l ¼ 532 nm) (Horiba Evolution, Horiba Jobin Yvon, France). Their surface elemental compositions were analyzed by an XPS (Escalab250Xi, Thermo Scientific, Britain).

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2.5. Antibacterial activity tests of PHMG and PBI-PHMG The antibacterial activities of PHMG and PBI-PHMG were tested against E. coli and S. aureus. Cryopreserved strains of the bacteria were transferred into LB broth, followed by shaking culture in a shaker at 37  C for 16 h until the bacterial concentration reached 109 CFU mL
1. The cultured bacterial cells were collected by centrifugation and washed with a phosphate-buffered saline solution (PBS) to remove residual growth media. These bacterial cells (105 CFU mL
1) were then incubated with PHMG and PBI-PHMG solution with a concentration of 6.25, 12.5, 25.0, 37.5, and 50.0 mg mL
1 at 37  C for 15 min, respectively. Afterward, the bacterial suspensions were plated on LB agar plates. The plates were allowed to grow for 16 h in a biological incubator at 37  C. The viabilities of E. coli and S. aureus were determined by the colony counting method. Colonies were counted and compared to those on control plates to calculate their loss of viability rates according to the following equation: Loss of viability % ¼ (counts of the control sample
counts of samples after incubation with bacterial suspensions)/counts of the control sample  100%. At least four runs were tested for each condition to determine the standard deviations. 2.6. Impedance-based electrochemical detection of bacteria Electrochemical measurements were carried out on an electrochemical workstation (CHI660E, Shanghai Chenhua Instrument Co., Ltd., China) using a three-electrode configuration. A platinum sheet (2.0 cm2) and an Ag/AgCl electrode were used as the counter and reference electrodes, respectively. A GC or PPG electrode with a size of 1.44 cm2 (6  24 mm) served as the working electrode. The electrodes were immersed in 0.9 wt% saline solutions containing different concentrations of bacteria at room temperature. The frequency was scanned from 0.1 to 10 kHz. The impedance and the time associated impedance variation were measured. The charge transfer resistant (Rct) was calculated from the diameter of semicircles in Nyquist diagrams, which were simulated by the ZSimpWin software. 2.7. Bacterial morphology on electrodes After electrochemical tests, GC and PPG electrodes were immersed into 2.5 wt% glutaraldehyde solution to fix bacteria on the electrodes. The fixed bacterial cells were gradually dehydrated with a series of ethanol/water mixed solutions with the ethanol concentrations ranging from 10, 30, 50, 70, and 90 vol%, to pure ethanol, and then freeze-dried under vacuum. The dried bacterial cells were examined by the SEM under the electron acceleration voltage of 10.00 kV. 3. Results and discussion 3.1. Synthesis of PBI-PHMG As illustrated in Fig. 1, PBI-PHMG was synthesized using a onestep reaction, in which the end amino groups of PHMG were amidated by the anhydride groups of PDA. By tuning the feeding amount of PDA from 1.25 to 15.0 wt%, five PBI-PHMG samples were obtained. Due to the high molecular weight of PHMG, all PBI-PHMG samples can be dissolved in water. Fig. 2 shows that they have the characteristic UV absorption and fluorescence emission peaks originated from PBI. Fig. 2a displays that the intensity of absorption peaks increases with the increase of the PDA feeding amount, and the rising trend slows down when the PDA feeding amount is higher than 5.0 wt%. In comparison, the intensity of fluorescence

Fig. 1. Schematic illustrations of (a) the synthesis of PBI-PHMB, (b) the preparation of PPG electrodes, and (c) the application of PPG electrodes as an impedance-based electrochemical sensor for bacterial detection. (A colour version of this figure can be viewed online.)

peaks in Fig. 2b first increases with the increase of the PDA feeding amount up to 10.0 wt% and then decreases when it goes up further. Notably, there are red-shifts in the fluorescent spectra of PBI10.0PHMB and PBI15.0-PHMB, suggesting the aggregation of PBI cores at higher PDA feeding amount [41,42]. These optical spectroscopy results indicate that PBI5.00-PHMB has a relatively high PBI content, and its PBI cores have limited aggregation. Next, we focused on characterizing the chemical properties of PBI-PHMG by 1H NMR in CD3OD in comparison with pure PHMB. As shown in Fig. 3, the peaks at around 1.16 ppm (c, d) and 1.43 ppm (b, e) can be ascribed to methylene protons in the hexamethylene segments of PBI-PHMB, while the peaks at around 3.22 ppm are contributed by the CH2 group connected with the guanidine group. The signal at 2.76 ppm comes from protons in the methylene group connected with the amino end-group in PHMB. After the amidation of the amino end-group, a new peak appears at 3.95 ppm in Fig. 3b, which can be assigned to methylene protons near the amide group, indicating the successful conjugation of the perylene structure into PHMB. The perylene-related signal at around 8.70 ppm is also observed in Fig. 3b, which further confirms the incorporation of PBI moieties into PHMB. The molecular weight of PHMB was calculated to be 5147 based on the ratio between the peak “f” and the peaks “c” and “d”. Fig. 3c shows the PBI content in the five PBI-PHMG samples calculated based on the ratio of peak areas at 8.70 and 1.16 ppm (see the detailed calculation method in SD). The PBI content continuously rises with the increase of the PDA feeding amount, and the rising trend slows down when the PDA feeding amount is more than 10.0 wt%. Considering the significant red-shift of PBI10.0-

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Fig. 2. Optical spectra of the five PBI-PHMB samples synthesized with different PDA feeding amounts. (a) UVevis absorption spectra and (b) fluorescence spectra of PBIPHMB in water. The concentration of all PBI-PHMB samples is at 0.25 mg mL
1. (A colour version of this figure can be viewed online.)

PHMG shown in Fig. 2b, the aggregation of PBI cores induced by their strong pi-pi stacking may have taken place in PBI10.0-PHMG, which is not desirable for the subsequent assembly of PBI-PHMG to GC materials. Thus, PBI5.00-PHMG was selected to modify GC electrodes considering its relatively high PBI content and highly dispersed PBI without significant pi-pi conjugations. We further compared the antibacterial activity of pure PHMG and PBI-PHMB against E. coli and S. aureus to ensure PBI-PHMB can retain the antibacterial activity of PHMG. E. coli and S. aureus cells at the density of 105 CFU mL
1 were incubated with PHMG and PBIPHMG at different concentrations ranging from 6.25, 12.5, 25.0, 37.5, to 50.0 mg mL
1 for 15 min, respectively. As shown in Fig. 4, the loss of viability of both types of bacterial cells increases with the increase of the polymer concentration with a similar trend, indicating that both of them can effectively inactivate bacteria. Notably, PBI-PHMG demonstrates a slightly higher bacterial inactivation activity than that of PHMG at the concentrations below 25.0 mg mL
1. PDA serves as a linker to combine multiple PHMG molecules into PBI-PHMG. There are more guanidinium groups in PBI-PHMG than those in PHMG. Besides, the electron-deficient perylene core is beneficial to attract negatively charged bacteria by PBI-PHMG [43]. Thus, the antibacterial activity of PBI-PHMG is higher than that of PHMG. Nevertheless, the observed antibacterial activity of PBI-PHMG indicates that it is applicable to modify the interface of GC electrodes.

Fig. 3. 1H NMR spectra of (a) pure PHMG and (b) PBI5.00-PHMG dissolved in CD3OD. (c) The correlation between the conjugated PBI content in PBI-PHMB samples and the PDA feeding content used in their synthesis.

3.2. Preparation of GC and PPG electrodes As illustrated in Fig. 1, porous Ni foams were first coated with a layer of shellac biopolymer and then dried in air. GC materials were grown on the Ni surface by carbonizing the coated biopolymer layer at 600  C in N2. PBI-PHMG was then self-assembled on the GC materials via pi-pi stacking interactions. Raman spectroscopy was employed to confirm the changes in surface properties. Fig. 5 shows that GC electrodes have characteristic D, G, and 2D peaks at 1343, 1596, and 2710 cm
1, respectively. The D and G peaks correspond to defects or structural disorders, and optical mode vibration of two carbon atoms on an sp2 hybridized carbon network, respectively, while the 2D peaks are the first overtone of the D peak, which is sensitive to structural defects. The Raman spectrum indicates that GC materials grown on Ni foams are not pristine graphene, and they contain defective graphene layers. After conjugated with PBIPHMG, the Raman spectrum of PPG electrodes in Fig. 5 shows typical peaks of PBI at 1304 and 1381 cm
1 [44]. The strong PBI characteristic peaks overlap with the D and G peaks of GC materials. The 2D peak of GC can be clearly identified in the Raman spectrum of PPG. Further, there is a slight red-shift from 1293, 1370 cm
1 to

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electrodes in Fig. 6b can be deconvoluted into three peaks at 284.7, 285.9, and 288.1 eV, corresponding to C atoms in CeC, CeO, and COOH functional groups [45]. After conjugated with PBI-PHMG, the intensity of the peak at 285.9 eV increases significantly while a new peak at 288.7 appears in Fig. 6c. Comparing with the C 1s spectrum of PBI-PHMG, the enhanced peak at 285.9 eV is the overlap of XPS peaks of CeO in GC materials and CeN in PBI-PHMG, and the new peak at 285.9 eV can be attributed to C atoms in guanidine groups. We cannot observe the weak peak at 288.1 eV in Fig. 6c, which may be overshadowed by the intense guanidine peak. Overall, the emerging N 1s peak and the changes in the C 1s spectra from GC to PPG electrodes indicate the successful conjugation of PBI-PHMG onto GC electrode surfaces. SEM was used to compare the surface structures of GC and PPG electrodes. Fig. 7a, c, and e show that multiple-layer graphene sheets finely adhered to the surface of porous Ni foams. Interestingly, after conjugated with PBI-PHMG, some graphene sheets stand vertically toward the surface of Ni foams and form a wrinkled structure. A previous study has used perylene tetracarboxylate as an assistant agent to exfoliate graphite into graphene nanosheets under ultrasonic oscillation, which is driven by pi-pi stacking interactions similar to this study [46]. The wrinkled structures observed in the SEM images (Fig. 7b, d, and f) of PPG electrodes are likely formed by graphene nanosheets exfoliated by PBI-PHMG. These structures are beneficial to increasing the surface area of PPG electrodes. 3.3. Bacterial detection using PPG electrodes

Fig. 4. The loss of viability of (a) E. coli and (b) S. aureus after incubating with PHMG and PBI-PHMG at different concentrations for 15 min. (A colour version of this figure can be viewed online.)

Fig. 5. Raman spectra of PBI-PHMG, GC, and PPG electrodes. (A colour version of this figure can be viewed online.)

1304, 1381 cm
1 when comparing the spectra of PBI-PHMG and PPG electrode, which can be attributed to pi-pi stacking interactions between PBI-PHMG and GC materials [44]. Next, XPS was used to examine the surface chemical characteristics of PBI-PHMG, GC, and PPG electrodes. C1s, O1s, and Ni 2p peaks are at around 285, 530, and 854 eV in the survey scan spectrum of GC electrodes (Fig. 6a). New peaks emerge at 197 and 400 eV in the spectrum of PPG electrodes, which can be attributed to N and Cl in the assembled PBI-PHMG. C1s spectrum of GC

The experimental setup for impedance-based electrochemical detection of bacterial cells is illustrated in Fig. 1. A modified Randles equivalent circuit consisting of solution-phase resistance (Rs), charge-transfer resistance (Rct), double-layer capacitance (Q), and Warburg impedance caused by mass transfer resistance (Zw), was used to describe the electrochemical process taking place on electrodes [27]. The diameter of semicircles of measured Nyquist plots is related to Rct. The Rct values of Ni foam, GC, and PPG electrodes in saline solutions without bacterial cells are shown in Fig. S1 in SD. GC electrodes display a much lower Rct than that of Ni foam. The better hydrophilicity of GC materials improves the interfacial compatibility between GC electrodes and saline solution as compared to Ni foams, leading to the lower Rct. After conjugated with PBI-PHMG, the Rct of PPG electrodes slightly increases, which may be caused by hydrophobic and non-conductive hexamethylene segments in PBI-PHMG. We observed that bacterial cells quickly accumulated on electrodes when electrodes were dipped in bacterial suspensions, and some of them were damaged and leaked cytoplasm materials, accompanied by changes in measured Rct values. Nyquist plots of PPG electrodes in bacterial saline suspensions with different concentrations ranging from 2 to 160 CFU mL
1 were recorded over 30 min to obtain stable signals. Upon interacting with bacterial cell suspensions, bacterial cells were captured and damaged on electrodes, and the cytoplasm leakage from bacterial cells would cause changes on electrode surfaces, resulting in different Rct values. Fig. 8a and b shows that the diameter of semicircles gradually decreases with the increase of bacterial concentration for both E. coli and S. aureus, which suggests that more bacterial cells may be captured at higher concentrations and release more conductive substances from the cytoplasm, leading to lower Rct values. Further, the results in Fig. 8a and b, and Fig. S2 in the supplementary data show that PPG electrodes are highly sensitive to both types of bacteria at low bacterial concentrations (<40 CFU mL
1) but become less sensitive when the bacterial concentration is higher than 40 CFU mL
1. The corresponding variation in Rct (DRct) shows a

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Fig. 6. (a) XPS survey scans of PBI-PHMG, GC, and PPG electrodes; (b) C1s spectrum of GC electrode; (c) C1s spectrum of PPG electrode; (d) C1s spectrum of PBI-PHMG.

Fig. 7. SEM images of GC (a, c, and e) and PPG (b, d, and f) electrodes at different magnifications. The insert of (f) shows GC layers in wrinkled structures.

linear correlation with bacterial concentrations from 2 to 20 CFU mL
1. We also evaluated the bacterial detection ability of the GC electrode as shown in Fig. S3 in SD. The Rct values of GC electrodes do not show a clear correlation with the bacterial concentrations, suggesting that the GC material surface cannot

effectively accumulate and inactivate bacterial cells. To further confirm that the measured DRct is due to the membrane damage-induced bacteria lysis, we carried out two control experiments. In the first control experiment, Rct of a PPG electrode in physiological saline solution (100 mL) was measured first. Next, an E. coli bacterial suspension (1 mL, 105 CFU mL
1) was sonicated with a probe sonicator to break bacterial cells, and the resulting suspension with damaged cells was added to the above saline solution. The Rct of the same PPG electrode was measured again. The Rct decreases significantly after adding the suspension with damaged bacterial cells as shown in Fig. S4 in SD. This result indicates that intercellular components of bacterial cells, such as proteins, DNA, RNA, Kþ, and Naþ, can effectively reduce the Rct of PPG electrodes. The bacterial cell breaking took place on the surface of PPG electrodes directly in our bacterial detection experiments. In the second control experiment, an E. coli bacterial suspension (10 mL, 109 CFU mL
1) was sonicated for 15 min using an ultrasonic cell pulverizer (JY92-2D, Ningbo Xinzhi Biotechnology Co., Ltd.) in an ice water bath. Then, insoluble particles were removed by centrifugation, and the supernatant was gradually diluted into a series of bacterial suspensions at different concentrations. The electrical conductivity of these bacterial suspensions was measured using a conductivity meter (DDS-307A, Shanghai INESA & Scientific Instrument Co., Ltd.). As shown in Fig. S5 in SD, the conductivity increases with the increase of bacterial suspension concentrations. This result supports that bacterial intracellular fluids released from damaged bacterial cells can significantly decrease the conductivity of physiological saline. These two control experiments support that our measured DRct is caused by the leakage of bacterial intracellular fluid from damaged bacterial cells. Overall, the high sensitivity to low concentration bacterial cells by the PPG electrodes can be used to detect bacteria accurately and enable efficient monitoring of microbiological contaminations. Compared with recently reported impedance-based bacterial

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Fig. 8. Impedance spectra of PPG electrodes measured upon incubation with (a) E. coli and (b) S. aureus saline suspensions at different concentrations for 30 min. Variation of charge transfer resistance (DRct) vs. bacterial concentration ((c) E. coli and (d) S. aureus) in the range from 2 to 20 CFU mL
1. (A colour version of this figure can be viewed online.)

detection sensors, the PPG electrodes have two significant advantages [15,20,27,47]. First, they have higher sensitivity and shorter detection time. For example, a microfluidic device developed by Yang et al. showed a bacterial detection limit of 300 CFU mL
1 and a detection time of 1 min [48]. Wu et al. reported a nano-decorated porous impedance electrode sensor that can detect bacterial cells with the concentration down to 10 CFU mL
1 within 20 min [20]. Second, the preparation method of the PPG electrodes is simple and the detection method is also straightforward. For example, recently, toll-like receptor-modified electrodes were reported, which responded selectively to Gram-negative bacteria between 1 and 105 lysed cells mL
1 while remained insensitive to Gram-positive bacteria up to 10 particles mL
1 [49]. However, these electrodes are less stable and more expensive than PPG electrodes. We also studied the changes in Rct values of PPG electrodes for E. coli or S. aureus at different incubation times. Fig. 9 shows that the Rct values of the electrodes incubated with E. coli decrease significantly from 807 to 463 U in the first 12.5 min and then slightly increased to an equilibrium value of around 625 U. This phenomenon may be explained by the quick release of conductive intracellular components in the cytoplasm upon bacterial attachment on PPG electrodes. In contrast, the Rct values of the electrodes incubated with S. aureus first increases with time and then decreases to an equilibrium value. The attachment of S. aureus cells increases the Rct value in the first 5 min. Afterward, the leakage of cytoplasm materials from damaged cells leads to lower Rct values. The observed delay in the first 5 min may be caused by thick peptidoglycan layers of gram-positive bacteria [50], which slow down the destruction of cytoplasmic membranes and the leakage of intracellular components. Previous studies reported that intact bacterial cells attached to electrodes might block the electrical current flow and increase the impedance of electrodes [51,52]. The result in Fig. 9d suggests that the adsorption rate of S. aureus cells on PPG electrodes is faster than their cell lysis rate in the first 5 min. Thus, intact S. aureus cells deposited on PPG electrodes form a

blocking layer, leading to increased Rct values. The different timedependent changes in Rct values may provide a potential method to differentiate gram-positive and gram-negative bacteria. Nevertheless, for both E. coli and S. aureus, the measured Rct values reach the final equilibrium within 25 min, suggesting a fast bacterial cell detection speed. We contribute the short equilibrium time and high sensitivity to the unique interfacial properties of PPG electrodes, which help to accumulate and inactivate bacterial cells efficiently. SEM was used to visualize morphological changes of E. coli and S. aureus cells upon interacting with GC and PPG electrodes for 5 min. Fig. 10a shows that E. coli cells have a rod shape with a smooth surface on GC electrodes, suggesting limited damages caused by GC electrodes. In contrast, upon incubating with PPG electrodes, most E. coli cells attached to the wrinkled graphene surface lose their integrity after incubation for 5 min. As shown in Fig. 10b, black arrows point to severe damages on E. coli cells. S. aureus cells on GC electrodes are regular round spheres (Fig. 10d), while those on PPG electrodes become oblate (Fig. 10e). After 30 min, more severe damages are observed in Fig. 10c and f for both types of bacterial cells. The E. coli cells lose their rod shape while the S. aureus cells do not retain their round shape. Overall, these SEM images show that PPG electrodes induce more severe damages to E. coli cells compared to S. aureus cells in the first 5 min, which is consistent with the slower inactivation of S. aureus cells indicated in Fig. 9. For practical applications, one concern is the impedance-based sensors are highly sensitive to surface morphology and shape of electrodes. We prepared five PPG electrodes using the same procedure with a similar size, and their impedance spectra were measured in a physiological saline solution and shown in Fig. S6 in SD. The five PPG electrodes gave similar impedance curves with different Rct values. Fig. S7 in SD shows the corresponding DRct of these five PPG electrodes, and they all display a linear correlation with bacterial concentrations from 2 to 20 CFU mL
1. Although it is challenging to prepare identical PPG electrodes, we can use DRct or

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Fig. 9. Impedance spectra of PPG electrodes measured from 100 to 0.1 Hz upon incubating with (a) E. coli and (b) S. aureus saline suspensions (20 CFU mL
1) for a different time. The corresponding changes in measured Rct (c and d). Experimental data are shown as dots, and the calculated results are presented as solid lines. (A colour version of this figure can be viewed online.)

Fig. 10. SEM images of E. coli cells on (a) GC, (b) PPG electrodes after incubation for 5 min, and (c) after 30 min. S. aureus cells on (d) GC, (e) PPG electrodes after incubation for 5 min, and (f) after 30 min.

DRct/Rct other than the exact Rct for detecting bacteria. Their high detection sensitivity of PPG electrodes is useful for many applications. We expect that they can also be calibrated first to obtain a linear correlation of a specific electrode if quantitative detection is required. The other concern is that dead cells deposited on PPG electrodes might affect their performance. Our preliminary tests show that PPG electrodes covered with some dead bacterial cells can still provide normal responses when the bacterial concentration is within the detection limit (i.e., less than 40 CFU mL
1). However,

when the bacterial concentration is higher than 40 CFU mL
1, their performance deteriorates. Dead cells can also be partially washed to regenerate PPG electrodes. The regeneration of used PPG electrodes is currently under investigation in our on-going project. 4. Conclusions A new strategy is presented to improve the performance of impedance-based bacterial sensors by interfacial engineering of GC electrodes using antibacterial materials. A commonly used

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antibacterial material, i.e., PHMG, was incorporated in a new PBIcontaining PHMG (PBI-PHMG) compound. Under the optimized loading, PBI5.00-PHMG can easily dissolve in water and retain the intrinsic antibacterial activity of PHMG. The PBI in PBI-PHMG helps its assembly on the GC material surface via pi-pi stacking interactions. The obtained new PPG electrodes have a positivecharged surface, which demonstrates much stronger affinity to bacterial cells. Bacterial cells can quickly accumulate and be inactivated on PPG electrodes. The release of cytoplasm materials from damaged bacterial cells significantly changes the electrical conductivity, leading to improved bacterial detection sensitivity. The bacteria in saline solutions can be detected at the concentration as low as 2 CFU mL
1 within 30 min. The facile and low-cost electrode preparation method and the low detection limit and short detection time of PPG electrodes make them promising for practical applications, such as portable devices for point-of-use bacterial detection. Authorship contribution statement Lingli Zhu: Validation, Methodology, Formal analysis, Writing original draft. Liang Wang: Conceptualization, Formal analysis, Writing - original draft, Writing - review & editing, Funding acquisition. Xinqi Zhang: Validation, Methodology, Formal analysis, Writing - original draft. Ting Li: Validation, Methodology. Yilei Wang: Validation, Methodology, Formal analysis, Writing - review & editing. Muhammad Adil Riaz: Validation, Methodology. Xiao Sui: Validation, Methodology. Ziwen Yuan: Validation, Methodology. Yuan Chen: Conceptualization, Formal analysis, Writing - review & editing, Funding acquisition.

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Declaration of competing interest

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All authors declare that there are no conflicts of interest in this work.

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Acknowledgments This work was supported by the National Natural Science Foundation of China Projects (51143003, 51203116, and 51602217) and the Natural Science Foundation of Tianjin City, China (18JCYBJC17300). Y. Chen acknowledges financial supports from the Australian Research Council under the Future Fellowship scheme (FT160100107) and a Tianjin city-sponsored short-term research visit program.

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