A structure-activity study of antibacterial poly(ester urethane)s with

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A structure-activity study of antibacterial poly(ester urethane)s with uniform distribution of hydrophobic and cationic groups Chao Peng, Apoorva Vishwakarma, Steven Mankoci, Hazel A. Barton, and Abraham Joy Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.9b00029 • Publication Date (Web): 07 Mar 2019 Downloaded from http://pubs.acs.org on March 10, 2019

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A Structure-Activity Study of Antibacterial Poly(Ester Urethane)s with Uniform Distribution of Hydrophobic and Cationic Groups Chao Penga, Apoorva Vishwakarmaa, Steven Mankocia, Hazel A. Barton*b, Abraham Joy*a aDepartment

of Polymer Science and bDepartment of Biology, The University of Akron,

Akron, Ohio 44325, United States

ABSTRACT. Infections associated with antibiotic-resistant bacteria have become a threat to the global public health. Antimicrobial polymers, which are synthetic mimics of antimicrobial peptides, have gained increasing attention as they may have a lower chance of inducing resistance. The cationic-hydrophobic balance and distribution of cationic and hydrophobic moieties of these polymers is known to have a major effect on antimicrobial activity. We studied the properties of a series of facially amphiphilic antimicrobial surfactant-like poly(ester urethane)s with different hydrophobic pendant groups (P1, P2, and P3) and cationic groups distributed uniformly along the polymer chain. These polymers exhibited bactericidal activity against Gram-negative Escherichia coli and Pseudomonas aeruginosa as well as Gram-positive Staphylococcus aureus and Staphylococcus

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epidermidis. Microscopy and dye release assays demonstrated that these polymers cause membrane disruption, which is dependent on the cationic-hydrophobic ratio in the polymer. Membrane permeability assays revealed that these polymers can permeabilize the outer membrane of E. coli and damage the cytoplasmic membrane of both E. coli and S. aureus. In addition, our results indicate that the three polymers exhibit different extent of membrane disruption against E. coli. P1 caused minor damage to the cytoplasmic membrane integrity, but it was able to dissipate the cytoplasmic membrane potential, leading to cell death. P2 and P3 depolarized the cytoplasmic membrane and also caused significant damage to the cytoplasmic membrane. Overall, we showed a new class of broad-spectrum bactericidal polymers whose membrane disrupting ability against E. coli correlates with the structural differences of the hydrophobic pendant groups.

INTRODUCTION Antimicrobial peptides (AMPs) are short cationic peptides present in various organisms that are an important part of the host innate response, and have received extensive attention due to their broad-spectrum bactericidal activity and reduced likelihood of resistance compared to traditional antibiotics.1-3 AMPs are believed to cause membrane lysis by binding to negatively charged surfaces on bacterial membranes to disrupt the cell membrane integrity.1,

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Although AMPs are efficient antimicrobial agents, the

commercialization of AMPs has been limited by their relatively high manufacturing cost


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and inherent proteolytic degradation.4, 7, 8 Consequently, the use of synthetic polymeric mimics of AMP has gathered increased relevance. These AMP analogs (antimicrobial polymers) present several advantages over natural AMPs, such as lower cost, long-term activity, metabolic stability, and the flexibility of chemical modification.9, 10 In recent years, a number of advancements in antimicrobial polymers have been reported based on various platforms, such as poly(meth)acrylates, polycarbonates, polyesters, polyamides, and poly(oxanorbornene)s.11-21 Kuroda et al. developed cationic random copolymers of methacrylates and methacrylamides, and studied the influence of different cationic groups as well as the hydrophilic-hydrophobic ratio on both antimicrobial activity and toxicity against eukaryotic cells.11, 12 Yang et al. also developed biodegradable antimicrobial polycarbonates that displayed high bactericidal efficiency against clinically isolated methicillin-resistant Staphylococcus aureus (MRSA) and showed high efficacy in vivo when tested in a mouse systemic infection model.15 There are several antimicrobial polymers reported in literature in which the cationic and hydrophobic moieties are randomly distributed along the polymer chain.11, 12, 14, 18, 22-27 Recent studies have shown that the distribution of the cationic and hydrophobic pendant groups in the polymer chain can play a role in the resultant antimicrobial property of the polymer.28, 29 For example, according to the study by Sambhy et al. polymers with cationic pyridines and hydrophobic tails in the same repeating unit exhibited much better


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selectivity to bacterial cells compared to polymers with cationic and hydrophobic groups distributed randomly along the polymer chain.29 In this work, we report a densely functionalized antimicrobial poly(ester urethane) system, in which the cationic and hydrophobic pendant groups are adjacent to each other in the same repeating unit, resulting in a cationic surfactant-like structure. This design provides a uniform local hydrophilic-hydrophobic balance and facilitates the electrostatic and hydrophobic interactions between the polymer and the negatively charged lipid molecules in the bacterial membrane. Three antimicrobial poly(ester urethane)s bearing different hydrophobic pendant groups were synthesized based on a previously developed unsaturated poly(ester urethane) platform.30 The antimicrobial activity of these polymers was investigated in model bacteria: Gram-negative Escherichia coli and Gram-positive Staphylococcus aureus. The mechanism of action of these polymers against bacteria was investigated by microscopy studies, dye release assays using a bacterial membranemimicking liposome model, and membrane permeability assays. The toxicity of these polymers against eukaryotic cells was also examined by a red blood cell hemolysis assay. Overall, the results show a strong correlation between the structure of the polymers and the resultant bacterial activity and mammalian cell toxicity. EXPERIMENTAL SECTION


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Materials. Tin(II) 2-ethylhexanoate (Sn(Oct)2), 2-hydroxy-4’-(2-hydroxyethoxy)-2methylpropiophenone (Irgacure 2959), sodium phosphate dibasic, potassium phosphate monobasic, potassium chloride, calcium chloride, HEPES, D-lactose monohydrate, and Triton X-100 were purchased from Sigma-Aldrich. Fluorescein isothiocyanate (FITC), hexamethylene diisocyanate (HDI) and 2-nitrophenyl-β-D-galactopyranoside (ONPG) were purchased from Acros Organics. Anhydrous methanol, triethylamine (TEA), and dimethylformamide (DMF) were purchased from EMD Millipore. Cysteamine hydrochloride, N,N-diisopropylethylamine (DIPEA), and 6-carboxyfluorescein were purchased from Chem-Impex International. Dimethyl sulfoxide (DMSO) and sodium chloride were purchased from BDH Chemicals. N-phenyl-1-naphthylamine (NPN), 3,3’dipropylthiadicarbocyanine iodide (DiSC3(5)) were purchased from TCI America. 1Palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine (POPE) and 1-palmitoyl-2-oleoylsn-glycero-3-phosphoglycerol (POPG) were purchased from Cayman Chemical. Mueller Hinton broth (MHB) was purchased from Himedia. Melittin, a membrane-pore forming peptide was purchased from Enzo Life Sciences. Defibrinated sheep blood was purchased from Hardy Diagnostics. Escherichia coli K12 (ATCC 10798), Staphylococcus aureus (ATCC 25923), and Staphylococcus epidermidis (ATCC 12228) were obtained from the American Type Culture Collection (ATCC), with Escherichia coli UB1005 from the E. coli Genetic Stock Center (CGSC) at Yale University. Sephadex G-50 was purchased from Pharmacia


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Fine Chemicals. Dichloromethane was dried by distilling over calcium hydride. Unless otherwise stated, all reagents were used as received. Analytical Methods NMR spectra were recorded on a Varian NMRS 300 MHz instrument. 1H NMR chemical shifts are reported in ppm relative to the solvent’s residual 1H signal. Size exclusion chromatography (SEC) analysis in DMF was performed on an HLC-8320 GPC from TOSOH equipped with an RI detector using polystyrene (PS) as the standard. Absorbance and fluorescence spectroscopy were performed using a BioTek Synergy H1 plate reader. The morphology of the bacteria was characterized by scanning electron microscope (SEM) using JEOL-JSM-7401F with operating voltage as 1 kV. Fluorescence imaging was performed on a U-TV1X-2 fluorescence microscope (Olympus Co., Japan) with standard FITC/TRITC filter set. Synthesis of Cationic Poly(Ester Urethane)s. The unsaturated poly(ester urethane) precursors were synthesized according to a reported procedure.30 As an example, the synthesis of P2 is as follows: Pre2 (0.21 g), cysteamine hydrochloride (0.57 g) and Irgacure 2959 (57 mg) were dissolved in a mixture of anhydrous methanol (4 mL) and anhydrous dichloromethane (2 mL). Once dissolved, the solution was irradiated at 350 nm for 30 minutes. The polymer was purified by dialysis in methanol to give a colorless solid (yield ≈90%).


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P1. 1H NMR (300 MHz, MeOD), δ (ppm): 0.91-1.00 (m, 3H), 1.35-1.70 (m, 12H), 2.84-3.21 (m, 11H), 4.27-4.35 (m, 4H), 5.02 (m, 1H). P2. 1H NMR (300 MHz, MeOD), δ (ppm): 1.32-1.66 (m, 10H), 2.59-3.18 (m, 13H), 4.24-4.29 (m, 4H), 5.02 (m, 1H), 7.20-7.28 (m, 5H). P3. 1H NMR (300 MHz, MeOD), δ (ppm): 0.88-0.93 (m, 3H), 1.30-1.64 (m, 24H), 2.82-3.11 (m, 11H), 4.31 (m, 4H), 5.02 (m, 1H). Preparation of FITC-Labeled Polymers. As an example, the synthesis of P1-FITC is as follows: P1 (10 mg) was dissolved in anhydrous DMF (0.5 mL). Then, anhydrous triethylamine (5 μL), FITC stock solution (0.5 mL, 1 mg/mL in anhydrous DMF), and DIPEA (5 μL) were added. The reaction was carried out at room temperature for 2 hours. The product was purified by dialysis in water for 24 hours with water changes at 1h, 2h, 4h, 6h, and 16h. Minimum Inhibitory Concentration (MIC) and Time-Kill Assays. Unless otherwise stated, all bacterial cultures were grown in MHB with shaking (200 rpm) at 37°C. Overnight cultures of either E. coli K12, S. aureus, S. epidermidis, or P. aeruginosa were diluted in MHB to an OD600nm 0.001 (measured by Hach DR 2800 Spectrophotometer), by initially diluting to 0.1 and then serially diluting 100 fold (~1×106 CFU/mL). For the MIC assays, this suspension was mixed with the polymer aqueous solution in a 1:1 ratio in a 96-well plate. The optical density (OD600nm) was measured and followed by incubating at


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37°C for 16-20 h, whereupon, optical density (OD600nm) was measured again using a BioTek Synergy H1 plate reader. The minimum inhibitory concentration (MIC) was determined as the lowest concentration of polymer that prevented bacteria growth after overnight incubation. For the time-kill assays, the working bacterial suspensions were mixed with the polymer aqueous solution in a 1:1 ratio. 100 L aliquots at different time points were serially diluted onto trypticase soy agar (TSA). The results were expressed as colony forming units (CFU)/mL over time. Hemolysis Assays. Defibrinated sheep red blood cells (RBC) were centrifuged at 500 x g for 10 min at 5 °C. The RBC pellet was washed with phosphate-buffered saline (PBS; pH 7.4) until the supernatant no longer contained any visually observable hemoglobin. The RBC suspension was then diluted 1:50 in PBS and mixed with polymer aqueous solution in a 1:1 ratio. Wells containing no polymer were used as 0% hemolysis control, and 1% Triton X-100 was added to the positive control for 100% hemolysis. After incubation at 37°C for 1 h, the plate was centrifuged at 500 x g for 10 min at 5°C and the hemoglobin release as a relative measure of hemolysis was determined at 450 nm on a BioTek Synergy H1 plate reader. Lactate Dehydrogenase (LDH) Assay for NIH-3T3 Cells. Mouse fibroblast NIH-3T3 cells were cultured using DMEM medium supplemented with 10% FBS and 1% penicillin streptomycin. The cells were grown at 37 °C in 5% CO2 until reaching ~90% confluence.


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The cells were harvested using 0.25% trypsin solution and seeded in a 96-well plate at a density of 1×104 cells/well. The cells were allowed to attach to the wells overnight before treatment with polymers. The polymer treated cells were incubated at 37 °C for 1 hour before determining the cell viability. The cell viability was determined using a Pierce LDH Cytotoxicity Assay Kit according to the instruction of the manufacturer. Outer Membrane Permeability Assay. A 20 mL culture of E. coli was incubated until the culture reached an OD600nm of 0.5. The culture was centrifuged at 2000 x g for 5 min, and the cell pellet was resuspended in 20 mL of 10 mM HEPES/150 mM NaCl (pH 7.4). 100 μL of this suspension was mixed with 50 μL of polymer solution and 50 μL of N-phenyl1-naphthylamine (NPN) solution (40 μM) in a 96-well plate. Wells containing no polymer were used as the negative control, and with 1% Triton X-100 as a positive control. The fluorescence at 350/420 nm (excitation/emission) was measured on a BioTek Synergy H1 plate reader. Cytoplasmic Membrane Permeability Assay. E. coli K12 was used to inoculate 3 mL of MHB supplemented with 2% lactose and incubated until mid-log phase (OD600nm~0.5). The bacterial suspension was then centrifuged at 2000 xg for 5 min, and the cell pellet was resuspended in PBS (pH 7.4) to an OD600nm of 0.05. 100 μL of this suspension was then added to each well of a 96-well plate along with 50 μL of polymer solution and 50 μL of ONPG solution (5 mM). Wells containing no polymer were used as the negative


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control, and melittin was used as the positive control. The UV absorbance at 420 nm was recorded at 37 °C for 60 min. Cytoplasmic Membrane Depolarization Assay. A culture of E. coli UB1005 or S. aureus was prepared as described for the cytoplasmic membrane permeability assay and resuspended in 10 mM HEPES/150 mM NaCl (1% DMSO) with 4 μM DiSC3(5) and incubated at 37°C for 60 min. The bacterial suspension was further diluted 2 times in 10 mM HEPES/150 mM NaCl (1% DMSO) and incubated at 37°C for an additional 60 min to give the working suspension. This suspension was then mixed with polymer solutions at a 1:1 ratio in a 96-well plate and the fluorescence at 610/660 nm (excitation/emission) was monitored over a period of 60 min at 37 °C. Wells containing no polymer were used as the negative control, and the wells containing melittin (100 μg/mL) were used as a positive control. SEM Sample Preparation. A 3 mL culture of E. coli K12 with an OD600nm of 0.5 was mixed with polymer aqueous solution (32 μg/mL) in a 1:1 ratio, and incubated at 37°C for 30 min. The mixture was then filtered through a 0.22 μm Durapore® Membrane Filter and fixed with 4% formaldehyde in PBS for 2 hours. The bacteria were then washed with DI water (3X) and dehydrated in a gradient ethanol/water mixture (50%, 60%, 70%, 80%, 90%, 95%, and 100%; 15 min for each step), followed by critical point drying and sputter


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coating.

The samples were observed using a JEOL-JSM-7401F scanning electron

microscope (SEM). Fluorescence Microscopy. The bacteria were resuspended in 10 mM HEPES with 150 mM NaCl, and diluted to an OD600nm value of 0.1. This suspension was then mixed with 100 μg/mL FITC-labelled polymer solution in a 1:1 ratio and incubated at 37 °C for 30 min. Fluorescence imaging was performed on a U-TV1X-2 fluorescence microscope (Olympus Co., Japan) with standard FITC/TRITC filter set. Liposome Preparation and Dye Release Assay. Liposomes were prepared using POPE/POPG (4:1). The lipid film was hydrated with 10 mM HEPES buffer with 50 mM 6-carboxyfluorescein and 150 mM NaCl, freeze-thawed 5 times, and placed on a shaker overnight. This lipid solution was then extruded 15 times through an Avanti MiniExtruder equipped with filters of 100 nm pore size and placed on a shaker for 1 hour. The unincorporated dye was removed from the liposome using a Sephadex G-50 column and lipid concentration was quantified by phosphorus assay.31 As calculated using a NaH2PO4 standard curve (Figure S5), the final lipid concentration was 30 μM. The liposomes were mixed with polymer solutions. The samples were incubated at 37 °C for 1 hour and fluorescence emission at 530 nm (excitation at 495 nm) was monitored over a period of 60 min at 37 °C on a BioTek Synergy H1 plate reader. Wells containing no


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polymer were used as 0% dye release control, and wells containing 1% Triton X-100 were the 100% dye release positive control. RESULTS AND DISCUSSION Polymer Design and Synthesis

Scheme 1. Synthetic route of the antimicrobial poly(ester urethane)s (P1, P2, and P3) The novel antimicrobial poly(ester urethane)s reported in this work were synthesized based on an alkene functionalized poly(ester urethane) platform previously reported by our group.30 The synthetic route of the antimicrobial poly(ester urethane)s is depicted in Scheme 1. Briefly, an alkene functionalized diol was first synthesized via a one-step Baylis-Hillman reaction of functionalized aldehyde with 2-hydroxyethyl acrylate in a mixture of 1,4-dioxane and water catalyzed by DABCO according to a reported procedure.30, 32 The resulting diol monomer was then polymerized with hexamethylene diisocyanate (HDI) in anhydrous CH2Cl2 catalyzed by tin(II) octoate to generate an


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unsaturated poly(ester urethane) precursor. As confirmed by SEC analysis, the molecular weight of these precursors was in the range of 5-9 kDa (Pre1~8.0 kDa, Ð~1.7; Pre2~5.1 kDa, Ð~1.9; Pre3~8.8 kDa, Ð~1.7). The unsaturated polymer precursors were then functionalized by reacting with cysteamine hydrochloride via thiol-ene reaction to provide the final cationic poly(ester urethane)s (P1, P2, P3). The chemical structure of the cationic polymers was confirmed by 1H NMR spectroscopy (Figure S1, S2, and S3). As shown by the polymer structure, the cationic and hydrophobic groups are presented in the same repeating unit, and they are adjacent to each other, giving a cationic surfactantlike structure. The rationale of using such polymer design is to provide a uniform local hydrophilic-hydrophobic balance and increase the affinity of the polymers to the negatively charged lipid molecules in the bacterial membrane, leading to enhanced antimicrobial efficacy. Antimicrobial Activity Determined by MIC Measurement and Time-Kill Test The antimicrobial activity of these poly(ester urethane)s was investigated using the Gram-negative E. coli and P. aeruginosa as well as Gram-positive S. aureus and S. epidermidis. These antimicrobial polymers showed low MIC values for E. coli (MIC = 8-16 μg/mL; Table 1), S. aureus (MIC = 16-32 μg/mL; Table 1), P. aeruginosa (MIC = 16-32 μg/mL; Table 1), and S. epidermidis (MIC = 4-16 μg/mL; Table 1) in MHB. The hydrophobicity of the pendant group does not have a significant influence on the MIC values, as the three


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polymers exhibit only slightly different MICs (1-2 dilution difference). Similar trend was observed when comparing the MIC results using molar-based concentrations. To understand whether these polymers are bacteriostatic or bactericidal, we monitored bacteria viability over 24 hours for E. coli and S. aureus, as proof-of-concept. As shown in Figure 1 (a), all three polymers are bactericidal toward E. coli, as evidenced by the ~5 log reduction in CFU after treatment with 16 μg/mL polymer. This reduction in CFU values for E. coli increased with increasing hydrophobicity in the pendant groups. The polymers were also bactericidal toward S. aureus, albeit at higher concentrations and lower killing rates (Figure 1(b)). At the concentration of 32 μg/mL, P2 and P3 reduced the number of viable S. aureus by ~5 log after 2 h and 6 h, respectively. P1 was not as effective against S. aureus, being bacteriostatic for up to 8 hours, followed by growth. Nonetheless, P1 was bacteriocidal at a concentration of 64 μg/mL. Similar trend in the killing efficiency would be observed when compared using the same molar-based concentration, as these polymers have similar molecular weight. Also, polymers with more hydrophobic pendant groups showed lower molar-based MIC values (P1~4.0 µM, P2~3.1 µM, P3~1.8 µM, against S. aureus). Table 1. Minimum inhibitory concentration (MIC) of the polymers against E. coli, S. aureus, P. aeruginosa, and S. epidermidis and HC10 of the polymers toward red blood cells

polymer

MIC Gram-negative bacteria

Gram-positive bacteria

HC10


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P1 P2 P3 Ampicilli n Melittin

E. coli µg/m µM L 16 2.0 8 1.6 8 0.9 1

2.9

25

8.8

P. aeruginosa µg/m µM L 32 4.0 16 3.1 16 1.8 128 371.2 50

17.6

S. aureus µg/m µM L 32 4.0 16 3.1 16 1.8 0.2 0.6 25

8.8

S. epidermidis µg/m µM L 16 2.0 4 0.8 16 1.8 4 11.6 0.8

µg/m L

µM

379.4 19.8 3.5

47.4 3.8 0.4

0.3

Figure 1. Time-kill test of (a) E. coli and (b) S. aureus over a period of 24 hours Toxicity of the Antimicrobial Polymers The toxicity of these polymers was evaluated by hemolysis assay, with the hemolytic activity quantified by percentage of hemoglobin released from lysed red blood cells relative to 100% lysis by 1% Triton X-100.33,

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HC10 is defined as the polymer

concentration that causes 10% hemolysis and it is commonly used as a parameter to reflect the toxicity of the polymers. The percentage of hemolysis induced by the


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antimicrobial poly(ester urethane)s is shown in Figure 2, with a lower HC10 value indicating higher toxicity. The polymers showed hydrophobicity-dependent toxicity (Figure 2 and Table 1). P2 and P3 with more hydrophobic pendant groups exhibited higher hemolytic activity than P1. Thus, while increased hydrophobicity leads to better antimicrobial efficacy, it also dramatically increased the hemolytic activity of the polymers, making P2 and P3 less selective to bacterial cells over mammalian cells (Figure 2 and Table 1). The same trend was also observed in the results of cell viability assay using NIH-3T3 fibroblast cells, in which polymers with more hydrophobic pendant groups exhibited higher toxicity (Figure S9).

Figure 2. Hemolytic activity of P1, P2, and P3 Fluorescence Microscopy Study of E. coli


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In this work, we performed mechanistic studies of these antimicrobial polymers against Gram-negative E. coli and Gram-positive S. aureus to understand their mechanism of action. To determine the mechanism of action of our polymers against the bacterial membrane, we carried out microscopy studies in E. coli, which has an outer (periplasmic) membrane (compared with the Gram-positive S. aureus surrounded by a peptidoglycan cell wall). FITC-labeled antimicrobial polymers were added to E. coli cells and observed via fluorescence microscopy. As shown in Figure 3 and S4, the E. coli cells were labelled with the fluorescent polymer, indicating that the polymer was bound to the cell.

Figure 3. Fluorescence microscopy images of E. coli before treatment (left) and after treatment with P1-FITC for 30 min (right) Membrane Permeabilization of E. coli To determine the location of binding by these polymers, an outer membrane permeability assay for E. coli was carried out using a membrane-impermeable fluorescent probe N-


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phenyl-1-naphthylamine (NPN), whose fluorescence is weak in an aqueous environment but strong in a hydrophobic environment. When the outer membrane is disrupted, NPN dye is able to enter the cell membrane and accumulate in a hydrophobic environment, leading to an increase in fluorescence intensity, which can be used as an indication of permeability. As shown in Figure 4, all three polymers permeabilized the outer membrane in a concentration-dependent manner, and polymers with more hydrophobic pendant groups caused greater permeability of the outer membrane (P3>P2>P1).

Figure 4. E. coli outer membrane permeability of P1, P2, and P3 measured using NPN as a fluorescent probe A cytoplasmic membrane permeability assay was performed using a colorimetric probe ortho-nitrophenol-β-galactoside (ONPG) to detect β-galactosidase enzymatic activity. In E. coli, β-galactosidase is the enzymatic product of lacZ, which catalyzes the hydrolysis of


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β-galactosides.35 Disruption of the cytoplasmic membrane would lead to the enzymatic reaction between β-galactosidase and ONPG (caused by the permeabilization of ONPG into the cells or the release of β-galactosidase into the medium), generating a yellow ortho-nitrophenol product, which has absorbance at 420 nm. As shown in Figure 5, P1 caused minimal activity as the absorbance at 420 nm was very close to the negative control even at concentrations significantly above its MIC. On the other hand, P2 and P3 showed significant β-galactosidase activity (P3>P2>P1), again in a concentration dependent fashion. These results indicate that P1 may exhibit weaker membrane disrupting ability compared to P2 and P3, indicating lower permeabilization of ONPG or comparatively lower release of β-galactosidase. However, the assay is not able to provide more detailed information on the mechanism (whether the enzymatic activity is a result of the permeabilization of ONPG or the release of enzyme) other than the overall membrane disrupting ability. Interestingly, Figure 5(d) showed that melittin exhibited very different kinetics compared to P2 and P3, in which the absorbance at 420 nm gradually increased over the test period, suggesting that melittin might have a different mechanism of action against E. coli compared to the described poly(ester urethane)s. Melittin can bind to and insert into the bacterial membrane, leading to the formation of toroidal pores. In the case of P2 and P3, the absorbance increased rapidly in the beginning and reached a plateau within 10 minutes. The results suggested that P2 and P3 would cause the formation of larger pores compared to melittin in the beginning, which led to


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the rapid increase in the absorbance. The presence of plateau could be caused by the blockage of the pores by the antimicrobial polymers or the potential deactivation of βgalactosidase by the antimicrobial polymers.

Figure 5. E. coli cytoplasmic membrane permeability assay of P1 (a), P2 (b), P3 (c), and melittin (d) over a period of 60 min measured using ONPG as a colorimetric probe Further SEM studies were performed to observe the effect of the polymers on the bacterial morphology. As shown in Figure 6, the polymer treated bacteria exhibited rough and wrinkled surfaces (b, c, d), while the untreated bacterial cells exhibited a relatively smooth surface (a), indicating that these polymers disrupted the bacterial membrane.


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Figure 6. SEM images of E. coli before treatment (a) and after treatment with 16 μg/mL of P1 (b), P2 (c), and P3 (d) for 30 min; scale bar is 1 μm Membrane Depolarization of E. coli Since P1 was an effective antimicrobial polymer, but did not cause sufficient cytoplasmic membrane permeabilization to allow leakage of the β-galactosidase enzyme, we wondered whether the extent of membrane disruption caused by P1 would affect the ability of the cell to maintain the proton-motive force necessary for cell energetics. To test


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this, we performed a cytoplasmic membrane depolarization assay for E. coli using DiSC3(5) probe, which is a membrane potential-sensitive cyanine dye. DiSC3(5) can readily enter cell membranes and accumulate in healthy polarized cytoplasmic membranes.36 Once accumulated in the cytoplasmic membrane, DiSC3(5) shows weak fluorescence due to self-quenching; if the antimicrobial polymers disrupt the cytoplasmic membrane and dissipates the membrane potential, DiSC3(5) will be released into the medium, leading to a corresponding increase of fluorescence intensity.37 As shown in Figure 7, all three polymers caused membrane depolarization, which confirmed the interactions between the polymers and the cytoplasmic membrane. This result also indicated that P1 was able to dissipate the membrane potential and cause cell death even though it did not cause sufficient damage to the cytoplasmic membrane to show any significant β-galactosidase activity (Figure 5(a)), since a functioning transmembrane potential is essential for metabolic activity and growth.38

Figure 7. E. coli cytoplasmic membrane depolarization by P1 (a), P2 (b), and P3 (c) over a period of 60 min measured using DiSC3(5) as a fluorescent probe


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Membrane Disrupting Ability Studied by Dye Release Assay To understand the effect of polymer structure on the membrane disrupting ability, a dye release assay was performed in which fluorescence detection of 6-carboxyfluorescein (CF) leakage from bacterial membrane-mimic liposomes was used as an indication of the membrane disrupting ability. The fluorescence intensity of CF is weak when encapsulated in the liposomes due to self-quenching. However, release of the dye from the liposomes due to liposome disruption would lead to an increase in the fluorescence intensity. In this study, a lipid composition that mimics the membrane of E. coli was used, which consists of 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine (POPE) and 1palmitoyl-2-oleoyl-sn-glycero-3-phosphoglycerol (POPG) in a ratio of 4:1. The dye encapsulated liposomes were incubated with the antimicrobial polymers and the fluorescence intensity at 530 nm was monitored (495 nm excitation). As shown in Figure 8, all three polymers disrupted the liposomes, leading to CF release. The dye release was concentration dependent (b, c, d) and polymers with more hydrophobic pendant group caused more dye release (P3>P2>P1) at the same concentration (a), which correlated well with the trend of MIC values and killing rates of the three polymers.


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Figure 8. (a) Summary of dye release from liposomes that were treated with the antimicrobial polymers at different concentration; dye release kinetics of the liposomes after treatment with P1 (b), P2 (c), and P3 (d) over a period of 60 min. Membrane Permeability of S. aureus In the absence of an outer (periplasmic) membrane in S. aureus, we wanted to determine whether our polymers could still affect the cytoplasmic membrane protected by the thick, peptidoglycan cell wall of this organism. Cytoplasmic membrane disruption of S. aureus was observed for all three polymers using the DiSC3(5) fluorescent probe (Figure 9), increasing with the hydrophobicity of the pendant groups (and correlated with


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antimicrobial activity), suggesting that these polymers could still access the cytoplasmic membrane, even in the presence of a thick cell wall.

Figure 9. Cytoplasmic membrane depolarization of S. aureus by P1 (a), P2 (b), and P3 (c) over a period of 30 min measured using DiSC3(5) as a fluorescent probe Conclusion A series of antimicrobial surfactant-like poly(ester urethane)s were designed and developed

via

step-growth

polymerization

followed

by

post-polymerization

modification using thiol-ene reaction. These novel antimicrobial polymers exhibited antimicrobial activity against E. coli, S. aureus, P. aeruginosa, and S. epidermidis. Microscopic analysis demonstrated that these polymers bind to and disrupt bacterial membranes, with their membrane disrupting ability depending on the hydrophilichydrophobic balance. The polymers with more hydrophobic pendant groups showed stronger tendency to disrupt the bacterial membrane, which correlated well with their antimicrobial activity. Membrane permeability assays showed that all three antimicrobial polymers were able to permeabilize the outer (periplasmic) and inner (cytoplasmic)


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membrane of E. coli, and that the cell wall of S. aureus did not prevent these polymers from disrupting the cytoplasmic membrane. P2 and P3 that showed significant damage to the cytoplasmic membrane of E. coli, while P1 caused minor damage to the integrity of cytoplasmic membrane, but even such an extent of disruption was able to dissipate the cytoplasmic membrane potential and cause cell death. The structure-activity relationships and mechanistic insights from these studies can help guide the design and development of new antimicrobial polymers with improved antimicrobial activity and cytocompatiblity. ASSOCIATED CONTENT Supporting Information Molecular weight information, NMR spectra, and fluorescence microscopy images. This material is available free of charge via the Internet at http://pubs.acs.org. AUTHOR INFORMATION Corresponding Authors * (A.J.) E-mail: [email protected] * (H.A.B.) E-mail: [email protected] Notes The authors declare no competing financial interest.


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Antimicrobial Poly(ester urethane)s 148x103mm (150 x 150 DPI)


A structure-activity study of antibacterial poly(ester urethane)s with

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