Article pubs.acs.org/Biomac
CO-Releasing Polymers Exert Antimicrobial Activity Diep Nguyen,†,‡ Thuy-Khanh Nguyen,†,‡ Scott A. Rice,§,⊥ and Cyrille Boyer*,†,‡ †
Australian Centre for Nanomedicine (ACN); ‡Centre for Advanced Macromolecular Design (CAMD), School of Chemical Engineering; and §Centre for Marine-Innovation, School of Biological, Earth, and Environmental Sciences, University of New South Wales, Sydney 2052, Australia ⊥ The Singapore Centre for Environmental Life Sciences Engineering and The School of Biological Sciences, Nanyang Technological University, Singapore 639798, Singapore S Supporting Information *
ABSTRACT: Infectious diseases remain one of the leading causes of death worldwide despite the tremendous effort devoted to the design and development of antimicrobial agents. However, the decrease in the effectiveness of some antibiotics is often associated with the development of drug resistance by pathogen. This leads to an urgent need for the development of new therapeutic approaches that can overcome the development of drug resistance. Recent evidence suggests that the biological signaling molecule carbon monoxide (CO) presents remarkable antimicrobial properties. Herein, we report the design and synthesis of a new type of water-soluble CO-releasing polymer with antimicrobial activity against Pseudomonas aeruginosa that is highly efficient at preventing biofilm formation.
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INTRODUCTION According to the World Health Organization (WHO), infectious diseases are the second leading cause of death worldwide and are responsible for approximately 15 million deaths annually.1 The decline in effectiveness of most current antibiotics is attributed to the development of drug resistance by pathogens. The emergence and widespread resistance to antibiotics in pathogenic bacteria poses a serious threat to global public health if drugs are no longer effective.2 According to the Centers for Disease Control and Prevention, each year in the United States, at least 2 million people are infected with bacteria that are resistant to antibiotics, and at least 23 000 people die each year as a direct result of these infections. In fact, a recent report published by Review on Antimicrobial Resistance predicts that drug-resistant infections could cost 10 million lives each year as well as $100 trillion by 2050.3 Thus, there is an urgent need to develop new classes of antimicrobial agents to combat antibiotic-resistant pathogens.4 Recently, the biological carbon monoxide (CO)5−9 and nitric oxide10,11 signaling molecules were reported to have promising effects on bacteria. CO was identified to be an effective inhibitor of the respiratory chain in Pseudomonas aeruginosa, which causes 30% of the nosocomial infections in the hospital.12 Furthermore, CO was shown to promote phagocytosis of Escherichia coli via p38-mediated surface expression of TLR413 and to act as an important mediator of the host defense response to microbial sepsis in mice.14 Wegiel et al.15 demonstrated that the bactericidal property of CO relies on the production of ATP. CO binds to heme-containing bacterial respiratory complexes, leading to the generation of ATP, which results in inflammation © XXXX American Chemical Society
and ultimately bacterial clearance by professional phagocytes. However, the practical clinical use of CO gas as an antibacterial chemotherapy poses problems associated with toxicity at high concentration and difficulties storing and delivering CO gas to the target tissue in a controlled fashion.16 To sidestep these limitations, a range of small molecules capable of releasing CO (CO releasing molecules, or CORMs, Scheme 1) under specific conditions has been explored.16−22 CORMs were documented to possess bactericidal properties against pathogens including E. coli, Staphylococcus aureus, and P. aeruginosa.6,23−26 For example, CORM-2 and CORM-3 were shown to have efficient antibacterial properties.27 Indeed, the addition of CORM-2 or CORM-3 to a solution led to a decrease in the viability of E. coli and S. aureus cells after 30 min.25 Furthermore, the cells were not able to resume growth after 4 h of exposure to these CORMs. CORM-3 was also reported to combat antibiotic-resistant strains of P. aeruginosa.23 Recently, Nagel et al.28 prepared a novel photoCORM [Mn(CO)3(tpa-κ3N)]Br that has the ability to prevent the growth of E. coli cultures. CORMs are proposed to exert their lethal effect on bacteria through the “Trojan Horse” mechanism.29 According to this mechanism, the antimicrobial action of CORMs is mediated by the inhibition of respiration. More specifically, CORMs are transported to the cell, and CO release is enhanced by a reaction with intracellular thiolcontaining molecules. Then, CO is accessible to membraneReceived: May 27, 2015 Revised: July 20, 2015
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DOI: 10.1021/acs.biomac.5b00716 Biomacromolecules XXXX, XXX, XXX−XXX
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Biomacromolecules Scheme 1. Examples of CO-Releasing Molecules Reported To Possess Antimicrobial Activities
polymers. Myoglobin assay showed that CO release from the polymers takes place on the same time scale as the release from the free complex. The resulting Mn(CO)3-polymer conjugates with distributed macromolecular weights, and sizes can be used for the passive delivery of CO to tumor tissue or sites of inflammation. In this work, we report a novel class of CO-releasing polymeric particles that releases CO in the presence of the bacteria P. aeruginosa, exhibiting antimicrobial activity and inhibiting biofilm formation. This paper describes for the first time the use of a polymer as a CO carrier for the prevention and treatment of biofilm formation and inhibition of P. aeruginosa cell growth. In contrast to CORM, CO-releasing polymer presents a longer action and higher antimicrobial activity.
bound heme targets and binds to terminal oxidase, leading to an inhibition of respiration that precedes a decrease in cell viability. In addition to their antibacterial activity, CORMs are also reported to possess antibiofilm activity.30 Recently, CORMs such as ALF492 were demonstrated to comprise a new class of drugs for the treatment of severe forms of malaria infection.31 Because their mode of action is not similar to currently used antibiotics, CORMs hold promise as a novel antimicrobial chemotherapy for the treatment of infectious disease. However, the clinical use of CORMs as preventative agents is limited by a lack of stability, solubility, and specificity. Many CORMs present a low solubility in water (often insoluble) and require the addition of organic solvents such as dimethyl sulfoxide (DMSO). For example, CORM-2, one of the most commonly used compounds in biological and medicinal studies, is not water soluble, and only soluble in DMSO. Although the use of small amount of DMSO can be performed by in vitro experiments to investigate the effect of CORMs in the cells, it is not practicable for in vivo and clinical application. One promising avenue to address these drawbacks is the development of CO releasing macromolecules for the controlled and sustained release of CO.32−35 The application of polymer-based delivery offers several advantages over small therapeutic molecules including improvements in pharmacokinetics and accumulation, a reduction in side effects, and a decrease in immunogenicity.36−39 Another advantage of polymeric carriers (such as polymeric nanoparticles) is their ability to deliver a drug directly to a target site.40,41 In addition, macromolecules offer a high CO-loading capacity due to possibility of several CO groups attaching to a single macromolecule, resulting in a significant increase in CO concentration and an improved clinical response. As such, there has been increasing interest in using nanomaterials as CO carriers, particularly CO-generating polymeric materials. However, there are few studies on COreleasing polymers. The conjugation of a polymer with CORMs was first applied in a micellar system in 2010. Hubbell and coworkers42 developed CO-releasing polymeric micelles with a decreased diffusion in tissue and consequently enhanced the ability to target distal tissue draining sites. These micelles were prepared from triblock copolymers composed of a hydrophilic and biocompatible poly(ethylene glycol) block, a poly(ornithine acrylamide) block bearing Ru(CO)3Cl(ornithinate) moieties, and a hydrophobic poly(n-butylacrylamide) block. CO-release studies showed that micelles release CO in the presence of thiol-containing compounds (e.g., glutathione), and the rate of CO release was slower than that of CORM-3. In addition, they were demonstrated to successfully attenuate the lipopolysaccharide-induced inflammation of human monocytes and to reduce the cytotoxicity of the Ru(CO)3Cl (amino acidate) moiety by the stealth feature of poly(ethylene glycol). Moreover, the linking of photoCORMs to copolymers was also established. Brückmann et al.43 generated a CORM-functionalized copolymer by conjugation of the manganese tricarbonyl moiety to 2-hydroxypropyl methacrylamide (HPMA)-based
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EXPERIMENTAL METHODS
Materials. 1-Thio-β-D-glucose sodium salt, 1-thioglycerol, and 2diethylaminoethanethiol hydrochloride were purchased from SigmaAldrich at the highest purity available. DMSO (Ajax Chemical), diethyl ether (Ajax Chemical), petroleum spirit (Ajax Chemical), acetonitrile (Ajax Chemical), and toluene (Ajax Chemical) were used as received. Monomer, oligo(ethylene glycol) methyl ether acrylate with average Mn of 480 g/mol, was received from Sigma-Aldrich and prior to reaction was deinhibited by passing the monomers through a column filled with basic alumina. Dialysis membrane (MWCO 6000) was obtained from Spectrum Laboratories (cellu SepT4, regenerated cellulose-tubular membrane). 2,2-Azobisisobutylronitrile (162 g/mol, AIBN) which was purchased from Sigma-Aldrich, was recrystallized from methanol before use. RAFT chain transfer agent, 2(((butylsulfanyl)carbothioyl)sulfanyl)propanoic acid (CTA), was synthesized according to the literature procedure.44 For the measurement of carbon monoxide release, myoglobin from equine skeletal muscle was purchased from Sigma-Aldrich. The BacTiter-Glo Microbial Cell Viability Assay (Promega, Alexandria, Australia) was used to measure cell viability. Characterization Methods. NMR Spectroscopy. 1H NMR spectra were recorded using Bruker DPX-300 (300 MHz) and DPX400 (400 MHz) spectrometers. d3-CD3CN and d6-DMSO were used as solvents. All chemical shifts are quoted in parts per million (ppm) and referenced to residual the residual solvent frequencies (1H NMR: d3-CD3CN = 1.94, d6-DMSO = 2.50). The monomer (i.e., OEGA) conversion was calculated by the following equation to give ∼76% conversion, where I5.9 ppm and I4.2 ppm correspond to the integral of the vinyl signal from the monomer at 5.9 ppm and the ester signal from the monomer/polymer at 4.1 ppm, respectively:
⎛ ⎞ I 5.9ppm ⎟ ⎜ α (OEGA conversion %) = ⎜1 − 4.1ppm ⎟ × 100 I ⎝ ⎠ 2 The theoretical molecular weight was calculated by the following equation: ⎛⎛ [M ] ⎞ ⎞ 0 M n (theoretical) = ⎜⎜⎜ ⎟ × α × MWM⎟⎟ + MWCTA ⎝⎝ [CTA]0 ⎠ ⎠ B
DOI: 10.1021/acs.biomac.5b00716 Biomacromolecules XXXX, XXX, XXX−XXX
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°C) to remove remaining solvent. By 1H NMR, the estimated DPP(OEGA) and DPP(VBC) values are 76 and 10 units, respectively. After removal of solvent and drying under vacuum, P(OEGA)-b-P(VBC) diblock copolymer with Mn,NMR = 38 200 g/mol, Mn,SEC = 26 600 g/ mol, and PDI = 1.25 was obtained. Conjugation of Thiol Compounds to Diblock Copolymer via Nucleophilic Reaction. General Procedure for Conjugation of Thiol Compound to Diblock Copolymer. A mixture of P(OEGA)-bP(VBC) (1 equiv.) and thiol compound (10 × 1.2 equiv.) was dissolved in DMSO. Then, triethylamine (15.6 eqv.) was added, and the reaction mixture was stirred at 50 °C for 24 h. The mixture then was dialyzed using cellulose-tubular membrane MWCO 6000 g/mol with water and methanol to remove unreacted thiol and triethylamine. Example of Conjugation of 1-Thio-β-D-glucose Sodium Salt to P(OEGA)-b-P(VBC) Copolymer. A mixture of 970 mg of P(OEGA)-bP(VBC) (2.5 × 10−2 mmol) and 65 mg of 1-thio-β-D-glucose sodium salt (0.39 mmol) was dissolved in 10 mL of DMSO. Fifty-four microliters of triethylamine was added subsequently, and the reaction mixture was stirred at 50 °C for 24 h. The mixture was then dialyzed using cellulose-tubular membrane MWCO 6000 g/mol with water and methanol as cosolvent (50:50 v/v). After drying, 920 mg of sugar functionalized P(OEGA)-b-P(VBGlu) diblock copolymer with Mn, NMR = 40 700 g/mol, Mn, SEC = 39 700 g/mol, and PDI = 1.34 was obtained. Attachment of CORM-2 to Thiodiblock Copolymer. General Procedure for Attachment of CORM-2 to Thiofunctionalized Copolymer. A MeOH solution of tricarbonyldichlororuthenium(II) dimer (CORM-2) (10 × 0.5 eqv.) was added dropwise to a MeOH solution of thio- functionalized copolymers (1 equiv.). The reaction was stirred for 24 h at room temperature under a nitrogen atmosphere in the absence of light. The reaction medium was precipitated in diethyl ether and centrifuged (8000 rpm for 5 min). Attachment of CORM-2 to P(OEGA)-b-P(VBGlu). A MeOH solution of tricarbonyldichlororuthenium(II) dimer (CORM-2) (42 mg, 1.04 mmol) (4 mL) was added dropwise to MeOH solution of P(OEGMA)-b-P(VBGlu) (650 mg, 16 × 10−2 mmol) (5 mL). The reaction was stirred for 24 h at room temperature under a nitrogen atmosphere in the dark. The reaction medium was precipitated in diethyl ether and centrifuged (8000 rpm for 5 min). Then ATR-FTIR analysis was performed to check the presence of a carbon monoxide peak in the block copolymer (signal of carbon monoxide in the region between 1800 and 2100 cm−1). After drying, 450 mg of P(OEGA)-bP(VBGlu-Ru com) (1) with Mn, NMR = 44 000 g/mol, Mn, SEC = 40 800 g/mol, and PDI = 1.24 was obtained. Studies of CO Release by Myoglobin Assay. Myoglobin Assay Procedure. A stock solution of myoglobin (Mb) from equine skeletal muscle (2 mg/mL) was freshly prepared in 0.1 M phosphate buffer solution (pH = 7.4) and was degassed by bubbling with nitrogen for at least 15 min. To this degassed solution was added a freshly prepared solution of sodium dithionite (24 mg/mL) at 1:10 dithionite/Mb (v/ v) to convert met-Mb to deoxy-Mb. Two controls were done in duplicate, the negative control (0% CO-Mb) a deoxy-Mb solution, and the positive control (100% CO-Mb) obtained by bubbling pure CO gas into deoxy-Mb solution. First, stock solutions of polymers 1, 2, and 3 were prepared by dissolving CO-releasing compounds in phosphate buffer solution. Then an aliquot of stock solutions was added to the deoxy-Mb solution to produce final concentrations of 20 μM. This solution was quickly transferred to a cuvette and then overlaid with 500 μL of light mineral oil (Sigma) to prevent CO escaping or the myoglobin being oxygenated. The absorption was recorded at room temperature at predetermined time points using a CARY 300 spectrophotometer, measuring between 500 and 600 nm with a step of 2 nm. The amount of MbCO formed can be calculated utilizing the formula as previously reported:38
where [M]0 is the initial monomer concentration; [CTA]0 is the initial RAFT agent concetration; α is the conversion of monomer; MWM is the molecular weight of monomer; and MWCTA is the molecular weight of RAFT agent. Infrared Spectroscopy. Attenuated total reflection Fourier transform infrared (ATR-FTIR) spectra of polymer samples were obtained using a Bruker Spectrum BX FTIR system using diffuse reflectance sampling accessories. The spectrophotometer was equipped with a tungsten halogen lamp and Si/Ca beam splitter. Spectra were obtained at regular time intervals in the MIR region of 4000−500 cm−1 at a resolution of 4 cm−1 (128 scans) and analyzed using OPUS software. UV−vis Spectroscopy. UV−vis measurements were performed on a CARY 300 spectrophotometer (Bruker) using a quartz cuvette. Size Exclusion Chromatography (SEC). SEC was performed using tetrahydrofuran (THF) or dimethylacetamide (DMAc) as the eluent. The SEC system was a Shimadzu modular system comprising an auto injector, a Phenomenex 5.0 μm beadsize guard column (50 × 7.5 mm2) followed by three Phenomenex 5.0 μm bead-size columns (105, 104, and 103 Å), and a differential refractive-index detector. The system was calibrated with narrow molecular weight distribution polystyrene standards with molecular weights ranging from 200−106 g mol−1. Dynamic Light Scattering (DLS). DLS was carried out on a Malvern Zetasizer Nano Series running DTS software (He−Ne laser, 4 mW, λ = 633 nm, angle 173°). The polydispersity index (PDI) was used to describe the width of the particle size distribution and calculated via the DTS software from a Cumulants analysis of the measured intensity autocorrelation function. It is related to the standard deviation of the hypothetical Gaussian distribution (i.e., PDI = σ2 /ZD2, where σ is the standard deviation, and ZD is the Z average mean size). Polymer samples were dissolved in Milli-Q grade water (1 mg/mL) and filtered through 0.45 μm pore size filter to remove dust prior to analysis. The samples were transferred to their respective disposable cuvettes for analysis. Inductively Coupled Plasma Optical Emission Spectroscopy (ICPOES). The percentage of ruthenium in the CO-releasing polymer was determined by ICP-OES technique using PerkinElmer OPTIMA 7300 ICP optical emission spectrometers. Briefly, 10 mg of CO-releasing polymer was dissolved in 10 mL of distilled water, and the solution was then analyzed by ICP-OES using a detection wavelength of 240.272 nm. Synthetic Procedures. Synthesis of P(OEGA). [OEGA480]/ [CTA]/[AIBN] = 100:1.0:0.1. OEGA (10.5, 22 × 10−3 mol), CTA (66 mg, 2.2 × 10−4 mol), AIBN (4.5 mg, 2.7 × 10−5), and toluene (40 mL) were prepared in a 100 mL round-bottom flask equipped with a magnetic stirrer bar. The reaction mixture was degassed with nitrogen for 30 min. (Note: toluene has been chosen due to its low transfer constant and its miscibility with petroleum ether, which is employed for the purification step of polymer by precipitation). The reaction mixture was then placed into an oil bath preheated to 70 °C, and the polymerization was run for 3 h. Upon completion, the reaction was quenched in an ice bath for 15 min, and two aliquots were sampled for SEC and 1H NMR analyses. The monomer conversion was determined by 1H NMR analysis. The polymer was purified via three precipitations with the mixture of petroleum spirit and diethyl ether (30:70 v/v) and centrifuging (5 min, 8000 rpm). After removal of solvent and drying under vacuum, polymer P(OEGA) with Mn, NMR = 36 700 g/mol, Mn, SEC = 21 700 g/mol, and PDI = 1.25 was obtained. Synthesis of Diblock Copolymer P(OEGA)-b-P(VBC). [P(OEGA)]/ [VBC]/[AIBN] = 1:40:0.1. P(OEGA) (5.6 g, 1.5 × 10−4), VBC (920 mg, 6 × 10−3), AIBN (2.5 mg, 1.5 × 10−5), and toluene (30 mL) were prepared in a round-bottom flask equipped with a magnetic stirrer bar. The reaction mixture was degassed with nitrogen for 15 min. The degassed solution was immersed in a preheated oil bath at 70 °C for 12 h. The reaction was then placed in an ice bath for about 15 min to terminate the polymerization, and two aliquots were collected for SEC and 1H NMR analyses. VBC conversion was determined by 1H NMR analysis. The reaction media was precipitated in diethyl ether and centrifuged (5 min, 8000 rpm). The purification process was repeated three times, and the reaction medium was dried in vacuum oven (40
⎞ ⎛ε A ⎞ ⎛ εiso [COMb] = ⎜ d542 − 542 ⎟ × ⎜ ⎟ [Mb] + [COMb] A iso ⎠ ⎝ εd542 − εCO542 ⎠ ⎝ εiso where A542 and Aiso are the measured absorbances at each time point at 542 and 552 nm, respectively. εd542, εCO542, and εiso are the extinction C
DOI: 10.1021/acs.biomac.5b00716 Biomacromolecules XXXX, XXX, XXX−XXX
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Biomacromolecules Scheme 2. Synthesis of CO-Releasing Polymers
coefficients of deoxyMb at 542 nm, COMb at 542, and of the isosbestic point at 552 nm, respectively. The concentration of MbCO was calculated and plotted against different time points. By applying a suitable fit function, the plateau level of the MbCO concentration was determined, and subsequent division by the initial concentration of CO-releasing compound gave equivalents of CO liberated. The half-life of CO release from polymers (1−3) was then estimated from the graph. Growth Inhibition and Biofilm Prevention Assays. The laboratory strain Pseudomonas aeruginosa PAO1 was used to characterize the effects of CO-releasing polymers on planktonic growth and biofilm formation. The procedures of growth were similar as previously published by our group45 with some modifications. Briefly, in all assays, overnight cultures in Luria−Bertani (LB) medium were diluted to an optical density (OD) at 600 nm of 0.01 in 1 mL of M9 minimal medium (containing 48 mM Na2HPO4, 22 mM KH2PO4, 9 mM NaCl, 19 mM NH4Cl, 2 mM MgSO4, 20 mM glucose, 100 μM CaCl2, pH 7.0) in tissue-culture treated 24-well plates (BD). Prior to incubation, the bacterial medium was inoculated with CO-releasing compounds to yield final concentrations of 10−200 μM, as indicated, while control wells were left untreated. Treatments were added to the wells, each from a 10 μL aliquot of a stock solution at the appropriate concentration previously sterilized by passing through a 0.22 μm pore size filter: CORM-2 (Sigma-Aldrich) made fresh in DMSO, COreleasing and non-CO polymers were made in phosphate-buffered saline (PBS). The plates were incubated at 37 °C with shaking at 180 rpm in an orbital shaker-incubator that does not stop agitation when the door is opened (model OM11, Ratek, Boronia, Australia), and the biofilms were allowed to grow for up to 7.5 h. After incubation at each time point, the planktonic biomass was quantified by removing the supernatant, and then its OD600 was measured. The remaining biofilm was washed once with PBS (1 mL) before the addition of 0.03% crystal violet stain made from a 1:10 dilution of Gram Crystal Violet (BD) in PBS. The plates were incubated on the bench for 20 min before the wells were washed twice with PBS (1 mL each). Photographs of the stained biofilms were obtained using a digital camera. The amount of the remaining crystal violet stained biofilm was quantified by adding 1 mL of 100% ethanol followed by measuring OD550 of the homogenized suspension. OD measurements of control
wells where no bacteria were added at the beginning of the experiment were subtracted from all values (i.e., OD600 = 0.03 and OD550 = 0.10). Cell Viability Assays. P. aeruginosa PAO1 was used to characterize the antibacterial effect of CO-releasing polymers on biofilm formation. Biofilms were grown as previously described46,47 with some modifications. Briefly in all assays, overnight cultures in LB medium were diluted to an OD600 of 0.005 in 1 mL of M9 minimal medium in tissue-culture treated 24-well plates (BD). The plates were incubated at 37 °C with shaking at 180 rpm in an orbital shaker-incubator that does not stop agitation when the door is opened, and the biofilms were allowed to grow for 6 h without any disruption. At this time, various treatments, including CORM-2, CO-releasing, and non-CO polymers at a final concentration of 100 μM, were added to the wells. Each treatment was added from a 10 μL aliquot of a stock solution at the appropriate concentration of the compound dissolved in either DMSO (for CORM-2) or PBS and previously sterilized by passing through a 0.22 μm pore size filter. The plates were incubated for a further 1 or 2 h before the biomass or viability was quantified of both planktonic and biofilm bacteria. For biomass quantification, the planktonic phase in the supernatant was removed and the OD600 measured by using a microtiter plate reader (Wallac Victor2, PerkinElmer). Biofilm biomass was measured by crystal violet staining. The biofilm on the well surfaces was first washed once with PBS (1 mL) before the addition of 0.03% crystal violet stain made from a 1:10 dilution of Gram Crystal Violet (BD) in PBS. The plates were incubated on the bench for 20 min before the wells were washed twice with PBS. Photographs of the stained biofilms were obtained using a digital camera. The amount of the remaining crystal violet stained biofilm was quantified by adding 1 mL of 100% ethanol followed by measuring OD550 of the homogenized suspension. OD measurements of control wells where no bacteria were added at the beginning of the experiment were subtracted from all values (i.e., OD600 = 0.03 and OD550 = 0.10). For viability measurements, the BacTiter-Glo Microbial Cell Viability Assay kit (Promega, Alexandria, Australia) was used. The measurement is based on the quantitation of the ATP present in the bacteria by using a thermostable luciferase. After the final 1 or 2 h of incubation with various treatments, the planktonic solution was directly mixed with BacTiter-Glo reagent. After 5 min, luminescence was measured by using a multimode microtiter plate reader (Wallac D
DOI: 10.1021/acs.biomac.5b00716 Biomacromolecules XXXX, XXX, XXX−XXX
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Figure 1. 1H NMR spectra of P(OEGA), P(OEGA)-b-P(VBC), P(OEGA)-b-P(VBGlu), and P(OEGA)-b-P(VBGlu-Ru com) (1) recorded in DMSO deuterated using 300 MHz. Victor2, PerkinElmer). To measure the viability of biofilm bacteria, biofilms on the interior surfaces of the wells were first washed twice with PBS before being resuspended and homogenized in PBS by incubating in an ultrasonication bath (150 W, 40 kHz; Unisonics, Australia) for 20 min. This resuspension method is used similarly for analyzing colony-forming units (CFU) from biofilms.47 Resuspended biofilm cells were then mixed with BacTiter-Glo reagent and their viability quantified by luminescence measurement. Statistical analysis: all assays included four replicates and were repeated in two independent experiments. Statistical analyses were
performed with GraphPad Prism 6 (GraphPad Software) using twoway analysis of variance (ANOVA) followed by Dunnett’s multiple comparison test comparing treated samples to the control.
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RESULTS AND DISCUSSION Synthesis and Characterization of CORM-Conjugated Polymers. The approach for the synthesis of CORMconjugated polymers is presented in Scheme 2. The synthesis of poly(oligoethylene glycol methyl ether acrylate)-blockE
DOI: 10.1021/acs.biomac.5b00716 Biomacromolecules XXXX, XXX, XXX−XXX
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Figure 2. ATR-FTIR spectra of polymers: P(OEGA), P(OEGA)-b-P(VBC), P(OEGA)-b-P(VBGlu), and P(OEGA)-b-P(VBGlu-Ru com) (1).
nethiol hydrochloride, by nucleophilic substitution in DMSO in the presence of triethylamine (TEA) as catalyst at 50 °C for 24 h. The mixture was then purified by dialysis against water and methanol, which resulted in P(OEGA)-b-P(VBGlu), P(OEGA)-b-P(VBGly), and P(OEGA)-b-P(VBAmine). The successful modification of 1-thio-β-D-glucose sodium salt was confirmed via 1H NMR by the presence of new signals assigned to the glycosidic anomeric proton at 4.6 ppm and hydroxyl groups from 4.96−5.12 ppm (Figure 1). In addition, ATR-FTIR spectroscopy revealed the presence of hydroxyl groups from 3300−3700 cm−1 (Figure 2). 1H NMR analysis also confirmed the successful incorporation of 1-thioglycerol and 2-diethylaminoethanethiol hydrochloride into the copolymer P(OEGA)-b-P(VBC) by the absence of −CH2Cl signal at 4.6 ppm (Supporting Information, Figure S2). The thiol ether-functionalized block copolymers were then reacted with tricarbonyldichlororuthenium(II) dimer (CORM2) in anhydrous methanol. The reactions were stirred for 24 h at room temperature under a nitrogen atmosphere in the absence of light. The reaction mediums were precipitated in diethyl ether, which resulted in P(OEGA)-b-P(VBGlu-Ru com) (1), P(OEGA)-b-P(VBGly-Ru com) (2), and P(OEGA)-bP(VBAmine-Ru com) (3). CORM-conjugated polymers 1−3 were stored at 4 °C for further studies. ATR-FTIR analysis was used to confirm the presence of a CO peak in the block copolymer by the presence of signals in the region between 1800 and 2100 cm−1 (Figure 2). ICP-OES technique was used to determine the Ruthenium content (wt %) in polymers (Supporting Information; Table 1). The amounts determined by ICP-OES were in good agreement with the theoretical values, which confirm the high yield of this reaction. Subsequently, we decided to test the solubility of these polymers in water. P(OEGA)-b-P(VBGlu-Ru com) (1), P(OEGA)-b-P(VBGly-Ru com) (2), and P(OEGA)-b-P(VBAmine-Ru com) (3) were easily dissolved in water to yield translucent solutions (Supporting Information, Figure S5). The solutions were also analyzed by DLS (Supporting Information, Figure S6 and Table S2), which showed a size distribution (in number) lower than 10 nm for the three polymers, confirming that the polymers are fully soluble in water and do not form aggregates.
poly(vinylbenzyl chloride) copolymers (P(OEGA)-b-P(VBC) was performed via reversible addition−fragmentation chain transfer polymerization.48,49 A P(OEGA) macro-chain transfer agent was first prepared using n-butyltrithiocarbonate isopropionate as the chain transfer agent and AIBN as the radical initiator with the ratio of [OEGA]0/[RAFT]0/[AIBN]0 = 100:1:0.1 in toluene at 70 °C. The conversion of the monomer was calculated using 1H NMR spectroscopy by comparing the intensity of vinyl proton peaks (at 5.6 and 6.1 ppm) and ester −OCH2 proton peaks (at 4.1 ppm). The resulting polymer was purified by precipitation in a mixture of petroleum spirit and diethyl ether (30:70 v/v). SEC analysis confirmed the synthesis of P(OEGA) with a narrow molecular weight distribution (PDI = 1.25) and an average molecular weight (Mn, SEC) of 21 700 g/ mol (Supporting Information, Figure S1), which is in good agreement with theoretical values. 1H NMR analysis indicated 76 repeating units of OEGA and Mn, NMR of 36 700 g/mol (Figure 1). The difference between SEC and NMR analyses was attributed to the polystyrene calibration used in the SEC analysis. Subsequently, the P(OEGA) homopolymer was chain extended in the presence of 4-vinylbenzyl chloride (VBC) in acetonitrile for 12 h. After purification by several precipitations, the P(OEGA)-b-P(VBC) copolymer was characterized by SEC, 1 H NMR, and ATR-FTIR. SEC confirmed the shift of the molecular weight distribution toward a higher molecular weight (from 21 700 to 26 600 g/mol) (Supporting Information, Figure S1). 1H NMR and ATR-FTIR spectroscopy also confirmed the incorporation of VBC units by the presence of characteristic signals at 4.6 ppm (−CH2Cl), 6.5−7.5 ppm (aromatic region), and 690 cm−1 (−CH2Cl), respectively (Figures 1 and 2). The number of repeating units of VBC in the copolymer was determined by 1H NMR analysis at 10 units by comparing the unchanged signal of −OCH2 of P(OEGA) at 4.1 ppm with the signal of −CH2Cl of VBC at 4.6 ppm using the following equation: DPnVBC = (I4.6 ppm/I4.1 ppm) × DPnOEGA, where I4.6 ppm and I4.1 ppm correspond to integrations of signal at 4.6 and 4.1 ppm, respectively. Then, the benzyl chloride group from P(OEGA)-b-P(VBC) was reacted with different thiol compounds, including 1-thio-βD-glucose sodium salt, 1-thioglycerol, and 2-diethylaminoethaF
DOI: 10.1021/acs.biomac.5b00716 Biomacromolecules XXXX, XXX, XXX−XXX
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Interestingly, the release rate depends on the nature of the thiol employed for the synthesis, as the polymers have a half-life ranging from 40−180 min (Table 2). More importantly, the
Table 1. Characteristics of CO-Releasing Polymers Determined Using SEC and 1H-NMR Analyses polymers P(OEGA) P(OEGA)-b-P(VBC) P(OEGA)-b-P(VBGlu) P(OEGA)-b-P(VBGly) P(OEGA)-b-P(VBAmine) P(OEGA)-b-P(VBGlu-Ru com) (1) P(OEGA)-b-P(VBGly-Ru com) (2) P(OEGA)-b-P(VBAmine-Ru com) (3)
Mn, NMR (g/mol)
Mn, SEC (g/mol)
PDI
36 700 38 200 40 700 40 000 39 500 44 200
21 700 26 600 39 700 24 000 23 000 40 800
1.25 1.25 1.34 1.20 1.19 1.24
44 200
32 200
1.50
44 500
41 300
1.76
Table 2. Number of CO Equivalents Released Per VBC Unit and the Half-Life of CO Releasea polymer
equivalents of CO
t1/2 (min)
P(OEGA)-b-P(VBGlu-Ru com) (1) P(OEGA)-b-P(VBGly-Ru com) (2) P(OEGA)-b-P(VBAmine-Ru com) (3)
0.80 0.84 0.78
∼180 ∼40 ∼90
a
Experiments done in duplicate using 24 mg/mL of sodium dithionite (Na2S2O4).
release rates for polymers (1−3) are slower than CORMs (halflives of CORM-353 and ALF49231 in vitro are approximately 1 min and less than 5 min, respectively), which clearly demonstrates the advantage of our approach. Indeed, we extended the half-life of CO release by a factor of 40−180 in comparison with CORM-3 and ALF492. This short half-life of small CORMs is a determining limiting factor for its bioapplication and future applications in clinic. To compare the half-lives between CO-releasing polymers and CORMS, we tried to synthesize the corresponding CORMs compounds obtained in the polymer via a nucleophilic substitution of benzyl chloride with thiol compounds, and then, the thiol ether compounds were reacted in the presence of CORM-2. However, these compounds were not soluble in water, and we could not perform further investigations. Indeed, to solubilize these compounds, they require the addition of large amount of solvents (DMSO), which could result by a significant change in the CO release rate. The differences in the half-life of CO-releasing polymers 1−3 can be attributed to different structures of thiol moiety conjugated to polymers. Among CO-releasing polymers, polymer 1 has the longest half-life, which is due to the presence of glucose moieties. The release of CO is provoked by a ligand exchange between CO and H2O molecules. This release rate depends strongly of water accessibility to Ruthenium complex, as previously demonstrated by Stupp and co-workers.52,54 In these studies, Stupp and co-workers functionalize polymeric fiber with CORM. After gelation of fiber, the author show more than eight-fold increase half-life
Study of CO Release by Myoglobin Assay. The release of CO from polymers 1−3 was investigated using the myoglobin assay as previously reported.50 In this assay, CORM-conjugated polymer is added to a solution of reduced deoxy-myoglobin (Deoxy-Mb), which can trap CO to form carbonmonoxymyoglobin (MbCO). Deoxy-Mb shows a characteristic absorption maximum at 557 nm, and there are new absorptions at 540 and 577 nm from MbCO. Therefore, using these signals, CO release was monitored by UV−vis spectroscopy (Figure 3). The myoglobin assay confirmed that polymers 1−3 all spontaneously release CO in buffer solution in the presence of sodium dithionite. The total amount of released CO is close to the theoretical values and was calculated using the number of VBC units present in the copolymer. Each unit of VBC can carry one molecule of CORM-2 and can release one CO molecule. The release rate of CO can depend on the amount of reducing agent, that is, sodium dithionite, used in the assay.29,51 We have performed the assay using a standard concentration of sodium dithionite (i.e., 24 mg/mL of buffer) described in previous publications.42,52 To confirm that sodium dithionite could affect the release rate, we varied the concentration of sodium dithionite from 20 mg/mL to 29 mg/mL and showed that the release could be slightly accelerated by an increase of sodium dithionite (Supporting Information, Figures S7 and S8).
Figure 3. (A) UV−vis spectra of a reduced horse skeletal muscle myoglobin solution in the presence of polymer 2 (20 μM) in phosphate buffer solution. (B) CO release from CO-releasing polymers 1−3 (20 μM) versus time in reduced myoglobin solution (the amount of CO was normalized per VBC unit, experiments done in duplicate, using 24 mg/mL of sodium dithionite (Na2S2O4). G
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Figure 4. Effect of P(OEGA)-b-P(Gly-CO) (2) on P. aeruginosa PAO1 growth and biofilm formation versus different concentrations of CO releasing CORM and P(OEGA)-b-P(VBGly-Ru com) (2). Note: bacterial biofilms were grown in multiwell plates for up to 7.5 h in the presence of P(OEGA)-b-P(VBGly-Ru com) (2) and CORM-2 from the beginning of growth. Planktonic biomass was determined by measuring the OD600 of the supernatant; biofilm biomass was determined by crystal violet staining (OD550). Error bars represent standard error (n = 2).
result is attributed to the sustained release of CO by the polymer in comparison to CORM-2. P(OEGA)-b-P(VBGlu-Ru com) (1) also affected biofilm formation, although its effect was less significant than that of the other two CO-releasing polymers (2 and 3). At 100 and 200 μM, P(OEGA)-bP(VBGlu-Ru com) (1) induced a 70% and 85% decrease in biofilm formation, respectively, compared to the untreated controls (Supporting Information, Figure S9). On the basis of its growth inhibition and biofilm prevention effects, P(OEGA)b-P(VBGly-Ru com) (2) was selected for further bactericidal study on pre-established P. aeruginosa biofilms. Bactericidal Effect of CO-Releasing Polymers on P. aeruginosa PAO1 Cell Viability. To further examine the effect of CO-releasing polymers on planktonic and biofilm viability of PAO1, the bactericidal activity of P(OEGA)-bP(VBGly-Ru com) (2) was tested against the pre-established biofilms. The bacterial biofilms were grown for a 6 h period and then were exposed to various additional treatments for 1 and 2 h. The CO releasing molecule, CORM-2, and the non-CO releasing polymers, P(OEGA)-b-P(VBGly), were used as positive and negative controls, respectively. The viability of planktonic and biofilm cells was determined by the Bactiter-Glo Microbial Cell Viability assay, in which the viability of the culture was assessed by measuring the ATP content of the cells. At 100 μM, the non-CO polymer P(OEGA)-b-P(VBGly) had no effect on the planktonic and biofilm viability as expected, but instead promoted a slight growth of the bacteria during the treatments. In contrast, both CORM-2 and the COreleasing polymer P(OEGA)-b-P(VBGly-Ru) (2) at 100 μM showed significant decrease in the viability of the bacteria for both planktonic and biofilm cells, thus clearly demonstrating the antibacterial activity of those compounds is dependent on the CO release. However, the CO-releasing polymer P(OEGA)-b-P(VBGly-Ru) (2) presented a higher efficiency to inhibit bacterial growth and prevent biofilm formation in comparison with CO-releasing molecule CORM-2. This result is attributed to the high CO-loading capacity of the polymer and the sustainable delivery of CO using polymers. Specifically, P(OEGA)-b-P(VBGly-Ru com) (2) showed an almost complete eradication of both planktonic and biofilm viable cells by 94% and 83% (P < 0.0001), respectively, after 2 h of treatment. The number of viable cells was only reduced by approximately 60% for both planktonic and biofilm cells after exposure to CORM-2 compared to untreated sample (Figure 5). Interestingly, there was no significant reduction in the
compared to the half-lives of both CORM-3 and PA 2 in solution. In our case, the steric hindrance around Ruthenium caused by the polymer and thiol compound could limit the accessibility and diffusion of water molecules, which results by a significant decrease of CO release rate. CO-Releasing Polymers Inhibit Bacterial Growth and Prevent Biofilm Formation. P. aeruginosa is responsible for 30% of nosocomial infections in intensive care units, which is due to the increase of its resistance against antibiotics.23,55 This situation motivated us to test our polymers against P. aeruginosa. The antibacterial activity of the three types of CO-releasing polymers was first examined by the ability to inhibit the growth of P. aeruginosa PAO1. The commercial carbon monoxide releasing molecule CORM-2, which has previously been shown to exhibit antibacterial properties against P. aeruginosa PAO1,56,57 was used as a positive control. The bacteria were grown in minimal M9 medium in the presence of 100 μM CORM-2 and 10−200 μM of the COreleasing polymers for a 7.5 h period. All three CO-releasing polymers showed strong inhibition of the bacterial growth at 100 μM, with a greater than 90% reduction in planktonic biomass compared to untreated culture wells over the incubation period (Figure 4 and Supporting Information, Figure S9). This result was comparable to that induced by the positive control using CORM-2. In contrast, thiol-conjugated polymers (non-CO releasing polymers used as a negative control) did not affect the growth of P. aeruginosa PAO1 at any testing concentration (Supporting Information, Figure S10), indicating that the polymeric carriers were not toxic, and the inhibition of growth was mainly due to CO released from the polymers. Among the three CO-releasing polymers, P(OEGA)b-P(VBGly-Ru com) (2) polymer was found to have the strongest effect on cell growth. At 50 μM, P(OEGA)-bP(VBGly-Ru com) (2) caused a 93% decrease in planktonic biomass, while P(OEGA)-b-P(VBAmine-Ru com) (3) displayed an approximately 84% reduction in planktonic biomass versus untreated controls. The effect on P. aeruginosa biofilm formation by COreleasing polymers was then assessed by crystal violet (CV) staining at OD550. At 50 μM, P(OEGA)-b-P(VBGly-Ru com) (2) and P(OEGA)-b-P(VBAmine-Ru com) (3) prevented biofilm formation over the incubation period, with a 95% reduction in biofilm biomass after 7.5 h compared to the untreated biofilms. This is slightly higher than that induced by CORM-2 at 100 μM (92% reduction in biofilm biomass). This H
DOI: 10.1021/acs.biomac.5b00716 Biomacromolecules XXXX, XXX, XXX−XXX
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Figure 5. Effect of P(OEGA)-b-P(VBGly-Ru com) (2) on P. aeruginosa PAO1 viability. Biofilms were grown in multiwell plates for 6 h and then treated further for 1 or 2 h in the presence or absence of 100 μM CORM-2, P(OEGA)-b-P(VBGly), or P(OEGA)-b-P(VBGly-Ru com) (2) before analysis of planktonic (left) and biofilm viability (right) by measuring the ATP content of bacteria. Error bars represent standard error (n = 4). Asterisks indicate statistically significant difference of treatments versus untreated culture (∗∗∗∗, P < 0.0001).
Figure 6. Effect of P(OEGA)-b-P(VBGly-Ru com) (2) on P. aeruginosa PAO1 biofilm formation. Biofilms were grown in multiwell plates for 6 h and then treated for 1 h in the presence of 100 μM of CORM-2, P(OEGA)-b-P(VBGly), or P(OEGA)-b-P(VBGly-Ru com) (2). (A) Planktonic biomass was determined by measuring the OD600 of the supernatant; biofilm biomass was determined by CV staining (OD550). (B) Stained biofilms treated with the indicated concentrations of CORM-2, P(OEGA)-b-P(VBGly), and P(OEGA)-b-P(VBGly-Ru com) (2). Error bars represent standard error (n = 4). Asterisks indicate statistically significant difference of treatments versus untreated culture (ns, not significant; ∗∗∗∗, P < 0.0001).
untreated controls. This result is in accord with those obtained in the viability assay using the Bactiter-Glo reagent (Figure 5). The decrease in biofilm biomass when cultures were treated by CO releasing compounds was not significantly different in comparison without treatment. At 100 μM, P(OEGA)-bP(VBGly-Ru com) (2) inoculated wells showed only an 11% reduction in biofilm biomass compared to the untreated controls (Figure 6A). Longer treatment with P(OEGA)-bP(VBGly-Ru com) (2) did not improve the decrease in biofilm staining by CV; rather, more staining was observed versus the untreated wells (Supporting Information, Figure S11). This result is attributed to the fact that both live and dead bacteria are stained by CV. While CV-stained biofilm images of the untreated control showed evenly distributed staining, wells treated with CORM-2 and P(OEGA)-b-P(VBGly-Ru com) (2) show some areas without staining (Figure 6B). The results of the viability and CV staining assays confirm the strong antibacterial property of CO polymers, but these polymers are not able to disperse biofilm.
viability of the bacteria between 1 and 2 h of treatment with CORM-2. However, a longer treatment of the CO-releasing polymer P(OEGA)-b-P(VBGly-Ru) (2) resulted in a prolonged inhibition of cell growth. This result could be attributed to the fact that the CO-releasing polymer has longer half-life in comparison with CORM-2. In addition to the antibacterial activity of CO, the ability of CO-releasing polymers to disperse biofilms was assessed on the biofilm-forming model organism P. aeruginosa PAO1. Preestablished biofilms were grown for a 6 h period and then were exposed to various treatments. Planktonic biomass was quantified by measuring the OD600, and biofilm biomass was determined by CV staining followed by a measurement at OD550. After 1 h of treatment, the polymers without COreleasing moieties had no effect on the planktonic, but showed a slight increase in the number of planktonic cells, suggesting their nontoxicity (Figure 6A). In contrast, P(OEGA)-bP(VBGly-Ru com) (2) and CORM-2 induced decreases in planktonic growth of 30% and 25%, respectively, relative to the I
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(7) Desmard, M.; Foresti, R.; Morin, D.; Dagouassat, M.; Berdeaux, A.; Denamur, E.; Crook, S. H.; Mann, B. E.; Scapens, D.; Montravers, P.; Boczkowski, J.; Motterlini, R. Antioxid. Redox Signaling 2012, 16, 153−163. (8) Davidge, K. S.; Motterlini, R.; Mann, B. E.; Wilson, J. L.; Poole, R. K. In Advances in Microbial Physiology; Robert, K. P., Ed.; Academic Press: Waltham, MA, 2009; Vol. 56, pp 85−167. (9) Motterlini, R.; Otterbein, L. E. Nat. Rev. Drug Discovery 2010, 9, 728−743. (10) Lu, Y.; Slomberg, D. L.; Schoenfisch, M. H. Biomaterials 2014, 35, 1716−1724. (11) Lu, Y.; Slomberg, D. L.; Shah, A.; Schoenfisch, M. H. Biomacromolecules 2013, 14, 3589−3598. (12) Parr, S. R.; Wilson, M. T.; Greenwood, C. Biochem. J. 1975, 151, 51−59. (13) Otterbein, L. E.; May, A.; Chin, B. Y. Cell. Mol. Biol. 2005, 51, 425−432. (14) Chung, S. W.; Liu, X.; Macias, A. A.; Baron, R. M.; Perrella, M. A. J. Clin. Invest. 2008, 118, 239−247. (15) Wegiel, B.; Larsen, R.; Gallo, D.; Chin, B. Y.; Harris, C.; Mannam, P.; Kaczmarek, E.; Lee, P. J.; Zuckerbraun, B. S.; Flavell, R.; Soares, M. P.; Otterbein, L. E. J. Clin. Invest. 2014, 124, 4926−4940. (16) Heinemann, S. H.; Hoshi, T.; Westerhausen, M.; Schiller, A. Chem. Commun. 2014, 50, 3644−3660. (17) Gonzalez, M. A.; Yim, M. A.; Cheng, S.; Moyes, A.; Hobbs, A. J.; Mascharak, P. K. Inorg. Chem. 2012, 51, 601−608. (18) Schatzschneider, U. Inorg. Chim. Acta 2011, 374, 19−23. (19) Zobi, F. Future Med. Chem., 20135, 175−188.10.4155/ fmc.12.196 (20) Chakraborty, I.; Carrington, S. J.; Mascharak, P. K. Acc. Chem. Res. 2014, 47, 2603−2611. (21) Pierri, A. E.; Pallaoro, A.; Wu, G.; Ford, P. C. J. Am. Chem. Soc. 2012, 134, 18197−18200. (22) Schatzschneider, U. Br. J. Pharmacol. 2015, 172, 1638−1650. (23) Desmard, M.; Davidge, K. S.; Bouvet, O.; Morin, D.; Roux, D.; Foresti, R.; Ricard, J. D.; Denamur, E.; Poole, R. K.; Montravers, P.; Motterlini, R.; Boczkowski, J. FASEB J. 2009, 23, 1023−1031. (24) Davidge, K. S.; Sanguinetti, G.; Yee, C. H.; Cox, A. G.; McLeod, C. W.; Monk, C. E.; Mann, B. E.; Motterlini, R.; Poole, R. K. J. Biol. Chem. 2009, 284, 4516−4524. (25) Nobre, L. S.; Seixas, J. D.; Romao, C. C.; Saraiva, L. M. Antimicrob. Agents Chemother. 2007, 51, 4303−4307. (26) Desmard, M.; Davidge, K. S.; Bouvet, O.; Morin, D.; Roux, D.; Foresti, R.; Ricard, J. D.; Denamur, E.; Poole, R. K.; Montravers, P.; Motterlini, R.; Boczkowski, J. FASEB J. 2009, 23, 1023−1031. (27) Motterlini, R.; Mann, B. E.; Foresti, R. Expert Opin. Invest. Drugs 2005, 14, 1305−1318. (28) Nagel, C.; McLean, S.; Poole, R. K.; Braunschweig, H.; Kramer, T.; Schatzschneider, U. Dalton Trans. 2014, 43, 9986−9997. (29) Wilson, J. L.; Jesse, H. E.; Hughes, B.; Lund, V.; Naylor, K.; Davidge, K. S.; Cook, G. M.; Mann, B. E.; Poole, R. K. Antioxid. Redox Signaling 2013, 19, 497−509. (30) Murray, T. S.; Okegbe, C.; Gao, Y.; Kazmierczak, B. I.; Motterlini, R.; Dietrich, L. E. P.; Bruscia, E. M. PLoS One 2012, 7, e35499. (31) Pena, A. C.; Penacho, N.; Mancio-Silva, L.; Neres, R.; Seixas, J. D.; Fernandes, A. C.; Romao, C. C.; Mota, M. M.; Bernardes, G. J.; Pamplona, A. Antimicrob. Agents Chemother. 2012, 56, 1281−1290. (32) Gonzales, M. A.; Han, H.; Moyes, A.; Radinos, A.; Hobbs, A. J.; Coombs, N.; Oliver, S. R. J.; Mascharak, P. K. J. Mater. Chem. B 2014, 2, 2107−2113. (33) Pierri, A. E.; Huang, P.-J.; Garcia, J. V.; Stanfill, J. G.; Chui, M.; Wu, G.; Zheng, N.; Ford, P. C. Chem. Commun. 2015, 51, 2072−2075. (34) Dordelmann, G.; Meinhardt, T.; Sowik, T.; Krueger, A.; Schatzschneider, U. Chem. Commun. 2012, 48, 11528−11530. (35) Dördelmann, G.; Pfeiffer, H.; Birkner, A.; Schatzschneider, U. Inorg. Chem. 2011, 50, 4362−4367. (36) De Jong, W. H.; Borm, P. J. A. Int. J. Nanomed. 2008, 3, 133− 149.
CONCLUSION In this work, three CORM-conjugated polymers were designed and synthesized for the sustainable delivery of CO. In comparison with small CO releasing molecules, these polymers present several advantages. First, CO releasing polymers present a good solubility in water and biological media, which is beneficial for bioapplications. Many CORMs, including CORM-2 and others, are not soluble in water and require the use of organic solvent such as DMSO. Second, the release rate of CO from these polymers is slower than that of CORM compounds. The polymer architecture reduces the water accessibility, which allows it to significantly extend the halflife of these compounds. Because of this sustainable release of CO, CO-releasing polymers display an higher inhibition planktonic growth and prevent biofilm formation against P. aeruginosa in comparison with CORM-2. The prepared COreleasing polymers hold considerable promise for targeted antimicrobial applications. In summary, the encapsulation or conjugation of CORMs into polymeric structure could allow to enhance their stability and significantly improve their water solubility. Further studies are in progress to investigate the effect of CO on the formation and the development of biofilms after treatment with CO-releasing polymers.
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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/acs.biomac.5b00716. Experimental procedure, NMR spectra, SEC traces, FTIR, and UV−vis myoglobin assay (PDF)
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AUTHOR INFORMATION
Corresponding Author
*E-mail:
[email protected]. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS C.B. acknowledges the Australian Research Council for his Future Fellowship (FT1200096) and DVCR Prof. Les Field UNSW−Australia for internal funding (SPF01). D.N. was supported by the Australia Awards Scholarship.
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REFERENCES
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K
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