The Pyroelectric Effect Enables Simple and Rapid Evaluation of

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Biological and Medical Applications of Materials and Interfaces

The Pyroelectric Effect Enables Simple and Rapid Evaluation of Biofilm Formation Oriella Gennari, Valentina Marchesano, Romina Rega, Laura Mecozzi, Filomena Nazzaro, Florinda Fratianni, Raffaele Coppola, Luca Masucci, Emanuela Mazzon, Alessia Bramanti, Pietro Ferraro, and Simonetta Grilli ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.8b02815 • Publication Date (Web): 20 Apr 2018 Downloaded from http://pubs.acs.org on April 20, 2018

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ACS Applied Materials & Interfaces

The Pyroelectric Effect Enables Simple and Rapid Evaluation of Biofilm Formation O. Gennari1, V. Marchesano1, R. Rega1, L. Mecozzi1, F. Nazzaro2, F. Fratianni2, R. Coppola3 L. Masucci4, E. Mazzon5, A. Bramanti1,5 P. Ferraro1and S. Grilli1* 1

Institute of Applied Sciences & Intelligent Systems, National Research Council (CNR-ISASI), Via Campi Flegrei 34, 80078 Pozzuoli (NA), Italy 2 Institute of Food Sciences, National Research Council (CNR-ISA), Via Roma 64, 83100 Avellino, Italy 3 DIAA – University of Molise, Via de Sanctis, snc 86100, Campobasso (Italy) 4 Institute of Microbiology, Catholic University of the Sacred Heart, “A. Gemelli” Foundation, Largo A. Gemelli 8, 00168 Rome, Italy 5 IRCCS Centre for Neuroscience Bonino-Pulejo, Strada Statale 113, 98124 Messina, Italy *[email protected]

ABSTRACT Biofilms are detrimental to human life and industrial processes due to potential infections, contaminations and deterioration. Therefore, the evaluation of microbial capability to form biofilms is of fundamental importance for assessing how different environmental factors may affect their vitality. Nowadays, the approaches used for biofilm evaluation are still poor in reliability and rapidity and often provide contradictory results. Here we present what we call Biofilm Electrostatic Test (BET) as a simple, rapid and high reproducible tool for evaluating in vitro the ability of bacteria to form biofilms through the electrostatic interaction with a pyro-electrified carrier. The results show how the BET is able to produce viable biofilms with a density 6-fold higher than the control, just after 2 h incubation. The BET could pave the way to a rapid standardization of the evaluation of bacterial resistance among biofilm-producing microorganisms. In fact, due to its simplicity and cost-effectiveness, it is well suited for a rapid and easy implementation into a microbiology laboratory.

Keywords: biofilm; rapid formation; bacterial adhesion; anti-microbial tests; surface potential; electrostatic attraction; cation-free; pyroelectric effect.

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INTRODUCTION The majority of bacteria are organized into structured microbial communities, encased in extracellular matrix known as biofilms.1-6 Virtually, every surface – animal, mineral, or vegetable – is appropriate for colonization and formation of biofilms by bacteria, including contact lenses, ship hulls, dairy and petroleum pipelines, rocks in streams, and all varieties of biomedical implants and transcutaneous devices.7-9 The bacteria growing in a biofilm matrix have markedly altered metabolism and enhanced cell-cell communications. The formation of biofilms causes the bacteria to become inherently more resistant to the environmental stresses and able to express a tolerance to antibiotics, which is up to 1000 times higher than that exhibited by the corresponding planktonic counterparty.10 Therefore, biofilms contribute to a broad range of problems in human health, such as tooth cavities, bio fouling of medical devices and lethal chronic infections indeed.11-19 Biofilms also impinge on a variety of industrial settings. They are responsible of increased hydrodynamic drag on ships and lead to augmented fuel consumption.20 Nevertheless not all biofilms are bad. Some bacteria that co-evolved and are accommodated to human niches are important for the establishment of the microbiome, as in case of the gastrointestinal tract. In this complex and diversified framework, knowledge about biofilm development is of essential importance in order to manipulate, control or eradicate them.3 Nowadays, different in vitro models are available for evaluating the ability of bacterial strains to form biofilms in susceptibility tests with antimicrobial agents.21 Conventional methods are based on plating and microscopic counting for detecting the amount of bacteria adhering onto eukaryotic cells22 or inanimate surfaces.23 However, conventional methods are laborious and require bacteria to remain culturable after the releasing process. Microscopic and colorimetric approaches can be used after fixation and Gram staining24,25 but these methods are also laborious. At present the crystal violet staining is the most widely used method for an in vitro quantification of biofilm, due to its relative simplicity and sensitivity.25,26 This method, however, shows important limitations. It usually requires at least

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24−48 h of incubation and repeated processing steps, with large standard deviations of the readouts and poor adaptability to large-scale screening. Recently, a new technology named BioFilmRing Test® (Saint Beauzire, France) has been proposed for the assessment of bacterial biofilms.27 The principle is based on the immobilization of magnetic beads by the growing biofilm matrix in vitro. Although having a great potential, the procedure has still some limitations for use in the clinical setting. It requires the additional step of treating the bacterial culture with the magnetic beads, thus introducing a further variable that might influence the biofilm formation and its repeatability. Moreover, the evaluation of biofilm formation is performed indirectly, through the observation of the beads accumulation by external magnetic forces. All of these methods are poorly standardized and, most of them require about 1 day incubation prior to start the assay. This is highly detrimental in particular for clinicians who desire to get microbiology test results as soon as possible in order to save lives. In summary, to the best of our knowledge, assays capable of evaluating the production of biofilm by bacteria, and subsequent bacterial susceptibility/resistance to antimicrobial drugs, are still difficult requests for clinical microbiology.28 Here we propose the Biofilm Electrostatic Test (BET) as a new tool for simple, rapid and costeffective evaluation of biofilm formation, through the electrostatic interaction of planktonic bacteria – namely bacteria suspended in a liquid medium29 – with a pyro-electrified carrier, that we call here BET-carrier, in the form of strip or fiber, according to the desired architecture. The BET-carrier is produced by a pyroelectric-based process,30 called pyro-electrification.31 It provides a polarization field able to immobilize the bacteria and test their ability to form live biofilms within 2 hours by direct microscope observation. This method avoids time-consuming and laborious incubations and/or intermediate chemical treatments. We introduced the pyro-electrification technique for the first time in 201631 for producing layers with a polarization field able to pattern the adhesion of

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eukaryotic cells. Here, the same kind of polarization field is used for governing the adhesion of prokaryotic cells, thus bringing the pyro-electrification to another family of applications. It is well known in microbiology that the bacteria have a cell wall that can be of two different types. Correspondingly, they are classified usually in gram-positive bacteria and gram-negative bacteria, according to the different response of these two kinds of cell wall to the so called ‘Gram staining’, in honour of the Danish bacteriologist Hans Christian Gram who first developed the technique in 1884.32-37 The gram-positive bacteria stain purple while the gram-negative bacteria stain pink. Both gram-positive and gram-negative bacterial cells exhibit a negative net charge at neutral pH.7,32-37 The initial adhesion of a microorganism cell to a substrate is a very complex process governed by a wide variety of physical and chemical conditions such as hydrophobicity, bacterial motility, outer membrane proteins and surface electrical potential.1 Even though these factors have been studied largely, still controversial results can be found in literature, probably due to a lack of standardization between the methods used in each procedure. Here we focus the attention on the role of the surface potential. This feature has been used very recently for immobilizing bacteria or for influencing cell adhesion38,39 and, most widely, for developing cationic surfaces with bactericidal properties. In fact, their net positive charge is able to attract and kill bacteria through ion exchange mechanisms.32-33,40,41 Here, the polarization charge of the BETcarrier interacts electrostatically with the bacterial cells without damaging their cytomembrane, thus providing biofilms that are mature within 2 h incubation and viable up to 24 h, thus allowing one to perform rapid and reliable biofilm evaluation. The results reported here demonstrate that BET might open the route to a reliable and cost-effective evaluation of biofilm formation and related antimicrobial susceptibility, with statistically relevant data in short time scales, thus leading to a significant impact in clinical microbiology protocols. The ability to get rapid biofilm formation is particularly relevant in all of those assays where low abundant samples of clinical isolates are available. It is noteworthy that the BET-carrier is produced in an environmentally friendly manner, avoiding cation-based chemistry. 4


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RESULTS The Biofilm Electrostatic Test with the carrier in the form of strip Two kinds of polysulfone (PSU) strips (about 100 µm thick and 2×2 cm2 sized) were produced and incubated for 24 h at 37°C in two different Petri dishes covered with planktonic Escherichia coli bacteria (gram-negative) in phosphate buffer saline (PBS): (1) bare strip that, from here on, represents the control; (2) pyro-electrified strip that, from here on, represents the BET-carrier in the form of strip. The Experimental Section contains details about the preparation of the control (the bare strip) and about the bacterial cultures. The Supporting Information describes the preparation of the BET-carrier. It is important to note that the BET-carrier was used with the positive face in contact with the bacteria and that the results concerning the biofilm formation on the negative face are not reported here because they demonstrated a negligible different behaviour compared to the control (see the Supporting Information for details). The lack of growth medium allowed us to verify the adhesion degree of the planktonic bacteria and their viability, minimizing the contribution from proliferation effects. The control and the BET-carrier were observed under a standard optical inverted microscope at different times (2 h, 8 h, 24 h), and Fig.1 (a) shows the corresponding typical images.

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Figure 1: (a-c) Optical microscope images of E. coli bacteria forming biofilms on the control, at three different time intervals; (d-f) optical microscope images of the E. coli forming biofilms on the BET-carrier, at the same time intervals; (g) distribution of the mean values of BD evaluated by ImageJ over three replicates of the experiments and over ten pictures recorded for each sample in case of E. coli; (h) linear functions that best fit the BD values reported in the histogram together with the results of the fit (R2 is the coefficient of determination). The contrast of the optical microscope images was enhanced digitally in order to make the bacteria more visible. The scale bars are 50 µm long.

These microscope images show the immobilized and biofilm forming bacteria on the control and BET-carrier at different time intervals. The number of adhesion bacteria on the BET-carrier appeared clearly higher than that on the control at each observation time. In order to demonstrate the reliability of the results, we performed three replicates of the experiments and we recorded ten images for both the control and the BET-carrier, at every time. We analyzed the images by ImageJ, an open source image-processing program developed at the National Institutes of Health (NIH). We integrated the grey scale intensities corresponding to the bacterial pixel positions and we considered the resulting values as a measure of the biofilm density (BD), in arbitrary units. We introduced the dimensionless parameter relative biofilm density (BDr) as the ratio between BD on the BET-carrier and BD on the control at a fixed incubation time, as follows: 6


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BDr =

BD(BET − carrier) BD(control)

This parameter BDr quantifies the ability of the BET-carrier in promoting biofilms related to the control, used as a reference. Therefore, values of BDr >> 1 correspond to a significant enhancement in biofilm formation. The histogram in Fig.1 (b) reports the mean values of BD obtained for the control and for the BET-carrier, at each observation time. Taking into account the absence of the culture broth, the behavior of BD gives information about the rate of immobilization of planktonic bacteria forming biofilm, thus neglecting the proliferation effects. These results show that the BD on the BET-carrier (green columns) is about 4-fold higher than that on the control (blue columns), at each observation time. This means that BDr ∼ 4, with a BD on the BET-carrier that, after just 2 h, was comparable to that obtained onto the control after 24 h. Figure 1(c) shows the lines that best fit the BD values in time for the control and the BET-carrier. We consider the angular coefficient of the fitting lines as a measure of the biofilm growth rate (BGR) in time and we call here the relative biofilm growth rate (BGRr) as the ratio between BGR on the BET-carrier and BGR on the control as follows: BGRr =

BGR(BET − carrier) BGR(control)

The parameter BGRr quantifies the rapidity of the BET-carrier in forming the biofilm, related to that of the control used as a reference. Therefore, values of BGRr >> 1 correspond to a significant enhancement in biofilm growth rate. The fit results reported in Fig.1(c) show a value BGRr ∼ 5, thus demonstrating a 5-fold faster biofilm formation on the BET-carrier, compared to the control. The same experiments were performed with a strain of gram-positive bacteria, the Staphylococcus epidermidis, and Fig.2 (a) shows the typical images recorded by a standard inverted optical microscope.

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Figure 2: (a-c) Optical microscope images of S. epidermidis bacteria forming biofilms on the control, at three different time intervals; (d-f) optical microscope images of the S. epidermidis forming biofilms on the BET-carrier, at the same time intervals; (g) distribution of the mean values of BD evaluated by ImageJ over three replicates of the experiments and over ten pictures recorded for each sample in case of S. epidermidis; (h) linear functions that best fit the BD values reported in the histogram together with the results of the fit (R2 is the coefficient of determination). The contrast of the optical microscope images was enhanced digitally in order to make the bacteria more visible. The scale bars are 50 µm long.

Also in case of S. epidermidis we observed that the BET-carrier exhibited a much stronger ability to form biofilm at each observation time, compared to the control. We performed three replicates of the experiments for both control and BET-carrier and we recorded ten pictures at every observation time. Again, we analysed the images by ImageJ and the histogram in Fig.2 (b) shows the resulting distribution of the mean values of BD. In this case BDr ∼ 6, thus demonstrating the ability of the BET-carrier to form biofilms of S. epidermidis with a density 6-fold higher than that on the control, at each observation time.

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Figure 2(c) shows the lines that best fit the behaviour of BD in case of the control and of the BET-carrier with S. epidermidis, together with the fitting results. These results show a value of BGRr ∼ 6, thus demonstrating a 6-fold faster biofilm formation on the BET-carrier in case of S. epidermidis. The histogram in Figure 3(a) compares the ability of the BET-carrier in forming biofilms for the two bacterial strains (E. coli and S. epidermidis), by reporting the values of BDr at each observation time for the two strains.

Figure 3: (a) Distribution in time of BDr for E. coli and S. epidermidis in case of BETcarrier in the form of strip; (b) linear functions that best fit the evolution in time of BD on the BET-carrier in the form of strip for E. coli and S. epidermidis.

This histogram shows clearly that the BET-carrier exhibited, in average, about 30% increment of BDr in case of the gram-positive bacteria (S. epidermidis), compared to the gram-negative (E. coli). In other words, the gram-positive bacteria formed on the BET-carrier a biofilm that, in average, had a density about 30% higher than that formed by the gram-negative. Figure 3(b) compares the linear regressions that best fit the variation of BD in time, for each bacterial strain, together with the resulting fitting parameters. The resulting values of BGRr demonstrate that the gram-positive bacteria (orange line) formed the biofilm onto the BET-carrier about 9% faster than in case of gram-negative (violet line). In order to verify the viability of the biofilms, both the control and the BET-carrier was mounted onto a glass slide in a well after 24 h incubation with planktonic bacteria, for performing the 9



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reaction with the live/dead staining kit (see Experimental Section for details). The same procedure was performed for E. coli and S. epidermidis. A conventional upright microscope (Axio Imager, Carl Zeiss, Germany) was used for observing the samples after staining and Fig.4(a-d) shows the typical fluorescence images.

Figure 4: (a,b) Fluorescence microscope images of the E. coli forming biofilms onto the control and the BET-carrier, respectively, after 24 h incubation and live/dead staining (scale bar 50 µm long); (c,d) fluorescence microscope images of the S. epidermidis forming biofilms onto the control and the BET-carrier, respectively, after 24 h incubation and live/dead staining (scale bar 50 µm long); (e,f) distribution of the BD values for live (green columns) and dead (red columns) bacteria for E. coli and S. epidermidis, respectively.

The green spots refer to viable bacteria, while the red ones to the dead bacteria. These images demonstrate clearly the viability of the high density biofilms formed onto the BET-carrier for both bacterial strains after 24 h incubation. The BET-carrier was able to immobilize planktonic bacteria more rapidly than the control and to favour biofilm formation without damaging the bacterial cytomembrane even after 24 h incubation. Three replicates of the experiments were carried out for each bacterial strain and ten pictures were captured all around each surface, in order to demonstrate the reliability of the biofilm vitality. Also in this case, the images were analyzed by ImageJ in order to evaluate the mean values of the signal intensities for the green (Syto 9) and the red (propidium iodide) channel, and hence to have information about the average amount of live and dead biofilm. 10


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Figure 4(e,f) report the resulting data where the green and red columns refer to the mean values of BD for red (dead) and green (live) channels for E. coli and S. epidermidis, respectively. These results show first of all that the amount of dead biofilm (red columns) is statistically negligible for both the control and the BET-carrier, thus demonstrating their biocompatiblity. The amount of live biofilm (green columns) immobilized onto the control appears similar for both strains, thus showing a negligible difference in the interaction mechanism of the control with the bacterial cytomembrane. Conversely, the values obtained for the BET-carrier confirm its ability to promote a biofilm with a density 4-fold and 6-fold higher than that on the control, for E. coli and S. epidermidis respectively. Moreover, the live/dead staining results show that about 98% of the bacteria forming the biofilms are alive on the BET-carrier for both bacterial strains, thus demonstrating definitely its ability to immobilize planktonic bacteria and to promote a rapid biofilm formation that is fully viable even after 24 h. Table 1 summarizes the data that characterize the biofilm formation on the BET-carrier in the form of strip for both bacterial strains.

Table 1. List of data that characterize the biofilm formation on the BET-carrier in the form of strip, for E. coli and S. epidermidis, in 24 h incubation. Bacterial strain

BDr

BGRr

Biofilm viability

E. coli

4.5 ± 0.2

4.6 ± 0.2

(97 ± 5) %

S. epidermidis

5.9 ± 0.3

5.8 ± 0.3

(98 ± 5) %

The Biofilm Electrostatic Test with the carrier in the form of fiber The solution of polystyrene (PS) with Mw∼350,000 was used for drawing three thin fibers manually (see the Supporting Information). Two fibers were cultured with planktonic E. coli in a Petri dish in PBS as well as in Luria-Bertani (LB) broth, and represented the controls. The third fiber was subjected to pyro-electrification (see the Supporting Information) and then cultured with E. coli, and represented the BET-carrier in the form of fiber. Both the controls and the BET-carrier were 11



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observed under the optical microscope at different times. The other solution of PS (Mw∼192,000) was used for drawing another BET-carrier in the form of fiber. For the sake of simplicity we call here BET-carrier/strong and BET-carrier/weak the pyro-electrified fibers obtained by using the PS with Mw∼350,000 and Mw∼192,000, respectively. The initial concentration of the bacteria was 1.4×107 cells/mL in PBS in all cases. Figure 5 reports the corresponding results.

Figure 5: (a-d) Microscope images of the control incubated with E. coli in PBS at different time intervals (scale bar 50 µm); (e-h) images of the control incubated with E. coli in LB at different time intervals (scale bar 50 µm); (j,k) images of the BET-carrier/weak incubated with E. coli at different time intervals (scale bars 50 µm and 20 µm); (l) large view of the BET-carrier/strong while attracting electrostatically silica beads under dry conditions in ambient air (scale bar 50 µm); (m-p) images of the BET-carrier/strong incubated with E. coli at different time intervals (scale bar 50 µm); (q) distribution of the mean values of BT (biofilm thickness) measured on the control in PBS, the BET-carrier/weak and the BETcarrier/strong, in the form of fiber, over three replicates of the experiment.

The dashed lines in Fig.5(k,m-p) highlight the edges of the biofilm adhering onto the fiber surface, while the red arrow in Fg.5(o) indicates the typical position where we measured the biofilm thickness (BT) to quantify the ability of the fiber in promoting the biofilm formation. The biofilm formed poorly onto the control both under PBS and LB culture, with BT ∼ 4 µm after 24 h incubation (see Fig.5(b)). Only after about 120 h incubation in PBS, we observed an appreciable biofilm onto the control fiber (see Fig.5(d)), regardless of its vitality that is negligible here since we 12


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are interested at short time formation of biofilms. Similar results were obtained in case of control in LB, with poor biofilm formation even after 48 h incubation (see Fig.5(h)), with highly non uniform distribution of the adhering bacteria and with poor repeatability between the replicated experiments. Conversely, a value of BT ∼ 10 µm was measured onto the BET-carrier/strong, after just 3 h incubation and grew up to about 110 µm after 24 h incubation. The behaviour of the BETcarrier/weak was similar to that of the control with negligible differences in BT. In order to test the reliability of the technique three replicates of the experiments were performed and the histogram in Fig.5(q) shows the mean values of BT, obtained by averaging the measures over ten images acquired along each fiber. The BET-carrier/strong promoted systematically a biofilm with BT up to about 30-fold higher than that on the control. These results demonstrate the ability of BET to have reproducible biofilms even on carriers in the form of flexible fibers where, compared to the strips, the bacteria have a reduced surface accessibility and are subject to higher fluctuations effects. This would allow microbiologists to investigate in vitro a wider range of biofilm architectures and environmental settings.

Comparison with cationic surfaces Cationic surfaces are obtained usually by the covalent attachment of different chemical compounds such as quaternary ammonium organo-silanes,42-44 antimicrobial peptides,45,46 polyethylenimines (PEI)47 and many others. Here we produced two kinds of cationic surfaces: (1) bare PSU strip and (2) bare PS fiber (Mw∼350,000), both functionalized with cationic PEI (see Experimental Section for details). E. coli cells were seeded and incubated in two different Petri dishes containing the PEIcoated strip and the PEI-coated fiber, in PBS. After 24 h incubation, they were stained by the live/dead kit and Figure 6(a,b) show the typical fluorescence images.

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Figure 6: (a,b) Fluorescence microscope images of the E. coli forming biofilms on the PEI coated strip and fiber, after 24 h incubation and successive live/dead staining. The scale bar is 50 µm long in (a) and 20 µm long in (b). The histogram in (c) shows the distribution of the mean values of BD for live (green columns) and dead (red columns) biofilm on the PEIcoated and BET-carrier under strip configuration, after 24 h incubation with E. coli and live/dead staining.

The cationic surfaces appear to promote the biofilm formation, but the fluorescence images after live/dead staining reveal unequivocally the net prevalence of the red channel, demonstrating the ability of a cationic surface to immobilize planktonic bacteria but to cause the cytomembrane damage. Three replicates of the experiments were performed with the PEI-coated strip in order to demonstrate their reliability, and the histogram in Fig.6 (c) shows the mean values of BD after 24 h 14


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incubation with E. coli and successive live/dead staining. The BD values were evaluated separately for the green channel (live biofilm) and for the red channel (dead biofilm), in the corresponding fluorescence images. Table 2 summarizes the data that characterize the biofilm viability on the BET-carrier in the form of strip with those on the PEI-coated strip, in case of E. coli. Table 2. List of data that characterize the biofilm viability on the BET-carrier (strip) and on the PEI-coated strip in case of E. coli, after 24 h incubation. Biofilm viability PEI-coated strip BET-carrier (strip)

(3.0± 0.2) % (97 ± 5) %

These results demonstrate definitely the ability of the BET of promoting the formation of biofilms in analogy with cation-based surfaces, but with the disruptive difference of being viable even after 24 h, thus opening the route to an easy to accomplish tool for rapid and reliable susceptibility tests in the field of microbiology.

DISCUSSION Figure 7 show the simplified views of the interaction mechanism between planktonic bacteria and each kind of functionalized surface: (a) the surface of the PEI-coated PSU strip; (b) the surface of the BET-carrier in the form of PSU strip.

(a)

(b)

Figure 7. Simplified schematic views of the typical interaction of the bacterial cells with (a) the PEI-coated strip, where an electrostatic bond occurs between the cytomembrane and the amino groups on the surface, and (b) the BET-carrier (strip), where no molecular bond occurs. P-face and N-face indicate the polarity of the BET-carrier faces. See the Supporting Information for more details.

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The schemes refer to the case of the strips for the sake of simplicity, but the discussion is valid as well for the fibers. Both kinds of surfaces exhibit a positive electrostatic polarity that exerts an electrostatic attraction (black arrows) towards the bacterial membrane that exhibits a net negative charge in both gram-positive and gram-negative bacteria (see the Introduction Section), but with a substantial difference. In case of PEI, the positive polarity is provided by the net positive charge of the cationic groups NH3+, while in case of the BET-carrier it is provided by the δ+ charge of the permanent dipole generated through pyro-electrification. Therefore, according to the fundamentals of electrostatics, the negative net charge onto the bacterial cytomembrane (COO- groups) of both gram-positive and gram-negative bacteria is attracted (black arrows) by the positive polarity on each surface, leading to bacteria immobilization and biofilm formation in both cases. However, the cationic groups NH3+ on the PEI surface, once attracted the bacteria, form an electrostatic bond with the COO- groups onto the cytomembrane, thus displacing the divalent cations forming the lipopolysaccharide network. This leads inevitably to the disruption of the cytomembrane and, as a direct consequence, to the microorganism death.29,48 Conversely, the BET-carrier attracts electrostatically the bacterial cytomembrane without involving net free charges. This ‘cation-free’ interaction is able to immobilize the bacteria avoiding molecular bindings and, as a consequence, preserving the integrity of the cytomembrane. The polarization field of the BET-carrier is intense enough to immobilize the planktonic bacteria very rapidly and to sustain a live biofilm even after 24 h incubation. The intensity of the polarization field depends on the strength of the δ+ charge and, hence, on the molecular weight of the polymer. In fact, the results in Fig.5(j,k) show clearly that a lower molecular weight provides a weaker polarization field and, as a consequence, a weaker bacterial immobilization. At the end we have a portable BET-carrier sustaining a vital biofilm ready to use for biofilm susceptibility tests with high reproducibility. Moreover, the 30% increment of BD observed in case of gram-positive bacteria (S. epidermidis), compared to the gram-negative (E. coli) (see the results in Fig.3), is in full agreement with the 16


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results presented recently by Ferrer et al.49 They measured the electric polarization properties of single bacteria through electrostatic force microscopy and they reported a value of the effective dielectric constant ε one order of magnitude higher in case of the gram-positive bacteria compared to the gram-negative. This means that the S. epidermidis bacteria, when in the vicinity of the same polarization field of the BET-carrier, polarized more strongly than E. coli and therefore were →

subjected to a higher electrostatic force F that, at first approximation, can be described as follows: →

→

F = −p ∇ E where p is a constant of proportionality involving, among other factors, the dielectric constant ε of the bacterium. The minus sign refers to the opposite direction of the electric field and the electrostatic force vectors. Therefore, at the same electric field, the bacteria with higher ε (i.e. S. epidermidis) are subjected to higher electrostatic force. This occurs regardless of the different motility attributed usually to bacteria with (e.g. E. coli) and without (e.g. S. epidermidis) flagella which role in biofilm formation is also controversial.50-56 Our results demonstrate unequivocally that the electrostatic factor has a predominant role in biofilm formation making the S. epidermidis to form biofilms with higher density systematically at each observation time (see Fig.3).

CONCLUSION In conclusion, we show here the ability of the BET of getting mature biofilms of both gramnegative and gram-positive bacteria within 2 hours and with 6-fold higher density than the control. Thanks to the cytomembrane integrity, the BET-carrier is able to sustain the growth of a vital biofilm even after 24 h incubation, and with a growth rate 6-fold higher than the control. The advantages of BET are manifold: 1) the 6-fold enhancement of BD and the high reproducibility would enable much faster and reliable antimicrobial tests, especially in those clinical cases where a fast test response is vital for the patience; 2) the 6-fold higher BGR would be of crucial importance in all of those cases where low abundant samples of clinical isolates are available; 3) the cost17



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effective and relative simplicity of the procedures would enable an easy implementation of the BET in biochemical and clinical laboratories, avoiding expensive equipment and too specialized personnel; 4) the availability of BET-carrier in the form of both strip and fiber would enable a deep investigation in vitro of biofilm architectures under different environmental and scaffold conditions; 5) the cation-free electrostatic interaction makes the BET an environment-friendly tool that avoids totally any chemical treatment. EXPERIMENTAL SECTION The preparation of the polymer solutions. Solid-state polymers were bought from Sigma Aldrich and were used as received, without further purification. Polysulfone (PSU) (Mw ∼ 35,000, transparent pellets) was dissolved at 80% w/w in anisole, and stirred at 70 °C for 3 h. Polystyrene (PS) with two molecular weights (192,000; 350,000) was dissolved at 60% w/w in anisole and stirred at 70 °C for 6 h. The resulting polymer solutions of PSU and PS were stored at 4 °C. The preparation of the bare strip used as control. The freestanding PSU bare strip was obtained by spin coating a (2×2) cm2 sized glass coverslip at 4000 RPM for 2 minutes with the PSU solution and by peeling off the slide accurately just after solvent evaporation. The cation functionalization. Cationic polyethylenimine (PEI, branched, average molecular weight 25000, water-free from Sigma Aldrich) was used for the cation functionalization of the PSU strip and PS fiber as follows. The strip and the fiber were produced as usual by spin-coating and manual spinning, respectively, and then immersed into a PEI solution (1.5 mg/mL in distilled water) for 20 minutes, rinsed with water and dried with a nitrogen flow. The bacterial cultures. The experiments were carried out against the gram-negative Escherichia coli DH5-alpha (kindly provided by Dr. Claudia Tortiglione), and against the gram-positive Staphylococcus epidermidis (kindly provided by Prof. Luca Masucci). The bacteria strains were plated and incubated on Luria-Bertani (LB) agar plates (10 g/l NaCl, 10 g/l tryptone, 5 g/l yeast extract, 15 g/l agar, Thermo Fisher Scientific, provided by Life Technologies Italia). The day before the beginning of the experiment, a single bacterial colony was picked up and cultured in LB broth 18


Page 19 of 24 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


medium at 37 °C in a shaker incubator at 225 RPM for 16−18 h to achieve saturation conditions. A 1:5 volumetric dilution of cell culture was then grown in LB until reaching the log phase. Then the growth was stopped and bacteria were harvested by centrifugation at 7.200 g (Beckman Coulter tj25 centrifuge, California, USA) for 10 min in order to separate the cells from the medium. Sterilized LB broth was measured (3 mL) into sterile tubes. The bacteria concentration was evaluated

by

the

spectrophotometric

measurement

(Bio-Rad

SmartSpecTM

Plus

Spectrophotometer, California, USA) of the suspension absorbance at 600 nm (Optical Density at 600nm, i.e. OD600), considering that 8×108 cell/mL have an OD600 =1. The motility of the bacteria was evaluated by phase-contrast microscopy according to microbiological laboratory procedures.57 The viability test. The viability of the bacterial strains (E. coli and S. epidermidis) was tested through the live/dead viability/cytotoxicity assay kit (Live/Dead BacLight bacterial viability kit, Thermo Fisher Scientific, Waltham, MA USA ). The live/dead kit is a convenient and easy-to-use kit for monitoring the viability of bacterial populations as a function of the membrane integrity of the cell. The live/dead BacLight Bacterial Viability Kits utilize mixtures of our SYTO® 9 greenfluorescent nucleic acid stain and the red-fluorescent nucleic acid stain, propidium iodide. These stains differ both in their spectral characteristics and in their ability to penetrate healthy bacterial cells. Cells with a compromised membrane that are considered to be dead or dying will stain red, whereas cells with an intact membrane will stain green. The cells were incubated on each carrier for 24 h. Following incubation, each carrier was immersed in 8 µL of 1000-fold diluted live/dead kit solution and was incubated for 15 min in the dark. The fluorescence micrographs were acquired by an inverted laser scanning confocal microscope (Zeiss LSM 700, Germany), equipped with a 20× objective.

SUPPORTING INFORMATION The Supporting Information is available as a separate document file.

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The Pyroelectric Effect Enables Simple and Rapid Evaluation of

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