Article pubs.acs.org/Biomac
Antimicrobial Polycarbonates: Investigating the Impact of Balancing Charge and Hydrophobicity Using a Same-Centered Polymer Approach Amanda C. Engler,†,§ Jeremy P. K. Tan,‡,§ Zhan Yuin Ong,‡ Daniel J. Coady,† Victor W. L. Ng,‡ Yi Yan Yang,*,‡ and James L. Hedrick*,† †
IBM Almaden Research Center, 650 Harry Road, San Jose, California 95120, United States Institute of Bioengineering and Nanotechnology, 31 Biopolis Way, Singapore 138669, Singapore
‡
S Supporting Information *
ABSTRACT: Biodegradable antimicrobial polymers are a promising solution for combating drug resistant microbes. When designing these materials, the balance between charge and hydrophobicity significantly affects the antimicrobial activity and selectivity toward microbes over mammalian cells. Furthermore, where the charge and hydrophobicity is located on the molecules has also proven to be significant. A series of antimicrobial homopolymer polycarbonates were synthesized, where the hydrophobic/hydrophilic balance was controlled by varying the spacer between the charged quaternary ammonium moiety and the polymer backbone (a “samecentered” structure where the hydrophobic moiety is directly attached to the charged moiety). These homopolymers were active against all microbes tested but depending on the spacer length some hemolytic activity was observed. To reduce the polymer hemolytic activity we systematically varied the polymer composition by copolymerizing the different monomers used in the “same center” homopolymers. By maintaining charge on each repeat unit but copolymerizing monomers having varied hydrophobic side chain lengths, polymers with high activity and selectivity were achieved. In addition, these macromolecules act via a membrane disruption mechanism, making them less likely to induce resistance.
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INTRODUCTION
Similar to AMPs, antimicrobial polymers have been designed to kill pathogens via a membrane disruption mechanism. When designing antimicrobial polymers, sufficient cationic charge must be incorporated into the macromolecule to promote adhesion to the microbe cell envelope and a hydrophobic moiety should be incorporated that will attach onto or integrate into the cellular membrane for lysing the membrane.10,11 These materials must selectively target and kill microbes without imparting toxicity on mammalian cells. There are many factors that influence antimicrobial activity and selectivity. However, hydrophilic/hydrophobic balance (amphiphilicity) of an antimicrobial polymer is particularly important because it significantly impacts how the polymer interacts with cellular membranes.10 When altering the amphiphilicity of antimicrobial polymers, there are several different approaches that are typically used by polymer chemists. In a recent review article, three different approaches were defined: segregated monomer, facially amphiphilic polymer, and same-centered polymer.11 In the “segregated monomer” approach, a relatively nonpolar monomer is randomly polymerized with a cationic monomer to create a statistical copolymer12 or polymerized in sequence
Antibiotics play an important role in healthcare and medical fields. However, growing emergence of antibiotic resistant bacteria is of great concern.1 For example Pseudomonas aeruginosa, a Gram-negative bacteria is a major cause of infections, causing 10 to 20% of hospital-acquired infections2 and has a naturally high level of antibiotic resistance due to restricted outer membrane permeability combined with multidrug efflux systems.3,4 Antimicrobial peptides (AMPs) are insensitive to drug efflux systems and are a promising alternative to small molecule antibiotics.4 These peptides kill a broad range of pathogenic microbes, including Gram-positive bacteria, Gram-negative bacteria and fungi.5 A common attribute of AMPs is that they are amphiphilic, containing cationic amino acids and amino acids with a hydrophobic side chain. They attack directly on microbial membranes using electrostatic interaction between cationic charge of AMPs and anionic charge on the membrane surface and disrupt the membrane by insertion of hydrophobic components into membrane lipid domains. The physical nature of the membrane disruption mechanism reduces the possibility of microbes developing resistance.6−8 Despite their therapeutic potential, applicability of AMPs is limited because of their cytotoxicity (e.g., hemolysis), short half-life in vivo (labile to proteases), and high manufacturing cost.9 © XXXX American Chemical Society
Received: August 19, 2013 Revised: October 22, 2013
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to create a block copolymer.13 The amphiphilicity is controlled by varying the monomer feed ratio. In the “facially amphiphilic” polymer approach, each polymer repeat unit has a nonpolar section and a separate cationic charged section.12,14 The amphiphilicity is controlled by varying the nonpolar section. In “same-centered” polymers, a hydrophobic alkyl chain is directly attached to a positively charged moiety; that is, the center is the atom that contains positive charge and has the hydrophobic group directly attached. Alkyl chain length is varied to adjust the amphiphilicity.15,16 All three of these strategies result in efficacious antimicrobial polymers. For some polymer backbones, one strategy often results in polymers with higher activity and better selectivity. Recently, we introduced biodegradable antimicrobial polycarbonates that self-assemble to form cationic micelles. While in micelle form, these structures have an increased local concentration of cationic charge leading to enhanced interactions with negatively charged microbe cell walls/ membranes, enhancing their antimicrobial activities and selectivity. Two different systems have been investigated (Scheme 1), both prepared via a segregated monomer
segregated monomer approach and further improvement in antimicrobial efficacy, we synthesized homopolymers with high antimicrobial activity using a same-centered approach. We hypothesized that these polymers would not need to be in the micelle form to be active. The hydrophobic/hydrophilic balance was controlled by varying the spacer between the charged quaternary ammonium moiety and the polymer backbone, as shown in Scheme 2a. We anticipated that these polymers would act through a membrane disruption mechanism where charged quaternary ammonium groups interact with the negative charge on the membrane lipids and the side chains snorkel (extend) into the acyl chain region of the lipid bilayer, also known as a snorkeling mechanism.19 To reduce polymer toxicity, we systematically varied the polymer composition by copolymerizing different monomers used in the “same center” homopolymers, as shown in Scheme 2b. By maintaining charge on each repeat unit and copolymerizing monomers with varied hydrophobic side chain lengths, we sought to achieve highly selective polymers with broad spectrum activity. CMC measurements were also performed to determine if the polymers were self-assembled when active, as with our previous systems. In addition, the antimicrobial mechanism of action was explored by visualizing the bacteria’s cell wall/membrane before and after antimicrobial polymer treatment.
Scheme 1. Antimicrobial Polycarbonates Obtained via a Segregated Monomer Approach
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EXPERIMENTAL SECTION
Materials. MTC-OC6F5 was obtained from Central Glass and used as received. The compounds 1-(3,5-bis(trifluoromethyl)phenyl)-3cyclohexyl-2-thiourea and benzyl 2,2-bis(methylol)propionate were prepared according to literature protocol.20 Dichloromethane was dried using activated alumina columns and stored over molecular sieves (3 Å). All other materials were used as received. Trypic soy broth (TSB) and yeast media broth (YMB) powder were purchased from BD Diagnostics (Singapore) and used to prepare the microbial broths according to the manufacturer’s instruction. MTC-OctCl and MTC-HexCl. These compounds were produced using known literature procedures.1 A flask was charged with MTCOC6F5 (5.9 g, 18.2 mmol), 8-chloro-1-octanol (2.5 g, 15.2 mmol), proton sponge (3.3 g, 15.2 mmol), and THF (8 mL). The reaction mixture was stirred for 12 h and excess ammonium acetate was added. After an additional 3 h of stirring, the reaction mixture was added directly to a silica gel column. The product was isolated using hexane/ ethyl acetate as the eluent to yield an oil (4.3 g, 71%). MTC-OctCl 1H NMR (400 MHz, CDCl3, 22 °C): δ 4.72 (d, 2H, -CH2OCOO), 4.23 (d, t 4H, -CH2OCOO, -OCH2CH2), 3.57 (t, 2H, -OCH2CH2(CH2)3CH2CH2CH2Cl), 1.84 (m, 2H, -OCH2CH2(CH2)3CH2CH2CH2Cl), 1.76 (m, 2H, -OCH2CH2(CH2)3CH2CH2CH2Cl), 1.45 (m, 2H, -OCH 2 CH 2 (CH 2 ) 3 CH 2 CH 2 CH 2 Cl), 1.35 (br m, 9H, -CH 3, OCH2CH2(CH2)3CH2CH2CH2Cl). MTC-HexCl 1H NMR (400 MHz, CDCl3, 22 °C): δ 4.72 (d, 2H, -CH2OCOO), 4.23 (d, t 4H, -CH2OCOO, -OCH2CH2), 3.57 (t, 2H, -OCH2CH2CH2CH2CH2CH2Cl), 1.84 (m, 2H, -OCH2CH2CH2CH2CH2CH2Cl), 1.76 (m, 2H, -OCH2CH2CH2CH2CH2CH2Cl), 1.45 (m, 2H, -OCH2CH2CH2CH2CH2CH2Cl), 1.39 (m, 2H, -OCH2CH2CH2CH2CH2CH2Cl), 1.35 (s, 3H, -CH3). Representative Polymerization. In a nitrogen-charged glovebox, benzyl 2,2-bis(methylol)propionate initiator (0.01 g, 0.045 mmol), MTC-HexCl monomer (0.38 g, 1.34 mmol), and 1-(3,5-bis(trifluoromethyl)-phenyl)-3-cyclohexyl-2-thiourea (TU) (0.033 g, 0.089 mmol) were dissolved in 0.90 mL of dichloromethane. The compound 1,8-diazabicyclo[5.4.0]undec-7-ene (DBU) (0.014 g, 0.089 mmol) was added and the reaction mixture was stirred at room temperature for 20 min. The polymer was terminated by adding several drops of acetyl chloride. The reaction catalyst was quenched
approach. The first system was a triblock system consisting of a quaternary ammonium polycarbonate (prepared from MTCPrCl) as the center block and poly(trimethylene carbonate) as the outer block (Scheme 1a).17 The polymers self-assembled into micelles and were active against Gram-positive bacteria and fungi at minimum inhibitory concentrations (MICs) higher than the critical micelle concentrations (CMCs). The second system was a dynamic micelle system consisting of random copolymers of MTC-PrCl and MTC-ethyl (Scheme 1b).18 These polymers were active against Gram-positive bacteria, Gram-negative bacteria, and fungi with MICs from 63 to 500 mg/L, which were higher than the polymer CMCs. In both systems, the hydrophobic/hydrophilic balance was accomplished using a cationic monomer (MTC-PrCl, quaternized) and a separate hydrophobic monomer (trimethylene carbonate or MTC-ethyl). This balance also drives self-assembly. Our previously published antimicrobial polycarbonates were made via a “segregated monomer” approach to obtain active antimicrobial agents. In both cases, the polymers acted at concentrations above the CMC, indicating that polymers were in the assembled form when active. For comparison with the B
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Scheme 2. (a) Homopolymers Studied, (b) Statistical Block Copolymers Studied, and (c) Polymer Synthesis
2.5 nm. The intensity (peak height) ratios of I337/I334 from the excitation spectra were analyzed as a function of polymer concentration. The CMC was taken from the intersection between the tangent to the curve at the inflection and tangent of the points at low concentrations. Particle Size and Zeta Potential Determination. The particle size and zeta potential measurements were conducted using the Zetasizer 3000 HAS (Malvern Instrument Ltd., U.K.), which is equipped with a He−Ne laser beam at 658 nm. The particle size and zeta potential of polymers in DI water was averaged over 5 runs. Minimal Inhibitory Concentration (MIC) Measurements. Staphylococcus epidermidis (ATC 12228), Staphylococcus aureus (ATCC 29737), Escherichia coli (ATC 25922), Pseudomonas aeruginosa (ATCC 9027), and Candida albicans (ATCC 10231) obtained from ATCC were reconstituted from its lyophilized form according to the manufacturer’s protocol. Microbial samples were cultured in TSB (bacteria) and YMB (fungi) solution at 37 °C and room temperature, respectively, under constant shaking of 300 rpm. The MICs of the polymers were measured using the broth microdilution method as reported. Briefly, 100 μL of TSB broth containing a polymer (with a fixed DI water concentration of 20% v/v) at various concentrations was placed into each well of a 96-well tissue culture plate. An equal volume of microbial suspension (3 × 105 CFU ml−1) was added into each well. Prior to mixing, the microbial sample was first inoculated overnight to enter its log growth phase. The concentration of microbial solution was adjusted to give an initial optical density (O.D.) reading of approximately 0.07 at 600 nm wavelength on microplate reader (TECAN, Switzerland), which corresponds to the concentration of McFarland Standard No. 1 (3 × 108 CFU ml−1), the microbial solution was further diluted by 1000 to achieve an initial loading of 3 × 105 CFU ml−1. The 96-well plate was kept in an
with an excess of benzoic acid and the crude polymer solution was precipitated into isopropanol, yielding a white solid. 1H NMR (400 MHz, CDCl3, 22 °C): δ 4.3 (br, 4H, CHCH2), 4.15 (t, 2H, -OCH2CH2), 3.57 (t, 2H, -OCH2CH2CH2CH2CH2CH2Cl), 1.80 (m, 2H, -OCH2CH2CH2CH2CH2CH2Cl), 1.65 (m, 2H, -OCH2CH2CH2CH2CH2CH2Cl), 1.47 (m, 2H, -OCH2CH2CH2CH2CH2CH2Cl), 1.39 (m, 2H, -OCH2CH2CH2CH2CH2CH2Cl), 1.27 (s, 3H, - CH3). Quaternization Reaction. PMTC-HexNMe3−polymers were added to a vial and dissolved in MeCN. The vial was then cooled to −78 °C and trimethylamine (TMA) gas was added. The vial was then heated to 50 °C and allowed to stir overnight or until reaction was complete. The reaction mixture was then precipitated into diethyl ether. For further purification, the polymers were dissolved in an acetonitrile/isopropanol mixture and dialyzed. 1H NMR (400 MHz, MeOD, 22 °C): δ 4.3 (br, 4H, CHCH2), 4.16 (t, 2H, -OCH2CH2), 3.38 (br, 2H, -OCH2CH2CH2CH2CH2CH2N), 3.15 (br, 9H, -NCH3), 1.80 (br, 2H, -OCH2CH2CH2CH2CH2CH2N), 1.68 (br, 2H, -OCH2CH2CH2CH2CH2CH2Cl), 1.45 (m, 4H, -OCH2CH2CH2CH2CH2CH2Cl), 1.27 (s, 3H, -CH3). Critical Micelle Concentration (CMC) Determination. The CMC of the polymers in deionized (DI) water was determined using pyrene as a probe. Fluorescence spectra were recorded by a LS 50B luminescence spectrometer (Perkin-Elmer, U.S.A.) at room temperature. Aliquots of pyrene in acetone solution (6.16 × 10−5 M, 10 μL) were added to containers and acetone was left to evaporate. Polymer solutions (1 mL) at varying concentrations were added into the containers and left to equilibrate for 24 h. The final pyrene concentration in each sample was 6.16 × 10−7 M. The excitation spectra were scanned from 300 to 360 nm at an emission wavelength of 395 nm. Both the excitation and emission bandwidths were set at C
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Table 1. Composition and Properties of Antimicrobial Polycarbonates GPC monomers PrCl30 HexCl15 HexCl30 HexCl60 OctCl15 OctCl30 OctCl60 75%OctCl 50%OctCl 25%OctCl 75%OctCl 50%OctCl 25%OctCl a
25%PrCl 50%PrCl 75%PrCl 25%HexCl 50%HexCl 75%HexCl
DP
Mn by NMR
Mn
Mw
PDI
CMC (mg/L)
30 15 30 60 15 30 60 30 30 30 30 30 30
7000 4000 8000 16000 4400 8800 18000 8400 7900 7400 8500 8300 8100
6800 4700 7100 11000 3600 4600 5600 5100 5400 6200 6200 6400 5900
8400 5700 8300 12000 4600 5800 6700 6000 6500 7700 7300 7500 7100
1.23 1.21 1.17 1.17 1.29 1.23 1.21 1.18 1.2 1.24 1.18 1.17 1.21
281.8 125.9 84.1 63.1 79.4 44.7 39.8 56.2 79.4 158.5 50.1 56.2 63.1
Dh (nm) 190 240 246 267 249 195 187 228 232 218 172 209 215
± ± ± ± ± ± ± ± ± ± ± ± ±
20 43 7 11 26 21 26 23 23 23 18 32 23
PDIa 0.43 0.50 0.41 0.49 0.40 0.42 0.41 0.44 0.45 0.43 0.45 0.47 0.46
± ± ± ± ± ± ± ± ± ± ± ± ±
0.02 0.06 0.03 0.01 0.03 0.04 0.03 0.03 0.03 0.03 0.03 0.03 0.03
zeta potential (mV) 53.8 44.1 53.7 63.1 44.2 59.8 64.3 53.9 51.8 59.1 57.0 54.0 55.5
± ± ± ± ± ± ± ± ± ± ± ± ±
3.7 3.4 5.3 3.2 5.7 4.1 3.3 5.7 1.8 4.2 2.7 1.9 4.6
Polydispersity index (PDI) obtained from dynamic light scattering.
Table 2. MIC (mg/L) and HC50 (mg/L) of Homopolymers polymer
S. aureus (Gram +)
S. epidermidis (Gram +)
E. coli (Gram −)
P. aeruginosa (Gram −)
C. albicans (Fungus)
hemolysis HC50
HexCl15 OctCl15 PrCl30 HexCl30 OctCl30 HexCl60 OctCl60
31 4 500 31 4 31 4
4 4 16 4 4 4 4
125 31 1000 125 16 125 16
1000 125 >1000 1000 125 1000 125
250 125 500 250 125 250 125
>1000 250−500 >1000 >1000 250 >1000 125−250
incubator at 37 °C (bacteria) and room temperature (fungi) under constant shaking of 300 rpm for 18 and 42 h, respectively. The MIC was taken as the concentration of the antimicrobial polymer at which no microbial growth was observed with unaided eyes and the microplate reader at the end of 18 h (bacteria) or 42 h (fungi) incubation. Broth containing microbial cells alone was used as negative control, and each test was carried out in 6 replicates. Agar Gel Assay. The microbes were inoculated and prepared according to the same procedure in the MIC measurement described earlier. The samples were treated with the polymer at various concentrations (none, 1/2× MIC, 1× MIC, and 2× MIC) and were incubated at 37 °C (bacteria) or room temperature (fungi) under constant shaking of 100 rpm. After 18 h (bacteria) and 42 h (fungi), the microbial samples were taken out from each well for a series of 10fold dilutions. Twenty microliters of the diluted microbial solution was streaked onto an agar plate (LB Agar from first Base). The plates were incubated for 24 and 42 h at 37 °C and room temperature, respectively, and counted for colony-forming units (CFU). Time-Kill and Killing Efficiency Test. The microbes were inoculated and prepared according to the same procedure in the MIC measurement described earlier. The samples were treated with the polymer at various concentrations (none, 1/2× MIC, 1× MIC, and 2× MIC) and were incubated at 37 °C (bacteria) or room temperature (fungi) under constant shaking of 100 rpm. At regular time intervals (0 h, 30 min, 1 h, 2 h, 4 h, 6 h, 8 h, and 18 h), microbial samples were taken out from each well for a series of 10-fold dilutions. Twenty microliters of the diluted microbial solution was streaked onto an agar plate (LB Agar from first Base). The plates were incubated for 24 and 42 h at 37 °C and room temperature, respectively, and counted for colony-forming units (CFU). For killing-efficiency studies, the microbial treated samples were taken after 18 or 42 h incubation and plated using the same protocol for viable counts. Hemolytic Activity Assay. The undesired biological activity of the polymers against mammalian cells was tested using freshly drawn rat red blood cells (rRBCs) obtained from Animal Handling Unit of Biomedical Research Centers, Singapore. Briefly, rRBCs were subjected to 25× volumetric dilutions in phosphate-buffered saline
(PBS) to achieve 4% blood content (by volume). Antimicrobial solutions were prepared by dissolving polymers in PBS at concentrations ranging from 0 to 2000 mg/L. Equal volumes of antimicrobial solutions (100 μL) were then mixed with the diluted blood suspension (100 μL). The mixtures were then incubated at 37 °C for 1 h to allow for the interactions between rRBC and the polymers to take place. Following incubation, the mixture was subjected to centrifugation (3000 g for 5 min) after which the supernatant (100 mL) was transferred into a 96-well microplate. The hemoglobin release was measured spectrophotometrically by measuring the absorbance of the samples at 576 nm using a microplate reader (TECAN, Switzerland). Two control groups were provided for this assay: untreated rRBC suspension (as the negative control), and rRBC suspension treated with 0.1% Triton-X (as the positive control). Each assay was performed in 4 replicates and repeated 3 times to ensure reproducibility of the experiments. Percentage of hemolysis was as follows Hemolysis (%) ⎡ (O.D. ⎤ 576nm of treated sample − O.D.576nm of negative control) ⎥ =⎢ ⎣ (O.D.576nm of positive control − O.D.576nm of negative control) ⎦ × 100%
Field-Emission Scanning Electron Microscopy (FE-SEM). Bacteria cells grown in TSB with or without polymer treatment were performed using similar protocol as MIC measurements but with only a 2 h incubation time. All samples were collected into a microfuge tube, pelleted at 4000 rpm for 5 min, and then washed with PBS twice. After this, fixing the samples with 2.5% glutaraldehyde for 60 min was conducted, followed by washing with DI water twice. Dehydration of the samples was performed by using a series of ethanol/water solution (35, 50, 75, 90, 95, and 100%). The dehydrated samples were dried at room temperature for 2 days before being mounted on carbon tape and coated with platinum for imaging using a JEOL JSM-7400F (Japan) field-emission scanning electron microscope. Confocal Microscopy. E. coli was inoculated and prepared according to the same protocol described in the MIC measurement. D
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One milliliter of microbial suspension (3 × 107 CFU ml−1) was added into each well of a 96-well plate and left shaking in the incubator at 37 °C for 4 h. The bacteria broth was then removed and replaced with 1 mL of 10 v/v % of DI water containing TSB with 2× MIC polymer concentration and 500 mg/L FITC-dextran (100k Da). The mixture was incubated with the attached bacteria for 0, 10, and 20 min at 37 °C under constant shaking of 100 rpm. The mixture solution was removed and washed 3 times with PBS before analysis by Zeiss LSM-510 META confocal microscope using a 63× magnification apochromate lens.
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Table 3. MIC (mg/L) and HC50 (mg/L) Polymers against Various Microbes as a Function of % OctCl Diluted with PrCl microbe S. aureus S. epidermidis E. coli P. aeruginosa C. albicans hemolysis, HC50
RESULTS AND DISCUSSION
Antimicrobial Polymer Synthesis and Self-assembly. The antimicrobial polymers were designed to have a samecentered structure where all repeat units contain a hydrophobic spacer (propyl, hexyl, or octyl) between the polymer backbone
100% OctCl
75%OctCl 25%PrCl
50%OctCl 50%PrCl
25%OctCl 75%PrCl
100% PrCl
4 4 16 125 125 250
4 4 16 125 125 500−1000
8 8 31 250 125 >1000
31 16 63 1000 125 >1000
500 16 1000 >1000 500 >1000
Table 4. MIC (mg/L) and HC50 (mg/L) Polymers against Various Microbes As a Function of % OctCl Diluted with HexCl microbe S. aureus S. epidermidis E. coli P. aeruginosa C. albicans hemolysis, HC50
100% OctCl
75%OctCl 25%HexCl
50%OctCl 50%HexCl
25%OctCl 75%HexCl
100% HexCl
4 4 16 125 125 250
4 4 16 125 125 250−500
4 4 31 250 125 500−1000
8 4 31 500 125 500−1000
125 4 125 1000 250 >1000
and quaternary ammonium group (Scheme 2a). Organocatalytic ring-opening polymerization (ROP) of functional carbonates was employed to synthesize homopolymers at three different degrees of polymerization (15, 30, and 60). Briefly, cyclic carbonates containing alkyl chloride were polymerized by ROP using a cocatalyst system (DBU/TU). Following polymerization, the reaction was quenched by addition of acetyl chloride and the alkyl chloride was quaternized postpolymerization using trimethylamine (Scheme 2c). Polymer length was dictated by the initial monomer, initiator feed ratios and verified by 1H NMR (Table 1). Polydispersity was determined by gel permeation chromatography. All polymers had a narrow molecular weight distribution with a polydispersity index of ∼1.2−1.3. For mitigation of polymer hemolytic activity, statistical copolymers containing a propyl or hexyl and octyl spacer were copolymerized (Scheme 2b). For the statistical polymer series, the mole percent of octyl was varied from 0 to 100% and the degree of polymerization (DP) was held constant at 30. 1H NMR of the polymers confirmed that the final polymer composition closely matched the feed ratio of the different monomers. All polymers are labeled by the spacer and DP (e.g., HexCl15 is a polymer with a DP of 15 and a hexyl spacer). Both homopolymers and statistical block copolymers were found to self-assemble with CMCs ranging from 39.8 to 281.8 mg/L (Table 1). For the homopolymers, CMCs increased as polymer molecular weight decreased. For the statistical block copolymers, as the hydrophobicity was decreased, not surprisingly, the CMC increased. From hydrodynamic diameter measured from dynamic light scattering (DLS), the sizes did not follow any trends and had broad size distributions. These homopolymers and statistical block polymers have very dynamic characteristics as they are able to associate and dissociate rapidly. Surface charge of both homopolymer and statistical copolymer micelles were measured and found to be positively charged ranging from 44.1 to 64.3 mV. In general,
Figure 1. Hemolysis of (a) homopolymers, (b) PrCl/OctCl statistical polymers, and (c) HexCl/OctCl statistical polymers. E
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Figure 2. Viable bacteria CFUs after 18 h of incubation with 75%OctCl 25%PrCl and 50%OctCl 50%PrCl at various concentrations (0, 1/2× MIC, 1× MIC and 2× MIC) (a) S. epidermidis, (b) S. aureus, (c) E. coli, (d) P. aeruginosa, and (e) C. albicans.
zeta potential for the homopolymers was found to increase with DP while it remained constant for the statistical polymers. Antimicrobial and Hemolytic Activities of Polymers. Polymer antimicrobial activity was evaluated against five different clinically relevant microbes, Staphylococcus epidermidis, Staphylococcus aureus, Escherichia coli, Pseudomonas aeruginosa, and Candida albicans. The effect of the polymers on bacterial growth inhibition was examined using a microdilution assay to determine the MIC of each polymer. MICs were taken at the lowest polymer concentration that inhibited bacterial growth after 18 h of incubation with an initial bacterial loading of 3 × 105 CFU/mL. Antimicrobial polymer toxicity was evaluated via hemolysis assays using rat red blood cells. MIC values for all homopolymers tested are shown in Table 2 and the results of the hemolysis assays are shown in Figure 1a. Three different degrees of polymerization were used to determine the effect of molecular weight on antimicrobial activity and hemolytic activity. The dependence of antimicrobial activity on molecular weight was minimal with the OctCl homopolymer showing higher activity against E. coli at DP 30 and 60 than DP 15. For the OctCl homopolymers, hemolytic activity increased with increasing molecular weight. For all polymers, the MIC values are lower than the CMC for S. epidermidis, S. aureus, and E.coli, indicating that self-assembly is not necessary for antimicrobial activity.
When comparing antimicrobial activity of polymers with different side chains, the PrCl homopolymer was inactive against all microbes except S. epidermidis and was nonhemolytic. The HexCl homopolymers were highly active against the tested Gram-positive bacteria and were nonhemolytic. There was good antimicrobial activity towards E. coli especially for HexCl and OctCl homopolymers; however, only the OctCl homopolymers were active against P. aeruginosa. By increasing side-chain length by two carbons (from octyl to hexyl), activity against P. aeruginosa was observed. Overall, the OctCl homopolymers were the most active polymer for all microbes tested. Unfortunately, hemolytic activity at concentrations approaching the MIC values of P. aeruginosa and C. albicans was observed. We suspect that deeper membrane penetration is possible with a longer spacer length and thus increased antimicrobial activity and hemolytic activity is observed. Other research groups have observed that variation in side chain length resulted in varying activity and selectivity toward different microbes.16,21,22 For example, Tew and coworkers16 observed that for their quaternary pyridinium functionalized polynorbornenes, a side-chain length of 8 carbons had MIC values less than 7 μg/mL for E. coli whereas a side-chain length of 6, 8, or 10 carbons had MIC values less than 7 μg/mL for B. subtilis. Furthermore, the HC50 values decreased with increasing side chain length, paralleling what is observed with the polycarbonate system. Similar trends were F
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Scheme 1b for E. coli was 125 mg/L,18 while the lowest MIC from Table 2 was 16 mg/L. However, when comparing hemolytic activity, the HexCl and OctCl homopolymers were significantly more hemolytic than those prepared via the segregated monomer approach. Minimal hemolytic activity was observed for the systems in Scheme 1a and for the system in Scheme 1b, the lowest HC50 value observed was 850 mg/L. Although high activity was observed for the OctCl polymers, their systemic application is limited due to their high hemolytic activity. There are several different strategies presented in the literature to reduce polymer hemolytic activity. Youngblood and co-workers added monomers with ethylene glycol chains and saw a reduction in hemolytic activity.23 Tew and coworkers copolymerized different facially amphiphilic monomers and observed significant reduction in hemolytic activity but also lost antimicrobial activity.14 For our system, statistical polymers were designed to increase charge density using the MTC-PrCl or MTC-HexCl monomer while maintaining the long hydrophobic tail (MTC-OctCl) for penetration into microbial membranes. We hypothesized that this approach would reduce hemolytic activity while maintaining antimicrobial activity. The OctCl polymers have higher antimicrobial activity but are hemolytic, whereas the PrCl and HexCl polymers have low hemolytic activity but have lower antimicrobial activity. The polymers prepared using MTC-OctCl and MTC-PrCl or MTCOctCl and MTC-HexCl maintained high antimicrobial activity even up to 75% MTC-PrCl or MTC-HexCl, respectively, when compared to the MTC-OctCl homopolymer (Table 3 and Table 4). The loss of activity with addition of MTC-PrCl is minimal; however, the hemolytic activity is significantly decreased (Figure 1b). The addition of MTC-HexCl also reduced the hemolytic activity but because it is more hydrophobic than the MTC-PrCl, the effect was not as drastic (Figure 1c). Most importantly, for P. aeruginosa and C. albicans the HC50 value for 75%OctCl/25%PrCl was four to eight times the MIC value, whereas for the OctCl homopolymer the HC50 was only one to two times the MIC. This method of balancing charge and hydrophobicity leads to polymers with high antimicrobial activity and minimal hemolytic activity. Antimicrobial Mechanism. From the MIC and HC50 values of all the polymers tested, each bacteria strain was treated with candidates that exhibited both high activity and selectivity, 75%OctCl/25%PrCl and 50%OctCl/50%PrCl using the agar gel assay. At both the MIC and 2× MIC concentrations, at least a 99.99% killing efficiency was achieved, suggesting a bactericidal mechanism (Supporting Information Figure S1). Furthermore, the colony forming units (CFU) counts after 18 h of incubation (Figure 2) showed the polymers were bactericidal to a wide spectrum of micro-organisms. At both 1× MIC and 2× MIC concentrations, there was a 8−14 log reduction in colony counts. The ability of both polymers to kill bacterial cells upon exposure for various time intervals was conducted using S. aureus and E. coli as model microbes (Figure 3). More than 50% of the microbes were killed after exposure to the polymers at 1× MIC and 2× MIC concentrations for 30 min. At both concentrations tested, killing efficiency of >99.99% was achieved within 4 h of treatment. This strong evidence indicates that the polymers are bactericidal instead of bacterial growth inhibiting. Using FE-SEM, imaging of microbial cell wall morphology was made possible. Figure 4 shows the highly distorted and corrugated bacterial cell surface and debris after incubating with 75%OctCl/25%PrCl at 2× MIC for 2 h, suggesting that the polymer killed bacteria via a
Figure 3. Fractional cell survival of (a) S. aureus and (b) E. coli after 30 min, 1 h, 2 h, 4 h, 6 h, and 8 h of incubation with 75%OctCl 25%PrCl and 50%OctCl 50%PrCl.
Figure 4. Field-emission scanning electron microscopy (FE-SEM) images of S. aureus (a,c) and E. coli (b,d) before (a,b) and after (c,d) 2 h treatment with 75%OctCl 25%PrCl at 2×MIC.
observed for quaternary amine-containing polypeptides and quaternary ammonium containing methacrylates.21,22 When comparing the same-centered polycarbonates to the polycarbonates prepared via the segregated monomer approach, the polymers with Hexyl and Octyl spacers were more active. For example, the system from Scheme 1a was inactive against E. coli17 and the lowest MIC observed system from G
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Figure 5. Confocal microscopic images of E. coli incubated with FITC-conjugated dextran (100k Da) in the absence (a) or presence of 75% OctCl25%PrCl for 10 min (b) and 20 min (c). Scale bar: 20 μm.
membrane-lytic mechanism. Damage of the E. coli cell membrane after exposure to the polymer was further studied by confocal microscopy. Before exposure to 75%OctCl25%PrCl at 2×MIC, FITC-dextran (100k Da) molecules were unable to enter the bacteria (Figure 5a). Upon exposure to the antimicrobial polymer, green fluorescence was seen in the bacteria, suggesting membrane rupture (Figure 5b,c). The fluorescence intensity became stronger, and the number of bacteria with green fluorescence increased after 20 min of incubation, implying that a longer exposure with 75%OctCl25% PrCl polymer resulted in more serve membrane damage. The membrane-lytic mechanism has been observed for other amphiphilic polymer systems17,18,24,25 including those with a same-centered structure.15,16
polymer backbone. These homopolymers were active against all microbes tested but the most active polymers had high hemolytic activity. To reduce hemolytic activity, we systematically varied polymer composition by copolymerizing different monomers used in the same center homopolymers. By maintaining charge on each repeat unit but copolymerizing varied hydrophobic side chain lengths, polymers with broad spectrum activity and high selectivity were achieved. These materials had CMC values higher than the MIC indicating selfassembly was unnecessary, as was previously observed. In addition, these macromolecules act via a membrane disruption mechanism, making them less likely to induce resistance.
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CONCLUSION A series of antimicrobial homopolymer polycarbonates with a same-centered structure were synthesized. Hydrophobic/ hydrophilic balance was controlled by varying the spacer between the charged quaternary ammonium moiety and the
ASSOCIATED CONTENT
* Supporting Information S
Additional MIC data. This material is available free of charge via the Internet at http://pubs.acs.org. H
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(24) Mowery, B. P.; Lee, S. E.; Kissounko, D. A.; Epand, R. F.; Epand, R. M.; Weisblum, B.; Stahl, S. S.; Gellman, S. H. J. Am. Chem. Soc. 2007, 129, 15474−15476. (25) Zhou, C.; Qi, X.; Li, P.; Chen, W. N.; Mouad, L.; Chang, M. W.; Leong, S. S. J.; Chan-Park, M. B. Biomacromolecules 2009, 11, 60−67.
AUTHOR INFORMATION
Corresponding Authors
*E-mail: (J.L.H.)
[email protected]. *E-mail: (Y.Y.Y.)
[email protected]. Author Contributions §
The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. A.C.E. and J.P.K.T. are co-first authors and contributed equally to this work Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS The authors would like to thank Central Glass for supplying MTC-OC6F5. This work was funded by IBM Almaden Research Center and the Institute of Bioengineering and Nanotechnology (Biomedical Research Council, Agency for Science, Technology and Research, Singapore).
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REFERENCES
(1) Taubes, G. Science 2008, 321, 356−361. (2) Bodey, G. P.; Bolivar, R.; Fainstein, V.; Jadeja, L. Rev. Infect. Dis. 1983, 5, 279−313. (3) Hancock, R. E. W.; Speert, D. P. Drug Resist. Updates 2000, 3, 247−255. (4) Poole, K. Clin. Microbiol. Infection 2004, 10, 12−26. (5) Wang, Z.; Wang, G. Nucleic Acids Res. 2004, 32, D590−592. (6) Som, A.; Vemparala, S.; Ivanov, I.; Tew, G. N. Biopolymers 2008, 90, 83−93. (7) Wiradharma, N.; Khoe, U.; Hauser, C. A.; Seow, S. V.; Zhang, S.; Yang, Y. Y. Biomaterials 2011, 32, 2204−2212. (8) Young, A. W.; Liu, Z.; Zhou, C.; Totsingan, F.; Jiwrajka, N.; Shi, Z.; Kallenbach, N. R. Med. Chem. Commun. 2011, 2, 308−314. (9) Marr, A. K.; Gooderham, W. J.; Hancock, R. E. W. Curr. Opin. Pharmacol. 2006, 6, 468−472. (10) Palermo, E. F.; Sovadinova, I.; Kuroda, K. Biomacromolecules 2009, 10, 3098−3107. (11) Engler, A. C.; Wiradharma, N.; Ong, Z. Y.; Coady, D. J.; Hedrick, J. L.; Yang, Y.-Y. Nano Today 2012, 7, 201−222. (12) Gabriel, G. J.; Maegerlein, J. A.; Nelson, C. F.; Dabkowski, J. M.; Eren, T.; Nüsslein, K.; Tew, G. N. Chem.Eur. J. 2009, 15, 433−439. (13) Oda, Y.; Kanaoka, S.; Sato, T.; Aoshima, S.; Kuroda, K. Biomacromolecules 2011, 12, 3581−3591. (14) Lienkamp, K.; Madkour, A. E.; Musante, A.; Nelson, C. F.; Nüsslein, K.; Tew, G. N. J. Am. Chem. Soc. 2008, 130, 9836−9843. (15) Sambhy, V.; Peterson, B. R.; Sen, A. Angew. Chem., Int. Ed. 2008, 47, 1250−1254. (16) Eren, T.; Som, A.; Rennie, J. R.; Nelson, C. F.; Urgina, Y.; Nüsslein, K.; Coughlin, E. B.; Tew, G. N. Macromol. Chem. Phys. 2008, 209, 516−524. (17) Nederberg, F.; Zhang, Y.; Tan, J. P.; Xu, K.; Wang, H.; Yang, C.; Gao, S.; Guo, X. D.; Fukushima, K.; Li, L.; Hedrick, J. L.; Yang, Y. Y. Nature Chem. 2011, 3, 409−414. (18) Qiao, Y.; Yang, C.; Coady, D. J.; Ong, Z. Y.; Hedrick, J. L.; Yang, Y.-Y. Biomaterials 2012, 33, 1146−1153. (19) Phoenix, D. A.; Dennison, S. R.; Harris, F. Antimicrobial Peptides; Wiley-VCH: Weinheim, Germany, 2013. (20) Pratt, R. C.; Nederberg, F.; Waymouth, R. M.; Hedrick, J. L. Chem. Commun. 2008, 114−116. (21) Engler, A. C.; Shukla, A.; Puranam, S.; Buss, H. G.; Jreige, N.; Hammond, P. T. Biomacromolecules 2011, 12, 1666−1674. (22) Dizman, B.; Elasri, M. O.; Mathias, L. J. J. Polym. Sci., Part A: Polym. Chem. 2006, 44, 5965−5973. (23) Allison, B. C.; Applegate, B. M.; Youngblood, J. P. Biomacromolecules 2007, 8, 2995−2999. I
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