Prevention of Bacterial Colonization on Catheters by a One-Step

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Prevention of Bacterial Colonization on Catheters by a One-Step Coating Process Involving an Antibiofouling Polymer in Water Hyeongseop Keum,† Jin Yong Kim,‡ Byeongjun Yu,† Seung Jung Yu,§ Jinjoo Kim,† Hyungsu Jeon,† Dong Yun Lee,‡ Sung Gap Im,§ and Sangyong Jon*,†,‡ †

KAIST Institute for the BioCentury, Department of Biological Sciences, ‡Graduate School of Medical Science and Engineering, and KAIST Institute for the NanoCentury, Department of Chemical and Biomolecular Engineering, Korea Advanced Institute of Science and Technology (KAIST), 291 Daehak-ro, Daejeon 34141, Republic of Korea §

S Supporting Information *

ABSTRACT: As reports of multidrug resistant pathogens have increased, patients with implanted medical catheters increasingly need alternative solutions to antibiotic treatments. As most catheter-related infections are directly associated with biofilm formation on the catheter surface, which, once formed, is difficult to eliminate, a promising approach to biofilm prevention involves inhibiting the initial adhesion of bacteria to the surface. In this study, we report an amphiphilic, antifouling polymer, poly(DMA-mPEGMA-AA) that can facilely coat the surfaces of commercially available catheter materials in water and prevent bacterial adhesion to and subsequent colonization of the surface, giving rise to an antibiofilm surface. The antifouling coating layer was formed simply by dipping a model substrate (polystyrene, PET, PDMS, or silicon-based urinary catheter) in water containing poly(DMA-mPEGMA-AA), followed by characterization by X-ray photoelectron spectroscopy (XPS). The antibacterial adhesion properties of the polymer-coated surface were assessed for Staphylococcus aureus (S. aureus) and Escherichia coli (E. coli) growth under static (incubation in the presence of a bacterial suspension) and dynamic (bacteria suspended in a solution under flow) conditions. Regardless of the conditions, the polymer-coated surface displayed significantly reduced attachment of the bacteria (antiadhesion effect > ∼8-fold) compared to the bare noncoated substrates. Treatment of the implanted catheters with S. aureus in vivo further confirmed that the polymer-coated silicon urinary catheters could significantly reduce bacterial adhesion and biofilm formation in a bacterial infection animal model. Furthermore, the polymer-coated catheters did not induce hemolysis and were resistant to the adhesion of blood-circulating cells, indicative of high biocompatibility. Collectively, the present amphiphilic antifouling polymer is potentially useful as a coating platform that renders existing medical devices resistant to biofilm formation. KEYWORDS: antibiofouling polymer, biofilms, catheters, healthcare related infection, medical devices, polymer films

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INTRODUCTION Medical device-related infections present a serious concern as life-threatening complications for patients who received medical implants, catheters, and endotracheal tubes. Medical devicerelated infections have contributed to an increase in morbidity and mortality, and they present a huge financial burden to patients.1 Among the common medical device-related infections, intravenous and urinary catheter-associated nosocomial infections are considered high-priority areas that account for more than 25% of all healthcare-associated infections in the United States.2 The formation of microbial biofilms on the © XXXX American Chemical Society

surfaces of medical devices is thought to be the main cause of the increased risks associated with life-threatening infections, because the bacteria in biofilms are protected by a matrix of extracellular polymeric substrates and develop multidrug resistance within these matrices. In fact, approximately 80% of all human infections are attributable to microbial biofilms.2−7 Received: May 16, 2017 Accepted: May 24, 2017

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DOI: 10.1021/acsami.7b06899 ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX

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ACS Applied Materials & Interfaces

Figure 1. Formation and characterization of the antifouling polymer-coated substrates. (A) A schematic illustration for the facile method of forming the poly(DMA-mPEGMA-AA) coating layer on a catheter substrate. The implantable medical devices (e.g., silicone urinary catheter) were coated simply by dipping into the poly(DMA-mPEGMA-AA) solution. (B) Static contact angles of the uncoated and polymer-coated surfaces of different substrates (polystyrene, PDMS silicone rubber, or PET). A droplet of purified water (10 μL) was dispensed onto the respective surface, and the measurements were immediately collected using a contact angle analyzer. (C) XPS survey scan spectra of the PET, PDMS (silicone rubber), polystyrene, and corresponding polymer-coated surfaces. The measurements were collected prior to applying the coating, immediately after applying the coating, and 7 days after applying the coating.

The majority of catheters are made using flexible, durable, hydrophobic polymer materials, including silicon rubber, polyurethane, and latex. A crucial problem with catheter materials is that they are prone to biofilm formation under pristine conditions, as their surfaces are rapidly fouled and coated with biological fluids, such as plasma proteins that facilitate bacterial attachment. Researchers have tested a variety of approaches for avoiding biofilm formation by incorporating functionalities into the catheter surfaces. Antimicrobials, such as antibiotics (e.g., rifampicin) and silver ion- or nanoparticlecoated catheters have performed well in inhibiting biofilm formation;8,9 however, silver compound (e.g., chlorhexidinesilver sulfadiazine)-coated catheters can induce hypersensitive reactions in patients.4,6,10−15 Antibiotic catheter coatings are less effective in preventing biofilm formation because the antibiotics do not work against all strains of bacteria; moreover, catheter-associated infections are mostly caused by antibioticsresistant pathogens.10,16,17 Biofilms are formed through a series

of steps that include reversible attachment, irreversible adhesion, colonization, and finally maturation of microbial communities within an extracellular polymeric substrate. Once adherence and subsequent colonization of bacteria takes place on a catheter surface, it is difficult to stop subsequent biofilm formation. Efforts have been made to intervene during the first bacterial adhesion step by coating the surface with an antifouling polymer; however, such antifouling surface coatings alone are not found to be as efficient as antimicrobial agentcoated surfaces, presumably due to the modest level of antibiofouling effect and the low or nonuniform surface coverage of the functionality. Furthermore, such coatings require manufacturing processes that are more complex than those of conventional medical catheters.1,11,18 As such, there is a need for a new antifouling coating material that enables the efficient and durable blocking of bacterial adhesion through a facile and reproducible manufacturing process. It would be ideal if the antiadhesion coating were ready-for-use in coating the B


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Table 1. Changes in the Atomic Concentrations on the PET, PDMS, and Polystyrene Substrates upon Polymer Coatinga materials PET

silicone (PDMS)

polystyrene

uncoated polymer-coated polymer-coated uncoated polymer-coated polymer-coated uncoated polymer-coated polymer-coated

(D0) (D7) (D0) (D7) (D0) (D7)

C (%)

O (%)

Si (%)

total (%)

73.37 94.41 87.37 29.00 88.60 79.35 91.11 83.52 81.82

26.63 5.59 12.63 44.10 9.66 17.47 6.32 14.32 14.18

0 0 0 26.90 1.74 3.18 2.57 2.16 4

100 100 100 100 100 100 100 100 100

a

Numerical values obtained from the XPS spectra collected from various substrates (PET, PDMS silicone rubber, and polystyrene) prior to coating, immediately after coating, and 7 days past coating. The chemical concentration differences between the bare and polymer-coated substrates are apparent from the data.

rendered the poly(DMA-mPEGMA-AA) product fairly watersoluble. Polydimethylsiloxane (PDMS), polystyrene, and polyethylene terephthalate (PET) substrates, as models of the medical devices, were tested to examine whether the amphiphilic polymer could coat the surfaces of the substrates. Immersion of the substrates in the polymer solution in water was predicted to result in the anchoring of numerous copies of the long dodecyl chain onto the hydrophobic surface of the substrates via hydrophobic or van der Waals interactions, thereby enabling the formation of a stable coating layer (Figure 1A). Multiple PEG chains on the surfaces of the polymercoated substrates were predicted to prevent the adsorption of biological species, such as proteins and cells. Negatively charged methacrylic acid may play a role in repelling negatively charged bacterial species. The formation of a polymer coating layer on the substrates was confirmed using static contact angle measurements in distilled water and X-ray photoelectron spectroscopy (XPS). Noticeable changes in the contact angles of the substrates were observed before and after applying the polymer coatings: 105.4 ± 1.3° versus 75.2 ± 0.8° for polystyrene, 101.9 ± 1.8° versus 81.1 ± 2.2° for silicone, and 67.2 ± 2.0° versus 72.9 ± 0.6° for PET (Figure 1B). Regardless of the type of substrate or specific contact angle, the resulting polymer-coated surfaces displayed similar contact angles, indicating that the amphiphilic, antibiofouling polymer coating layer was successfully formed as described in Figure 1A. These coating layers changed the surface wettability in a consistent manner. XPS survey scans further revealed the significant changes in the chemical compositions and atomic concentrations (C, O, and Si) present on each surface before and after formation of the polymer coating on the three substrates (PET, PDMS, and polystyrene) (Figure 1C). The numeric values of the atomic concentrations in each surface are listed in Table 1. For the PET surface, the atomic concentration of carbon increased from 73.37% to 94.41%, whereas the atomic concentration of oxygen decreased from 26.63% to 5.59% after applying the poly(DMA-mPEGMA-AA) polymer coating, which contained a higher percentage of carbon atoms than the pristine PET surface. Unlike the polymer-coated PET substrate, the polymercoated polystyrene surface showed a decrease in the atomic content of carbon from 91.11% to 83.52% and an increase in the oxygen content from 6.32% to 14.32% because poly(DMAmPEGMA-AA) contained a higher percentage of oxygen atoms than the pristine polystyrene surface. Lastly, the polymercoated PDMS silicone elastomer showed a dramatic increase in the atomic concentration of carbon from 29.00% to 88.60% and

surface of an existing commercial catheter using a simple process; moreover, aqueous solutions of such coatings are most desirable to avoid concerns around hazard chemicals or solvent remaining in the catheter. Recently, we reported that an amphiphilic polymer synthesized from a long alkyl chain dodecyl methacrylate (DMA), a short poly(ethylene glycol) methacrylate (PEGMA), and an acrylic acid (AA), designated as a poly(DMAmPEGMA-AA), could effectively coat the surfaces of hydrophobic materials, such as carbon nanotubes and plastic surfaces (i.e., polystyrene and cyclic olefin copolymer substrates). The resulting polymer-coated surfaces then effectively prevented the nonspecific adsorption of plasma proteins.19−21 In this study, we hypothesized that the amphiphilic water-soluble, antifouling polymer poly(DMA-mPEGMA-AA) could facilely coat the surfaces of commercial catheter materials in water, and the resulting polymer-coated catheters could prevent bacterial adhesion and subsequent bacterial colonization that tends to give rise to biofilm formation.19,22 It would be advantageous if the overall coating process could be rapidly completed in onestep in an environmentally friendly hygienic aqueous solution; moreover, the process should not alter the designed mechanical properties of the existing catheters. In this report, we examined whether the amphiphilic, antibiofouling polymer could coat the surfaces of several typical catheter materials. The resulting polymer-coated surfaces were shown to prevent the adhesion and subsequent colonization of pathogenic bacteria using a variety of characterization means, including in vitro and in vivo performance tests. The antibiofilm effect was methodically studied under static or dynamic conditions. The biocompatibility of the polymer were examined for their physiological suitability. Finally, in vivo experiments were carried out to validate the effect of the polymer coating on bacterial colony formation.

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RESULTS AND DISCUSSION Formation and Characterization of the Antibiofouling Polymer-Coated Substrates. The antibiofouling polymer, poly(DMA-mPEGMA-AA), was synthesized through radical polymerization of the corresponding monomers with a molar feed ratio of 3.5 (DMA):3.5 (PEGMA):3 (AA), as reported previously.19 The actual ratio of each monomer unit in the polymer, which was calculated based on the integration values in the 1H NMR spectrum, was proven to be 33.45% for DMA, 33.66% for PEGMA, and 32.89% for AA, close to the initial molar feed ratio (see Figure S1 of the Supporting Information). The hydrophilicity of the PEG and acrylic acid residues C


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Figure 2. Antiadhesion effects of the poly(DMA-mPEGMA-AA)-coated substrates (polystyrene/PDMS) against bacteria. (A) Microscopy images of the antiadhesion tests conducted using Gram-stained S. aureus grown on the poly(DMA-mPEGMA-AA)-coated polystyrene surfaces. S. aureus (3.0 × 106 CFU/mL) was incubated on the poly(DMA-mPEGMA-AA)-coated polystyrene surfaces (20 mg/mL) for 12 h at 37 °C, followed by washing (3 times with 1X PBS). The scale bar indicates 120 μm. (B) The antiadhesion effects of the PDMS substrates coated using poly(DMA-mPEGMA-AA) solutions in various concentrations (0, 1, 2, 5, 10, 20, mg/mL) and incubated in the presence of S. aureus in LB media (3.0 × 106 CFU/mL) for 8 and 24 h, respectively. The WST-1 color intensity of the S. aureus film grown on the polymer-coated surfaces was measured at 440 nm using a plate reader, and the antiadhesion percentages were denoted relative to a control uncoated PDMS sample. Fluorescence images (C) and the percentage (D) of adhesion of mCherry-labeled E. coli (3.0 × 106 CFU/mL) in each well containing the polymer-coated PDMS are indicated relative to a control uncoated PDMS sample.

positive bacteria, as it is commonly associated with catheter biofilms.10 The surface of a polystyrene-based 96-well plate was coated with a solution of poly(DMA-mPEGMA-AA) in water (20 mg/mL). Gram-stained S. aureus bacteria (stained prior to seeding) were used to test biofilm formation on the polymercoated wells to facilitate detection by color. As shown in Figure 2A, a microscopic image at 400× magnification displayed a significant level of adhesion of S. aureus onto the surface of the bare uncoated polystyrene substrate, even after 3 harsh washes with PBS. By contrast, few Gram-stained bacteria were detected on the polymer-coated polystyrene surface after the washing step. These results indicated that the antibiofouling properties of the poly(DMA-mPEGMA-AA) against proteins may be applied to construct an antiadhesion surface against bacteria. PDMS (a silicon elastomer) was next tested as a model substrate for medical catheters. A series of antibiofouling polymer-coated silicone substrates were prepared in water containing different amounts of poly(DMA-mPEGMA-MA), ranging from 1 (0.1 wt %) to 20 mg/mL (2 wt %). The substrates were then tested for their antiadhesion effects against bacteria. S. aureus and Escherichia coli (E. coli), selected as typical Gram-positive and -negative bacteria, respectively. A WST-1 assay was used to visually observe the mitochondrial dehydrogenase activities of the cells using a formazan dye. Each substrate was immersed in a solution containing S. aureus (3.0 × 106 CFU/mL) for 8 and 24 h, respectively. Figure 2B plots the percentage color intensity relative to the control bare surface in a proliferation assay. All of the polymer-coated silicone surfaces, regardless of the polymer concentration used,

a significant decrease in both the oxygen content, from 44.10% to 9.96%, and the Si content, from 26.90% to 1.74%, as the predominant siloxane (OSiO) groups in the pristine silicone surface became much less exposed as a result of the coverage of poly(DMA-mPEGMA-AA). The XPS surface characterization data clearly indicated that the polymer coating layers were successfully formed on the three different substrates in a one-step environmentally friendly process in water. We next assessed the durability and stability of the polymer coating layer on each substrate. An XPS survey scan of the polymer-coated substrate surfaces submerged in PBS for 7 days revealed that most of the coating layers remained present on day 7 (Figure 1C and Table 1). Although some variations in the chemical content and atomic concentrations in each surface were observed upon incubation in PBS for 7 days, the atomic concentrations remained closer to the values obtained from the freshly coated surfaces than from the noncoated pristine substrates. Although the polymer coating layer on each substrate was formed entirely through noncovalent interactions, it persisted for at least 7 days, suggesting that the coating was highly durable. Antiadhesion Effects of the Poly(DMA-mPEGMA-AA)Coated Polystyrene/PDMS against Bacteria. In earlier works, we showed that poly(DMA-mPEGMA-AA) could coat a polystyrene surface and prevent nonspecific protein adsorption.19 On the basis of these previous findings, we examined whether the polymer-coated polystyrene could prevent the adhesion and subsequent colonization of bacteria. Staphylococcus aureus (S. aureus) was selected as a typical GramD


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ACS Applied Materials & Interfaces displayed only 1−6% adhesion of S. aureus relative to the control uncoated silicone surface. Notably, even the lowest concentration of polymer (1 mg/mL) displayed excellent antiadhesion properties against S. aureus, with nearly 95.5% prevention relative to the control. This value was comparable to that obtained from the highest concentration of the polymer. Repeated independent experiments (n = 3) provided the same conclusion, indicative of good reproducibility. We next examined whether the polymer-coated silicone was effective against Gram-negative bacteria. Silicone-molded black 24 well plates were incubated for 24 h in the presence of a standardized mCherry-tagged E. coli solution, to enable fluorescence detection.23 As shown in Figure 2C, a high fluorescence intensity of mCherry was observed in the well of the uncoated bare silicon surface; however, little fluorescence was observed in the polymer-coated wells, indicating few bacterial adhesions. As observed with Gram-positive bacteria, the lowest polymer concentration (1 mg/mL) was sufficient to prevent bacterial adhesion, displaying a 10-fold lower fluorescence intensity than the control (Figure 2D). Taken together, these results indicated that the poly(DMA-mPEGMAAA) coating on a silicone elastomer surface formed readily and was effective in preventing the adhesion of both Gram-positive and -negative bacteria. Long-Term Prevention of Bacterial Colonization on a Poly(DMA-mPEGMA-AA)-Coated PDMS Surface under Static Conditions. Medical catheters, such as intravenous or urinary catheters, are used over days or even weeks; therefore, the durability of the antiadhesion polymer coating is a key requirement for commercial success. The antiadhesion durability was tested by incubating the poly(DMA-mPEGMAAA)-coated PDMS substrate for 7 days in the presence of S. aureus. SEM images of the bare and polymer-coated PDMS surfaces after 7 days revealed a stark difference between the surfaces in terms of the number of bacteria present on each surface (Figure 3). Most of the bare PDMS surface was covered with multiple layers of bacteria in biofilms, whereas few bacteria were observed on the surface of the polymer-coated PDMS.

These results suggested that the polymer coating layer on the PDMS surface was stable for at least 7 days and maintained its ability to inhibit adhesion and the subsequent colonization of S. aureus. Antibiofilm Effect of the Poly(DMA-mPEGMA-AA)Coated Urinary Catheter under Solution Flow Conditions. The antibacterial adhesion effect of the polymer coating on a commercial silicone-based urinary catheter commonly used in the clinic was evaluated under conditions similar to those observed in the clinical setting. As shown in Figure 4A, a solution containing S. aureus was allowed to flow over the surfaces of the bare or polymer-coated urinary catheters for 2 days using a peristaltic pump. After washing, bacterial adhesion to each catheter was visualized by Gram staining using crystal violet (CV) dye. Although the bare urinary catheter displayed a clear intense violet color distributed across its surface, the polymer-coated catheter displayed a dramatically less intense color within the tube (Figure 4B). The optical density (O.D.) in each Gram-stained catheter was measured to reveal that the polymer coating on the urinary catheter resulted in a ∼ 77% reduction in the S. aureus adhesion relative to the bare catheter (0.44 versus 3.26 O.D.) under the flow conditions (Figure 4C). These results clearly indicated that the one-step postcoating of the commercial urinary catheter was effective enough to prevent bacterial colonization and biofilm formation on the catheter surface. Evaluation of the Biocompatibility of the Poly(DMAmPEGMA-AA)-Coated PDMS. The hemolytic activity of the polymer-coated surface was assessed using sheep blood agar plates.24 As shown in Figure 5A, neither the pristine nor the polymer-coated PDMS silicone elastomers showed signs of inducing red blood cell hemolysis in the agar plates. We next examined whether the polymer coating could inhibit the adhesion of the blood-circulating cells, such as platelets or immune cells, because medical catheters, particularly intravenous catheters, are exposed to a variety of bodily fluids, including blood. The bare and polymer-coated PDMS substrates were exposed to freshly collected rat blood for 0.5 h, followed by washing. FE-SEM images of the surfaces revealed that blood-circulating cells, including platelets (the largest such cells) adhered substantially onto the bare PDMS surface, whereas few cells were observed on the polymer-coated surface (Figure 5B). The antiadhesion effects of the polymer-coated surface against blood cells were attributed to the inhibition of nonspecific plasma protein adsorption, which mediates the attachments of blood cells onto abiotic surfaces. These hemolysis and cell attachment results suggested that the polymer coating layer on the abiotic catheter surfaces was biocompatible and potentially suitable for preparing antibiofilm surfaces. Antibiofilm Efficacy of the Poly(DMA-mPEGMA-AA)Coated Urinary Catheters under In Vivo Conditions. We next examined whether the poly(DMA-mPEGMA-AA)-coated silicon catheters inhibited biofilm formation in vivo by implanting the bare (noncoated) or polymer-coated urinary silicon catheters subcutaneously at the right flank of a mouse. After 2 days, either PBS or S. aureus (5.0 × 106 CFU in 100 μL PBS) was injected into the site harboring the catheters. Mice were divided into four groups (n = 4/group), receiving bare or polymer-coated catheters, each treated with either PBS or S. aureus. Over the 4-day period after either PBS or bacteria injection, the control groups displayed relatively neat surgical sites, whereas severe pus or byproducts from inflammatory

Figure 3. Long-term prevention of bacterial colonization of the poly(DMA-mPEGMA-AA)-coated PDMS under static conditions. The biofilm-inhibiting efficacy of poly(DMA-mPEGMA-AA) under static conditions was demonstrated using FE-SEM imaging. S. aureus grown in LB media (3.0 × 106 CFU/mL) was incubated for 7 days on the poly(DMA-mPEGMA-AA)-coated or bare PDMS surfaces with periodic media additions every 24 h to avoid dehydration. The scale bars indicate 250X: 200 μm and 5000X: 10 μm. E


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Figure 4. Antibiofilm effects of the poly(DMA-mPEGMA-AA)-coated urinary catheter under flow conditions. (A) Schematic representation of the experimental setup used to conduct the biofilm control assay under S. aureus solution flow conditions. An S. aureus solution was flowed through the silicone tube and the (poly(DMA-mPEGMA-AA)-coated or uncoated urinary catheters) at a rate of 2 mL/min. The S. aureus culture was stirred continuously to avoid bacteria sedimentation. (B) Image of the CV-stained urinary catheters after a 48-h incubation period at room temperature. The excess CV stain was removed by immersing each catheter into distilled water 3 times. (C) Quantitative analysis of the destained CV dyes obtained from the respective catheters. The O.D. was measured at an absorbance of 580 nm using a plate reader.

efficacy of the polymer coating (Figure 6C). In the case of PBS treatment, both the bare and coated catheters provided mean O.D. values of 0.045 and 0.039, respectively, indicative of little differences between the surfaces. By contrast, S. aureus treatment of the polymer-coated catheter displayed a significantly lower O.D compared to the pristine bare catheters (0.102 versus 0.770), indicative of potent antibiofilm efficacy. These results could be interpreted as follows. Although the commercial as-manufactured bare silicone catheters were prone to bacterial adhesion and biofilm formation that protected the bacteria from the host defense, the antibiofouling polymercoated catheters tested here effectively resisted bacterial adhesion, rendering the planktonic bacteria vulnerable to immune attack.25

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CONCLUSIONS In this study, we reported a new use of an amphiphilic, antibiofouling polymer that effectively, durably, and reproducibly inhibited adhesion and colonization of pathogenic bacteria to a catheters’ surface. The antiadhesion coating could be readily formed on the surfaces of commercial catheters using a simple one-step dipping process. The aqueous dipping solution precluded concerns surrounding the presence of residual hazardous chemicals or solvents that may remain in the catheters. The antibiofouling poly(DMA-mPEGMA-AA)coated silicon catheters (urinary catheters and PDMS) effectively inhibited the adhesion of S. aureus and E. coli and the subsequent colonization or biofilm formation on the catheter surfaces under both static and dynamic conditions. It should be further noted that the antibacterial adhesion properties of the polymer coating were maintained for at least 7 days. Moreover, treatments of the implanted catheters with S. aureus in vivo confirmed that the poly(DMAmPEGMA-AA)-coated silicon catheters could significantly reduce bacterial adhesion and biofilm formation in a bacterial

Figure 5. Biocompatibility evaluation of the poly(DMA-mPEGMAAA)-coated PDMS. (A) Hemolysis activities of uncoated- and polymer-coated PDMS measured on a sheep blood agar plate after incubation at 37 °C for 16 h. (B) FE-SEM images of the attachment of blood circulating cells on the bare and poly(DMA-mPEGMA-AA)coated PDMS surfaces that were incubated with freshly collected rat plasma at 37 °C for 30 min, followed by washing using sterilized 1X PBS. The scale bar indicates 200 μm.

complications were visible across all mice injected with bacteria. The harvested catheters were Gram-stained using crystal violet dye to reveal clear differences in the colorization and transparency of the bare and polymer-coated groups. Substantial numbers of violet spots were observed across the surfaces of the bare catheters, indicating the presence of bacterial aggregates or biofilms; however, all of the polymercoated catheters displayed little colorization and appeared transparent (Figure 6B). The transparency of each catheter was measured in terms of the O.D. value to quantify the antibiofilm F


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Figure 6. Antibiofilm efficacy of the poly(DMA-mPEGMA-AA)-coated urinary catheters under in vivo conditions. (A) Overview of the in vivo experimental schedule. The entire set of in vivo experiments was carried out over 7 days. On day 7, mice were humanely sacrificed, and the inserted catheters were retrieved. (B) The CV staining results obtained from the urinary catheters (1 cm each) retrieved after the subcutaneous injection of S. aureus (5.0 × 106 CFU in 100 μL PBS). Excess CV stain was eliminated by immersing each urinary catheter piece into sterile DW 3 times. (C) The average O.D. values of the CV stained catheters with treatments of PBS and S. aureus. The measurements were conducted at an absorbance of 580 nm using a plate reader. 16 h. The prepared bulk silicone in the Petri dish was later uniformly sectioned (1.5 × 1.5 cm2). All prepared silicone rubber surfaces were washed with 100% ethanol and distilled water three times each, followed by moist evaporation under nitrogen gas stream.27 Polystyrene Surface Preparation. To prepare polystyrene surfaces for the polymer coating and contact angle measurements, Petri dishes were cut into uniform sizes (1.5 cm × 1.5 cm). Sectioned Petri dish surfaces were washed with 100% ethanol and distilled water three times each. Sterile polystyrene 96-well plates were used for the antiadhesion tests of the poly(DMA-mPEGMA-AA)-coated polystyrene against bacteria. PET Surface Preparation. A PET sheet was kindly provided by Dooroo Celltech (Kyounggi-do, Korea). The PET sheet was cut to a uniform size (1.5 × 1.5 cm2) and washed with 100% ethanol and distilled water three times each for further experiments. Coating of the Polymer onto the Surfaces. Two-hundred milligrams of the viscous polymer concentrate were dissolved in 10 mL distilled water to a 20 mg/mL concentration. The incised polystyrene, silicone rubber, and PET sections (1.5 × 1.5 cm2) were immersed in the well containing 200 μL poly(DMA-mPEGMA-AA) in water (20 mg/mL) at 4 °C for 16 h. After incubation, the surfaces were washed with distilled water three times. Formation and Characterization of the Antibiofouling Polymer-Coated Substrates. The polystyrene, silicone rubber, and PET surfaces were prepared for the static contact angle measurements. Each material surface was coated with polymer at 4 °C for 16 h. Once the surfaces had been coated, a droplet (10 μL) of distilled water was pipetted onto the bare and polymer-coated substrates, respectively. The contact angles were immediately measured using a contact angle analyzer (Phoenix 150, SEO, Kyounggi-do, Korea). The contact angles were measured three times, and the statistical average was calculated. The chemical compositions of bare or polymer-coated polystyrene, silicone rubber, and PET substrates (1 × 1 cm2) were characterized using X-ray photoelectron spectroscopy (Sigma Probe Multipurpose XPS; K-alpha, Thermo VG Scientific). A monochromatic Al Kα radiation X-ray source (12 kV, KE = 1486.6 eV) was used to obtain the survey scan spectra while maintaining a base pressure of 2.0 × 10−9 mb. The XPS survey scan spectra were recorded three times over the range 100−1100 eV. The atomic concentration was calculated using

infection animal model. Lastly, the polymer-coated catheters did not induce hemolysis and were resistant to adhesion by blood cells, indicative of a high degree of biocompatibility. Taken together, the ease with which the aqueous poly(DMAmPEGMA-AA) could coat a hydrophobic surface renders this postmodification process applicable to a variety of medical devices in an effort to improve their resistance to lifethreatening pathogen infections. This coating approach holds considerable potential as an antibiofilm platform.

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MATERIALS AND METHODS

Polymer Synthesis. 3.5 mmol dodecyl methacrylate (DMA) (Sigma-Aldrich, St. Louis, U.S.A.), 3.5 mmol poly(ethylene glycol) methyl ether methacrylate (mPEGMA, Mn ≈ 475) (Sigma-Aldrich, St. Louis, U.S.A.), 3.0 mmol acrylic acid (AA) (Sigma-Aldrich, St. Louis, U.S.A.), 0.1 mmol 2,2′-azobis(2-methylpropionitrile) (AIBN) (SigmaAldrich, St. Louis, U.S.A.), and 10 mL THF (Sigma-Aldrich, St. Louis, U.S.A.) were quantitatively measured. Prior to conducting the polymer synthesis, the neat PEGMA inhibitor was removed using an inhibitor removal column (Sigma-Aldrich, St. Louis, U.S.A.). THF was degassed for 15 min under N2. DMA, mPEGMA, AA, and AIBN were carefully added to the degassed THF, and the solution was firmly sealed. The chemical mixture was polymerized with stirring for 24 h in a 70 °C oil bath, followed by overnight evaporation to obtain the polymer in the absence of solvent.19,20 The synthesis was confirmed by 1H NMR spectroscopy using a 400 MHz NMR spectrometer (Bruker 400, Billerica, U.S.A.). Silicone Surface Preparations. The silicone rubber surface was prepared by thoroughly mixing the silicone elastomer base and SYLGARD 184 kit silicone curing agent (Dow Corning, Midland, U.S.A.) in a 10:1 ratio (w/w) to form the bulk silicone rubber. The silicone mixture was cast in Petri dishes (contact angle measurements, XPS measurements, long-term prevention of bacterial colonization, and biocompatibility tests were conducted using these samples) (SPL Life Science, Kyounggi-do, Korea), in 96-well plates (Gram-positive bacteria antiadhesion test) (SPL Life Science, Kyounggi-do, Korea),26 and in black 24-well plates (Gram-negative bacteria antiadhesion test) (Cellvis, Mountain View, U.S.A.), followed by heat-curing at 60 °C for G


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ACS Applied Materials & Interfaces the “Avantage” software (Thermo Scientific). The durability of the polymer coating over long periods of time was tested by immersing the newly coated substrates in sterile 1X PBS and incubating the substrates at ambient temperatures for 7 days on a FMS3 shaking incubator (FinePCR, Gunpo, Korea) maintained at 30 rpm. The incubated substrates were washed with distilled water three times and air-dried ahead of the XPS survey scan. The XPS analysis was carried out under the conditions described above. Antiadhesion Effects of the Poly(DMA-mPEGMA-AA)Coated Polystyrene and PDMS Substrates against Bacteria. Each well of polystyrene-based 96-well tissue culture plate was polymer-coated in the presence of 100 μL of the polymer solution (20 mg/mL) at 4 °C overnight and washed to remove excess polymer solution. S. aureus (ATCC 25923) was grown in Luria broth (LB) media (Conda, Madrid, Spain) for 18 h at 37 °C and was normalized to 3.0 × 106 CFU/mL. Prior to use of the bacteria, S. aureus pellets were Gram-stained using a Gram-stain kit (Becton, Dickinson and Company, New Jersey, U.S.A.) to facilitate identification by color.28 The stained bacteria were distributed among each well (100 μL/well) and were incubated for 12 h at 37 °C. The wells were then washed with sterile 1X PBS three times, and the remaining cells were observed under a microscope at 400× magnification (Nikon, ECLIPSE TE300, Japan). For the antibacterial adhesion tests of PDMS, a silicon rubbercasted 96-well microtiter plate was prepared and the corresponding wells were coated with different concentrations (0, 1, 2, 5, 10, 20 mg/ mL) of poly(DMA-mPEGMA-AA) (100 μL/well), followed by incubation at 4 °C for 16 h. S. aureus was grown in fresh LB media and normalized to 3.0 × 106 CFU/mL. Twenty microliters of the bacterial suspension was incubated in the presence of the bare and polymer-coated silicone surfaces for 8 and 24 h, respectively, at 37 °C. After incubation, the wells were washed with sterile 1X PBS three times, and the WST-1 reagent (Roche, Mannheim, Germany) in sterile 1X PBS (1:10) was added (100 μL/well), followed by 2 h incubation.29 The O.D. values were measured using a microplate reader (Molecular devices, Sunnyvale, U.S.A.) at 440 nm, and the background intensity was determined at 690 nm. A silicone-cast black 24-well plate was prepared to carry out the antiadhesion test of the poly(DMA-mPEGMA-AA) against Gramnegative bacteria, E. coli. mCherry-E. coli kindly provided by Dr. Byung-Ha Oh (Department of Biological Sciences, Korea Advanced Institute of Science and Technology) were grown overnight in LB media and diluted to 3.0 × 106 CFU/mL. Five-hundred microliters of the mCherry-E. coli suspension were added to the polymer-coated wells. After incubating the plate for 24 h at 37 °C, the wells were washed with sterile 1X PBS three times and then air-dried. The surface-bound mCherry-labeled E. coli was visualized under a Xenogen IVIS Lumina 100 (PerkinElmer, Waltham, U.S.A.), and the radiance within a region of interest (ROI) was measured using the Live Imaging Software (Version 2.6). The relative image intensity was normalized relative to background signal. Long-Term Prevention of Bacterial Colonization on Poly(DMA-mPEGMA-AA)-Coated PDMS Substrates under Static Conditions. The bare and polymer-coated PDMS blocks were prepared separately. One-hundred microliters of S. aureus bacteria grown in LB media (3.0 × 106 CFU/mL) were pipetted onto the silicone elastomer surfaces and incubated at 37 °C for 7 days. Every 24 h, fresh LB media was added to prevent the bacteria from drying out.30 After incubation, the surfaces were washed with sterile 1X PBS three times. The samples were fixed with 4% formaldehyde (w/v) (SigmaAldrich, St. Louis, U.S.A.) for 2 h, and the biofilm was dehydrated by successively immersing them in solutions containing increasing ethanol concentrations (25%, 50%, 75%, 95%, and 100%) for 10 min each. The samples were air-dried and platinum-coated (40 s) prior to conducting field-emission scanning electron microscopy (FE-SEM, Magellan 400, Hillsboro, U.S.A.) measurements. Antibiofilm Effect of the Poly(DMA-mPEGMA-AA)-Coated Urinary Catheter under Flow Conditions. Tygon experimental silicone tubing (Hanmi Rubber & Plastics, Seoul, Korea) and a medical silicone urinary catheter (SEWOON MEDICAL, Cheonan,

Korea) were sterilized (121 °C, 15 psi, 30 min) and dried overnight at 60 °C. The silicone urinary catheter was cut in half (15 cm each), and only one piece was coated with the polymer layer. After polymer coating, the bare and polymer-coated catheters were attached to the Tygon silicone tube outlets. A colony of S. aureus was cultured in 15 mL sterile LB media at 37 °C for 16 h in a shaking incubator. Overnight cultured S. aureus was diluted 100-fold into 500 mL fresh LB media and was stirred continuously to avoid bacterial sediment. The Tygon silicone tube was mounted on a peristaltic pump (EYELA, Tokyo, Japan) that maintained a flow rate of 2 mL/min for 48 h.31 Following a 48-h incubation under flow conditions, each catheter (the bare or polymer-coated catheter) was cut equally to normalize the catheter inner gross area. Catheters were dipped into the distilled water three times each to remove unbound bacteria. The washed catheters were immersed in a 0.1% crystal violet (CV) solution (Becton, Dickinson, and Company, New Jersey, U.S.A.) for 15 min. The stained biofilm was washed with distilled water three times to eliminate excess stain. The catheters were left to air-dry prior to conducting further assays. Once dried, the catheters were filled with decolorizer (Becton, Dickinson and Company, New Jersey, U.S.A.) for 15 min. Two-hundred microliters of the decolorized CV dye from the biofilm were transferred into a 96-well plate, and the O.D. was measured at 580 nm.31,32 Evaluation of the Biocompatibility of the Poly(DMAmPEGMA-AA)-Coated PDMS. Hemolytic activity of the poly(DMA-mPEGMA-AA) was measured using a blood agar plate (Komed, Seongnam, Korea). A piece of bare or polymer-coated PDMS was placed at a specified region on the agar plate The blood agar plate was incubated at 37 °C for 16 h to observe signs of hemolysis. For the antiadhesion effect of the polymer-coated PDMS against circulating blood cells, 10 mL fresh rat blood were collected using a heparin-coated syringe (Orient bio, Seongnam, Korea) to avoid coagulation.33 The blood was centrifuged at 1000 rpm (210 g value) for 10 min, and the supernatant was carefully collected to avoid disturbing the RBC layer. The gathered plasma was incubated in an RBC lysis buffer (BioLegend, San Diego, U.S.A.) at 4 °C for 5 min and then centrifuged at 3000 rpm (1910 g value) for 10 min. After centrifugation, pellets were thoroughly mixed with the lower third of the supernatant. The combined blood plasma with blood circulating cells (pellet) was pipetted onto the uncoated and polymer-coated PDMS silicone rubber surfaces and incubated at 37 °C for 0.5 h. After incubation, the samples were washed three times with sterilized 1X PBS, followed by dehydration of the surface-bound cells by serially immersing the substrates into solutions containing increasing concentrations of ethanol (25%, 50%, 75%, 95%, and 100%) for 10 min each.30 The dehydrated samples were air-dried and platinumcoated for 40 s prior to SEM imaging. In Vivo Experiments. Balb/c mice (Orient bio, Seongnam, Korea) were shaved before the experiments. One centimeter-long urinary catheter pieces were autoclaved and polymer-coated. The mice were anesthetized, and the catheter pieces were subcutaneously inserted (bare or polymer-coated) into the right flanks of the mice. The mice were given 2 days to heal at the incision site. Subsequently, sterile 1X PBS (100 μL) or S. aureus (5.0 × 106 CFU in 100 μL 1X PBS) were subcutaneously inoculated into the catheter-harboring site using an insulin syringe (SUNGSHIM MEDICAL, Kyounggi-do, Korea).34 After PBS or bacteria inoculation, the mice were observed over a 4-day period. On day 4, the mice were humanely euthanized, and the inserted catheters were harvested. The collected urinary catheter pieces were analyzed using a CV biofilm assay. The optical densities of the decolorized CV stain were measured at 580 nm using a microplate reader.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsami.7b06899. H


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(12) Bayston, R.; Fisher, L. E.; Weber, K. An Antimicrobial Modified Silicone Peritoneal Catheter with Activity against Both Gram Positive and Gram Negative Bacteria. Biomaterials 2009, 30 (18), 3167−3173. (13) Stevens, K. N. J.; Crespo-Biel, O.; van den Bosch, E. E. M.; Dias, A. A.; Knetsch, M. L. W.; Aldenhoff, Y. B. J.; van der Veen, F. H.; Maessen, J. G.; Stobberingh, E. E.; Koole, L. H. The Relationship between the Antimicrobial Effect of Catheter Coatings Containing Silver Nanoparticles and the Coagulation of Contacting Blood. Biomaterials 2009, 30 (22), 3682−3690. (14) Dave, R. N.; Joshi, H. M.; Venugopalan, V. P. Novel Biocatalytic Polymer-Based Antimicrobial Coatings as Potential Ureteral Biomaterial: Preparation and in Vitro Performance Evaluation. Antimicrob. Agents Chemother. 2011, 55 (2), 845−853. (15) Tran, P. A.; Webster, T. J. Antimicrobial Selenium Nanoparticle Coatings on Polymeric Medical Devices. Nanotechnology 2013, 24 (15), 155101. (16) Arias, C. A.; Murray, B. E. Antibiotic-Resistant Bugs in the 21st Century – a Clinical Super-Challenge. N. Engl. J. Med. 2009, 360 (5), 439−443. (17) Gutierrez-Gonzalez, R.; Boto, G. R. Do Antibiotic-Impregnated Catheters Prevent Infection in Csf Diversion Procedures? Review of the Literature. J. Infect. 2010, 61 (1), 9−20. (18) Huh, A. J.; Kwon, Y. J. ″Nanoantibiotics″: A New Paradigm for Treating Infectious Diseases Using Nanomaterials in the Antibiotics Resistant Era. J. Controlled Release 2011, 156 (2), 128−145. (19) Sung, D.; Park, S.; Jon, S. Facile Method for Selective Immobilization of Biomolecules on Plastic Surfaces. Langmuir 2009, 25 (19), 11289−11294. (20) Park, S. J.; Lee, K. B.; Choi, I. S.; Langer, R.; Jon, S. Y. Dual Functional, Polymeric Self-Assembled Monolayers as a Facile Platform for Construction of Patterns of Biomolecules. Langmuir 2007, 23 (22), 10902−10905. (21) Park, S.; Yang, H. S.; Kim, D.; Jo, K.; Jon, S. Rational Design of Amphiphilic Polymers to Make Carbon Nanotubes Water-Dispersible, Anti-Biofouling, and Functionalizable. Chem. Commun. (Cambridge, U. K.) 2008, 25, 2876−2878. (22) Park, J.; Yu, M. K.; Jeong, Y. Y.; Kim, J. W.; Lee, K.; Phan, V. N.; Jon, S. Antibiofouling Amphiphilic Polymer-Coated Superparamagnetic Iron Oxide Nanoparticles: Synthesis, Characterization, and Use in Cancer Imaging in Vivo. J. Mater. Chem. 2009, 19 (35), 6412−6417. (23) Larina, I. V.; Shen, W.; Kelly, O. G.; Hadjantonakis, A. K.; Baron, M. H.; Dickinson, M. E. A Membrane Associated Mcherry Fluorescent Reporter Line for Studying Vascular Remodeling and Cardiac Function During Murine Embryonic Development. Anat. Rec. 2009, 292 (3), 333−341. (24) Sovadinova, I.; Palermo, E. F.; Huang, R.; Thoma, L. M.; Kuroda, K. Mechanism of Polymer-Induced Hemolysis: Nanosized Pore Formation and Osmotic Lysis. Biomacromolecules 2011, 12 (1), 260−268. (25) Hook, A. L.; Chang, C. Y.; Yang, J.; Luckett, J.; Cockayne, A.; Atkinson, S.; Mei, Y.; Bayston, R.; Irvine, D. J.; Langer, R.; Anderson, D. G.; Williams, P.; Davies, M. C.; Alexander, M. R. Combinatorial Discovery of Polymers Resistant to Bacterial Attachment. Nat. Biotechnol. 2012, 30 (9), 868−U99. (26) Ding, X.; Yang, C.; Lim, T. P.; Hsu, L. Y.; Engler, A. C.; Hedrick, J. L.; Yang, Y. Y. Antibacterial and Antifouling Catheter Coatings Using Surface Grafted Peg-B-Cationic Polycarbonate Diblock Copolymers. Biomaterials 2012, 33 (28), 6593−6603. (27) Voo, Z. X.; Khan, M.; Narayanan, K.; Seah, D.; Hedrick, J. L.; Yang, Y. Y. Antimicrobial/Antifouling Polycarbonate Coatings: Role of Block Copolymer Architecture. Macromolecules 2015, 48 (4), 1055− 1064. (28) Beveridge, T. J. Use of the Gram Stain in Microbiology. Biotech. Histochem. 2001, 76 (3), 111−118. (29) Ngamwongsatit, P.; Banada, P. P.; Panbangred, W.; Bhunia, A. K. Wst-1-Based Cell Cytotoxicity Assay as a Substitute for Mtt-Based Assay for Rapid Detection of Toxigenic Bacillus Species Using Cho Cell Line. J. Microbiol. Methods 2008, 73 (3), 211−215.

H NMR spectra of the poly(DMA-mPEGMA-AA) (Figure S1) (PDF)

AUTHOR INFORMATION

Corresponding Author

*Tel: (+82)-42-350-2634. Fax: (+82)-42-350-4450. E-mail: [email protected] (S.J.). ORCID

Sung Gap Im: 0000-0001-7562-2929 Sangyong Jon: 0000-0002-6971-586X Author Contributions

The manuscript was written with contributions from all authors. S.J. conceived the project; S.J. and H.K. wrote the paper; H.K. performed most of experiments; S.J., H.K., and S.G.I. analyzed data; J.Y.K., B.Y., S.J.Y., J.K., D.Y.L., and H.J. carried out certain experiments. All authors have given approval to the final version of the manuscript. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was supported by Samsung Research Funding Center of Samsung Electronics under Project Number SRFCMA1501-01.

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