Biofilm Formation of Escherichia coli on Hydrophobic Steel Surface Provided by Laser-Texturing
Biofilm Formation of Escherichia coli on Hydrophobic Steel Surface Provided by Laser-Texturing
Investigation of modified surfaces to prevent biofilm formation
Lozenge-patterned surfaces obtained with laser texturing can reduce the risk of infection by preventing or delaying biofilm formation of Escherichia coli. To investigate this aspect, the biofilm formation ability of E. coli on both lozenge-patterned and untreated surfaces of 630 stainless steel coupons was examined over 48 h. Biofilm on the coupons was analysed for bacterial enumeration and total carbohydrates concentration and was observed using scanning electron microscopy (SEM). The surface modification by texturing caused a 6 h delay in the attachment of E. coli and an approximately 99% decrease in the number of adhered bacteria. However, it was determined that E. coli produced more extracellular polymeric substances (EPS) (p<0.01) to attach to the lozenge-patterned surface and formed a multi-layered biofilm. In conclusion, lozenge-patterned surfaces can be applied to reduce bacterial count and induce a delay in attachment, but the increased amount of EPS limits its use.
Healthcare-associated infections (HAIs) affect millions of people worldwide annually and are linked with increased morbidity and mortality (1, 2). Many types of HAI occur due to invasive procedures to treat patients. Therefore surgical instruments and medical devices are considered critical for the transmission of microorganisms. A wide variety of microorganisms have been isolated from surgical instruments and medical devices (3–5), most of which are pathogens. Among these, E. coli under certain conditions is responsible for many human infections (6–8).
Biofilm formation increases bacteria’s virulence and ability to survive by adhering to surgical instruments, medical devices or patients’ tissues (4, 9). Once attached to a surface, the bacteria begin to grow and produce EPS that promote the development of a biofilm structure, allowing planktonic cells to assume a multicellular lifestyle (10, 11). In most biofilms, the microorganisms account for less than 10% of the dry mass while the EPS can account for over 90% (12, 13). Carbohydrates form the majority of EPS (14, 15) which comprises a network of diverse macromolecules (16, 17). Secreted carbohydrates have been involved in the attachment of bacteria to a surface and are recognised as key elements that shape and provide structural support for the biofilm (18). However, the concentration of carbohydrates and other macromolecules can be influenced by cellular, surface and environmental factors. These factors include microbial species, the availability of nutrients, surface composition and roughness, cell motility, temperature, hydrodynamics and hydrophobicity (19).
Biofilm-caused infections are responsible for up to 60% of HAIs (20) because bacteria embedded in a biofilm are less sensitive to antimicrobials and to removal treatments such as disinfection or sterilisation (21–23). If the bacterial number adhering to a metal surface can be reduced or eliminated, the effectiveness of decontamination techniques will increase due to reduced microbial load. In this context, it is of great importance that the metal preferred in the production of medical equipment has the properties of easy processing, modifiability, corrosion-resistance and non-adhering (anti-adhesive) surfaces. For this reason, 630 stainless steel is widely used in medical equipment production. Even more important is the ability to create surface properties that can reduce or eliminate biofilm formation on the preferred metal. Thus, in recent years, much research in medical equipment production has focused on metals with surface properties designed for different antimicrobial strategies (24, 25). One of these strategies is to provide anti-adhesive surface properties and to achieve this, properties such as surface charge, free energy, zeta potential and wettability are taken into account (26, 27).
Laser texturing is a promising technique for the modification of surface roughness, chemistry and wettability, features that have been reported to affect bacterial adhesion to a surface (28–30). Laser texturing allows the creation of surface roughness from nano- to micro-scale with its single-step processing capability (31–33). Roughness modification can be used to change the wettability that plays an essential role in bacterial adhesion. Similarly, hydrophobic and superhydrophobic surface treatments have been developed as new strategies for designing anti-adhesive materials (26). Hydrophobic (contact angle >90°) materials are generally considered less prone to bacterial adhesion than hydrophilic (contact angle <90°) surfaces. However, optimal wettability to reduce bacterial adhesion is still controversial (24, 34, 35). Studies have been carried out to develop different geometric patterns with hydrophobic properties to prevent bacterial adhesion (36–39). Inspiration for a suitable surface pattern was taken from the superhydrophobic lotus leaf, which has a water contact angle of more than 150° (40). The lozenge pattern is more effective than geometric patterns such as square, triangle and strip (41).
The exact relationship between surface properties of the metal and bacterial adhesion has not been fully elucidated and therefore different inferences can be drawn from the literature. However, the effect of laser texturing on the development of anti-adhesive surface properties has been emphasised. Recently in the literature, it has been reported that surfaces with micron-, submicron- and nano-scale structure reduce bacterial adhesion by up to 90% compared to a smooth surface in the same environment (42). Studies on hydrophobicity generally focus on the bacterial number after a certain contact period; however, this approach is constrained to reporting the initial attachment time and the number of bacteria associated with it. Moreover, no study was found investigating changes in the carbohydrate amounts in EPS produced by bacteria on a hydrophobic surface over time. Thus, the data obtained in this study are important to improve the literature.
The objective of this study was to investigate adhesion and biofilm formation abilities of the potential pathogen E. coli on hydrophobic 630 stainless steel surfaces provided by laser texturing. For this purpose, experiments were performed with lozenge-patterned and untreated steel surfaces. Lozenge-patterned and untreated surfaces were exposed to E. coli culture over 48 h of exposure to compare bacterial adhesion.
2. Materials and Methods
2.1 Material Characterisation
In the study, rod-shaped 630 stainless steel (630/17-4 PH) with 50 mm diameter and 200 nm height was used as the base metal. The nominal elemental composition of the 630 stainless steel was nickel (3.69 wt%), chromium (14.13 wt%), carbon (0.022 wt%), silicon (0.362 wt%), manganese (0.878 wt%), phosphorus (0.023 wt%), sulfur (0.025 wt%) and iron (76.7 wt%).
2.2 Preparation of Test Specimens
The lozenge-patterned surfaces were provided by texturing the steel surface using a fibre laser (redENERGY G4 Pulsed Fiber Laser 20W EP-S (SPI Lasers, UK)) with 1064 nm wavelength and 60 kHz repetition rate (Teknik Form Plastic Mold Food and Construction Industry Trade Ltd Co, Turkey). Coupons with untreated surfaces were used as controls. The lozenge-patterned and control coupons were cut in the size of 10 × 10 × 2.5 mm. The untreated coupons were abraded by 1200-grade silicon carbide metallurgical paper to a smooth surface, polished with aluminium oxide and washed with sterile distilled water. The lozenge-patterned and untreated surfaces were degreased using ethanol, washed with distilled water and dried with a Pasteur oven at 70°C (43). They were kept in a desiccator until use.
2.3 Surface Morphology and Wettability Measurement
Surface morphologies of the lozenge-patterned and untreated coupons were examined by SEM (Figure 1). The structures in the form of lozenge patterns created by the laser irradiation on the laser-textured coupons are clearly shown in Figure 1(b).
The hydrophobic properties of each surface were determined by measuring the wetting angle. The wetting angles of both lozenge-patterned and untreated surfaces were measured using the OCA 20 (DataPhysics Instruments GmbH, Germany) with sessile drop technique, which is a type of characterisation method used to examine the water retention on the surface of solid samples. Droplets of distilled water used as the probe liquid were deposited at different positions on the lozenge-patterned and untreated surfaces. The profiles were captured and analysed by a force tensiometer K100 (Krüss Optronic, Germany). As shown in Figure 2, the surface wetting angle of the untreated surfaces was 67.40° ± 1.50°, exhibiting hydrophilic properties. In contrast, the contact angle of the lozenge-patterned surfaces was measured as 120.12° ± 2.62°, exhibiting hydrophobic properties.
2.4 Attachment and Biofilm Experiments
The effect of laser texturing on the 630 stainless steel surface’s anti-adhesive properties was investigated by operating a closed laboratory-scale system. The laboratory-scale system was set up using the coupons with lozenge-patterned and untreated surfaces immersed in a pure culture of E. coli (ATCC® 25922TM) in a 2 l glass beaker and operated over 48 h at 24°C. Before the coupons were placed in the system, both sides were sterilised by ultraviolet irradiation for 2 h. The E. coli culture was prepared in tryptic soy broth (TSB) (Merck KGaA, Germany) and adjusted to a starting inoculum of 1.5 × 105 colony forming unit (CFU) ml–1. When the sterilised coupons were immersed into the E. coli culture, the time was assumed to be 0 h. The system was operated over 48 h. Prolonged surgical operation times were taken into consideration when determining the experiment duration. The culture in the system was stirred magnetically at 150 rpm during the experiment and incubated at 24°C under aseptic conditions. The laboratory-scale system was set up in duplicate.
Coupons with the lozenge-patterned and untreated surfaces were removed from the laboratory-scale system after certain exposure times (2 h, 4 h, 6 h, 8 h, 10 h, 12 h, 24 h, 32 h and 48 h), sessile (from biofilm) and planktonic (from culture) E. coli were counted and CFU per millilitre was determined. The total amount of carbohydrates in EPS obtained from the biofilm layers on the coupon surfaces was measured at each sampling hour. The biofilm layers formed on the lozenge-patterned and untreated surfaces were examined with SEM.
2.4.1 Enumeration of E. coli
Both the sessile and the planktonic E. coli counts were determined simultaneously at each sampling time. E. coli in culture were planktonic bacteria (E. coli ml–1) and those obtained from the biofilm were sessile (E. coli cm–2). For the enumeration of sessile E. coli, one biofilm-coated coupon of both the lozenge-patterned and untreated surfaces was removed from the system. The coupon was dip-rinsed in sterile TSB to remove unattached cells. The coupon was then placed in a beaker containing 20 ml of sterile TSB and kept in an ultrasonic water bath (Alex Machine, Turkey) at a 40 kHz frequency for 5 min, allowing cells in the biofilm layer to pass into the TSB completely. The complete removal from the surfaces of the bacteria was controlled by SEM examination. Thus, a suspension of bacteria was obtained from sessile E. coli. Both the suspension and the E. coli culture were diluted to 10–10 in 10-fold serial increments at each sampling hour. 100 μl of each dilution was inoculated in tryptic soy agar (TSA) (Merck KGaA) in three replicates. After the incubation of Petri dishes at 37°C for 24 h, colonies on each plate were counted and CFU per millilitre was determined. The CFU numbers of sessile E. coli were calculated by dividing the determined bacterial number by the surface area of the coupon, and the bacterial counts were given as CFU cm–2.
2.4.2 Extracellular Polymeric Substances Extraction and Total Carbohydrate Analysis
A biofilm-coated coupon of both the lozenge-patterned and untreated surfaces was removed from the test system at each sampling time. The coupon was placed in a beaker containing 10 ml of sterile bidistilled water, and the biofilm layer in 1 cm2 surface area was transferred entirely by the ultrasonication method. The obtained 10 ml suspension was centrifuged at 6,000 g for 10 min, and the supernatant was decanted to a sterile centrifuge tube. The pellet was resuspended in a 10 ml aqueous solution containing 8.5% NaCl (Merck KGaA) and 0.22% formaldehyde (Merck KGaA) and it was vortexed at high speed for 60 s to recover the capsule-bound EPS. The recovered supernatant was combined with the first solution and 20 ml volume of the combined solution was centrifuged at 11,227 g for 30 min. After centrifugation, the supernatant (total extracellular material) was subjected to a filtration (0.22 μm) step (44). Total carbohydrate concentration in EPS extraction was determined using the phenol-sulfuric acid method (45). The carbohydrate analysis was performed in four replicates.
2.4.3 Scanning Electron Microscopy Analysis
The morphologies of biofilm layers formed on the lozenge-patterned and untreated surfaces and the changes in these layers during the experiment were examined by SEM analysis. Biofilm-coated coupons of both the lozenge-patterned and untreated surfaces were used for each sampling time. After fixation in 2.5% glutaraldehyde, the coupons were gradually dehydrated in 30%, 50%, 80% and 95% ethanol, then air-dried (46). The coupons were then coated with a thin layer of gold (30 nm) before imaging in the electron microscope (QuantaTM 450 FEG (FEI, USA)).
2.5 Statistical Evaluation
Statistical analyses were performed using the IBM® SPSS® version 21.0 software program. Standard deviations of means were calculated for all measurements. Spearman’s correlation coefficient test was used for statistical evaluation of the results. The application of the Mann-Whitney U test determined the differences between the means of the two types of surfaces (lozenge-patterned and untreated). Statistical significance for all analyses was set at a p value of 0.05 unless otherwise stated.
3. Results and Discussion
3.1 Biofilm Formation
Biofilm formation ability of E. coli on lozenge-patterned and untreated surfaces was evaluated by microbiological and microscopic analyses over 48 h. SEM analyses revealed that biofilm layers formed on both types of surfaces by the end of the experiment, but with different structures and morphologies. It was observed that while sessile E. coli on the untreated surface embedded in a dense monolayer biofilm covering the entire surface (Figure 3(a)), on the lozenge-patterned surfaces they formed a multi-layered biofilm with denser EPS (Figure 3(b)). Moreover, E. coli tended to locate in recessed parts of the lozenge-patterned surface, as reported by Helbig et al. (38), and developed many nanowires (Figure 3(c) and Figure 3(d)). Changes of surface conditions can cause physiological changes in the structure of the biofilm since the EPS production of bacteria depends on environmental stress (47).
3.2 Bacterial Count
The changes in the bacterial numbers for both the planktonic and sessile bacteria are shown in Figure 4. The planktonic E. coli entered the stationary phase after 12 h (5.38 × 108 ± 2.91 × 107 CFU ml–1) and the bacterial number was 1.46 × 109 ± 4.70 × 108 CFU ml–1 at the end of the experiment. Unlike the planktonic bacteria, sessile E. coli numbers on both lozenge-patterned and untreated surfaces increased during the experiment (p<0.05 and p<0.01, respectively). However, it was determined that the attachment time of E. coli to lozenge-patterned and untreated surfaces as well as the bacterial density on the surfaces were different (Figure 4).
Sessile E. coli on the untreated surface were detected for the first time after 6 h of exposure with 1.40 × 103 ± 6.16 × 102 CFU cm–2. A permanent increase was then observed in the bacterial count, reaching 9.53 × 106 ± 1.02 × 105 CFU cm–2 at the end of the experiment (Figure 4). It was also established that the number of planktonic bacteria was significantly higher than sessile bacteria (p<0.01). There was a significant positive relationship between sessile and planktonic E. coli counts (p<0.01). Indeed, in a biofilm there is continuous transition between sessile and planktonic states and these two forms can be considered as integrated components of prokaryotic life (48).
As regards the lozenge-patterned surface, E. coli biofilm was first detected after 12 h of exposure with 1.67 × 103 ± 2.66 × 102 CFU cm–2 (Figure 4). However, after 8 h of exposure, SEM analysis showed that very few (one or two) bacterial cells were present on the lozenge-patterned surface (Figure 5). Up to 12 h of exposure, E. coli on the lozenge-patterned surface can be assumed to be in a stage of reversible attachment. Bacterial adhesion to a material surface can be described as a two-phase process, including an initial phase of reversible physical contact and a time-dependent phase of irreversible molecular and cellular adherence (49, 50). When planktonic cells approach a surface, the bacterial movement is under the influence of van der Waals forces, electrostatic forces and hydrophobic interactions between the bacterial cell and the surface. During the reversible attachment stage, bacteria still show Brownian motion and can be easily removed from the surface (51, 52). While bacteria try to attach a surface, some can be easily detached. Bacterial cells which are still attached to the surface after a certain period can irreversibly adhere to the surface. Once attached irreversibly, they begin to multiply and form a structurally developed biofilm layer.
In the present research, sessile bacterial numbers during 12 h of exposure showed that the hydrophobic lozenge-patterned surface retarded the adhesion of E. coli by 6 h compared to the untreated surface. Villapun et al. (53) reported in their study using 304 stainless steel surfaces with different patterns that there were no differences between the treated and untreated surfaces in terms of the initial attachment time of the bacteria, and even after the first 30 min of exposure, bacteria were concentrated on the textured surfaces. Different results regarding the initial attachment may be caused by differences in the type of stainless steel, the structure of the pattern created on the steel surface and the medium in which the experiment was carried out. Indeed, bacterial adhesion is a complicated process influenced by many factors, such as the bacterial characteristics, the metal surface properties and environmental conditions (49, 50).
In order to completely prevent or reduce bacterial colonisation on a surface, it is necessary to prevent or minimise the attachment of bacteria. Recent studies with surface treatment methods have generally focused on increasing the surface’s hydrophobicity. However, the surface characteristics of bacteria are also important. Hydrophobic bacteria show a higher affinity towards hydrophobic surfaces, while hydrophilic bacteria prefer hydrophilic surfaces. E. coli has been reported as a hydrophilic bacterium (54, 55). However, the surface hydrophobicity of bacteria varies depending on the growth medium and bacterial age, as well as bacterial species (49). Hassan and Frank (56) compared the hydrophobicity of E. coli in TSB and nutrient broth and reported that E. coli in TSB was more hydrophilic. Therefore, the use of TSB medium in this study may be a factor in the delay of E. coli ’s attachment to the hydrophobic lozenge-patterned surface. As a consequence, the varied surface characteristics of bacteria in different media raises the question of how the bacteria will behave in their real environment such as body fluids and blood.
It was detected that the E. coli counts on the lozenge-patterned surface significantly increased with time (p<0.01) and the bacterial number was detected as 5.90 × 104 ± 6.17 × 103 CFU cm–2 at the end of the experiment. It was determined that the bacterial number was 161 times lower than that on the untreated surface at the end of the experiment. Several studies have reported that patterning of the surface for hydrophobicity reduced E. coli numbers (39, 54, 57, 58). The anti-fouling benefits of hydrophobic or superhydrophobic surfaces have also been reported in the literature (26, 59, 60). Approximately 99% decrease in E. coli number on the lozenge-patterned surface compared to the untreated surface could be considered an indicator of the effectiveness of laser texturing to reduce bacterial adhesion. However, reduced bacterial count does not mean that infection risk can be eliminated. Although the exact threshold of bacterial CFU per millilitre producing clinical symptoms is unclear and species-dependent, at levels as low as 10 CFU ml–1, sepsis becomes a threat (61). Surface patterning with a wetting angle around 120° as tested in the present research may be adequate to eliminate or reduce the risk of infection for up to 10 h since no bacteria were detected on the lozenge-patterned surface at this time.
3.3 Carbohydrate Concentrations
The amounts of total carbohydrate on the lozenge-patterned and untreated surfaces varied over the experiment and some fluctuations were observed (Figure 6). The total carbohydrate concentrations on the untreated surface increased up to 6 h of exposure and reached their highest value at 200.65 ± 9.86 μg cm–2. The concentrations then reduced until 10 h of exposure and the lowest value was detected as 84.55 ± 3.53 μg cm–2. Afterward, no significant changes were observed until the end of the experiment (Figure 6). Like the untreated surface, the total amount of carbohydrates in the lozenge-patterned surfaces was observed to peak (249.61 ± 11.55 μg cm–2) after 6 h of exposure and then decreased (177.19 ± 4.39 μg cm–2) until 10 h of exposure. However, an increase (228.71 ± 6.64 μg cm–2) after 12 h of exposure and then a decrease (152.99 ± 1.68 μg cm–2) after 24 h of exposure were detected. Afterward it remained stable until the end of the experiment (Figure 6). The fluctuations in carbohydrate amounts can be associated with the metabolic activity of the bacteria. It may be an indication that the bacteria in the biofilm produced carbohydrates, probably to create durable conditions, and were able to degrade self-produced carbohydrate to survive. Similar cases of self-produced EPS consumption have been observed in previous bacteriological experiments (62, 63).
The increase in total carbohydrate amounts on lozenge-patterned and untreated surfaces up to 6 h of exposure may be due to the accumulation of media-derived carbohydrates on the surfaces (Figure 4). It was observed that the amounts of culture-derived carbohydrates were higher on the lozenge-patterned surface than on the untreated ones. This may be due to the larger surface area of the lozenge-patterned surface compared to the untreated ones. Unfortunately, this situation raises the question of whether the texturing process will increase the adhesion of sugar in blood to the surgical equipment. In order to find an answer to this hypothesis, experiments must be carried out in blood. In addition, the decrease in carbohydrate amounts both on the lozenge-patterned and untreated surfaces after 8 h of exposure may be an indication that the bacteria contacted the surfaces and started consuming the carbohydrates on the surfaces to survive and attach.
During the experiment, the total carbohydrate amounts on the lozenge-patterned surface were significantly higher than those on the untreated surfaces (p<0.01). This was also supported by SEM micrographs (Figure 3). These findings may indicate that E. coli produced more EPS to attach to the lozenge-patterned surface to overcome the surface hydrophobicity and also to form a biofilm on the surface. It is known that bacteria secrete EPS to support stronger surface attachment and biofilm formation (64, 65). Therefore, the dense EPS layer on the lozenge-patterned surface may lead to an infection by causing an increase in the biofilm density and thus in the bacterial number with time.
Besides its protective and structural roles (66), EPS produced by E. coli can cause the development of phenotypic resistance and even the formation of permanent infections, owing to its components (67, 68). However, this is an indirect effect that plays in the persistence of bacteria rather than being directly implicated in the infective mechanism itself. Biofilms constitute reservoirs of bacteria that are potentially virulent, and the transition from biofilm to virulence and vice versa might be controlled by several regulators in E. coli (69).
This article highlights the differences in biofilm formation of E. coli on hydrophobic lozenge-patterned and hydrophilic untreated surfaces of 630 stainless steel. Although bacterial attachment was observed both on the lozenge-patterned and untreated surfaces, the texturing caused changes in the behaviour of E. coli on surfaces in terms of their attachment duration, EPS production and bacterial number. In light of these results, the effects of a hydrophobic surface due to increased wetting angle can be summarised as follows. Surface patterning with a wetting angle of around 120° delayed the attachment of E. coli for 6 h and resulted in an approximate 99% reduction in the number of bacteria attached to the surface. Contrary to the findings of the bacterial count, surface patterning caused E. coli to produce more EPS and form a multi-layered biofilm. Regarding the number of bacteria and retardation of attachment time, it can be recommended to use lozenge-patterned surfaces for short-term operations up to 10 h. However, the increased EPS development during the experiment limits the use of lozenge-patterned surfaces. The EPS finding obtained from this study reveals the importance of EPS analysis in surface modification studies aimed to delay or prevent biofilm formation.
‘Healthcare-Associated Infections (HAIs)’, Centers for Disease Control and Prevention (CDC), Atlanta, Georgia, USA: https://www.cdc.gov/hai/index.html (Accessed on 20th September 2021)
B. Allegranzi, S. B. Nejad, C. Combescure, W. Graafmans, H. Attar, L. Donaldson and D. Pittet, Lancet, 2011, 377, (9761), 228 LINK https://doi.org/10.1016/s0140-6736(10)61458-4
R. Donlan, Emerg. Infect. Dis., 2001, 7, (2), 277
C. J. Sanchez, K. Mende, M. L. Beckius, K. S. Akers, D. R. Romano, J. C. Wenke and C. K. Murray, BMC Infect. Dis., 2013, 13, 47 LINK https://doi.org/10.1186/1471-2334-13-47
S. de Souza Evangelista, S. G. dos Santos, M. A. de Resende Stoianoff and A. C. de Oliveira, Am. J. Infect. Control, 2015, 43, (5), 522 LINK https://doi.org/10.1016/j.ajic.2014.12.018
J. B. Kaper, J. P. Nataro and H. L. T. Mobley, Nat. Rev. Microbiol., 2004, 2, (2), 123 LINK https://doi.org/10.1038/nrmicro818
J. Jang, H.-G. Hur, M. J. Sadowsky, M. N. Byappanahalli, T. Yan and S. Ishii, J. Appl. Microbiol., 2017, 123, (3), 570 LINK https://doi.org/10.1111/jam.13468
M. A. Croxen and B. B. Finlay, Nat. Rev. Microbiol., 2009, 8, (1), 26 LINK https://doi.org/10.1038/nrmicro2265
K. Vickery, H. Hu, A. S. Jacombs, D. A. Bradshaw and A. K. Deva, Healthc. Infect., 2013, 18, (2), 61 LINK https://doi.org/10.1071/hi12059
J. W. Costerton, Z. Lewandowski, D. E. Caldwell, D. R. Korber and H. M. Lappin-Scott, Annu. Rev. Microbiol., 1995, 49, (1), 711 LINK https://doi.org/10.1146/annurev.mi.49.100195.003431
M. E. Davey and G. A. O’toole, Microbiol. Mol. Biol. Rev., 2000, 64, (4), 847 LINK https://doi.org/10.1128/mmbr.64.4.847-867.2000
H.-C. Flemming and J. Wingender, Nat. Rev. Microbiol., 2010, 8, (9), 623 LINK https://doi.org/10.1038/nrmicro2415
S. Fulaz, S. Vitale, L. Quinn and E. Casey, Trends Microbiol., 2019, 27, (11), 915 LINK https://doi.org/10.1016/j.tim.2019.07.004
B. Frølund, R. Palmgren, K. Keiding and P. H. Nielsen, Water Res., 1996, 30, (8), 1749 LINK https://doi.org/10.1016/0043-1354(95)00323-1
J. Wingender, M. Strathmann, A. Rode, A. Leis, and H.-C. Flemming, ‘Section Extracellular VI. Polymers: Isolation and Biochemical Characterization of Extracellular Polymeric Substances from Pseudomonas Aeruginosa’, in “Microbial Growth in Biofilms – Part A: Developmental and Molecular Biological Aspects”, ed. R. J. Doyle, Methods in Enzymology Book Series, Ch. 25, Vol. 336, Elsevier, Cambridge, USA, 2001, pp. 302–314 LINK https://doi.org/10.1016/s0076-6879(01)36597-7
A. W. Decho and T. Gutierrez, Front. Microbiol., 2017, 8, 922 LINK https://doi.org/10.3389/fmicb.2017.00922
H.-C. Flemming, J. Wingender, U. Szewzyk, P. Steinberg, S. A. Rice and S. Kjelleberg, Nat. Rev. Microbiol., 2016, 14, (9), 563 LINK https://doi.org/10.1038/nrmicro.2016.94
I. Sutherland, Trends Microbiol., 2001, 9, (5), 222 LINK https://doi.org/10.1016/s0966-842x(01)02012-1
X. Z. Li, B. Hauer and B. Rosche, Appl. Microbiol. Biotechnol., 2007, 76, (6), 1255 LINK https://doi.org/10.1007/s00253-007-1108-4
J. D. Bryers, Biotechnol. Bioeng., 2008, 100, (1), 1 LINK https://doi.org/10.1002/bit.21838
J. W. Costerton, L. Montanaro and C. r. Arciola, Int. J. Artif. Organs, 2007, 30, (9), 757 LINK https://doi.org/10.1177/039139880703000903
M. Kostakioti, M. Hadjifrangiskou and S. J. Hultgren, Cold Spring Harb. Perspect. Med., 2013, 3, (4), a010306 LINK https://doi.org/10.1101/cshperspect.a010306
W. A. Rutala and D. J. Weber, Clin. Infect. Dis., 2004, 39, (5), 702 LINK https://doi.org/10.1086/423182
M. Lorenzetti, I. Dogša, T. Stošicki, D. Stopar, M. Kalin, S. Kobe and S. Novak, ACS Appl. Mater. Interfaces, 2015, 7, (3), 1644 LINK https://doi.org/10.1021/am507148n
J. Hasan, S. Raj, L. Yadav and K. Chatterjee, RSC Adv., 2015, 5, (56), 44953 LINK https://doi.org/10.1039/c5ra05206h
X. Zhang, L. Wang and E. Levänen, RSC Adv., 2013, 3, (30), 12003 LINK https://doi.org/10.1039/c3ra40497h
S. Wu, B. Zhang, Y. Liu, X. Suo and H. Li, Biointerphases, 2018, 13, (6), 060801 LINK https://doi.org/10.1116/1.5054057
L. C. Hsu, J. Fang, D. A. Borca-Tasciuc, R. W. Worobo and C. I. Moraru, Appl. Environ. Microbiol., 2013, 79, (8), 2703 LINK https://doi.org/10.1128/aem.03436-12
K. A. Whitehead, J. Colligon and J. Verran, Coll. Surf. B: Bioint., 2005, 41, (2–3), 129 LINK https://doi.org/10.1016/j.colsurfb.2004.11.010
R. S. Friedlander, H. Vlamakis, P. Kim, M. Khan, R. Kolter and J. Aizenberg, Proc. Natl. Acad. Sci., 2013, 110, (14), 5624 LINK https://doi.org/10.1073/pnas.1219662110
F. Chen, D. Zhang, Q. Yang, J. Yong, G. Du, J. Si, F. Yun and X. Hou, ACS Appl. Mater. Interfaces, 2013, 5, (15), 6777 LINK https://doi.org/10.1021/am401677z
A. Y. Vorobyev and C. Guo, Laser Photonics Rev., 2012, 7, (3), 385 LINK https://doi.org/10.1002/lpor.201200017
A. Dunn, J. V. Carstensen, K. L. Wlodarczyk, E. B. Hansen, J. Gabzdyl, P. M. Harrison, J. D. Shephard and D. P. Hand, Opt. Lasers Eng., 2014, 62, 9 LINK https://doi.org/10.1016/j.optlaseng.2014.05.003
F. Song, H. Koo and D. Ren, J. Dent. Res., 2015, 94, (8), 1027 LINK https://doi.org/10.1177/0022034515587690
T. Wassmann, S. Kreis, M. Behr and R. Buergers, Int. J. Implant Dent., 2017, 3, 32 LINK https://doi.org/10.1186/s40729-017-0093-3
C. Díaz, M. C. Cortizo, P. L. Schilardi, S. G. G. de Saravia and M. A. F. L. de Mele, Mat. Res., 2007, 10, (1), 11 LINK https://doi.org/10.1590/s1516-14392007000100004
A. H. A. Lutey, L. Gemini, L. Romoli, G. Lazzini, F. Fuso, M. Faucon and R. Kling, Sci. Rep., 2018, 8, (1), 10112 LINK https://doi.org/10.1038/s41598-018-28454-2
R. Helbig, D. Günther, J. Friedrichs, F. Rößler, A. Lasagni and C. Werner, Biomater. Sci., 2016, 4, (7), 1074 LINK https://doi.org/10.1039/c6bm00078a
Q. Pan, Y. Cao, W. Xue, D. Zhu and W. Liu, Langmuir, 2019, 35, (35), 11414 LINK https://doi.org/10.1021/acs.langmuir.9b01333
A. de Bruin, Johnson Matthey Technol. Rev., 2018, 62, (3), 259 LINK https://technology.matthey.com/article/62/3/259-262/
M. Ayazi, N. G. Ebrahimi and E. J. Nodoushan, Int. J. Adhes. Adhes., 2019, 88, 66 LINK https://doi.org/10.1016/j.ijadhadh.2018.10.017
L.-C. Xu and C. A. Siedlecki, Acta Biomater., 2012, 8, (1), 72 LINK https://doi.org/10.1016/j.actbio.2011.08.009
H.-H. Ge, G.-D. Zhou and W.-Q. Wu, Appl. Surf. Sci., 2003, 211, (1–4), 321 LINK https://doi.org/10.1016/s0169-4332(03)00355-6
X. Zhang, P. L. Bishop and B. K. Kinkle, Water Sci. Technol., 1999, 39, (7), 211 LINK https://doi.org/10.2166/wst.1999.0361
M. DuBois, K. A. Gilles, J. K. Hamilton, P. A. Rebers and F. Smith, Anal. Chem., 1956, 28, (3), 350 LINK https://doi.org/10.1021/ac60111a017
C. Campanac, L. Pineau, A. Payard, G. Baziard-Mouysset and C. Roques, Antimicrob. Agents Chemother., 2002, 46, (5), 1469 LINK https://doi.org/10.1128/aac.46.5.1469-1474.2002
T.-F. C. Mah and G. A. O’Toole, Trends Microbiol., 2001, 9, (1), 34 LINK https://doi.org/10.1016/s0966-842x(00)01913-2
P. Stoodley, K. Sauer, D. G. Davies and J. W. Costerton, Annu. Rev. Microbiol., 2002, 56, 187 LINK https://doi.org/10.1146/annurev.micro.56.012302.160705
Y. H. An and R. J. Friedman, J. Biomed. Mater. Res., 1998, 43, (3), 338 LINK https://doi.org/10.1002/(sici)1097-4636(199823)43:3<338::aid-jbm16>3.0.co;2-b
M. Katsikogianni and Y. F. Missirlis, Eur. Cells Mater., 2004, 8, 37 LINK https://doi.org/10.22203/ecm.v008a05
T. R. Garrett, M. Bhakoo and Z. Zhang, Prog. Nat. Sci., 2008, 18, (9), 1049 LINK https://doi.org/10.1016/j.pnsc.2008.04.001
R. M. Goulter, I. R. Gentle and G. A. Dykes, Lett. Appl. Microbiol., 2009, 49, (1), 1 LINK https://doi.org/10.1111/j.1472-765x.2009.02591.x
V. M. Villapún, A. P. Gomez, W. Wei, L. G. Dover, J. R. Thompson, T. Barthels, J. Rodriguez, S. Cox and S. González, APL Mater., 2020, 8, (9), 091108 LINK https://doi.org/10.1063/5.0017580
N. Chik, W. S. Wan Md Zain, A. J. Mohamad, M. Z. Sidek, W. H. Wan Ibrahim, A. Reif, J. H. Rakebrandt, W. Pfleging and X. Liu, IOP Conf. Ser.: Mater. Sci. Eng., 2018, 358, 012034 LINK https://doi.org/10.1088/1757-899x/358/1/012034
Z. A. Mirani, A. Fatima, S. Urooj, M. Aziz, M. Khan and T. Abbas, Iran J. Basic Med. Sci., 2018, 21, (7), 760 LINK https://doi.org/10.22038/IJBMS.2018.28525.6917
A. N. Hassan and J. F. Frank, Int. J. Food Microbiol., 2004, 96, (1), 103 LINK https://doi.org/10.1016/s0168-1605(03)00160-0
F. H. Rajab, C. M. Liauw, P. S. Benson, L. Li and K. A. Whitehead, Food Bioprod. Process., 2018, 109, 29 LINK https://doi.org/10.1016/j.fbp.2018.02.009
D. Patil, S. Aravindan, M. K. Wasson, V. P. and and P. V. Rao, J. Micro Nano-Manuf., 2018, 6, (1), 011002 LINK https://doi.org/10.1115/1.4038093
P. V. Mahalakshmi, S. C. Vanithakumari, J. Gopal, U. K. Mudali and B. Raj, Curr. Sci., 2011, 101, (10), 1328 LINK https://www.currentscience.ac.in/Volumes/101/10/1328.pdf
J. Chapman and F. Regan, Adv. Eng. Mater., 2012, 14, (4), B 175 LINK https://doi.org/10.1002/adem.201180037
W. G. Pitt, M. Alizadeh, G. A. Husseini, D. S. McClellan, C. M. Buchanan, C. G. Bledsoe, R. A. Robison, R. Blanco, B. L. Roeder, M. Melville and A. K. Hunter, Biotechnol. Prog., 2016, 32, (4), 823 LINK https://doi.org/10.1002/btpr.2299
S. Arkan-Ozdemir, N. Cansever and E. Ilhan-Sungur, Water Sci. Technol., 2020, 82, (5), 940 LINK https://doi.org/10.2166/wst.2020.396
E. Ilhan-Sungur and A. Çotuk, Corros. Sci., 2010, 52, (1), 161 LINK https://doi.org/10.1016/j.corsci.2009.08.049
H. Rohde, E. C. Burandt, N. Siemssen, L. Frommelt, C. Burdelski, S. Wurster, S. Scherpe, A. P. Davies, L. G. Harris, M. A. Horstkotte, J. K.-M. Knobloch, C. Ragunath, J. B. Kaplan and D. Mack, Biomaterials, 2007, 28, (9), 1711 LINK https://doi.org/10.1016/j.biomaterials.2006.11.046
E. A. Izano, M. A. Amarante, W. B. Kher and J. B. Kaplan, Appl. Environ. Microbiol., 2008, 74, (2), 470 LINK https://doi.org/10.1128/aem.02073-07
G. Sharma, S. Sharma, P. Sharma, D. Chandola, S. Dang, S. Gupta and R. Gabrani, J. Appl. Microbiol., 2016, 121, (2), 309 LINK https://doi.org/10.1111/jam.13078
G. G. Anderson, J. J. Palermo, J. D. Schilling, R. Roth, J. Heuser and S. J. Hultgren, Science, 2003, 301, (5629), 105 LINK https://doi.org/10.1126/science.1084550
S. S. Justice, C. Hung, J. A. Theriot, D. A. Fletcher, G. G. Anderson, M. J. Footer and S. J. Hultgren, Proc. Natl. Acad. Sci., 2004, 101, (5), 1333 LINK https://doi.org/10.1073/pnas.0308125100
C. Beloin, A. Roux, and J.-M. Ghigo, ‘ Escherichia coli Biofilms’, in “Bacterial Biofilms”, ed. T. Romeo, Current Topics in Microbiology and Immunology Series, Vol. 322, Springer-Verlag, Berlin, Germany, 2008, pp. 249–289 LINK https://doi.org/10.1007/978-3-540-75418-3_12
Simge Arkan-Ozdemir is a PhD candidate in Fundamental and Industrial Microbiology at Istanbul University, Turkey. She obtained a BA degree in Biology and an MSci degree in Fundamental and Industrial Microbiology from the same institution. The focus of her PhD is to investigate the microbiologically induced corrosion behaviours of anaerobic and aerobic bacteria and whether changes in the EPS contents in biofilm formed by bacteria on a metal surface affect the microbiologically induced corrosion. Her current research interests are microbial corrosion, bacteriology, biofilms and microbial ecology.
Nurhan Cansever holds a PhD degree in Metallurgical and Materials Science and currently works as a full-time professor at Yildiz Technical University in Istanbul, Turkey. A few of her research interests may be listed as materials science: atomic structure of materials, structure-property relations; corrosion: stress corrosion cracking, microbiologically influenced corrosion; surface treatments: electroless nickel coatings, thin films and anodic oxidation. She has worked as a visiting scientist in the University of Salford, UK, for a short period of time. She is a member of the Corrosion Association in Turkey.
Esra Ilhan-Sungur is professor in the Biology Department at Istanbul University, Turkey, since 2018. A key focus of her research is microbiologically induced corrosion and its prevention. Further research interests lie in the areas of anaerobic bacteria (especially sulfate-reducing bacteria), petroleum microbiology, microbial diversity and ecology, microbial genetics and biofilm. She was awarded a postdoctoral research scholarship by the Scientific and Technological Research Council of Turkey (TUBITAK-BIDEB) and worked as a guest researcher at Delft University of Technology, The Netherlands.