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Bactericidal Specificity and Resistance Profile of Poly(Quaternary Ammonium) Polymers and Protein-Poly(Quaternary Ammonium) Conjugates Weihang Ji, Richard R. Koepsel, Hironobu Murata, Sawyer Zadan, Alan S Campbell, and Alan J Russell Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.7b00705 • Publication Date (Web): 28 Jun 2017 Downloaded from http://pubs.acs.org on June 29, 2017

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Bactericidal Specificity and Resistance Profile of Poly(Quaternary Ammonium) Polymers and Protein-Poly(Quaternary Ammonium) Conjugates Weihang Ji,† Richard R. Koepsel,†,‡ Hironobu Murata,†,‡ Sawyer Zadan,‡ Alan S. Campbell,†, § Alan J. Russell*,†,‡,§,ǁ †

§

Center for Polymer-Based Protein Engineering, ‡Disruptive Health Technology Institute,

Department of Biomedical Engineering, and ǁDepartment of Chemistry, Carnegie Mellon University, Pittsburgh, Pennsylvania 15213, United States

ACS Paragon Plus Environment

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ABSTRACT: Antibacterial polymers are potentially powerful biocides that can destroy bacteria on contact. Debate in the literature has surrounded the mechanism of action of polymeric biocides and the propensity for bacteria to develop resistance to them. There has been particular interest in whether surfaces with covalently coupled polymeric biocides have the same mechanism of action and resistance profile as similar soluble polymeric biocides. We designed and synthesized a series of poly(quaternary ammonium) polymers, with tailorable molecular structures and architectures, to engineer their antibacterial specificity and their ability to delay the

development

of

bacterial

resistance.

These

linear

poly(quaternary

ammonium)

homopolymers and block copolymers, generated using atom-transfer radical polymerization, had structure dependent antibacterial specificity toward Gram positive and negative bacterial species. When single block copolymers contained two polymer segments of differing antibacterial specificity, the polymer combined the specificities of its two components. Nanoparticulate human serum albumin-poly(quaternary ammonium) conjugates of these same polymers, synthesized via “grafting from” atom-transfer radical polymerization, were strongly biocidal and also exhibited a marked decrease in the rate of bacterial resistance development relative to linear polymers.

These protein-biocide conjugates mimicked the behavior of surface-presented

polycationic biocides rather than their non-proteinaceous counterparts.

KEYWORDS: Poly (quaternary ammonium), Polymeric biocide, Bacterial resistance, Atomtransfer radical polymerization, Hydrophobicity, Protein-polymer conjugates


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INTRODUCTION The prevention and treatment of bacterial infections has been a focus of scientists for centuries1,2. Antimicrobial polymers, sometimes referred to as polymeric biocides, can be particularly effective with broad specificity and low residual toxicity relative to similar low molecular weight compounds2. Indeed, biology itself has generated a broad array of antimicrobial polypeptides. The development of polymeric biocides stemmed from combined research into polymeric disinfectants and antimicrobial peptides in the 1980s. This work resulted in the production of antimicrobial polymeric materials with widely varying molecular structures and architectures2-5. The production of new materials that combine high antimicrobial activity with a low impact on eukaryotic cells, all while delaying the onset or appearance of bacterial resistance, has represented the Holy Grail of this field for some time. Quaternary ammonium (QA) and quaternary phosphonium-containing polymers are some of the most widely studied polymeric biocides1. The mechanism by which these cationic polymers kill bacterial cells begins with the adsorption of the cationic biocide onto negatively charged bacterial cell surfaces, followed by disruption of the cytoplasmic membrane leading to cell death2, 6-8. The efficiency of each step in this process, and thus the biocidal activity as a whole, has been shown to depend on polymer molecular weight (MW), chain length, the alkyl spacer length between the cationic charge center and the polymer backbone, hydrophobicity, and the counterions present1,

2, 9

. Differences in biocidal activity between small molecule, polymeric,

dendritic, nanoparticulate and surface-coupled biocides have been attributed to the differences in each step of the killing mechanism6, and the structure and net charge of the bacterial or eukaryotic cell2, 3, 10-13. Ideally, a single polymeric biocide could be tailored to selectively target


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particular cell types, thereby destroying harmful bacteria while maintaining the viability of beneficial bacterial species and eukaryotic cells14, 15. Atom-transfer radical polymerization (ATRP), a widely studied controlled radical polymerization technique, has proven to be a powerful and facile means of preparing tightly defined polymer structures16-19. We have previously generated antibacterial surfaces covalently functionalized with non-leaching poly(quaternary ammonium) biocides using ATRP20-23. ATRP is a particularly attractive tool for the synthesis of biocidal surfaces because it enables facile control of chain length and MW, narrow MW distributions and a large library of available architectures16, 23-26. Extensive and sometimes inappropriate use of disinfectants and antibiotics has led to the emergence of antibiotic resistant species of bacteria, with approximately 2 million people infected annually with antibiotic-resistant bacteria in the United States alone27, 28. This trend is a result of the environmental pressure asserted by constant antibiotic use, coupled with the short life cycles and lateral gene transfer mechanisms of bacterial populations3. The continual discovery of new, resistant species of bacteria has led to interest in the development of efficient, broad-spectrum antimicrobial materials that delay or prevent resistance generation1. Whether and how bacteria generate resistance to polymeric biocides in solution and on surfaces is often referred to but not well understood29-32. Some researchers have presented exciting results and conclusions that imply that surface-bound biocidal polymers could delay or even prevent the generation of resistance3, 6.

Nevertheless, bacteria could theoretically become resistant to

polymeric biocides by modifying their cell wall/cellular membrane to decrease net negative charge (thereby inhibiting cationic polymer binding) or through increased activity of efflux pumps and biocide degrading enzymes3, 29-34. Poly(quaternary ammonium)-based materials have


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been shown to kill bacteria that are resistant to other cationic biocides35. Herein, we report on the synthesis and biologic activity of a series of cationic homopolymers and block copolymers with poly (quaternary ammonium) (pQA)-containing side-chains using ATRP. We hypothesized that antibacterial activity and resistance development could be tailored by fine-tuning the polymer structure. We were particularly interested in whether co-polymers combined the specificity of their individual components or whether they had their own unique bacterial specificity. Each putative polymeric biocide was thoroughly characterized and cell death was determined with four bacterial species, human erythrocytes and a human cell line (HEK293). This systematic approach provided insight into the relationship between polymer properties and membrane disruption. We also developed an assay to determine the speed and degree to which Gram negative and Gram positive bacteria could generate resistance to the polymeric biocides. Polymeric biocides on surfaces have unique properties, particularly from a resistance profile perspective. ATRP can grow polymers from solid surfaces as well as from the surface of proteins. Polymer-based protein engineering (PBPE) with ATRP has been used to generate protein-polymer conjugates with native activity, enhanced stability and additional properties imparted by the polymer itself19, 36-38, 39-44. To more fully explore how polymeric biocides might be optimized for activity and resistance profile, we decided to grow pQA’s from the surface of a model blood protein, human serum albumin (HSA). We discovered that the nanoparticulate HSA-pQA conjugates (that included a poly(oligo(ethylene glycol) methyl ether methacrylate) (pOEGMA) spacer) behaved more like polymer-coated surfaces than soluble polymers and exhibited important differences in antibacterial activity and resistance profile relative to their linear polymer counterparts.


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MATERIALS AND METHODS Materials and general methods. Oligo(ethylene glycol) methyl ether methacrylate (OEGMA, Mn = 475) was purchased from Sigma-Aldrich (St. Louis, MO) and passed through a basic aluminum oxide column to remove inhibitors prior to use. 2-(dimethylethylammonium)ethyl methacrylate (QA-C2), 2-(dimethylbutylammonium) ethyl methacrylate (QA-C4), and 2(dimethylhexylammonium) ethyl methacrylate (QA-C6) were synthesized as described previously (Scheme S1)41,

45, 46

. QA monomers bearing other alkyl side chains (i.e. QA-C3

(propyl) and QA-C5 (pentyl)) were prepared similarly. N-2-bromo-2-methylpropionyl-β-alanine and N-2-bromo-2-methylpropionyl-β-alanine N’-oxysuccinimide ester (NHS-Br) ATRP initiator compounds were prepared as described previously43. Human serum albumin (HSA), copper(I) bromide

(Cu(I)Br),

1,1,4,7,10,10-hexamethyltriethylenetetramine

(HMTETA),

and

fluorescamine were purchased from Sigma-Aldrich (St. Louis, MO) and used without further purification. 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide (MTT), Dulbecco’s phosphate buffered saline (DPBS), Dulbecco’s Modified Eagle’s Medium (DMEM), fetal bovine serum (FBS) and trypsin were purchased from Invitrogen (Carlsbad, CA). Tryptic soy broth (TSB) was purchased from Becton, Dickinson and Company (Franklin Lakes, NJ). HEK293 cells were purchased from ATCC (Manassas, VA). Dialysis tubing (Spectra/Por, Spectrum Laboratories Inc.) was purchased from Fisher Scientific (Pittsburgh, PA). Leuko reduced, packed human red blood cells (RBCs) were kindly provided by the Institute for Transfusion Medicine (Pittsburgh, PA). All other chemicals and solvents were of analytic grade and used as received. 1

H-NMR spectra were recorded on a 300 MHZ Bruker Avance in the NMR facility located in

the Center for Molecular Analysis, Carnegie Mellon University, Pittsburgh, PA. UV-Vis spectra were recorded using a UV-Vis spectrometer (Lambda 45, Perkin Elmer) with a temperature-


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controlled cell holder. Matrix-assisted laser desorption ionization time-of-flight mass spectroscopy (MALDI-TOF) was carried out using a PerSeptive Biosystems Voyager Elite MALDI-TOF mass spectrometer in the Center for Molecular Analysis, Carnegie Mellon University, Pittsburgh, PA. High-throughput absorbance measurements were made using a BioTek Synergy HT plate reader. Molecular weights were estimated by gel permeation chromatography (GPC) on a Waters 2695 Series equipped with three columns (Waters Ultrahydrogel Linear, 500 and 250), using 300 mM sodium sulfate buffer with 0.01% (mass/volume) sodium azide as the eluent at a flow rate of 1.0 mL/min and detection by a refractive index (RI) detector. All sample solutions were filtered (0.22 µm pore size) before analysis. Poly(ethylene glycol) standards were used for GPC calibration. Average hydrodynamic diameters and zeta potentials were determined using a NanoPlus-3 dynamic light scattering (DLS) analyzer (Micromeritics Instrument Corporation, Norcross, GA). Synthesis of pQA homopolymers by ATRP. For each QA monomer, four types of pQA homopolymers with different targeted degrees of polymerization (TDP = 25, 50, 100 and 200) were prepared by aqueous ATRP, resulting in a library of 20 pQA homopolymers. In a typical polymerization, (pQA-C2, TDP = 100), QA-C2 monomer (1.0 g, 3.76 mmol) was added to N-2bromo-2-methylpropionyl-β-alanine initiator (8.9 mg, 0.038 mmol) in deionized (DI) water (10 mL), sealed and bubbled with argon in an ice bath for 30 min. Next, deoxygenated catalystligand solution containing HMTETA (20.4 µL, 0.075 mmol) and Cu(I)Br (10.8 mg, 0.075 mmol) in ice cold DI water (10 mL) was added to the initiator-monomer solution, sealed and stirred at 4 °C for 4 h. The reaction was terminated by opening the reaction flask to air and an aliquot was removed for 1H-NMR analysis. Conversion was estimated by comparing integrated areas of residual vinyl protons (~5.5 ppm) and methyl group protons (~3.1 ppm) on the ammonium


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group. The homopolymer was isolated by dialysis using 3.5 kDa molecular weight cutoff tubing against DI water for 36 h and lyophilized. pQA-C2 homopolymers with TDP of 25, 50 and 200 were similarly synthesized by adjusting the molar ratio of monomer to initiator while maintaining [I]:[Cu(I)Br]:[HMTETA] = 1:2:2. Other pQAs with varying TDPs were synthesized according to the same methodology. It should be noted that dialysis filtration of all pQA homopolymers with TDP = 25 was carried out using 1.0 kDa molecular weight cutoff tubing. Synthesis of pQA block copolymers by ATRP. pQA-C2 and pQA-C3 macroinitiator were synthesized as previously described. Each reaction was terminated after 1 h (i.e. pQA-C2, TDP = 60, 120; pQA-C3, TDP = 60). In a typical co-polymerization (pQA-C255-b- pQA-C6; C6 TDP = 60), QA-C6 monomer (363.1 mg, 1.13 mmol) was added to pQA-C255 macroinitiator (274.9 mg, 0.019 mmol) in DI water (5 mL), sealed and bubbled with argon in an ice bath for 30 min. Then, deoxygenated catalyst-ligand solution containing HMTETA (10.2 µL, 0.038 mmol) and Cu(I)Br (5.4 mg, 0.038 mmol) in ice cold DI water (5 mL) was added to the initiator-monomer solution, sealed and stirred at 4 °C for 16 h. The reaction was terminated by opening the reaction flask to air and an aliquot was removed for 1H-NMR analysis as mentioned above. The copolymer was purified by dialysis using 3.5 kDa molecular weight cutoff tubing against DI water for 36 h and then lyophilized. Other copolymers were similarly synthesized by adjusting the molar ratio of monomer to initiator. Synthesis of HSA-pOEGMA-block-pQA conjugates by ATRP. To synthesize HSA-Br macroinitiator, NHS-Br and HSA (molar ratio 3:1; NHS-Br : primary amines of HSA) were dissolved in sodium phosphate buffer (0.1 M, pH 8.0) and stirred at room temperature for 3 h. Chemical structure of HSA-Br was confirmed by 1H-NMR and degree of initiator modification were determined via fluorescamine assay and MALDI-TOF, respectively (Supplementary


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information). To synthesize HSA-pOEGMA-b-pQA conjugates, OEGMA monomer (1784.1 mg, 3.76 mmol) and HSA-Br (301.4 mg, 0.19 mmol Br) were dissolved in DI water (30 mL), sealed and bubbled with argon in an ice bath for 30 min. Next, deoxygenated catalyst-ligand solution containing HMTETA (102.2 µL, 0.38 mmol) and Cu(I)Br (53.9 mg, 0.38 mmol) in DI water (10 mL) was added to the initiator-monomer solution, sealed and stirred in an ice bath for 20 min. HSA-pOEGMA conjugate was isolated by dialysis using a 25 kDa molecular weight cutoff tubing against DI water at 4 °C for 36 h and then lyophilized. pQA was then grown from HSA-pOEGMA macroinitiator (pQA-C2, TDP = 50, 100; pQA-C4, TDP = 50; pQA-C6, TDP = 100). In a typical polymerization (HSA-pOEGMA20-b-pQA-C2; TDP = 100), QA-C2 monomer (1000 mg, 3.76 mmol) and HSA-pOEGMA (417.3 mg, 0.038 mmol) were dissolved in DI water (50 mL), sealed and bubbled with argon in an ice bath for 30 min. Then, deoxygenated catalyst-ligand solution containing HMTETA (20.4 µL, 0.075 mmol) and Cu(I)Br (10.8 mg, 0.075 mmol) in DI water (10 mL) was added to the initiator-monomer solution, sealed and stirred at 4 °C for 18 h. HSA-pOEGMA-b-pQA-C2 conjugate was isolated by dialysis using 25 kDa molecular weight cutoff tubing against DI water and then lyophilized. Other conjugates were similarly synthesized by adjusting the monomer type and molar ratio of monomer to initiator. Grown polymers were cleaved from HSA through acid hydrolysis and purified as previously described39. Hemolysis rate determination. A serial dilution of each polymer was prepared in DPBS in a 96-well plate (100 µL per well) and mixed with DPBS diluted, twice washed RBCs (100 µL, 1.2 ×107 cells/mL initial). The plate was then incubated for 1 h at 37 °C with mild shaking. A complete hemolysis positive control was performed by adding cells to 2% v/v Triton-X. DPBS was used as the negative control. After incubation, the 96-well plate was centrifuged at 800 rcf


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for 5 min followed by transfer of 100 µL of supernatant to a fresh 96-well plate. Hemoglobin release was determined through absorbance measurements at 405 nm. Degree of hemolysis was defined as: % lysis = 100× (Apolymer – Ablank)/(ATriton-x – Ablank). Each sample was tested in triplicate. Polymer cytotoxicity determination. Polymer cytotoxicity was evaluated by the MTT assay. HEK293 cells were incubated in DMEM (10% FBS, 10 mM HEPES, 100 U/mL penicillin/streptomycin) at 37 °C in a humidified atmosphere containing 5% CO2. HEK293 cells were first cultured overnight in 96-well plates (20,000 cells/well initial). Varying polymer concentrations were then added to the wells and incubated for 24 h with 5% CO2 at 37 °C. MTT in DPBS (20 µL, 5 mg/mL) was added to each well and incubated for 4 h, at which time unreacted MTT was removed via aspiration before DMSO (150 µL) was added to each well. Absorbance was measured at 570 nm and cell viability calculated using the ratio of polymer exposed cell to control cell absorbance. Bacterial Culture and Polymer MIC determination. The minimum inhibitory concentration (MIC) of each polymer was determined in turbidity-based microdilution assays as described previously.15 Assays were performed in TSB for all bacteria. Escherichia coli (ATCC 8739), Staphylococcus aureus (ATCC 43300, MRSA), Staphylococcus aureus (ATCC 6538), Acinetobacter baumannii (ATCC 19606) and Bacillus cereus (ATCC 14579) were examined. Polymer was dissolved in DPBS and diluted by TSB in 2-fold serial dilutions. Bacterial cultures were prepared by incubating 1 to 2 bacterial colonies in TSB (5 mL) overnight at 37 °C with shaking. The cultures were then diluted with TSB to OD595 = 0.1 and incubated at 37 °C until the mid-log growth phase was reached (OD595 between 0.5 and 0.6; 1.5 to 4.5 h depending on bacterial strain). Cultures were next diluted to OD595 = 0.001 (approximately 5 x 105 cfu/mL)


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and used as stock solutions. Stock solution (100 µL) was mixed with polymer solution (100 µL) in a sterile 96-well plate, covered with sterile, breathable sealing tape (ThermoFisher Scientific, Waltham, MA) and incubated for 18 h at 37 °C. Plates were visually examined to determine bacterial growth indicated by increased turbidity. Polymer MIC was defined as the lowest polymer concentration at which no turbidity increase was observed for at least 2 out of 3 wells relative to the negative control (TSB). All assays were performed in triplicate in at least 3 independent experiments. Polymer serum compatibility was examined similarly to MIC determinations, except TSB was supplemented with FBS (10% or 30%). The resulting serum compatibility MICs were recorded and compared to calculated MICs in TSB alone. Polymer-induced rate of bacterial cell death determination. Determination of antibacterial kinetics was performed by inoculation of the polymers with bacteria in TSB medium and colony counting on 3M petrifilm rapid aerobic count plates (Saint Paul, MN). An overnight culture of MRSA was regrown to the mid-log growth phase (OD595 between 0.5 and 0.6) and diluted to OD595 = 0.001 in a centrifugation tube. Polymer solution was then added to reach a final concentration equal to twice the determined MIC of each respective polymer. Tubes were incubated at 37 °C with shaking at 250 rpm and aliquots (100 µL) taken at varying time points. A serial dilution in TSB medium was transferred to count plates and incubated overnight at 37 °C prior to cell counting. Each experiment was performed in duplicate. Bacterial resistance development assay. Bacteria samples from triplicate wells (100 µL from each; polymer concentration equal to one-half the calculated MIC) were added to fresh TSB (5.0 mL) and cultured at 37 °C to mid-log growth phase (OD595 between 0.5 and 0.6). The resulting culture was then used to determine new polymer MIC as described above. This procedure was


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repeated for a total of 8 successive passages and performed in duplicate. Bacterial resistance development in E. coli was similarly determined with a total of 14 successive passages. Statistical analysis. All statistical analyses were carried out using a 2-sample Student’s t-test with unequal variance. Values of p < 0.05 were deemed to be statistically significant. RESULTS AND DISCUSSION Synthesis and characterization of pQA linear polymers. Among cationic polymeric biocides, poly(2-(dimethylamino)ethyl methacrylate (pDMAEMA) and its QA-containing derivatives have attracted extensive interest due to their effectiveness and facile synthesis20-23, 47, 48

. The use of ATRP to synthesize these polymers gives us a uniquely powerful tool to tune the

properties of these biocides and determine structure function relationships. However, no comprehensive, systematic study has been performed to fully investigate the structure-bioactivity relationship of pQAs homo- and co-polymers. We therefore generated a library of pQA linear polymers with varying alkyl side chain lengths and MWs (Scheme 1), and determined their bioactivities toward bacteria and human cells. Scheme 1. Synthetic route of linear pQA polymers.

A

R = C2H5, C3H7, C4H9, C5H11, or C6H13

B


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R1 = C2H5, R2= C6H13 or R1 = C3H7, R2=C5H11

In preliminary experiments, we discovered that ATRP of amino monomers was less efficient than the polymerization of quaternized monomers. We therefore synthesized 5 QA monomers with alkyl side chain lengths varying from 2 to 6 carbons (i.e. QA-C2, QA-C3, QA-C4, QA-C5 and QA-C6) (Scheme S1). In total, we synthesized 20 unique pQA homopolymers (Scheme 1A). We also synthesized 4 block copolymers with different alkyl side chain lengths of each block by combining pQA-C2 and pQA-C6 as well as pQA-C3 and pQA-C5 (Scheme 1B). The TDP, monomer conversion percentage (estimated by 1H-NMR), average MW (calculated by both 1HNMR and GPC) and PDI for the polymers were determined (Table S1). The polymers had PDIs of approximately 1.4, which were typical for water-based ATRP49. It should be noted that we could not accurately measure the MW and PDI of pQA-C6-containing homopolymers and block copolymers by GPC, likely due to non-specific interactions of the relatively hydrophobic polymers with the GPC column. Cellular and bacterial toxicity of pQA linear polymers. To elucidate the relationship between polymer properties and the bioactivity of the putative pQA biocides with bacteria and human cells, we determined the ability of pQA linear polymers to irreversibly disrupt the membranes of erythrocytes, HEK293 cells and four species of bacteria (E. coli, A. baumannii, S. aureus and B. cereus) (Table 1).


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Table 1. Molecular weight, hydrophobicity and bioactivity of pQA linear polymers. Avg. Mna (x103)

HC50b (µg/mL) (Hmaxc (%))

IC50 (µg/mL)

pQA-C224

6.7

>2130 (3.41)

pQA-C248

12.6

pQA-C296

Polymer

MICe (µg/mL) (Selectivity Index)

d

E. coli

A. baumannii

MRSA

B. cereus

133

1250 (0.1)

5000 (0.03)

5 (26.6)

10 (13.30)

>2130 (1.98)

133

625 (0.21

5000 (0.03)

5 (26.6)

5 (26.6)

27.0

>2130 (1.03)

33

625 (0.05)

5000 (0.01)

5 (6.6)

5 (6.6)

pQA-C2180

48.5

>2130 (1.38)

33

625 (0.05)

5000 (0.01)

5 (6.6)

2.5 (13.2)

pQA-C323

7.7

>2242 (11.07)

140

625 (0.22)

5000 (0.13)

pQA-C340

11.5

>2242 (0.30)

280

312 (0.90)

2500 (0.11)

pQA-C394

24.3

>2242 (5.45)

35

156 (0.22)

5000 (0.01)

pQA-C3184

44.0

>2242 (3.41)

35

312 (0.11)

5000 (0.01)

pQA-C424

6.3

>2355 (0.52)

147

312 (0.47)

156 (0.94)

312 (0.47)

20 (7.35)

pQA-C442

11.2

>2355 (8.95)

147

40 (3.68)

2500 (0.06)

10 (14.7)

10 (14.7)

pQA-C490

26.1

>2355 (7.64)

18

40 (0.48)

5000 (0.01)

10 (1.8)

2.5 (7.2)

pQA-C4176

53.0

>2355 (1.35)

18

20 (0.96)

2500 (0.01)

10 (1.8)

5 (3.6)

pQA-C524

2.8

>2465 (1.45)

154

40 (3.85)

10 (15.4)

625-312 (0.25-0.5)

154)

pQA-C542

4.0

>2465 (0.36)

154

10 (15.4)

10 (15.4)

pQA-C596

10.0

>2465 (0.22)

19

10 (1.9)

10 (1.9)

pQA-C5190

31.7

>2465 (1.92)

19

10 (1.9)

80 (0.24)

pQA-C623

7.4

>2580 (11.84)

20

10 (2)

2.5 (8)

pQA-C644

14.2

>2580 (4.75)

20

10 (2)

2.5 (8)

pQA-C690

29.0

>2580 (4.71)

20

10 (2)

5 (4)

pQA-C6188

60.6

>2580 (10.28)

20

20 (1)

10 (2)

80-40 (0.250.5)

5 (4)

pQA-C255-b-C646

29.5

>2330 (2)

73

40 (1.82)

156 (0.26)

20 (7.80)

78 (0.26)

pQA-C298-b-C656

44.1

>2310 (4)

36

78 (0.46)

312 (0.12)

20 (1.81)

78 (0.46)

5 (28) 5-2.5 (56112) 35) 35)

156 (0.97) 10 (0.48) 10 (0.48) 156 (0.01) 156 (0.01) 156 (0.01)

5 (28) 10-5 (28-56) 5 (7) 35)

154) 19) 2.5 (7.6) 20) 20) 2.5 (8)


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pQA-C298-b-C6102

58.9

>2397 (6)

19

156 (0.12)

312 (0.06)

20 (0.94)

39 (0.48)

pQA-C355-b-C550

30.8

>2345 (0.6)

73

78 (0.94)

250 (0.03)

20 (3.67)

39 (1.88)

a

1

Average MW was determined by: GPC for pQA-C2, pQA-C3, pQA-C4 and pQA-C5 homopolymers;

H-NMR for pQA-C6 homopolymers and all block copolymers. bHC50 was defined as the polymer

concentration at which 50% hemolysis was observed against RBCs. cHmax was defined as the percent hemolysis at maximum polymer concentration tested. dIC50 was defined as the polymer concentration at which 50% viability was observed for HEK293 cells. eMIC was defined as the minimum inhibitory concentration of target bacteria. Selectivity index was defined as the ratio of IC50 to MIC.

We examined the hemolysis profile of each pQA linear polymer against RBCs by measuring hemoglobin release from RBCs when incubated with increasing polymer concentrations. The HC50 of each sample was defined as the polymer concentration at which 50% hemolysis was observed (Table 1). Overall, pQA linear polymers were not hemolytic at the concentrations tested. The majority of experiments showed less than 5% hemolysis at even the highest tested concentrations (Table 1). Lower polymer concentrations resulted in decreased hemolysis for each polymer (data not shown). However, two pQA-C6 linear polymers (pQA-C623 and pQA-C6188) yielded over 10% hemolysis. Our data agreed with prior observations that hydrophobic polymers damage RBC membranes50. Our data show an interesting cutoff in hydrophobicity. The C2-C5 polymer side chains are sufficiently hydrophilic so as not to disrupt the erythrocyte membrane, but the C6 polymer side chain is hydrophobic enough to burst the red blood cells. To further investigate the biocompatibility of the library of pQAs, we determined the cytotoxicity of each pQA linear polymer on a monolayer of adherent HEK293 cells using the MTT assay. The IC50 of each sample was defined as the polymer concentration at which 50%


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cell viability was observed (Table 1). We found that cytotoxicity gradually increased with increasing polymer concentration (data not shown). For polymers with the same alkyl side chain length, shorter polymers were generally less cytotoxic than their longer counterparts. This trend was evident from the IC50 values of pQA-C2, pQA-C3, pQA-C4 and pQA-C5 homopolymers, where DP’s of around 25 and 50 yielded greater IC50 values than DP’s of around 100 and 200 (Table 1). However, pQA-C6 homopolymers exhibited identical IC50 values for all DPs, again implying that significant hydrophobicity dominated the bioactivity of pQAs on eukaryotic cells. For pQA block copolymers, variations in polymer MW exhibited similar trends in cytotoxicity. The two block copolymers with average MWs around 30 kDa (pQA-C255-b-C646 and pQA-C355b-C550) had almost double the IC50 of pQA-C298-b-C656 (Avg. Mn = 44.1 kDa) and nearly 4-fold greater IC50 than pQA-C298-b-C6102 (Avg. Mn = 58.9 kDa) (Table 1). The cytotoxicity of each block copolymer was also generally between the individual values of their corresponding components with similar chain lengths. Naturally, we explored whether similar concentrations of mixtures of C2 and C6 PQAs would be effective broad spectrum biocides, or whether they may interfere with each other. Separate homopolymers were indeed effective in mixtures (data not shown), but the advantage of a copolymer is that it is a single molecular entity. We next investigated the ability of each pQA linear polymer to irreversibly disrupt the membrane (and kill) four bacterial species: E. coli, A. baumannii, S. aureus and B. cereus. MICs were determined for the Gram-negative bacteria (E. coli and A. baumannii) and the Grampositive bacteria (S. aureus and B. cereus).

MIC was defined as the minimum polymer

concentration at which no bacterial growth was detected in a turbidity assay. In general, the bactericidal efficiency of each polymer increased with increasing MW (MICs decreased with increasing DP). Longer polymers would be expected to interact with larger cell surface areas. We


16

Biomacromolecules

have also shown previously that on a surface, high charge densities were necessary to induce membrane destabilization6,23. In solution, we hypothesized that longer polymer chains would present more charges to the microbial surface. Our data showed that increased MW (carrying more charge per molecule) increased the efficiencies of initial adsorption, membrane binding and membrane disruption, while likely decreasing efficiency of biocide diffusion past the cell wall1,2. Interestingly, the quaternized monomers alone were not biocidal, which emphasized the benefit of localized charge density (charges per molecule) in the polymeric biocides (Table S2). Polymers with shorter alkyl side chains, such as pQA-C2, showed preferential bactericidal activity toward Gram-positive bacteria (MICs were as low as 5 µg/mL for MRSA and B. cereus, and up to 1250 µg/mL and 5000 µg/mL for E. coli and A. baumannii, respectively). Polymers with longer hydrophobic alkyl side chains such as pQA-C6 were more efficient toward killing of Gram-negative bacteria (MICs were as low as 10 µg/mL and 2.5 µg/mL for E. coli and A. baumannii, respectively), but had diminished activity toward MRSA (MIC as high as 156 µg/mL) (Table 1). Using the longest homopolymers, we determined the rate of bacterial cell lysis to further explore the structure-bioactivity relationships. pQA-C6188 killed more than 99.8% of E. coli within 3h (Figure 1A) and pQA-C298 killed more than 99.9% of MRSA within the same time period (Figure 1B). 9

10

10

10

A 10

B 10

10

10

9

8

8

7

Cell concentration (cfu/ml)


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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24

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Time (h)


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Figure 1. Rate of bacterial cell death induced by pQA linear polymers. (A) Rate of E. coli cell death in the absence (filled square) and presence of pQA-C6188 (filled circle) in TSB medium. (B) Rate of MRSA cell death in the absence (filled square) and presence of pQA-C298 (filled circle) and pQA-C298-b-C6102 (open circle) in TSB medium. Polymer concentrations were equal to twice the calculated MIC for each polymer. Gram-positive bacteria possess a thick, peptidoglycan cell wall, while Gram-negative bacteria have an outer membrane covering a much thinner cell wall with fewer porin channels for the transport of hydrophilic molecules51-54. It has been proposed that these differences play an important role in how effectively pQAs kill cells. That said, hydrophobic polymers could also destabilize the outer membrane. For Gram-negative bacteria, it has been suggested that linear polymers with an increased degree of hydrophobicity had higher rates of diffusion through the cell wall and membrane51, 54. Specifically, pQA homopolymers with alkyl side chain lengths of 4 carbons and higher (i.e. ≥ pQA-C4) effectively killed E. coli while those with alkyl side chain lengths of 5 carbons and higher (i.e. ≥ pQA-C5) effectively killed A. baumannii (Table 1). We also observed the importance of hydrophobicity in the biocidal activity of QA-C6 monomer, which was more active toward E. coli than the other monomers (Table S2). Since each pQA preferentially killed certain bacteria, we sought to discover if a single polymer, which combined two different bacteria-specific regions, would have broad specificity. Merging two narrow specificity polymeric biocides into a single polymer with combined specificity could be a general strategy to broaden the bactericidal spectrum of polymeric biocides. Taking the C2 and C6 pQAs, we synthesized a series of block copolymers. We were


18

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excited to observe that the block copolymer biocides retained the biocidal activity of each block, exhibiting biocidal activity against both Gram-negative and Gram-positive bacteria (Table 1). Specifically, pQA-C298-b-C6102 eliminated more than 99.8% of MRSA within 3 h (Figure 1B) while also killing E. coli effectively (Table 1). One potential drawback of polycationic biocides could be the efficacy of the polymer in the presence of serum (that might increase ionic strength above a threshold necessary to diminish efficacy). In addition to ionic strength issues, serum anionic proteins could bind to pQAs electrostatically, thereby inactivating them. We therefore decided to determine whether the pQA biocides would function in fetal bovine serum (FBS). We discovered that most of the pQAs had MIC values in FBS that were similar to those in serum-free TSB medium. (Figure S2 and Figure S3). The FBS proteins neither inactivated the pQA biocides by an ionic strength effect nor a direct binding. Overall, we were able to successfully tune the specificity of pQA linear polymers toward varying bacteria types by altering polymer molecular structure and chain length. Hydrophobicity and charge density had profound influences on the specificity of the biocides, which were consistent with previous studies1, 2, 55, 56. Next, we extended our analysis by determining whether pQAs could kill bacteria without harming eukaryotic cells. We measured the selectivity index of each polymer (defined as the ratio of IC50 to MIC) relative to HEK293 cells. Our results showed that a blanket assumption that polycations are harmless to eukaryotic cells could not be made. Data sets such as those in Table 1 should inform rational design of an optimal polymer for a given targeted polymeric biocide. Generation of bacterial resistance to pQA linear polymers. The propensity for bacteria to develop resistance to polymeric biocides in solution and on surfaces is non-trivial to determine.


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We investigated the resistance profiles of E. coli and MRSA against pQA linear polymers through repeated determinations of MIC over successive passages (Figure 2). We incubated cells from a prior passage at pQA concentrations equal to half of the MIC that had been measured from that prior passage. This allowed for a new MIC to be determined for cells exposed to, but not killed by, the selected polymeric biocide. Resistance profiles against antimicrobial peptides have been examined extensively, but less so for polymeric biocides29-32. Taking E. coli and MRSA as test cases, we found that E. coli did not develop significant resistance against the hydrophobic pQA homopolymer (pQA-C6188) or a block copolymer (pQAC298-b-C6102) that contained a hydrophobic segment (Figure 2A). E. coli MIC’s increased only 2-fold over 14 passages. Perhaps not surprisingly, given the multiple drug resistant strain of S. aureus that was used, MRSA developed resistance against the hydrophilic pQA homopolymer (pQA-C298), or block copolymer (pQA-C298-b-C6102), after several passages (Figure 2B). After only 5 passages, MRSA MIC’s for both of these polymeric biocides increased more than 250fold. This resistance generation could have been caused by the bacteria mutating to partially neutralize the negatively charged cell surface/cellular membrane of Gram-positive MRSA through a mechanism similar to what has been reported for resistance generation by cationic antimicrobial peptides3,

29-34, 57

. Partial neutralization of the cell surface would result in a

decreased efficiency of initial cationic biocide adsorption2,6-8. It has been suggested that polymeric biocides, and to an even greater extent, dendritic or nanoparticulate biocides were less susceptible to this means of resistance generation3, 6. Our findings that pQA-based polymeric biocides yielded resistance development in MRSA, but at a lower rate, led us to extend our investigation to the potential of surface-mimicking polymer-protein cationic conjugates to delay resistance generation.


20

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2.4

300

A2.2

B 250 Fold increase of MIC

2.0

Fold increse of MIC

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1.8 1.6 1.4 1.2

200 150 100 50

1.0 0 0.8 2

4

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12

14

1

Passages tested

2

3

4

5

Passages tested

Figure 2. Bacterial resistance generation with pQA linear polymers. (A) Propensity of E. coli to develop resistance to killing by pQA-C6188 (filled circle) and pQA-C298-b-C6102 (open circle) as a function of passage. (B) Propensity of MRSA to develop resistance to killing by pQA-C298 (filled circle) and pQA-C298-b-C6102 as a function of passage.


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Synthesis and characterization of HSA-pOEGMA-b-pQA conjugates. It has been reported previously that bacteria develop resistance to polycationic surfaces less effectively than they do to soluble polycationic biocides8. We were interested in whether nanoparticulate cationic biocides would behave more like molecules or surfaces, from a resistance profile perspective. In order to synthesize nanoparticulate pQAs, we decided to grow long-chain pQAs from the surface of a human protein (human serum albumin) by protein surface-initiated ATRP (Scheme 2). HSA was chosen as a model protein core for cationic biocide-functionalization due to its inherent biocompatibility and well-characterized structure for predictive modification58. We first conjugated accessible primary amines (i.e. lysine residues and N-terminus) with a water-soluble NHS-functionalized ATRP initiator prepared as previously described (Scheme 2A)43. 1H-NMR spectra (Figure S4) and MALDI-TOF (Figure S5) analyses were used to determine the success and extent of initiator modification, respectively. We determined that 48 of the 59 primary amino groups within the HSA structure were successfully modified. This density of initiating sites was higher than reported for polycationic star-polymers made via ATRP, allowing us to increase the number of charges per molecule than with a star polymer59, 60. We next grew a short pOEGMA block directly from the initiator sites on the HSA surface via “grafting from” ATRP to shield the HSA surface from electrostatic interaction with pQA (Scheme 2B). Finally, we extended the HSA-pOEGMA polymers with a long pQA block using ATRP (Scheme 2C). To our knowledge, the resulting HSA-pOEGMA-b-pQA conjugates were the first examples of polycationic biocidal polymers that were “grafted from” a protein. We hypothesized that the ability of ATRP to generate protein-based nanoparticles with extraordinarily high cationic charged surfaces would provide unique properties when compared to fixed charge centers on previously reported polycationic dendritic or nanoparticulate surfaces6, 61-63.


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Scheme 2. Synthetic route for HSA-pOEGMA-b-pQA conjugates.

R = C2H5, C4H9, or C6H13

To determine the length of polymers grown from the surface of HSA polymer, we cleaved the polymer from HSA through acid hydrolysis. The TDP, conversion percentage (estimated by 1HNMR) and average MW (calculated by 1H-NMR) of cleaved polymer were determined (Table S3). Increases in hydrodynamic diameter, measured by DLS, further confirmed successful polymer growth from the HSA core and the generation of substantial polycationic nanoparticles. Growth of longer pQA chains resulted in increased conjugate hydrodynamic diameter, as expected (Table 2).

Table 2. Physiochemical properties of HSA-pOEGMA-b-pQA conjugates.


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Conjugate

Avg. Mna

HSApOEGMA20b-pQA-C244 HSApOEGMA20b-pQA-C442 HSApOEGMA20b-pQA-C663 HSApOEGMA20b-pQA-C272 a

Volume charge density (+/nm3)

Surface charge density (+/cm2)

14.12 ± 3.72

1.13E-02

1.33 x 1013

106.6 ± 1.06

12.88 ± 2.29

3.14E-03

5.61 x 1012

1.507

159.5 ± 1.73

13.83 ± 2.78

1.41E-03

3.76 x 1012

1.466

146.4 ± 3.45

32.40 ± 9.81

2.41E-03

5.61 x 1012

(x10 )

Particle size (nm)

Zeta potential (mV)

1.095

71.0 ± 0.53

1.126

6

Average MW determined by 1H-NMR.

Impact of HSA-pOEGMA-b-pQA conjugates on prokaryotic and eukaryotic cells. We next repeated a complete assessment of the biocidal activity of the unique HSA-pOEGMA-bpQA conjugates by determining their impact against erythrocytes, HEK 293 cells and four species of bacteria (E. coli, A. baumannii, MRSA and B. cereus) (Table 3).

Table 3. Effect of HSA-pOEGMA-b-pQA conjugates on human and bacterial cells. HC50 (µg/mL) (Hmax (%))

IC50 (µg/mL)

HSA-pOEGMA20b-pQA-C244

>4150 (2.45)


Conjugate

MIC (µg/mL) (Selectivity Index) E. coli

A. baumannii

MRSA

B. cereus

32

2500 (0.01)

20 (1.6)

20 (1.6)

5 (6.4)

>4470 (1.43)

17

2500 (0.01)

20 (0.85)

20 (0.85)

5 (3.4)


>4000 (20.86)

8

80 (0.1)

65 (0.12)

65 (0.12)

10 (0.8)


>3347 (4.47)

16

2500 (0.01)

10 (1.56)

10 (1.56)

5 (3.12)


24

Biomacromolecules

The HSA-pOEGMA-b-pQA conjugates (at concentrations that were relative to total mass of conjugate, not polymer alone) were generally not hemolytic, causing less than 5% hemolysis, even the highest tested concentrations.

That said, the C6-containing conjugates yielded over

20% hemolysis at the maximum concentration, similar to their linear polymer counterparts (Table 3). The HSA-pOEGMA-b-pQA conjugates had more cytotoxicity against the HEK293 cells (Table 3) than their linear counterparts. We hypothesized that this was likely caused by the increased local charge density of these dramatically dense cationic conjugates. We next determined the impact of the HSA-pOEGMA-b-pQA conjugates on bacterial cells. (Table 3). OEGMA alone and HSA-pOEGMA20 conjugates showed no antibacterial activity (Table S2). All the HSA-pOEGMA-b-pQA conjugates effectively killed both Gram-positive bacteria, with the highest Gram-positive MIC value recorded for the C6-containing conjugate against MRSA, consistent with the pQA-C6 homopolymer. Further, the C6-containing conjugate was the most effective antibacterial agent toward the killing of E. coli, similar to trends shown by pQA linear polymers. Clearly, the same trends in hydrophobicity and charge density that influenced biocidal activity in linear pQAs, were at play with the conjugates.

The conjugates

also acted quickly on bacterial cells, with HSA-pOEGMA20-b-C272 capable of killing 99.8% of MRSA within 3h (Figure 3). The rate of cell lysis was not determined for C6-containing conjugates due to their particularly high cytotoxicity with HEK cells.

9

10

8

10

Cell concentration (cfu/mL)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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Figure 3. Rate of bacterial cell death induced by selected HSA-pOEGMA-b-pQA conjugates. Rate of MRSA cell death in absence (filled square) and presence of pQA-C298 (filled circle) and HSA-pOEGMA20-b-pQA-C272 (open circle) in TSB medium. Polymer concentrations were equal to twice the calculated MIC for each polymer. As was the case for the linear polymeric biocides, the HSA-pOEGMA-b-pQA conjugates maintained their function in serum (Figure S2 and Figure S7). In TSB supplemented with either 10% or 30% FBS, the conjugates were still effective against bacteria (with only a 2-fold increase in MIC). Generation of bacterial resistance to HSA-pOEGMA-b-pQA conjugates. We were interested in comparing the resistance profiles of the HSA-pOEGMA-b-pQA conjugates and linear polymer. We hypothesized that the very high charge density in the conjugates could possibly delay the onset of resistance by enabling them to behave more like surface biocides. We therefore examined the relative rates of resistance generation in MRSA and S. aureus (6538) induced by HSA-pOEGMA-b-pQA conjugates as described above (Figure 4). HSA-pOEGMA20b-pQA-C272 was selected as our model conjugate in this complex experiment because it was so effective at killing S. aureus. We were excited to see that the development of resistance to conjugate killing was significantly delayed, relative to resistance generation in bacteria exposed to the linear polymer. After 5 passages, the MIC of HSA-pOEGMA20-b-pQA-C272 against MRSA increase merely 4-fold compared to the more than 250-fold increase in pQA-C298 MIC. It took fully 8 passages for the MRSA to develop resistance (Figure 4A). Similarly, the conjugate


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MIC against a second strain of S. aureus increased 96-fold, whereas the linear polymer MIC increased more than 380-fold after 5 passages (Figure 4B). Combining all our data with published hypotheses on mechanism of action, we have formulated a hypothesis as to how the conjugates delay resistance.

Figure 4. Bactericidal resistance generation of selected HSA-pOEGMA-b-pQA conjugates. (A) Propensity of MRSA to develop resistance to killing by pQA-C298 (filled circle) and HSApOEGMA20-b-pQA-C272 (open circle) as a function of number of passages. (B) Propensity of S. Aureus 6538 to develop resistance to killing by pQA-C298 (filled circle) and HSA-pOEGMA20-bpQA-C272 as a function of number of passages. Experiments performed in TSB medium. Our data supported the idea that MRSA became resistant to the conjugates through natural selection of cells that had partial neutralization of their negatively charged cell surface/cellular membrane. It has been suggested that dendritic biocides were less susceptible to this mechanism of resistance development because the biocides had a sufficient number of charges per molecule to overcome the bacterial neutralization defense3, 6. We expected that pQA linear polymers and HSA-pOEGMA-b-pQA conjugates would both interact strongly with the negatively charged cell surface and cellular membrane before resistance developed. Upon successive passages of


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Biomacromolecules

bacteria exposed to, but not killed by the polymeric biocide, partial neutralization of these surfaces would result in a decreased ability of the polymeric biocide to kill the bacteria. S. aureus could generate a more neutralized cell wall by incorporating of D-alanine into teichoic acids and a more neutralized membrane incorporating D-lysine29,32. In considering why resistance generation by bacteria against HSA-pOEGMA-b-pQA might be delayed, we hypothesized that the increased size of the conjugate, relative to the linear polymer, would be able to draw membrane anions together from a larger surface area and still kill cells with partially neutralized membranes. The hydrodynamic diameter of HSA-pOEGMA20-b-pQA-C272 was approximately 140 nm, with a zeta potential of +32 mV. The calculated charge density (48 chains each with 72 QA repeat units) was approximately 5.6 x 1012 charges/cm2. On surfaces, we have shown that a threshold of 2.5 x 1012 charges/cm2 was needed to kill E. coli. Our data were consistent with the idea that the conjugates disrupted cell membranes by counter ion exchange rather than by membrane penetration23.

CONCLUSIONS We investigated in detail the bactericidal properties of a library of pQA linear polymers and their protein-polymer conjugates. We observed the dependence of killing efficiency on polymer structure and architecture, affording us the opportunity to generate structure-function relationships for polymeric cationic biocides. Polymerization through ATRP allowed the facile synthesis of pQA-based polymeric biocides with varying alkyl side chain lengths, MWs and hydrophobicity. pQA linear polymers with shorter alkyl side chains efficiently killed Grampositive bacteria, while those with longer alkyl side chains were more effective against Gramnegative bacteria, with varying levels of hemolytic and cytotoxic activity. HSA-pOEGMA-b-


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pQA conjugates generated using polymer-based protein engineering provided a nanoparticulate biocide with extraordinarily high charge density. The conjugates were biocidal and also dramatically slowed the development of bacterial resistance. E. coli killing by pQA linear polymers resulted in minimal development of resistance, while MRSA became resistant to these polymeric biocides after several passages. However, utilization of HSA-pOEGMA-b-pQA conjugates reduced the rate of resistance generation in MRSA. As we learn more about structurefunction properties in protein-polymeric biocides, we will be well placed to synthesize broad specificity, resistance-preventing and biocompatible biocides.


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ASSOCIATED CONTENT Supporting Information. Synthetic route of QA monomer, serum compatibility of pQA linear polymers and HSA-pOEGMA-b-pQA conjugates, 1H-NMR spectra of native and initiatormodified HSA, 1H-NMR spectra of representative pQA polymers, MALDI-TOF MS spectra of native and initiator-modified HSA, polymerization conditions and resulting characteristics of pQA linear polymers and HSA-pOEGMA-b-pQA conjugates, and MIC values for QA monomers and pOEGMA controls. This material is available free of charge via the Internet at http://pubs.acs.org.

AUTHOR INFORMATION Corresponding Author *E-mail: [email protected]; Tel: +1 412 268 9607; Fax: +1 412 268 1173 ORCID Alan J. Russell: 0000-0001-5101-4371 Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Funding The authors would like to thank the Institute of Transfusion Medicine (ITxM, Pittsburgh, PA) and the Center for Polymer-based Protein Engineering (Carnegie Mellon University, Pittsburgh,


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PA) for funding this work. We also gratefully acknowledge the Central Blood Bank (Pittsburgh, PA) for their supply of healthy erythrocytes.

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Biomacromolecules

9

10

10

10

A 10

B 10

9

8

8

10

7



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48

7

10

6

10

5

10

4

10

3

10

2

10

1

10

10

6

10

5

10

4

10

3

10

2

10

1

10

0

10

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10

-1

-1

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0

1

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3

4

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6

24

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2.4

A 2.2


2.0

Fold increse of MIC

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1.8 1.6 1.4 1.2 1.0

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0.8 2

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Cell concentration (cfu/mL)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48

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300 400


A 250 Fold increase of MIC

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48

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2

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Bactericidal Specificity and Resistance Profile of ... - ACS Publications

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