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Antimicrobial Activity of Amphiphilic TriazoleLinked Polymers Derived from Renewable Sources Michael Christopher Floros, Janaina Freitas Bortolatto, Osmir Batista de Oliveira, Sergio Luiz Souza Salvador , and Suresh S. Narine ACS Biomater. Sci. Eng., Just Accepted Manuscript • DOI: 10.1021/acsbiomaterials.5b00412 • Publication Date (Web): 27 Jan 2016 Downloaded from http://pubs.acs.org on February 1, 2016

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ACS Biomaterials Science & Engineering

Antimicrobial Activity of Amphiphilic TriazoleLinked Polymers Derived from Renewable Sources Michael C. Floros1, Janaína F. Bortolatto2, Osmir B. Oliveira Jr2, Sergio L. Salvador3 and Suresh S. Narine1* 1

Trent Centre for Biomaterials Research, Departments of Physics & Astronomy and Chemistry, Trent University, Peterborough, ON, Canada K9J 7B8 2

Department of Restorative Dentistry, Araraquara School of Dentistry, UNESP, Univ Estadual Paulista, Araraquara, SP, Brazil 3

Department of Clinical Analyses, School of Pharmaceutical Sciences, University of São Paulo, Ribeirão Preto, SP, Brazil

* Corresponding Author: Fax: 705 750 2786; Tel: 705 748 1011; E-mail: [email protected]

ACS Paragon Plus Environment


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Abstract

Conventional engineered polymers are strong, stable and can interact desirably within the human body in implants and medical devices. However, bacterial colonization of medical devices and implants constructed from these materials results in numerous hospital acquired infections (HAI) and deaths each year. Polytriazole based plastics containing triazole rings and fatty acid derivatives have been synthesized from biological sources without catalysts or solvents. In this study, three amphiphilic polytriazoles with varying triazole density and hydrophilic/hydrophobic segments demonstrated broad spectrum, contact antimicrobial properties against both Gram positive and negative bacteria. SEM analysis of bacteria killed by these polymers evidence membrane damage, indicating that these polymers act by direct contact with bacterial membranes. Surface hydrophobicity of these polymers increased with increasing triazole group density, which also improved the antimicrobial efficacy. This work demonstrates amphiphilic polytriazoles have antimicrobial properties and future utilization of triazole modified polymers may produce self-sterilizing materials which resist bacterial contamination and formation of antibiotic resistant organisms - ideal characteristics for medically relevant biomaterials.

Keywords: Biomimetic, Antimicrobial Polymer, Amphiphilic, Renewable, Click Chemistry, Biomaterial


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Introduction

Thermoplastic polymers are widely used in medical biomaterials, and possess desirable physical and mechanical properties for numerous applications. However, most polymers are highly susceptible to bacterial colonization, with complete biofilm cover occurring in as little as 24 hours of initial contamination.1-2 Bacterial colonization of medical devices is particularly troubling, with as many as 500 000 patients per year acquiring bloodstream infections from intravascular catheters, resulting in high mortality rates and billions of dollars in medical expenses.1,3 Ventilators, urinary catheters, implants, sensors and countless other biomaterials are vulnerable to bacterial infections.1,4-5 Concerns over antibiotic use to treat biomaterial associated infections, and this relation to antibiotic resistant bacteria “superbugs” in hospitalized patients must also be addressed.6-7 Biofilms are extremely resistant to antibiotic therapy, and infected devices often require removal.8-9 The inadequacy of current materials to resist habitation of bacteria and refrain from infecting the patient has become financially, socially and medically unacceptable.10 As increasing life expectancies prompt expanding biomaterial use, and antibiotic resistance threatens treatment of infections associated with these materials, new strategies must be developed to prevent biomaterial associated infections.11-13 Natural and synthetic amphiphilic compounds have demonstrated contact antimicrobial activity against pathogenic microbes.14-19 Amphiphilic compounds inhibit bacteria through membrane disruption caused by their distinct lipophilic and hydrophobic regions.20-21 Strategies to inhibit microbes through membrane disruption, sometimes referred to as the ‘Achilles heel’ represent an ideal target for preventing resistance, as membranes are considered too complex to alter.21-22 In this work, contact inhibition refers to the ability of a material to kill or inactivate



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microorganisms which directly contact the material. The mechanisms of contact inhibition in amphiphilic agents relies on lipophilic driven association, followed by damage or disruption to the microbial membrane.14,16,23 A major weakness of antibiotics is that they may not reach a contaminated biomaterial site or fall to concentrations below the minimum inhibitory concentration when administered or released from materials, which is a contributor to antibiotic resistance.24 Recent advances in lipid derived materials have produced polymers containing fatty acid derived moieties with comparable properties to conventional polymers. Vegetable oil derived materials contain endogenous lipids, are biocompatible, and provide a renewable and greener substitute for petrochemicals.25-26 Furthermore, oleic acid has inhibitory properties against bacteria, causing structural alterations to their membranes27-28, including against resistant strains.29 Oleic acid is also a

major constituent of many bacterial membranes, such as

Staphylococcus aureus 30, already prompting its use in a drug delivery agents.31 Triazoles formed by azide-alkyne Huisgen cycloaddition “CLICK” chemistry have become immensely prominent within the last decade, owing to their ease of synthesis and favourable properties.32 Triazole containing triazole moieties have demonstrated high biocompatibility in a variety of biological systems33-34, as well as antibacterial, antifungal and anticancer properties.35-37 Furthermore, these hydrophilic triazole groups are known to interact with lipids in microbial cellular membranes, altering their orientation and causing leakage.38 Polymerization of triazole-linked polymers can be conducted without catalysts or solvents, greatly reducing the presence of toxic compounds in the final polymers. These polytriazoles have properties comparable to commercial thermoplastics such as polyethylene.39 The need for new biomaterials resistant to infection is readily apparent, motivating our investigation of


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amphiphilic, thermoplastic polymers containing varying densities of triazole groups and lipid segments derived from oleic acid. The antimicrobial properties of a series lipid derived amphiphilic polymers are investigated herein.



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Materials and Methods

Materials Azelaic acid (98%), succinic acid (99%), sodium azide (99%), propargyl alcohol (99%), 4-toluenesulfonyl chloride (98%), triethyl amine (99%), Hoveyda-Grubbs Catalyst 2nd Generation (97%), Lithium Aluminum Hydride (95%), N,N’-Dicyclohexylcarbodiimide (99%), and 4-dimethylaminopyridine (99%) were purchased from Sigma-Aldrich and used as received. Oleic acid (85%) was purchased from Sigma-Aldrich and purified by fractional vacuum distillation. Polyethylene pellets were purchased from Sigma-Aldrich and processed by the method described below. The fatty acid derived diazide (E)-1,18-diazidooctadec-9-ene, and 3 dicarboxylic derived dialkynes; (E)-di(prop-2-yn-1-yl)octadec-9-enedioate, di(prop-2-yn-1yl)nonanedioate and di(prop-2-yn-1-yl)succinate were prepared by a published method.39 Bacillus atrophaeus (ATCC 9372), Escherichia coli (ATCC 8739) and Staphylococcus aureus (ATCC 6538) were purchased from the American Type Culture Collection. Polymer Preparation Three thermoplastic polytriazoles with different triazole densities were synthesized using a solvent and catalyst-free polymerization procedure.39 Equal molecular ratios of diazide and dialkyne monomer, previously prepared from fatty acid derivatives, were added to a PTFE round bottom flask with a magnetic stir bar and heated gradually to 110 °C under nitrogen, then stirred for 20 hours at this temperature. Polymers were cast from the melt into films 0.60 mm thick with dimensions of 17.5 X 12.0 mm in a heated hydraulic press at 170 °C. For antimicrobial analysis, the polymer films were cut into 6.0 mm diameter disks using a circular metal punch on a hydraulic press. Films were cleaned and sterilized for at least 24 hours in ethanol and stored in a


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sterile vacuum desiccator prior to testing. Polymers are named by the number of carbon atoms in the diazide and diacid segment of the monomers – i.e., C18C4 is a polymer from the monomers (E)-1,18-diazidooctadec-9-ene (18 carbons) and di(prop-2-yn-1-yl)succinate (4 carbons). Contact Angle Measurements were performed on a Ramé-hart model 200 goniometer (Ramé-hart, Succasunna, NJ, USA). A 15 µL sessile drop of deionized water was deposited on the surface of the polymer film using an automated micro-syringe. A photograph of the drop on the surface was captured within 2 seconds of deposition and used to calculate the contact angle and 3 different films per polymer were analyzed to account for any surface differences. Bacterial Preparation Bacillus atrophaeus (ATCC 9372) and Escherichia coli (ATCC 8739) were grown aerobically in Nutrient Broth/Agar (Difco) at 30 ºC and 37 ºC, respectively. Staphylococcus aureus (ATCC 6538) was grown aerobically in Tryptic Soy Agar/Broth (Difco) at 37 ºC. Bacteria were in the logarithmic growth stage when harvested with optical densities >0.6 at 600 nm. They were initially washed with a 0.9 % saline solution, and diluted in saline until they contained approximately 5.0 x 108 – 1.0 x 109 CFU/mL. Bacterial Morphology and Adhesion Harvested E. coli, B. atrophaeus and S. aureus were grown on solid medium for 24 - 48 h at 37 °C. Then, circular disks 6.0 mm in diameter of each polymer were placed onto the surface of the medium and incubated for 24 h at 37 °C in direct contact with the bacteria. Zones of inhibition were analyzed after 24 h by measuring any visible rings around the polymers disks. After incubation, the polymer disks were removed from the medium with forceps and inverted, such that the side in contact with the bacteria was upright. Adhered bacteria on the polymer disks



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were immediately fixed with glutaraldehyde (2.5 %), and washed with saline. Samples were dehydrated by graded treatment with increasing concentrations of ethanol, hexamethyldisilazane (HMDS) and were finally dried in a vacuum desiccator for a minimum of one week. Samples were gold sputter coated at 50 mTorr in a Desk V HP sputter coater (Denton Vacuum, Moorestown, New Jersey, USA) after mounting on an aluminum sample holder with conductive tape. Morphological changes to the bacteria between test samples and control polyethylene were observed with a JEOL scanning electron microscope, model JSM-6610/LV (Peabody, Massachusetts, USA) at an accelerating voltage of 12 kV. Composite SEM images of the uncoated polymer surfaces were resolved with a Phenom ProX, (Phenom-World, The Netherlands) scanning electron microscope at an accelerating voltage of 15 kV. Confocal Viability Characterization The antimicrobial efficacy of each polymer was determined against B. atrophaeus, E. coli, and S. aureus. Suspensions of bacteria was prepared by diluting B. atrophaeus, E. coli and S. aureus in their logarithmic growth phase directly from their respective liquid growth media described in the bacterial preparation step with saline until they reached a concentration of 5.0 x 108 – 1.0 x 109 CFU/mL. From each saline diluted bacterial suspension, 20 µL was pipetted onto the surface of each polymer disk (28 mm2) in triplicates to mimic a concentrated infectious secretion. Test samples were placed in a hydrated chamber40 to prevent dehydration and incubated at room temperature (~25 °C) for 1 or 4 h. Bacteria were liberated from the test surface by vortexing with 80 µL of freshly prepared BacLight™ L7012 LIVE/DEAD (Molecular Probes, Eugene, Oregon, USA) stock solution in 0.9 % saline. This dye contains two stains: SYTO 9 and propidium iodide (PI). The fluorescent agent in SYTO 9 stains all bacterial cells, while the dye in propidium iodide stains only dead cells with membrane damage. Samples were incubated in the


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dark for 15 minutes at room temperature, and images were acquired with a Leica Confocal TCS SP5 (Leica Microsystems GmbH, Heidelberg, Germany) equipped with a HCX PL APO lambda blue 63.0 x 1.40 oil objective and a motorized scanning stage. Images were captured with a resolution of 2048 x 2048 pixels per image. Fluorescent dyes were excited with the 488 nm argon emission for SYTO 9 and the 543 nm HeNe laser emission for propidium iodine. Fluorescence emissions were filtered with a tunable Acousto-Optical Beam Splitter (AOBS) into two channels: 500-550 nm for SYTO 9 and 599-702 nm for propidium iodide. Quantification of the fluorescence intensities of each signal were aided by the NIH developed freeware software ImageJ v1.4841, and the percent of dead bacteria was calculated as the ratio of the red fluorescence signal over the sum of the total fluorescent signal (red + green), in a method adapted from literature.42 Test sample was challenged in triplicate for each duration and bacteria type.



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Results and Discussion

Three triazole containing polymers derived from oleic acid were synthesized in this study. The structures of the polymers are represented in Scheme 1. A 1,5-substituted isomer of the triazole, was also formed as a minor product (only the major product is shown for clarity). These melt cast polymers contain triazole linkages in their backbone, advantageous as the antimicrobial properties will be present in the bulk as well as at the surface. Thermoplastic polymers are extremely versatile, and can be melt processed into numerous shapes and architectures for countless applications. In contrast, surface modifications can be worn off due to abrasion, such as scratching, causing loss of the desirable properties added through surface modification.

Scheme 1: Simplified repeating units of the triazole containing polymers with fatty acid derived lipophilic segments highlighted in red.


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Contact Angle Water contact angle is an important factor affecting bacterial adhesion to surfaces.43 Static water contact angles aided determination of the relative hydrophilic/hydrophobic characteristics of each polymer. The hydrophilic characterises are related in part to triazole groups density, as demonstrated in Figure 1. As the fraction of the lipophilic hydrocarbon region increases from C18C4 to C18C18, so does the water contact angle. An addition of 5 hydrocarbons per repeating unit between C18C4 and C18C9 resulted in a water contact angle increase from 68.1 ± 2.1 ᴼ to 80.6 ± 0.1 ᴼ. Surprisingly, C18C18 exhibited much more hydrophobicity, with a water contact of 123.7 ± 0.3 ᴼ, much higher than the other polymers, including polyethylene (~100 ᴼ). Significant surface micro-roughness on only C18C18 was revealed by microscopy analysis (Figure 1), and a composite stitch of multiple SEM images (Figure 2) reveals the uniformity of this behaviour over a large area. Surface micro-roughness is a key factor influencing hydrophobicity44, and similar behaviour has been demonstrated in other amphiphilic polymers.45 The formation of microstructured roughness on the surface of C18C18 may be related to the high similarities of the monomer units, each containing an 18 carbon transunsaturated chain. Previous approaches have used amphiphilic diblock monomers to promote microphase segregation and self-assembling morphologies in polymers.46-47



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Figure 1: Representative SEM images of the polymer surfaces with corresponding water contact angles. Errors represent standard deviations from triplicates. Scale bar = 5 µm.

Figure 2: Composite SEM image of C18C18, demonstrating widespread microroughness. Field of view is 150 µm horizontally.


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Bacterial Morphology and Adhesion Microbial contamination of medical devices, such as catheters generally occur within 24 hours of insertion48-49, by contact with contaminated fluids or colonization of the biomaterial.50 To simulate aggressive contamination and biofilm forming conditions, polymers were challenged in established planktonic bacterial colonies in growth medium for 24 h. No zones of inhibition (ZOI) were visible for any of the polymer/bacteria systems tested, as expected for surface contact killing polymers.51 Photographs from the zone of inhibition tests for each polymer and bacteria combination are included in the supporting information. Visualization of adhered bacteria after incubation at physiological temperature for 24 h was aided by SEM. Test polymers successfully resisted contamination after this challenge, exhibiting only non-viable adhered cells. Adhered bacteria displayed significant membrane damage, dehydration, and non-viable characteristics (Figure 3, i), similar to the appearance of bacteria killed by antimicrobial peptides.52 Membrane damage is especially apparent on E. coli (Figure 3, C ii), with apparent pore-like structures and scattered cellular components visible. In contrast, bacteria adhered to polyethylene (Figure 3, ii) display normal morphological appearances, thick film formations, and integration or adhesion to the polymer surface. The polyethylene surfaces were almost completely covered in bacteria, and most of the films were too thick to visualize the pristine surface. Biofilms require surface attachment during initiation stages, and the inhibition of adhered bacteria on biomaterial surfaces may prevent biofilm formation.53 In contrast, the densely packed bacteria on polyethylene already show multilayer biofilm formation.



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Figure 3: Representative SEM micrographs of (A) S. aureus, (B) B. atrophaeus and (C) E. coli on (i) polyethylene and (ii) C18C4. Scale bar = 1 µm.

Confocal Bacterial Viability To quantify the efficacy of polymers at killing bacteria over different durations, polymers were challenged by aqueous solutions of bacteria to simulate surface contamination by an infectious secretion. Efficacy of inhibition was determined by a Live/Dead BacLight fluorescence viability kit. Confocal microscopy analysis of bacteria directly on the polymer surfaces was not possible due to strong background fluorescence. Fluorogenic characteristics of triazoles have been previously described in literature.54 Bacteria were liberated from the polymer surface just prior to confocal analysis to enable fluorescent quantification. For each type of bacteria tested, C18C4 displayed the highest inhibition (Figure 4). Representative confocal images of the control polyethylene and C18C4 are displayed for both contact durations in Figure


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5. Contact times of 4 hours demonstrate near complete inhibition of all tested organisms, in contrast to >95% viability on polyethylene controls. The higher inhibition of C18C4 may be related to its higher wettability, with more of the polymer surface in contact with the aqueous bacterial challenge, or to a higher triazole density. In the aqueous test environment, segments of the polytriazoles may diffuse into the solution, enabling greater contact area and more effective inhibition, analogous to brush-like and surface grafted polymer architectures. This may also explain why the more hydrophilic polytriazoles have improved inhibition rates. Bacterial inhibition by a membrane damage mechanism, also observed in the SEM analysis, is further supported by the Live/Dead fluorescent viability testing. Propidium iodide can only stain cells with ruptured membranes.55



100

A Polyethylene C18C18 C18C9

Dead Bacteria (%)

80

C18C4

60

40 20 0

Dead Bacteria (%)

100

1h

Contact Time

4h

Contact Time

4h

Contact Time

4h

B Polyethylene C18C18 C18C9 C18C4

80 60 40 20 0

100 80

Dead Bacteria (%)

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1h

C Polyethylene C18C18 C18C9 C18C4

60 40 20 0

1h

Figure 4: Control and polymer kill rates for 1 or 4 h contact time on (A) E. coli, (B) B. atrophaeus and (C) S. aureus, as determined from relative BacLight™ LIVE/DEAD fluorescent PE – E. coli 1 h signals. Error bars represent standard deviations from triplicates.


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Figure 5: Representative Live/Dead fluorescent image of A – E. coli after 1 h i and 4 h ii contact time and in B – B. atrophaeus bacteria after 1 h (i) and 4 h (ii) contact time and C - S. aureus after 1 h (i) and 4 h (ii) on polyethylene (PE) and C18C4.

Mechanism of Action The exact mechanisms responsible for contact inhibition are still not fully understood. Experiments by which water insoluble polymers kill bacteria have shed considerable insight into a generalized mechanism. Microscopy analysis of polymer killed dead bacteria show membrane



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disordering and disruption from association with surface hydrophobic and hydrophilic groups on the polymer.56-57 Permeabilization of the membrane leading to leakage has been detected and described in many amphiphilic polymers as a result of these membrane effects.58 Fatty acids are a major constituent of bacterial membranes, but external membrane contact with fatty acids is also known to cause alterations in bacterial membrane fluidity,27-28 resulting in cell death.59 Gram-positive organisms are known to be especially susceptible to damage from unsaturated fatty acids, due to their membrane composition30,60 and may be related to the high efficacy of C18C4 against B. atrophaeus and S. aureus, which display near complete inhibition after 4 hours despite containing more robust Gram positive membranes. The polytriazoles prepared in this study were high molecular weight, and due to the solvent free melt condensation polymerization, the polytriazoles were formed without dilution or solvation and have a high degree of polymer chain entanglements. Soaking polymers in water and common organic solvents for extended time periods (>7 days) did not affect their dry weight. An additional test was performed by incubating a solution of E. coli in PBS with C18C4 (the most effective polymer) or polyethylene for 4 hours. After incubation, the number of CFU/mL were calculated to determine if the antimicrobial polymer displayed activity in solution. No differences in bacterial growth were observed between C18C4 and polyethylene, further supporting the contact inhibition mechanism proposed (Supporting Information S5). Several plausible explanations regarding the mechanism of amphiphilic contact inhibition by polymers have been described within the literature. Partial or full penetration of the bacterial membrane by polymer segments is often used to explain the action of contact active polymers.61-63 Polymer segments with sufficient length and flexibility may be able to directly penetrate through the cell wall or through gaps in the cell wall to reach the bacterial membrane,64-65 causing rupture


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through physical penetration or inducing lipid reorganizations and membrane disruption. C18C4 demonstrated the highest antimicrobial activity, being the most hydrophilic of the polymers tested. This effect may be due to partial solubility of segments of the entangled polymer, leading to protrusion of polymer chains from the surface which would occur the most in the hydrophilic C18C4, producing more polymer surface area and higher kill efficacies. The varying density of amphiphilic characteristics caused by different ratios of hydrophilic triazole groups to lipophilic segments in the polytriazoles may also interact differently with bacterial membrane lipids. Some studies have shown antimicrobial activities are improved as amphiphilic polymer molecular weights increase, supporting a direct interaction/penetration mechanism.66-68 However, antimicrobial properties of covalently attached quaternary ammonium functionalized silane coatings as thin as 2.5 nm have also been reported,53,69-71 significantly less than the 30 nm cell wall thickness of S. aureus,64 suggesting that alternative mechanisms may also be involved. Most contact active amphiphilic polymers previously reported were prepared through the addition of cationic or anionic groups to a base polymer.56 The synthetic techniques to accomplish this generally use free radical polymerizations, resulting in polymers with charged groups attached by a spacer group, limiting the potential polymer backbone and mechanical properties of these polymers.19,72 Cationic and anionic polymers also display toxicity to aquatic life73-75 and human cells.67,76-77 To the best of our knowledge, this is the first example of an antimicrobial amphiphilic polymer containing triazoles without cationic or anionic groups, and may alleviate the biocompatibility and environmental issues found in charged amphiphilic polymers. These polytriazoles are also thermoplastics, in contrast to many of the crosslinked alternatives, enabling applications where amphiphilic antimicrobial polymers can be injection molded or melt processed and used for applications requiring strong, flexible materials. A



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representation of a bacterial cell interacting with a polymer segment is displayed in Figure 6. Interactions with surface groups on the polymer alter the membrane structure, eventually leading to disruption through formation of transmembrane pores or rupture, killing the organism.

i

ii

Figure 6: Schematic representation of bacterial interaction (i) with the amphiphilic polymer (blue), and the bacterial membrane (brown), resulting in membrane disruption and inactivation (ii). Insert depicts segments of the polytriazole disrupting the structure of a simplified bacterial cell envelope.


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Conclusions

Lipid derived triazole containing polymers have broad spectrum antimicrobial activity against Gram-negative and positive bacteria. Concentrated aqueous solutions of bacteria underwent >90 % inhibition after 4 hours contact. Bacterial adhesion testing demonstrated that these polymers killed all adhering organism after 24 hours. Microscopy analysis supported by Live/Dead fluorescent staining exhibited membrane damage as the mechanism of activity, possibly due to interactions between membranes and polymer lipids. Control of hydrophobic characteristics was accomplished by varying the lipophilic segment length of one monomeric unit. C18C4, which had the highest hydrophilicity, demonstrated the highest bacterial kill rates. Increasing the density of triazole groups in the polymers increased corresponding hydrophilicity. The strategy adopted herein utilizes endogenous, biologically derived fatty acids and dicarboxylic acids as the building blocks for triazole-linked polymeric biomaterials. Furthermore, using natural lipids as hydrophobic segments improves the renewability of these materials while aiding in inactivation of bacteria. We hope this work will contribute to further studies on the use of triazole groups in different amphiphilic architectures and result in improved biomaterials which prevent bacterial contamination.



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Acknowledgements

We thank Marina Del Arco for her technical assistance with preparation of microbiological samples. The financial support of Elevance Renewable Sciences, NSERC, Grain Farmers of Ontario, GPA-EDC, CAPES/DFATD, Industry Canada, and Trent University is gratefully acknowledged.


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Supporting Information

Supporting Information Available: The following files are available free of charge: Bacterial Zone of Inhibition photographs Bacterial viability test in solution



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References

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For Table of Contents Use Only

Antimicrobial Activity of Amphiphilic Triazole-Linked Polymers Derived from Natural Sources Michael C. Floros1, Janaína F. Bortolatto2, Osmir B. Oliveira Jr2, Sergio L. Salvador3 and Suresh S. Narine1* 1

Trent Centre for Biomaterials Research, Departments of Physics & Astronomy and Chemistry, Trent University, Peterborough, ON, Canada K9J 7B8 2

Department of Restorative Dentistry, Araraquara School of Dentistry, UNESP, Univ Estadual Paulista, Araraquara, SP, Brazil 3

Department of Clinical Analyses, School of Pharmaceutical Sciences, University of São Paulo, Ribeirão Preto, SP, Brazil


Antimicrobial Activity of Amphiphilic Triazole ... - ACS Publications

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