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Non-toxic Cationic Coumarin Polyester Coatings Prevent Pseudomonas aeruginosa Biofilm Formation Elaheh A. Chamsaz, Steven Mankoci, Hazel A. Barton, and Abraham Joy ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.6b12610 • Publication Date (Web): 02 Feb 2017 Downloaded from http://pubs.acs.org on February 3, 2017

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ACS Applied Materials & Interfaces

Non-toxic Cationic Coumarin Polyester Coatings Prevent Pseudomonas aeruginosa Biofilm Formation Elaheh A. Chamsaz1, Steven Mankoci1, Hazel A. Barton2*, and Abraham Joy1* 1

Department of Polymer Science, The University of Akron, Akron, Ohio, USA 44325

2

Department of Biology, The University of Akron, Akron, Ohio, USA 44325

Corresponding authors *Email: [email protected], [email protected] Keywords: Antimicrobial polyester, antimicrobial coating, cationic polymer coating, Pseudomonas aeruginosa biofilm, non-toxic antimicrobial polymer, functionalized polyester Abstract: The rapid increase in bacterial infections and antimicrobial resistance is a growing public health concern. Infections arising from bacterial contamination of surgical tools, medical implants, catheters and hospital surfaces can potentially be addressed by antimicrobial polymeric coatings. The challenge in developing such polymers for in vivo use is the ability to achieve high antimicrobial efficacy while at the same time being non-toxic to human cells. Although several classes of antimicrobial polymers have been developed, many of them cannot be used in the clinical setting due to their non-selective toxicity towards bacteria and mammalian cells. Here we demonstrate that coumarin polyesters with cationic pendant groups are very effective against Gram negative Pseudomonas aeruginosa. Coumarin polyesters with pendant cationic amine groups were coated onto glass coverslips and tested for their antimicrobial activity against P. aeruginosa colonization of the surface. The results demonstrate that the cationic coumarin polyester kills the surface attached bacterial cells preventing biofilm formation but does not show any hemolytic activity or discernible toxicity towards mammalian cells. The antimicrobial polyesters described in this work have several advantages desired in antimicrobial coatings such

ACS Paragon Plus Environment


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as high antimicrobial activity, low toxicity towards mammalian cells, visualization and ease of synthesis and fabrication, all of which are necessary for translation to the clinic. 1. Introduction: Hospital acquired (nosocomial) infections are an emergent healthcare issue where routine hospital procedures can expose patients to antibiotic resistant infections.1-3 Annually there are about 1.7 million nosocomial infections in the US, of which about 100,000 are fatal, with medical costs in the range of $30 billion annually.4-6 These problems are further compounded by an alarming rise in antibiotic resistant microorganisms coupled with a decrease in the number of new antibiotics being approved.7-9 Polymeric antimicrobial coatings play a critical role in the prevention of microbial contamination in healthcare, food packaging and textile applications.10-12 Several types of polymers have been designed to control microbial infections, classified either as carriers of antimicrobial agents, or containing structural components that confer antimicrobial activity.11 Polymeric coatings that leach antimicrobial agents, such as silver, are widely used in clothing and in medical implants,13 however their effectiveness decreases over time due to the finite amount of antimicrobial agent. Polymeric coatings that are inherently antimicrobial are therefore advantageous as their efficacy results from the composition of the polymer itself, which can be designed to include chemical structures with known antimicrobial activity.5 Surface grafted or surface coated polymeric antimicrobials include polyvinylpyridines alkylated with long aliphatic chains or alkylated polyethylenimines.14-16 It is proposed that the presence of both the cationic charge and the hydrophobic chain account for the antimicrobial action of such coatings.17, 18 N-Halamine containing coatings exhibit antimicrobial activity due to the strong oxidative potential of such halogen containing polymers.19 Polydopamine and


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polysaccharide based antimicrobial coatings have also been reported.20,

21

Dip-coating and

polyelectrolyte multilayer assemblies are further methods for fabrication of novel antimicrobial coatings.22-24 Chitosan derivatives are widely used in several antimicrobial applications and the presence of the cationic amine functionality confers antimicrobial activity.12,

25

The potential

hemolytic activity of polymers with aromatic cationic residues and structural design of NHalamine type polymers limits their in-vivo use.26,

27

In light of the increasing incidence of

nosocomial infections, there is a heightened need for antimicrobial polymeric coatings for surgical tools, implantable devices and hospital surfaces and gowns.28 Such antimicrobial coatings need to have high molecular weights to fabricate uniform and stable coatings, show minimal toxicity to mammalian cells, demonstrate consistent antimicrobial activity throughout its lifetime, and very importantly – have manufacturing reproducibility and scale-up feasibility. Although several such coatings have been developed in the past, there is a strong need for antimicrobial polymers that show minimal in vivo toxicity to mammalian cells. The need is greater for antimicrobial polymers effective against Gram negative bacteria due to the smaller arsenal of small molecule antibiotics or polymeric antimicrobials toward this class of microbes.29, 30 Gram negative bacteria, which do not retain the crystal violet stain, have a very different cell envelop from the Gram positive bacteria. Gram negative bacteria have an inner membrane, a thin peptidoglycan cell wall and an outer membrane, which is negatively charged due to the presence of lipopolysaccharides.31 Pseudomonas aeruginosa is an ESKAPE (Enterococcus faecium, Staphylococcus aureus, Klebsiella pneumoniae, Acinetobacter baumanii, Pseudomonas aeruginosa, and Enterobacter species) pathogen, which can cause serious hospital acquired infections, including ventilator associated pneumonia, catheter associated sepsis and treatment resistant wound and burn



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infections.32 In the current work, we describe the development of a cationic amine polyester coating with very high efficacy against P. aeruginosa and minimal toxicity against mammalian cells. The polyester contains cationic pendant groups and aromatic coumarin units in the backbone to enable visualization of the coating and to decrease delamination of the film. We infer that the antimicrobial activity arises from the cationic pendant groups, which are structurally similar to lysine, and not from any small molecules or oligomers leaching from the polymer. The key attributes of the described work are its very high activity against Gram negative P. aeruginosa, resulting in the loss of biofilm formation and the lack of any observable toxicity to mammalian cells. Polyesters were chosen as the platform for this work as they are more hydrophilic than chain growth polymers and their degradable nature makes them suitable for applications where a temporary antimicrobial environment is beneficial. 2. Materials and characterizations: 2.1. Materials. All the solvents, tert-Butyldimethylchlorosilane, di-tert-butyl dicarbonate, casamino acid, live/dead BacLight Bacterial Viability Kit and LIVE/DEAD BacLight mammalian Vitality Kit were purchased from Thermo Fisher Scientific (Waltham, MA) and were used as received. Acetone was distilled prior to use. Dichloromethane was dried by distilling over anhydrous calcium hydride (CaH2). Sodium bisulfate (NaHSO4), potassium carbonate (K2CO3), 18-crown-6, p-toluenesulfonic acid (PTSA; CH3C6H4SO3H), 4 Å molecular sieves, 3-bromo-1-propanol, trimethylamine, and succinic acid were purchased from Acros Organics (Morris Plains, New Jersey). Ethyl 4-chloroacetoacetate and hydroquinone bis(2hydroxyethyl)

ether

was

purchased

from

Alfa

Aesar

(Haverhill,

MA),

and

diisopropylcarbodiimide (DIC) was purchased from Oakwood Chemicals (Estill, SC). 4(Dimethylamino)pyridinium-4-toluenesulfonate (DPTS) was prepared according to literature


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methods.33 4N HCl solution in 1,4-dioxane, DABCO, triton X-100, agar, Mueller-Hinton broth, calcium chloride (CaCl2), Fe-EDTA, ammonium sulfate ((NH4)SO4), disodium hydrogen phosphate (Na2HPO4), monopotassium phosphate (KH2PO4), sodium chloride (NaCl), magnesium chloride (MgCl2), sodium citrate, Sodium thiosulfate (Na2S2O3) and 6aminohexanoic

acid

were

purchased

from

Sigma

Aldrich

(St.

Louis,

MO).

Dimethylaminopyridine, t-butyl alcohol, dimethylaminopyridine, diethanolamine, thionyl chloride, N-Hydroxysuccinimide, iodine and sheep blood were purchased from VWR (Radnor, PA). Silica gel (40−63 µm, 230 × 400 mesh) for column chromatography was purchased from Sorbent Technologies, Inc (Norcross, GA). 2.2. Characterization. 1H Nuclear magnetic resonance (NMR) spectra were recorded on a Varian Mercury 300 MHz spectrometer spectrophotometer (Palo Alto, CA, USA). Chemical shifts are reported in ppm (δ) relative to residual solvent signals. Size exclusion chromatography (SEC) was performed on a TOSOH EcoSec HLC-8320, with two PSS Gram Analytical SEC Columns in series, using 25 mM LiBr in DMF as eluent at a flow rate of 0.8 mL/min. The column and detector temperatures were maintained at 50°C. Molecular weights were obtained relative to PMMA standards using the refractive index (RI) signal. Thermal transitions were analyzed using a differential scanning calorimeter (DSC) TA Q2000 with a liquid N2 cooling unit using a cooling cycle of 10°C/min and heating rate also at 10°C/min. TA Q500 thermogravimetric analyzer (TGA) was used to collect 5% decomposition temperature data from 25°C to 600°C at a heating rate of 10°C/min in a N2 atmosphere. Coating thickness was measured on an UV−visible−NIR (240−1200 nm) variable angle spectroscopic ellipsometer (VASE M-2000, J.A. Woollam Co.). Atomic force microscopy (AFM) images were obtained on DI MultiMode SPM AFM (Tapping mode). Imaging was done on an IX51 Epifluorescence



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Microscope (Olympus Co., Japan) using the DAPI, FITC and TRITC filters. Crosslinking of the polymer was done in a Rayonet® RMR-200 reactor at 350 nm wavelength. Contact angles were measured on Ramé–Hart Instruments Advanced Goniometer 500 F1 and analyzed with Drop Image Advanced software. 3. Experimental: 3.1. Synthesis of coumarin diol monomer: The coumarin diol was synthesized as previously reported in Scheme S1 (Supporting Information).34 In brief, resorcinol (10g, 91 mmol) was reacted with 4-chloromethyl acetoacetate (17 g, 103 mmol) in the presence of p-toluenesulfonic acid (3.6 g, 19 mmol) in toluene (150 mL) and refluxed at 110°C for 45 minutes. The reaction mixture was purified by extraction in water and ethyl acetate and dried under vacuum. Further purification was done via silica gel liquid column chromatography (ethyl acetate: hexane1:9) to yield 7-hydroxy-4-(chloromethyl)coumarin (1) (12 g, 65% yield). Then, compound (1) (2.95 g, 14 mmol) was refluxed over a stirring solution of water (350 mL) for 3 days to hydrolyze the chloromethyl into a hydroxymethyl group. The reaction mixture was filtered while hot, and cooled to room temperature over 12 h to yield 7-hydroxy-4-(hydroxymethyl)coumarin (2) (2.7 g). The product was used further without purification. The final 7-(hydroxypropoxy)-4(hydroxymethyl)coumarin (3) was synthesized by combining (2) (1.0 g, 5.2 mmol) with potassium carbonate (2 g, 14.5 mmol), 18-crown-6 (0.7 g, 2.65 mmol), and 3-bromo-1-propanol (1.5 g, 10.8 mmol) and dissolving in dry acetone (15 mL) and refluxing at 55°C for 2.45 hours in a CEM Discover microwave reactor. The reaction mixture was filtered and solvent was removed under reduced pressure and purified via silica gel liquid column chromatography using a mixture of ethyl acetate and hexane as the mobile phase. The product was dried under reduced pressure to yield pure (3) (1.18 g, 91%). The coumarin monomers were characterized by 1

HNMR.


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3.2. Synthesis of t-butyl protected carboxylic acid monomer (6): The pendant t-butyl protected carboxylate diol monomer was synthesized as previously reported in Scheme S2 (Supporting Information).35 In brief, succinic anhydride (10 g, 100 mmol) was reacted with anhydrous t-butyl alcohol (17 mL) in anhydrous toluene (150 mL) in the presence of dimethylaminopyridine (1.8 g, 15 mmol), N-hydroxysuccinamide (3.45 g, 30 mmol), and trimethylamine (4.2 mL, 30 mmol) to yield the ring opened mono-tert-butylsuccinate product (4). The product was purified by ethyl acetate and water extractions followed by recrystallization of the isolated products of the organic phase in 1:3 ether:hexanes (77%). The pure compound (4) (5.57 g, 31.9 mmol) was then reacted with tertButyldimethylchlorosilane (TBDMS) protected diethanolamine (24.6 mmol) in a 1-ethyl3-(3-dimethylaminopropyl) carbodiimide (EDC) (6.60 g, 34.4 mmol) mediated coupling reaction in anhydrous N,N-dimethylformamide (DMF) (30 mL) overnight. The resulting product (5) was purified by removing the DMF via rotovap, performing ethyl acetate and water extractions, and further purifying the product by performing silica gel liquid column chromatography using ethyl acetate and hexanes as the mobile phase (69.8%). Deprotection of the TBDMS groups was performed by reacting the purified product from the previous step with iodine in methanol (2% w/v) at reflux for three hours. After reaction, Na2S2O3 was added until the reaction mixture became clear and was concentrated on rotary evaporator. To purify the t-butyl protected carboxylate diol monomer (5), methylene chloride and water extractions were performed followed by silica gel liquid column chromatography using methylene chloride and methanol as the mobile phase to purify the products extracted into the organic phase. The product was



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dried under reduced pressure to yield pure monomer (6) (61%). Synthesis of the monomer was confirmed by 1HNMR. 3.3. Synthesis of Boc-protected amine monomer (9): The pendant Boc-protected amine monomer was synthesized as previously reported in Scheme S3 (Supporting Information)35. In brief, 6-aminohexanoic acid (10 g, 76.2 mmol) was esterified into its corresponding methyl ester by dissolving it in a large excess of anhydrous methanol (125 mL) and adding thionyl chloride (22.9 g, 193 mmol) dropwise. After reacting overnight and removing the excess methanol via rotary evaporator, the methyl-6-aminohexanoate (7) was dissolved in 50 mL of a 50/50 (by volume) mixture of water and dioxane and triethylamine (19.2 g, 190 mmol) was added to make the solution alkaline. Then, di-tertbutyl dicarbonate (20.5 g, 91 mmol) dissolved in 50 mL of 50/50 water and dioxane was added dropwise to the solution of methyl-6-aminohexanoate. The resulting methyl-6(amino-N-tert-butoxycarbonyl)hexanoate (8) was purified via ethyl acetate and water extractions and dried under vacuum (17.8 g, 95.5%). Compound (8) was reacted with diethanolamine (15.9 g, 151 mmol) neat at 80°C under vacuum to yield the desired Bocprotected amine diol monomer (9). The monomer was purified via silica gel liquid column chromatography using a mixture of methylene chloride and methanol as the mobile phase. The product was dried under reduced pressure to yield pure monomer (9) (17.5g, 75.7%). Synthesis of the monomer was confirmed with 1HNMR. 3.4. Synthesis of coumarin polyesters: The synthesis of the coumarin polyesters described here are based on our earlier published method.36 In brief, the coumarin polyesters with protected carboxylic acid (C-CO2-P) or amine (C-NH2-P) functional groups were synthesized by the room temperature carbodiimide-mediated polymerization of diol and diacid and shown in Scheme 1.


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Coumarin diol (3) (0.5g, 1.99 mmol, 0.75 equiv), pendant t-butyl protected carboxylate diol monomer (6) (0.17 g, 0.06 mmol, 0.25 equiv) or pendant Boc-protected amine diol monomer (9) (0.21 g, 0.06 mmol, 0.25 equiv) , and succinic acid (0.31 g, 2.66 mmol, 1 equiv) were taken in an evacuated and N2 filled round bottomed flask along with 4-(N,N-dimethylamino) pyridinium-4toluenesulfonate (DPTS)(0.31 g, 1.06 mmol, 0.25 equiv). Dichloromethane (1 mL for every 2 mmol diacid) was then added and stirred until monomer dissolved in the solvent. Then the solution was cooled over ice bath to 0°C and N,N-diisopropylcarbodiimide (DIC)(1.25 mL, 3 equiv) was added dropwise via syringe. The reaction was allowed to come to room temperature and stirred for 48 h under nitrogen. The polymer was precipitated in methanol twice and dried under vacuum to yield a solid (85%). The polymers were characterized by 1HNMR, SEC, TGA and DSC. 3.5. Deprotection of coumarin polyesters: The protected carboxylic acid (C-CO2-P) (0.35g) and/or protected amine (C-NH2-P) (0.35 g) polyesters were dissolved in 1.5 mL of CH2Cl2 in a round bottomed flask equipped with stir bar. This solution was cooled to 0°C and 0.7 mL of 4N HCl in 1,4-dioxane was added and stirred for 45 minutes at room temperature (Scheme 1). Then, HCl/Dioxane was removed from reaction mixture under reduced pressure and the polymer was dissolved in 2 mL CH2Cl2 and precipitated from diethyl ether and dried. Anionic carboxylic acid (C-CO2-) and cationic amine (C-NH3+) polyesters were characterized by 1H NMR, TGA and DSC. 3.6. Coating preparation: 3.6.1. Spin coating of polymers on glass coverslips: Circular coverslips (12 mm), thickness number 1 were cleaned by sonicating in methanol for 15 minutes and acetone for another 15 minutes. Each coverslip was dried under argon and visually



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inspected to ensure they were free of any debris or imperfections after solvent washing. Right before spin coating, the coverslips were etched in plasma cleaner for 10 mins. Spin coating was performed by preparing 2% (w/v) solutions of polymers in chloroform. For the cationic amine polyester, spin coating was performed using 2% (w/v) polymer solution in a 9:1 CHCl3: DMF mixture. The polymer solution was spin coated on the coverslips at 2000 rpm for 60 seconds and dried and annealed at 65°C in a vacuum oven. 3.6.2. Spin coating of polymers on silicon wafer: Silicon wafers were cleaned with piranha solution (H2SO4: H2O2 - 7:3 (v/v)) at 90°C for 45 minutes, rinsed with water and dried by argon prior to spin coating. Polymer solution was spin coated on silicon wafer in the same manner as with the glass coverslips. 3.7. Crosslinking of polymeric coating: Coated coverslip containing cationic amine polyester was placed on a Petri dish and irradiated from top at 350 nm for 10 minutes in a Rayonet® RMR-200 reactor. 3.8. Surface roughness and thickness measurement: The spin coated polymer on silicon wafer was used for roughness and thickness measurements. Surface roughness was measured by AFM and surface thickness was measured by ellipsometry. 3.9. Contact angle measurement: Circular coverslips (12 mm) were cleaned and spin coated as indicated above. The static contact angle of Milli-Q water was measured on a Ramé–Hart Instruments Advanced Goniometer 500 F1 and analyzed with Drop Image Advanced software. 10 µL of Milli-Q water was deposited on the surface and immediately the contact angles were measured over a period of 120 sec at 25°C. Three measurements were taken for each sample. 3.10. Coating stability measurement: The cationic amine polyester coated coverslips (noncrosslinked and crosslinked) were incubated at 37°C in phosphate-buffered saline (PBS), pH 7.4,


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for a period of 1 month. The surface of each spin coated sample was analyzed before and after incubation using a IX51 Epifluorescence Microscope (Olympus Co., Japan) under 10X magnification with DAPI filter. 3.11. Microbiology: The strain Pseudomonas aeruginosa PAO1 containing a stably integrated, GFP gene under a constitutive promoter (pTdK-GFP) was used for the colonization/biofilm assays based on the method of De Kievit et al.37 Briefly, an overnight culture of P. aeruginosa grown in TSB was adjusted to an OD600 of 1.0, and diluted 1:1,000 in FAB medium (0.1 mM CaCl2, 0.01 mM Fe-EDTA, 0.15 mM (NH4)SO4, 0.33 mM Na2HPO4, 0.2 mM KH2PO4, 0.5 mM NaCl, 0.5% (wt/vol) casamino acids, 1 mM MgCl2, 10 mM sodium citrate) for the working bacterial solution. FAB medium was used for this experiment to induce biofilm formation. Each polymer coated coverslip was sterilized with a 70% ethanol solution and drying in a sterile 24well culture plate. Negative controls demonstrated that this method of sterilization did not inhibit downstream assays (data not shown). Sterile, polymer coated coverslips were then inoculated with 2 mL of bacterial solution and incubated at 37°C for 5 hours, rinsed gently in FAB media (3X) to remove non-adhered bacteria, imaged on a IX51 Epifluorescence Microscope (Olympus Co., Japan), under 10X magnification with 1 µL of 20 mM propidium iodide used as an indicator of cell death. In order to determine both the number of attached cells and cell viability, cell coverage was calculated using Image J software on five random fields-of-view. 3.12. Bactericidal plating assay: The bactericidal activities of polymeric coatings were evaluated with overnight broth culture of bacteria adjusted to yield approximately 1×109 CFU/ml. The polymer coated coverslips were placed into a 12 well plate and 2 mL of bacteria broth was added to each well plate and kept in the incubator at 37°C for 24 hours. Then, 1 mL of bacteria solution was taken from each well plate and serially diluted in sterile TSB (pH 7.4) and



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spread onto TSA plates and were incubated at 37°C to determine the number of colony forming units (CFUs). All determinations were performed in triplicate including the growth controls. 3.13. Disk diffusion assay: 100 µL of overnight broth cultures adjusted to yield approximately 1×109 CFU/ml was spread onto TSA plates. The polymer coated coverslips were placed on the inoculated agar surface and incubated at 37°C 24 h. All tests were performed in triplicate and the antibacterial activity was expressed as the mean of inhibition diameters (mm) produced by polymeric coatings 3.14. Hemolysis assay: All assays were performed using defibrinated sheep blood washed in 6-8 rinses of 150 mM NaCl, phosphate buffered saline (PBS; pH 7.4) by centrifugation at 500 x g before diluting 1:50 in PBS for the red blood cell (RBC) working solution. The hemolysis assay was carried out by placing polymer coated coverslips into 12 well plates and inoculation with 2 mL of RBC solution at 37°C for 1 hr. To measure cell lysis via hemoglobin release, the RBC solution was centrifuged at 500 x g for 10 minutes with absorbance measured at 450 nm, and compared with a non-polymer (0%) and 1% Triton X-100 (100%) control. All determinations were performed in triplicate. 3.15. In vitro cytotoxicity: NIH-3T3 mouse embryonic fibroblast cells, were grown on sterilized glass coverslips coated with the polymers or a glass-only control at ~5,000 cells/cm2. Cells were grown in DMEM/10% fetal bovine serum (Hyclone) and 1% penicillin-streptomycin (10,000 U/mL, Thermo Fisher Scientific) at 37°C in 5% CO2. LIVE/DEAD staining for mammalian cells were performed on days 1 and 3 after seeding the cells onto the surface, using Calcein AM and ethidium homodimer-1 stains. 250 µL each of 2 µM calcein AM and 4 µM ethidium homodimer1 were added to each well. After incubating the cells for 10-15 minutes, the well plate was


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removed and the cells were imaged on the IX51 Epifluorescence Microscope. All determinations were performed in triplicate. 4. Result and Discussion 4.1. Synthesis and characterization of coumarin polyesters. In this work, a series of functionalized coumarin polyester coatings were developed to test the effect of either carboxylic acid or amine pendant groups to prevent P. aeruginosa colonization on surfaces. The composition of the polyesters were chosen to optimize antimicrobial activity and stability of the polymer film over the testing period. The polyesters are designed with pendant groups (25 mole%) that mimic the side chains of aspartic acid (anionic) or lysine (cationic). The aromatic coumarin unit (75 mole%) in the polyester enables visualization of the coating and decreases delamination of the film. In addition, the fidelity of the polymer coating can be monitored by the fluorescence of the coumarin units. Photoresponsive coumarin diol (3), protected carboxylic acid or amine monomers (6, 9) and

succinic

acid

polyesterification.34,

were 35

polymerized

by

room

temperature

carbodiimide-mediated

The resultant protected carboxylic acid polyester (C-CO2-P) and

protected amine polyester (C-NH2-P), were deprotected to provide anionic carboxylic acid (CCO2-) and cationic amine (C-NH3+) polyesters (Scheme 1). These polymers were characterized by 1H NMR (Supporting Information Figure S1-S4) and size exclusion chromatography (SEC) (Table 1). The number average molecular weight of the t-butyl protected carboxylic acid polyester (Mn = 24 kDa) and Boc-protected amine polyester (Mn = 28 kDa) are relatively similar allowing comparisons between the four polymers used in this study (Supporting Information Figure S5, and S6). As shown in Table 1, the glass transition temperature (Tg) for all polyesters range from 46.5°C to 55.1°C indicating that the different functional groups do not cause large differences in thermal properties of the polymers.



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Scheme 1. Synthetic Route for the Synthesis of Coumarin Polyesters

Reagents and conditions: a) DPTS/DIC/CH2Cl2/ Room temperature, 48h b) HCl/Dioxane/CH2Cl2, 45 minutes.

Table 1. Characterization of Coumarin Polyesters Mna Mwa Tg b Polyester Ða (°C) (kDa) (kDa) 27.9 44.2 1.58 55.1 C-CO2-P 51.5 C-CO2 24.4 38.7 1.58 49.8 C-NH2-P + 46.5 C-NH3

Thicknessc Roughnessd (nm) (nm) 219.9 5.32 112.4 5.8 198.1 4.89 113.3 5.21

a

Determined by DMF SEC relative to PMMA standards. Ð = polydispersity index bDetermined by DSC. cDetermined by Ellipsometer. dDetermined by AFM.

4.2. Polymer coating fabrication and characterization. The polymers were spin coated onto a glass coverslips and subsequently dried and annealed to provide homogenous thin coatings. The thickness of coatings was measured by ellipsometry and their surface roughness was evaluated by AFM. The thickness of the protected coatings (C-CO2-P and C-NH2-P) are in the range of 200 µm, while deprotected coatings (C-CO2- and C-NH3+) are in the range of 100 µm (Table 1). In all cases the surface roughness, which indicates the deviation from uniform film thickness across the film surface, was measured between 4.8-5.8 nm, which is significantly less than the


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size of P. aeruginosa (0.5-0.8 µm x 1.5-3.0 µm) and hence will not a significant factor in bacterial attachment on these films.38-40 The stability of the coatings was evaluated by incubation at 37°C in phosphate buffered saline (PBS), pH 7.4, for a period of a month. As shown by the fluorescence of the coumarin moieties (Supporting Information Figure S7), both the non-crosslinked and crosslinked cationic amine coatings (C-NH3+) were not substantially different from each other over this time period. 4.3. Inhibition of P. aeruginosa colonization on polymer coated surfaces. To determine the effect of chemical functionality on bacterial colonization on the surfaces, P. aeruginosa was plated onto the polymer coatings in static culture. After 5 h, bacterial colonization and viability were quantified by live/dead fluorescent microscopy. As shown in (Figure 1), there is a strong influence of cationic or anionic surface charge on the ability of P. aeruginosa to form biofilms and cell viability. The protected carboxylic acid polyester (C-CO2-P) and the corresponding anionic carboxylic acid polyester (C-CO2-) do not show a significant difference in the survival of P. aeruginosa; however, there were significantly less bacterial colonies on the anionic carboxylic acid surface compared to the protected carboxylic acid surface. Live/dead microscopy indicated that most of the attached bacteria were viable and retain the ability to form biofilm. P. aeruginosa is able to colonize the protected amine polyester (C-NH2-P) surface and retains the ability to form biofilm, but more dead bacteria were observed on this surface relative to the carboxylic acid surfaces (Figure 1). In contrast to all the other surfaces, the cationic amine polyester (C-NH3+) shows a remarkable ability to kill the attached P. aeruginosa cells, resulting in the absence of biofilm formation. Live/dead microscopy showed that there were no live bacteria and hence biofilm formation is not seen. The plating density was maintained at a high level so as to provide the bacteria maximum



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ability to survive under the experimental conditions. In order to test the influence of crosslinked coatings on bacterial attachment and biofilm formation, the cationic amine polyester (C-NH3+) was crosslinked for 10 min, upon which the coumarin units undergo a [2+2]-photocycloaddition reaction to form cross-linked coatings.34 P. aeruginosa colonization on the crosslinked coatings showed very similar behavior to that of the non-crosslinked cationic amine polyester. On both the coatings there were no surviving bacteria and hence biofilm formation was not observed.

Figure 1. Fluorescence microscopy images (10X magnification) of P. aeruginosa biofilms on Glass (A), C-CO2-P (B), C-CO2- (C), C-NH2-P (D), C-NH3+ (E) and C-NH3+ (crosslinked) (F). Green indicates viable bacteria, red indicates dead bacteria. Amine containing polyesters are known to target bacterial cell surface due to the presence of negatively charged phospholipids.41 Calculated percentage area covered by bacteria on all the polymeric surfaces 5 h post-inoculation are shown in (Figure 2). The percentage area covered on glass was 30.8±4.94. For C-CO2-P and C-NH2-P polyesters the percentage area covered was 25.1±5.08 and 23.1±9.31 respectively. In the case of deprotected polyesters, the percentage area


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covered for the anionic carboxylic acid polyester was 8.29±5.51 and 4.55±2.98 for cationic amine polyester. This observation concurs with literature showing lower bacterial adhesion on hydrophilic surfaces.42

40

30

20

10

NH 3+ C-

NH 2 -P C-

CO 2C-

CO 2 -P C-

Gl

as s

0

CNH + 3 Cr os s li nk ed

% Area coverage

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


Figure 2. Percentage area covered by P. aeruginosa over different surfaces. To understand the reason for P. aeruginosa survival and biofilm formation on the various polyester surfaces, the contact angles of the thin coatings were measured. All the surfaces show a decrease of contact angle with time, indicating polymer chain rearrangement upon contact with water. When the functional groups are protected, the C-CO2-P and C-NH2-P, surfaces have contact angles around 74° (Supporting Information Figure S8). The anionic carboxylic acid polyester (C-CO2-) surface has a contact angle of 70°, while cationic amine polyester (C-NH3+) surface has a lower contact angle of 60°. Interestingly, crosslinking of the cationic polyester coatings results in an increase in contact angle relative to the non-crosslinked cationic polyester, which could be explained by decreased mobility and availability of the NH3+ groups on the polymer surface resulting in formation of a more hydrophobic surface. However, the surface



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energies of the various polyester coatings are not the reason for the ability of the amine containing polyesters to kill the attached bacteria and inhibit their ability to form biofilms. Both the crosslinked and non-crosslinked cationic amine polyesters show this ability although the crosslinked coatings have a higher contact angle than the non-crosslinked coatings. The protected carboxylic acid (C-CO2-P), protected amine (C-NH2-P) and anionic carboxylic acid (C-CO2-) polyesters show contact angles in the same range as the crosslinked cationic amine polyester, but they do not exhibit the ability to kill the attached P. aeruginosa. 4.4. Bactericidal activity. In order to investigate if the antimicrobial activity of the cationic amine polyester coating is resulting from any residual small molecules or oligomers leaching out of the polymer coatings, the bactericidal activity of the various surfaces were evaluated by incubating bacteria in well plates containing polymer coated glass coverslips for 24 h. Subsequently bacteria from the supernatant were plated on agar plates and the colony forming units were counted. As shown in (Figure 3), there is no significant bactericidal activity in any of the supernatant solutions, indicating the absence of any small molecules or oligomeric species leaching from the coatings. This suggests that the observed bactericidal activity most likely results from contact of P. aeruginosa with the cationic amine polyester surface. In addition to the above experiment, a zone-of-inhibition study was carried out to eliminate the possibility of the anti-bacterial activity arising from bactericidal leachates. As shown in (Supporting Information Figure S9), after 24 h of incubation, there was no discernible effect on bacterial growth and no observation of any zone-of-inhibition, indicating again the need for bacterial contact with the cationic surface for bactericidal activity.


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1E9 1E8 1E7 1000000

CFU/mL

100000 10000 1000

os sl

-N H + r 3

C

C

C -N H 3 +

H 2 -P C -N

O 2 C -C

C

-C O 2 -P

10

in ke d

100

G la ss

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


Figure 3. Bactericidal activity of polymer coatings toward P. aeruginosa. 4.5. Hemolysis activity measurement. Hemolysis refers to the damage of red blood cells leading to the release of intracellular erythrocyte content into blood plasma.43 Therefore all medical devices and drugs that come into contact with blood need to be tested for potential hemolytic activity. In order to determine if the polymer surfaces are potentially harmful to the membranes of mammalian cells, we tested the polymer coatings for the ability to lyse red blood cells, which can be readily assayed by hemoglobin release. The hemolytic activity of the various polymer coatings were analyzed and compared to the known hemolysis agent, the surfactant Triton X-100 (Figure 4). Interestingly all the coatings, had almost no hemolytic activity after treatment with red blood cells.



120 100

% Hemolysis

80 60 40 20

U V)

+

C l

-P

in m

3

3

C -N H

+

C l

-

(1 0

-N H

2

C

H2

C -N H

-P 2

C -C O

-C O C

la ss G

Tr ito n

+

-

0

1%

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

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Figure 4. Hemolysis activity of polymer coatings toward defibrinated sheep blood. 4.6. Cytotoxicity evaluation. Toxicity of the polymer coatings was evaluated toward NIH-3T3 cell growth on spin coated coverslips of the various polyesters after 24 h and 72 h. LIVE/DEAD staining was used to evaluate the viability of the cells. The polymeric coatings do not exhibit any observable cytotoxicity on NIH-3T3 mouse embryonic fibroblast cells after 24 and 72 hours study (Figure 5). This is very significant in the case of the cationic polymers that show very high antimicrobial activity against P. aeruginosa, but do not have any observable cytotoxicity on NIH-3T3 mouse embryonic fibroblast cells. This selectivity is possibly due to the fact that the surfaces of bacteria possess more negative charges compared to mammalian cells31, leading to stronger interactions between the cationic polyesters and the bacteria. The low toxicity could be a result of the relatively hydrophilic polyester backbone and lack of any pendant hydrophobic groups that can insert into the mammalian cell membranes.35


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Figure 5. Fluorescence microscopy images (10x magnification) of NIH-3T3 mouse embryonic fibroblast cells after 24h on TCPS (A), C-CO2-P (B), C-CO2- (C), C-NH2-P (D), C-NH3+ (E) and C-NH3+ (crosslinked) (F) surfaces. After 72h on TCPS (A’), C-CO2-P (B’), C-CO2- (C’), CNH2-P (D’), C-NH3+ (E’) and C-NH3+ (crosslinked) (F’) surfaces after LIVE/DEAD staining. Green indicates viable cells, red indicates dead cells. 5. Conclusion We have synthesized and prepared a new class of functionalized coumarin polyester coatings with cationic amine pendant groups that demonstrate excellent inhibition of P. aeruginosa colonization of polymer coated surfaces. As shown by comparison to protected carboxylic acid and amine polyesters, the antimicrobial activity of the cationic coumarin polyesters arises from the pendant cationic charge and is not due to leaching of small molecules or oligomers from the polymeric matrix. Even with such high antimicrobial activity the above cationic amine polyester does not show any hemolytic activity or significant cell toxicity toward NIH-3T3 cells. Due to their composition and high molecular weight these polyesters can be fabricated into coatings, an important aspect for applications such as for coating surgical devices, catheters or medical implants. This platform represents a significant step forward within the field of stable antimicrobial coating and shows promise as a system for inhibition of P. aeruginosa colonization on surfaces. ASSOCIATED CONTENT



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Supporting Information Schemes for synthesis of monomers, 1H NMR spectra for protected and deprotected polyesters, SEC analysis of polyesters, ﬂuorescent microscopy images of noncrosslinked and crosslinked cationic polyester films, static contact angle and zone of inhibition assay performed for this study. This material is available free of charge at http://pubs.acs.org Acknowledgments The P. aeruginosa PAO1 (pTdK-GFP) strain was kindly provided by Dr. Matthew Parsek at the University of Washington, WA. The mouse embryonic fibroblast NIH-3T3 cells were kindly donated by Prof. Matthew Becker, University of Akron. The work described here was funded in part by an NSF CAREER grant to AJ (NSF DMR # 1352485). References (1)

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Nontoxic Cationic Coumarin Polyester Coatings ... - ACS Publications

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