Lipopolysaccharide Neutralization by Cationic-Amphiphilic Polymers

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Lipopolysaccharide Neutralization by Cationic-amphiphilic Polymers through Pseudo-aggregate Formation Divakara S S M Uppu, and Jayanta Haldar Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.5b01567 • Publication Date (Web): 03 Feb 2016 Downloaded from http://pubs.acs.org on February 8, 2016

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Lipopolysaccharide Neutralization by Cationic-amphiphilic Polymers through Pseudo-aggregate Formation Divakara SSM Uppu and Jayanta Haldar* Chemical Biology & Medicinal Chemistry Laboratory, New Chemistry Unit (NCU), Jawaharlal Nehru Centre for Advanced Scientific Research (JNCASR), Jakkur, Bengaluru, Karnataka, India-560064.

ABSTRACT Synthetic polymers incorporating the cationic charge and hydrophobicity to mimic the function of antimicrobial peptides (AMPs) have been developed. These cationic-amphiphilic polymers bind to bacterial membranes that generally contain negatively charged phospholipids and cause membrane disintegration resulting in cell death. However, cationic-amphiphilic antibacterial polymers with endotoxin neutralization properties, to the best of our knowledge, have not been reported. Bacterial endotoxins such as lipopolysaccharide (LPS) cause sepsis that is responsible for a great amount of mortality worldwide. These cationic-amphiphilic polymers can also bind to negatively charged and hydrophobic LPS and cause detoxification. Hence, we envisaged that cationic-amphiphilic polymers can have both antibacterial as well as LPS binding properties. Here, we report synthetic amphiphilic polymers with both antibacterial as well as endotoxin neutralizing properties. Levels of pro-inflammatory cytokines in human monocytes caused by LPS stimulation were inhibited by more than 80% when co-incubated with these polymers. These reductions were found to be dependent on concentration and more importantly, on the side chain chemical structure due to variations in the hydrophobicity profiles of these polymers. These cationic-amphiphilic polymers bind and cause LPS neutralization and detoxification. Investigations of polymer interaction with LPS using fluorescence spectroscopy and dynamic light scattering (DLS) showed that these polymers bind but neither dissociate nor promote LPS 1 ACS Paragon Plus Environment

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aggregation. We show that polymer binding to LPS leads to sort of a pseudo-aggregate formation resulting in LPS neutralization/detoxification. These findings provide an unusual mechanism of LPS neutralization using novel synthetic cationic-amphiphilic polymers.

INTRODUCTION Flourishing bacterial resistance to antibiotics coupled with languishing antibiotic discovery creates an urgent need for the development of new antimicrobial agents which exert novel mechanisms of action.1 In the past two decades, antimicrobial peptides (AMPs) have received increasing attention due to their broad-spectrum activities and ability to combat multidrug-resistant microbes.2-7 Apart from broad spectrum antimicrobial activity, AMPs also possess anti-endotoxin/anti-inflammatory properties. For e.g. AMPs such as LL-37, Indolicidin human lactoferrin-derived peptide8 and Buforin II bind to bacterial endotoxins (pathogen associated molecular patterns (PAMPs)) such as lipopolysaccharide (LPS) from Gram-negative bacteria that otherwise result in uncontrollable and harmful inflammation leading to sepsis.9-14 This uncontrolled inflammation is due increased production of pro-inflammatory cytokines such as TNF-α (tumor necrosis factor- α), IL-6 (Interleukin-6) from macrophages/monocytes that cause septic shock.9, 10 Globally, an estimated 18 million cases of sepsis occur each year resulting in 8 million deaths.15 LPS, a major component of outer membrane of Gram-negative bacteria, is a negatively charged hydrophobic molecule that spontaneously self-assembles to form aggregates in aqueous solutions.9,

10

Recognition of LPS by either soluble or cell surface receptors need the native

aggregate structure of LPS.9, 10 Most of the amphiphilic molecules (including AMPs) reported till now interact with LPS aggregates accompanied by aggregate dissociation12,

2 ACS Paragon Plus Environment

16-23

whereas

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Polymyxin B (PMB),24 rBPI2125 and ultrashort AMPs26 have been shown to promote LPS aggregation and hence inhibit cytokine secretion. Small molecular antibacterials have been shown to possess anti-inflammatory27-31 or immunomodulatory properties.32-34 Polyvalent dendrimer glucosamine conjugates with immunomodulatory properties have also been reported.35 In some of the cases the molecular mechanism of inhibiting cytokine secretion has not been investigated. Cationic-amphiphilic polymers mimicking the AMPs with potent antibacterial activity and low toxicity have extensively been reported.36-55 However, cationic and amphiphilic antibacterial polymers with anti-endotoxin properties, to the best of our knowledge, have not been reported. The advantage of using polymers is due their multivalent (hydrophobic, electrostatic and hydrogen bonding interactions) nature. We envisaged that amphiphilic polymers with cationic charge, hydrophobicity and amide moieties can efficiently bind to negatively charged hydrophobic LPS thereby prevent its binding to the receptors and hence inhibit cytokine secretion. Here, we report amphiphilic polymers based on poly(isobutylene-alt-N-alkyl maleimide) backbone with both antibacterial and anti-endotoxin properties. The advantages of using maleic anhydride copolymers are due to their biocompatibility, easy functionalization, low cost and US FDA (United States Food and Drug Administration) approved for human use.56,57 We observe that the side chain chemical structure of these amphiphilic polymers not only plays role in their antibacterial and cytotoxic activities but also in their LPS neutralization and detoxification properties leading to inhibiting cytokine secretion (TNF-α and IL-6) in human monocytes. Interestingly, using fluorescence spectroscopy, dynamic light scattering (DLS) and electron microscopy, we show that these amphiphilic polymers interact with LPS aggregates but rather do not lead to aggregate dissociation.


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MATERIALS AND METHODS All the solvents were of reagent grade and dried prior to use wherever required. Poly(isobutylene-alt-maleic anhydride) (Mw ~ 6000 Da, Catalog no. 531278) purchased from Sigma-Aldrich and used as received. All other reagents were procured from the commercial suppliers and used as received. Culture media and the antibiotics were from HIMEDIA (India) and Sigma-Aldrich respectively. NMR spectra were recorded using Bruker AMX-400 (400 MHz for 1H and 100 MHz for

13

C) spectrometer. The chemical shifts (δ) are reported in parts per

million downfield from the peak for the internal standard TMS for 1H-NMR. Infrared (IR) spectra of the solid compounds were recorded on Bruker IFS66 V/s spectrometer using KBr pellets. IR spectra of the compounds soluble in low-boiling solvents were recorded with the same instrument using NaCl crystal. Mass spectra were recorded on a Micromass Q-ToF micromass spectrometer. Optical density and absorbance were measured by Tecan InfinitePro series M200 Microplate Reader. Bacterial strains S. aureus (MTCC 737) and E. coli (MTCC 443 equivalent to ATCC 25922) were purchased from MTCC (Chandigarh, India). MRSA (ATCC 33591), vancomycin resistant E. faecium (VRE) ((OrlaJensen) Schleifer and Kilpper-Balz, ATCC 51559) were obtained from ATCC (Rockville, Md). E. coli was cultured in Luria Bertani broth (10 g of tryptone, 5 g of yeast extract, and 10 g of NaCl in 1000 mL of sterile distilled water while S. aureus and MRSA were grown in nutrient broth (1 g of beef extract, 2 g of yeast extract, 5 g of peptone and 5 g of NaCl in 1000 mL of sterile distilled water). VRE was cultured in Brain Heart Infusion broth (BHI). For solid media, 5% agar was used along with above mentioned composition. The bacterial samples were freeze dried and stored at -80 °C. 5 μL of these stocks were added to 3 mL of the nutrient broth and the culture was grown for 6 h at 37 °C prior to the experiments.


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Synthesis and characterization. Synthesis and characterization of side chain alkylating agents are provided in supporting information. Synthesis

of

polymeric

derivatives.

Poly(isobutylene-alt-N-(N',N'-dimethylaminopropyl)-

maleimide) (PIBMI). To a solution of 10 g of poly(isobutylene-alt-maleic anhydride) (PIBMA) (Avg. Mw = 6000 g/mol) in 60 mL of DMF, 7.96 g of 3-aminopropyldimethylamine (1.2 equivalents with respect to the monomer weight of the polymer (154 g/mol)) was added and stirred at 120 °C for 48 h in a screw-top pressure tube. The reaction mixture was cooled, precipitated with 200 mL of distilled water and was centrifuged at 10,000 rpm for 15 min. The polymer was dried at 55 °C for 24 h under vacuum to give a pale yellow solid with 100% yield (complete conversion of the anhydride to imide was confirmed by complete disappearance of peaks at 1850 cm-1 (C=O asym. str.) and 1785 (C=O sym. str.) for the anhydride ring and appearance of peaks 1767 cm-1 (C=O asym. str.), 1696 cm-1 (C=O sym. str.) for the imide ring by FT-IR). PIBMA: FT-IR: 2950-2850 (C-H str.), 1850 cm-1 (C=O asym. str.), 1785 cm-1 (C=O sym. str.), 1470-1410cm-1 (C-C str.), 1290-1110 (C-O str.); 1H-NMR (400 MHz, (CD3)2SO): δ/ppm 0.7–1.2 (br CH2C(CH3)2, 6H), 1.7 (br CH2C(CH3)2, 2H), 2.2 (br, CHCH, 1H), 3.1 (br, CHCH, 1H). PIBMI: FT-IR: 2950-2850 cm-1 (C-H str.), 1767 cm-1 (C=O asym. str.), 1696 cm-1 (C=O sym. str.), 1470-1410 cm-1 (C-C str.), 1290-1110 cm-1 (C-O str.); 1H-NMR (400 MHz, CDCl3): δ/ppm 0.7–1.2 (br CH2C(CH3)2, 6H), 1.7 (br CH2C(CH3)2, 2H), 1.86 (br NCH2CH2CH2N(CH3)2, 2H), 2.2-2.5

(br

NCH2CH2CH2N(CH3)2,

8H),

2.7–3.1

(br,

CHCH,

2H),

3.6

(br

NCH2CH2CH2N(CH3)2, 2H). Synthesis of alkyl chain quaternized polymers. To a solution of 0.5 g of PIBMI in 20 mL of DMF/CHCl3 (1:1), 1.04 g of 1-bromopentane/ 1.15 g of 1-bromo-2(2-methoxyethoxy) ethane (3


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equivalents with respect to the monomer weight of PIBMI (238.18 g/mol)) was added and stirred at 75 °C for 96 h in a screw top pressure tube. The solution was cooled, precipitated with 40 mL of n-hexane/diethyl ether and filtered. The white solid was washed with n-hexane (4 × 40 mL)/diethylether and dried at 40 °C for 12 h under vacuum (yield: 100%). Qn-PenP: FT-IR: 2950-2850 (C-H str.), 1767 cm-1 (C=O asym. str.), 1696 cm-1 (C=O sym. str.) 1470-1410cm-1 (C-C str.), 1290-1110 (C-O str.); 1HNMR (400 MHz, D2O): δ/ppm 0.85-0.9 (br terminal –CH3, 3H), 0.95–1.2 (br CH2C(CH3)2, 6H), 1.3-1.5 (br CH2CH2CH2, 6H) 1.7 (br CH2C(CH3)2, 2H), 2.0 (br NCH2CH2CH2N(CH3)2, 2H), 2.7–3.1 (br CHCH, 2H), 3.1-3.3 (br NCH2CH2CH2NCH2(CH3)2, 10H), 3.6 (br NCH2CH2CH2N(CH3)2, 2H). Qn-BEGP: FT-IR: 2950-2850 (C-H str.), 1767 cm-1 (C=O asym. str.), 1696 cm-1 (C=O sym. str.) 1470-1410cm-1 (C-C str.), 1290-1110 (C-O str.); 1HNMR (400 MHz, D2O): δ/ppm 0.95–1.2 (br CH2C(CH3)2, 6H), 1.7 (br CH2C(CH3)2, 2H), 2.0 (br NCH2CH2CH2N(CH3)2, 2H), 2.7–3.1 (br CHCH, 2H), 3.1-3.2 (br NCH2(CH3)2, 6H), 3.45 (s, terminal –CH3), 3.55-3.8 (br, NCH2CH2CH2NCH2(CH3)2 and OCH2CH2O, 10H), 4.0 (br OCH2CH2N(CH3)2, 2H). Synthesis of amide or ester quaternized polymers. To a solution of 0.5 g of PIBMI in 20 mL of dry DMF : dry CHCl3 (1:1), 2 equivalents (with respect to the monomer weight of PIBMI) of Nalkyl-1-bromoethanamide or N-propyl-1-bromoethanoate was added and stirred at 75 °C (amide) or 65 °C (ester) for 96 h in a screw top pressure tube. The solution was cooled, precipitated with 40 mL of diethylether and filtered. The white solid was washed with diethylether (4 × 40 mL) and dried at 40 °C for 6 h under vacuum. The percentage of conversion given by the degree of quaternization was calculated from 1H NMR and was found to be in the range of 90-95% for all the quaternized polymers.


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Qn-prAP: FT-IR: 3250 cm-1 (amide N-H str.), 2950-2850 (C-H str.), 1767 cm-1 (imide C=O asym. str.), 1696 cm-1 (imide C=O sym. str.) 1680 cm-1 (amide I, C=O str.), 1560 cm-1 (Amide II, N-H ben.), 1470-1410cm-1 (C-C str.), 1290-1110 (C-O str.); 1HNMR (400 MHz, D2O): δ/ppm 0.878 (br, terminal –CH3, 3H), 0.95–1.2 (br, CH2C(CH3)2, 6H), 1.543 (br, -CONHCH2CH2CH3, 2H), 1.7 (br, CH2C(CH3)2, 2H), 2.0 (br, NCH2CH2CH2N(CH3)2, 2H), 2.7–3.1 (br, CHCH, 2H), 3.1-3.3

(br,

NCH2CH2CH2N(CH3)2,

8H),

3.5

(br,

-CONHCH2-,

2H),

3.6

(br,

NCH2CH2CH2N(CH3)2, 2H), 3.8 (br, -N(CH3)2CH2CO, 2H). Qn-prEP: FT-IR: 2950-2850 (C-H str.), 1767 cm-1 (imide C=O asym. str.), 1696 cm-1 (imide C=O sym. str.), 1735 cm-1 (ester C=O str.) 1470-1410cm-1 (C-C str.), 1290-1110 (C-O str.); 1

HNMR (400 MHz, D2O): δ/ppm 0.85 (br, terminal –CH3, 3H), 0.95–1.2 (br, CH2C(CH3)2, 6H),

1.57 (br,-COOCH2CH2CH3, 2H), 1.7 (br, CH2C(CH3)2, 2H), 2.0 (br, NCH2CH2CH2N(CH3)2, 2H), 2.7–3.1 (br, CHCH, 2H), 3.1-3.3 (br, NCH2CH2CH2N(CH3)2, 8H), 3.6 (br, NCH2CH2CH2N(CH3)2, 2H), 3.7 (br, -N(CH3)2COCH2, 2H) 4.0 (br, -COOCH2-, 2H). Qi-prAP: FT-IR: 3250 cm-1 (amide N-H str.), 2950-2850 cm-1 (C-H str.), 1767 cm-1 (imide C=O asym. str.), 1696 cm-1 (imide C=O sym. str.) 1680 cm-1 (amide I, C=O str.), 1560 cm-1 (Amide II, N-H ben.), 1470-1410 cm-1 (C-C str.), 1290-1110 cm-1 (C-O str.); 1H-NMR (400 MHz, D2O): δ/ppm 0.92–1.4 (br, CH2C(CH3)2, 6H), 1.19 (br, CH(CH3)2, 6H), 1.7 (br, CH2C(CH3)2, 2H), 2.1 (br, NCH2CH2CH2N(CH3)2, 2H), 2.8–3.2 (br, CHCH, 2H), 3.2-3.4 (br, NCH2CH2CH2N(CH3)2, 6H), 3.6 (br, NCH2CH2CH2N(CH3)2, 2H), 3.6 (br, NCH2CH2CH2N(CH3)2, 2H), 4.06 (m, – CH(CH3)2, 1H), 4.1 (br, -N(CH3)2CH2CO, 2H). Qn-prenAP: FT-IR: 3250 cm-1 (amide N-H str.), 2950-2850 cm-1 (C-H str.), 1767 cm-1 (imide C=O asym. str.), 1696 cm-1 (imide C=O sym. str.) 1680 cm-1 (amide I, C=O str.), 1560 cm-1 (Amide II, N-H ben.), 1470-1410 cm-1 (C-C str.), 1290-1110 cm-1 (C-O str.); 1H-NMR (400


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MHz, D2O): δ/ppm 0.92–1.4 (br, CH2C(CH3)2, 6H), 1.7 (br, CH2C(CH3)2, 2H), 2.1 (br, NCH2CH2CH2N(CH3)2, 2H), 2.8–3.2 (br, CHCH, 2H), 3.2-3.4 (br, NCH2CH2CH2N(CH3)2, 6H), 3.6 (br, NCH2CH2CH2N(CH3)2, 2H), 3.6 (br, NCH2CH2CH2N(CH3)2, 2H), 3.85 (br, N(CH3)2CH2CO, 2H), 4.15 (br, -CONHCH2-, 2H), 5.30 (br, -CH2CH=CH2, 2H), 5.85 (br, CH2CH=CH2, 1H). QCyprAP: FT-IR: 3250 cm-1 (amide N-H str.), 2950-2850 cm-1 (C-H str.), 1767 cm-1 (imide C=O asym. str.), 1696 cm-1 (imide C=O sym. str.) 1680 cm-1 (amide I, C=O str.), 1560 cm-1 (Amide II, N-H ben.), 1470-1410 cm-1 (C-C str.), 1290-1110 cm-1 (C-O str.); 1H-NMR (400 MHz, D2O): δ/ppm 0.64 (br, cyCH2, 2H), 0.86(br, cyCH2, 2H), 0.92–1.4 (br, CH2C(CH3)2, 6H), 1.7 (br, CH2C(CH3)2, 2H), 2.1 (br, NCH2CH2CH2N(CH3)2, 2H), 2.73 (m, cyCH, 1H), 2.8–3.2 (br, CHCH, 2H), 3.2-3.4 (br, NCH2CH2CH2N(CH3)2, 6H), 3.6 (br, NCH2CH2CH2N(CH3)2, 2H), 3.6 (br, NCH2CH2CH2N(CH3)2, 2H), 4.1 (br, -N(CH3)2CH2CO, 2H). QCybuAP: FT-IR: 3250 cm-1 (amide N-H str.), 2950-2850 cm-1 (C-H str.), 1767 cm-1 (imide C=O asym. str.), 1696 cm-1 (imide C=O sym. str.) 1680 cm-1 (amide I, C=O str.), 1560 cm-1 (Amide II, N-H ben.), 1470-1410 cm-1 (C-C str.), 1290-1110 cm-1 (C-O str.); 1H-NMR (400 MHz, D2O): δ/ppm 0.92–1.4 (br, CH2C(CH3)2, 6H), 1.7 (br, -CH2C(CH3)2, 2H), 1.77 (br, cyCH2, 2H), 2.0 (br, cyCH2, 2H), 2.1 (br, NCH2CH2CH2N(CH3)2, 2H), 2.29 (br, -CH(CH2)3, 2H), 2.8–3.2 (br, CHCH, 2H), 3.2-3.4 (br, NCH2CH2CH2N(CH3)2, 6H), 3.6 (br, NCH2CH2CH2N(CH3)2, 2H), 3.6 (br, NCH2CH2CH2N(CH3)2, 2H), 4.06 (br, -N(CH3)2CH2CO, 2H), 4.24 (br, cyCH, 1H). Qn-buenAP: FT-IR: 3250 cm-1 (amide N-H str.), 3049 cm-1 (C-H str.), 1767 cm-1 (imide C=O asym. str.), 1696 cm-1 (imide C=O sym. str.),1680 cm-1 (Amide I, C=O str.), 1642 cm-1 (C=C str.), 1560 cm-1 (Amide II, N-H ben.), 1470-1410 cm-1 (C-C str.), 1290-1110 cm-1 (C-O str.); 1H-


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NMR (400 MHz, D2O): δ/ppm 0.96–1.27 (br, CH2C(CH3)2, 6H), 1.7 (br, CH2C(CH3)2, 2H), 2.0 (br, NCH2CH2CH2N(CH3)2, 2H), 2.3 (br, -CONHCH2CH2-, 2H), 2.7–3.1 (br, CHCH, 2H), 3.29 (br,

NCH2CH2CH2N(CH3)2,

6H),

3.35

(br,

-CONHCH2CH2-,

2H),

3.6

(br,

NCH2CH2CH2N(CH3)2, 2H), 3.6 (br, NCH2CH2CH2N(CH3)2, 2H), 4.10 (br, -N(CH3)2CH2CO, 2H), 5.17 (br, CH2CH2CH=CH2, 2H), 5.85 (m, CH2CH2CH=CH2, 1H). Qn-buynAP: FT-IR: 3250 cm-1 (amide N-H str.), 3310 cm-1 (≡C-H str.), 2857-2942 cm-1 (-C-H str.), 2119 cm-1 (C≡C str.), 1767 cm-1 (imide C=O asym. str.), 1696 cm-1 (imide C=O sym. str.), 1680 cm-1 (Amide I, C=O str.), 1642 cm-1 (C=C str.), 1560 cm-1 (Amide II, N-H ben.), 14701410 cm-1 (C-C str.), 1290-1110 cm-1 (C-O str.); 1H-NMR (400 MHz, D2O): δ/ppm 0.96–1.27 (br, CH2C(CH3)2, 6H), 1.7 (br, CH2C(CH3)2, 2H), 1.82 (br, -CH2C≡CH, 1H), 2.1 (br, NCH2CH2CH2N(CH3)2, 2H), 2.29 (br, -CONHCH2CH2-, 2H), 2.7–3.1 (br, CHCH, 2H), 3.2-3.4 (br,

NCH2CH2CH2N(CH3)2,

6H),

3.37

(br,

-CONHCH2CH2-,

2H),

3.6

(br,

NCH2CH2CH2N(CH3)2, 2H), 3.6 (br, NCH2CH2CH2N(CH3)2, 2H), 4.16 (br, -N(CH3)2CH2CO, 2H). Gel Permeation Chromatography (GPC) GPC of the water soluble Poly(isobutylene-alt-maleic acid) obtained after hydrolysis of Poly(isobutylene-alt-maleic anhydride) was performed. Experiments were carried out on a Shimadzu-LC 20AD instrument with refractive index (RI) detector using Polysep-GPC-P Linear (300×7.8 mm, Phenomenex, Catalogue no. 00H-3147-K0, Separation Range: 1 KDa - 10 MDa) column in sodium acetate buffer (0.2 M, pH = 5.3) with a flow rate of 0.8 mL min-1. Pullulan standards (1.3 KDa, 6 KDa, 10 KDa, 22 KDa and 50 KDa) were used for the experiment to determine the polydispersity index (PDI = Mw/Mn) of the polymer.


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Calculation of degree of quaternization and molecular weight. The degree of quaternization was calculated by 1H NMR analysis of the tertiary nitrogen of PIBMI and quaternized derivative, QAPIBMI (Fig. 1). The NCH2(CH3)2 protons shift downfield after quaternization and by integrating the protons corresponding to the unquaternized NCH2(CH3)2, the degree of quaternization was calculated (Fig. 1). Degree of quaternization of the polymeric derivatives was calculated using 1H NMR analysis (Fig. 1). Degree of quaternization (x) = (1- y) × 100 % Wherein

y = {([CH2N(CH3)2]8)/ ([CH2CH2]/2)} y = {(m/8)/ (n/2)}, m = [CH2N(CH3)2] and n = [CH2CH2]

For e.g. QCyprAP, Degree of quaternization (x) = 1-{(0.589/8) / (2.075/2)} × 100 % = 93% Wherein, [CH2CH2] is the integrals of the hydrogens, a & a and [CH2N(CH3)2] is the integrals of the hydrogens, b that are bold and italicized. The molecular weight (number average molecular weight, Mn) (Table 1) of all the derivatives is calculated based on the molecular weight of the precursor polymer, poly(isobutylene-alt-maleic anhydride) (average Mw ~ 6000 Da, n~39 and PDI ~ 1.2 from GPC) and the obtained degree of quaternization (% of DQ) as described above. For non-quaternized polymer, PIBMI: Mn = m y × n = 238.18 × 39 = ~9300 Da


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For quaternized polymer, e.g. QCyprAP Mn = [(mx × 0.93) + {my × (1-0.93)}] n = [(415.16 × 0.93) + (238.18 × 0.07)] 39 = 15.68 kDa Where mx and my are the molecular weights of the quaternized and non-quaternized repeating units of the polymer (Fig. 1) respectively. Antibacterial activity58-60. Antibacterial activity of the polymers was measured and MIC was calculated as per CLSI guidelines. Antibacterial activity of polymers was assayed in a modified micro-dilution broth format. Stock solutions were made by serially diluting the test compounds using autoclaved Millipore water. Bacteria, to be tested, grown for 6 h in the suitable media contained ~109 CFU mL-1 (determined by spread plating method), which was then diluted to ~105 CFU mL-1 using cation adjusted Mueller-Hinton broth (CAMHB) as per CLSI guidelines. 50 μL of serially diluted compound was added to a 96 well plate (Polystyrene) containing 150 μL bacterial solutions. Two controls were made; one containing 150 μL of media and 50 μL of compound and the other containing 150 μL of bacterial solution and 50 μL water . The plate was then incubated at 37 C for a period of 24 h and the O.D. value was measured at 600 nm using a Tecan InfinitePro series M200 Microplate Reader. MIC value was determined by taking the average of triplicate O.D. values for each concentration and plotting it against concentration using Origin Pro 8.0 software. The data was then subjected to sigmoidal fitting. From the curve the MIC value was determined, as the point in the curve where the O.D. was similar to that of control having no bacteria. MIC curves for individual agents are representative data from the two independent experiments and each experiment was performed in triplicates.


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Hemolytic activity. The hemolytic activity was determined against human erythrocytes. Erythrocytes were isolated from freshly drawn, heparanized human blood and resuspended to 5 % v/v in PBS (pH 7.4). In a 96-well micro titer plate, 150 μL of erythrocyte suspension was added followed by 50 μL of serially diluted compound to give a final solution of 3.75 % v/v erythrocytes. PBS buffer was added instead of polymer solution as negative hemolysis control and Triton X-100 (1% v/v) was used as positive hemolysis control. The plate was incubated for 1 h at 37 °C and was then centrifuged at 3,500 rpm for 5 min. 100 μL of the supernatant was then transferred to a fresh micro titer plate and absorbance at 540 nm was measured using a Tecan InfinitePro series M200 Micro plate Reader. Percentage of hemolysis was determined as (A - A0)/ (Atotal -A0) x 100, where A is the absorbance of the test well, A0 the absorbance of the negative controls, and Atotal the absorbance of 100% hemolysis wells, all at 540 nm. Hemolysis was plotted as a function of polymer concentration and the HC50 was defined as the polymer concentration, which causes 50% hemolysis relative to the positive control. In some cases, hemolysis did not reach 50% up to the highest polymer concentration tested and the HC50 was not determined. Hemolysis curves for each polymer are representative data from two independent experiments and each experiment was performed in triplicates. In-vitro cytotoxicity. CytoTox 96 Non-Radioactive Cytotoxicity Assay (Promega) kit was used for determining the cytotoxicity of the compounds. In brief, HEK293 cells that were maintained in complete DMEM media (Gibco) supplemented with 10% FBS (Gibco) and PenicillinStreptomycin solution (Gibco) were seeded in 96 well plates at a concentration of 104 cells/well. They were allowed to adhere to the plate overnight. 0.5% Triton-X and media were used as positive and untreated controls respectively. The cells were treated with respective test compound solutions. After 24 hrs of treatment, the plates were centrifuged at 1100 rpm for 5


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min. The supernatants from respective wells were transferred and the assay was performed according to the manufacturer’s instructions. 100 μL of the supernatant was then transferred to a fresh micro titer plate and absorbance at 490 nm was measured using a Tecan InfinitePro series M200 Micro plate Reader. Percentage of cell death was determined as (A - A0)/ (Atotal -A0) x 100, where A is the absorbance of the test well, A0 the absorbance of the negative controls, and Atotal the absorbance of triton-X treated wells, all at 490 nm. Percentage of LDH release was plotted as a function of polymer concentration and the IC50 was defined as the polymer concentration, which causes 50% LDH release relative to the positive control. In some cases, LDH release did not reach 50% up to the highest polymer concentration tested and the IC50 was not determined. Membrane-active properties Cytoplasmic membrane depolarization assay. Bacteria were harvested, washed with 5 mM HEPES and 5 mM glucose and resuspended in 5 mM glucose, 5 mM HEPES buffer and 100 mM KCl solution in 1:1:1 ratio (~108-9 CFU mL-1). Measurements were made in a Corning 96 well black plate with clear bottom with 150 µl of bacterial suspension and 2 µM of DiSC3(5). For Gram-negative bacteria, 0.2 mM of EDTA was used to permeabilize the outer membrane and allow the dye uptake. The fluorescence of the dye was monitored using a Tecan InfinitePro series M200 Micro plate Reader at excitation wavelength of 622 nm and emission wavelength of 670 nm. Dye uptake, and resultant self quenching, was modulated by the membrane potential. After reaching the maximum uptake of the dye by bacteria, which was indicated by a minimum in dye fluorescence, polymer solution was added to the cells, and the decrease in potential was monitored by increase in fluorescence. All the other test compounds were dissolved in water at 4 mg mL-1 and DiSC3(5) dissolved in DMSO were further diluted in the above 5 mM glucose, 5


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mM HEPES buffer and 100 mM KCl solution in 1:1:1 ratio. A control without the polymers was served as negative control. Cytoplasmic membrane permeabilization assay. Bacteria were harvested, washed, and resuspended in 5 mM HEPES and 5 mM glucose buffer of pH 7.2 (~108-9 CFU mL-1). Then, 150 µl of bacterial suspension, 10 μM propidium iodide (PI) and polymer solution were added to the cells in a Corning 96 well black plate with clear bottom. Stock solutions of PI and the polymers were made in water and further diluted in HEPES. Excitation wavelength of 535 nm and emission wavelength of 617 nm were used. The uptake of PI was measured using a Tecan InfinitePro series M200 Microplate Reader by the increase in fluorescence of PI for 30 min as a measure of membrane permeabilization. Release of ATP levels. Mid-log phase bacteria (~108-9 CFU mL-1) were harvested and washed twice with 10 mM TRIS buffer (pH = 7.5) and were resuspended in the same buffer. Then, 150 µl of bacterial suspension and 50 µl of test drugs were added to the micro centrifuge tube and incubated at 37 °C for 15 min. After 15 min, the bacterial suspension was centrifuged and 50 µl of the supernatant was transferred into a Corning 96 well black plate with clear bottom to find out the released ATP levels using ATP Bioluminescence Assay Kit (Sigma Aldrich) as per the manufacturer’s instructions. A standard curve for ATP levels was generated using the ATP standards provided in the kit in the range of 1E-6 to 1E-11 moles of ATP levels. Stock solutions of the polymers were made in water and further diluted in water. Relative ATP levels both in the standard curve and the test sample measurement were measured by subtracting the background ATP levels from the test sample ATP levels as per the manufacturer’s instructions. All measurements were performed in duplicates using Tecan InfinitePro series M200 Microplate Reader.


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Reverse-phase high performance liquid chromatography (RP-HPLC). Chromatographic profiles were analyzed by reverse phase HPLC using 0.1% trifluoroacetic acid (TFA) in water/acetonitrile (0−100%) as mobile phase. HPLC analysis was performed on a Shimadzu-LC 8 Å liquid chromatography instrument (C18 column, 10 mm diameter, 250 mm length) with UV detector monitoring at 220 nm. The data was acquired from 0-40 min and presented from 5-30 min. In-vitro cytotoxicity against human PBMCs. Fresh human blood was drawn and human peripheral blood mononuclear cells (PBMCs) were isolated using a standard Ficoll-Hypaque density centrifugation technique and the number of PBMCs (and viability) was determined by trypan blue exclusion. More than 95% of cells were viable. All the media components used were certified and contained low endotoxin levels. After isolation, human PBMCs were resuspended in RPMI 1640 growth medium (with L-glutamine and sodium bicarbonate, Gibco, Life Technologies) supplemented with 10% of heat inactivated and low endotoxin fetal bovine serum (FBS, Life Technologies) and 1% penicillin-streptomycin at 37 C in a humidified-air atmosphere (5% CO2/95% air). Human PBMCs (2×104) were seeded into 96 well plates and incubated overnight at 37 C in a humidified-air atmosphere (5% CO2/95% air). The release of cytosolic LDH was then assessed after 24 h of incubation with polymeric derivatives using concentrations up to 100 μg mL-1. Stimulation of human PBMCs with LPS. Freshly isolated human PBMCs were seeded into 24well plates (1×106 cells) in 1 mL of RPMI 1640 complete medium. After 3 hr of resting, the cells were stimulated with 20 ng mL-1 of E. coli 0111:B4 LPS (Sigma Aldrich) either in the presence or absence of polymeric derivatives (different concentrations). A control experiment was performed using HBSS (Life Technologies) as vehicle control. The cells were incubated for 18-


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24 hours and then cell culture supernatants were analyzed for cytokines such as tumor necrosis factor (TNF-α) and interleukin- 6 (IL-6) using the human ELISA kits (BD Biosciences) following the manufacturer’s instructions. LPS binding and dissociation. Bodipy conjugated LPS (from E. coli, Molecular Probes, Life Technologies) was used for studying the interaction with polymers. Stock solution of bodipyLPS (100 µg/mL) was prepared and diluted to 10 µg/mL in 1X PBS (pH= 7.4). Bodipy-LPS stock solutions were sonicated for 2 min before use. 2 mL of solution was taken in a quartz cuvette containing bodipy-LPS (500 ng mL-1) and the test compounds in PBS. Experiments were performed using a λS55 Fluorescence Spectrophotometer (Perkin Elmer) at an excitation wavelength of 485 nm and emission was collected from 500-700 nm (excitation slit width = 5 nm, emission slit width = 15 nm) at room temperature. Dynamic light scattering (DLS). E. coli 0111:B4 LPS and polymers were dissolved in Hank’s balanced salt solution (HBSS without Ca2+ and Mg2+). Experiments were performed using Zetasizer Nano Z (Malvern Instruments) at room temperature. LPS and the polymers were taken in disposable cuvettes in 2 mL of solution. Data were acquired for 200 scans.

RESULTS AND DISCUSSION Synthesis and characterization of cationic-amphiphilic polymers. A series of water soluble amphiphilic polymers were synthesized based on quaternized poly(isobutylene-alt-N-alkyl maleimide) (PIBMI) using facile synthetic methodology of post-functionalization (Scheme 1). The conventional approaches use cationic and hydrophobic monomers as precursors to synthesize amphiphilic antibacterial polymers. On the other hand, our post-functionalization approach incorporates cationic charge and hydrophobicity to a precursor polymer to generate 16 ACS Paragon Plus Environment

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amphiphilic antibacterial polymers. The advantage of this approach is that the molecular properties of the polymer can be controlled just by changing the appended side chain chemical structure. The nucleophilic ring opening and closure of highly reactive anhydride of poly (isobutylene-alt-maleic anhydride) (PIBMA) with 3-aminopropyldimethylamine was achieved in a single step by heating at 120 °C for 48 h (Scheme 1). The complete conversion of anhydride to imide, poly(isobutylene-alt-N-(N,N-dimethylaminopropylmaleimide) (PIBMI) (Scheme 1) was investigated by FT-IR and was confirmed by complete disappearance of peaks at 1850 cm-1 (C=O asym. str.) and 1785 (C=O sym. str.) for the anhydride ring and appearance of peaks 1767

Scheme 1. General scheme for the synthesis of cationic-amphiphilic polymers. 17 ACS Paragon Plus Environment

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cm-1 (C=O asym. str.), 1696 cm-1 (C=O sym. str.) for the imide ring. The complete conversion from PIBMA (soluble in DMSO) to PIBMI (soluble in CHCl3) has also been confirmed by 1H NMR (Fig. S1). Gel permeation chromatography (GPC) has been performed and found that PIBMA has an average molecular weight (Mw) of ~ 6000 Da and polydispersity index (PDI) of ~1.2. Because of the complete conversion from PIBMA to PIBMI (confirmed by FT-IR & 1H NMR), the molecular weight of PIBMI has been calculated to be ~ 9300 Da. The second step was the quaternization of tertiary nitrogen of PIBMI with the alkylating agent at 65-75 °C for 96 h to give water soluble QAPIBMI derivatives. Water soluble quaternized derivatives were characterized by FT-IR and 1H NMR and the NMR spectra are provided in the supporting information. Quaternized polymeric derivatives containing alkylating agents with variable hydrophobic/hydrophilic balance (amide, ester and bisethylene glycol (BEG) moieties) were synthesized for optimization of selective toxicity towards bacteria. The quaternized polymeric derivatives are denoted according to the type of alkylating agent used. Qn-penP (n-pentyl), QnprAP (n-propyl amide), Qn-prEP (n-propyl ester), Qn-BEGP (bisethyleneglycol, n-BEG) represent the PIBMI derivatives quaternized with alkylating agents containing different alkyl, amide, ester and bis ethylene glycol moieties respectively (Scheme 1). Keeping the amide moiety constant, further variations such as cyclization (QCybuAP, QCyprAP), isomerization (Qi-prAP) and unsaturation (Qn-prenAP, Qn- buenAP and Qn-buynAP) of the side chains were incorporated in the amphiphilic polymers. Degree of quaternization and molecular weight has been calculated from 1H NMR analysis as described in the methods section.61-63 The cationic charge density of the polymers given by the degree of quaternization was found to be nearly constant (92-95%) facilitating a better comparison between them (Table 1). The molecular weight (Mn) of all the polymers was found to be in the range of 15-18 KDa (Table 1).


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Figure 1. 1H NMR of QCyprAP (in D2O) indicating the peaks used for the calculation of degree of quaternization (δ/ppm): (a) for 2.15 (br, -NCH2CH2CH2-, 2H) and (b) for 2.2-2.5 (br, NCH2CH2CH2N(CH3)2, 8H).

Table 1. Degree of quaternization and molecular weight of cationic-amphiphilic polymers. S. No.

Polymer

Degree of

Molecular Weight a

Quaternization (DQ)

a,b

(Mn) (KDa)b

1

Qn-PenP

96 %

17.9

2

Qn-prAP

95%

16.1

3

QCybuAP

91%

17.6

4

Qn-buenAP

93 %

16.2

5

QCyprAP

93%

15.7

6

Qi-prAP

95%

18.0

7

Qn-buynAP

92%

16

8

Qn-prenAP

94%

18.2

9

Qn-prEP

96%

19.0

10

Qn-BEGP

97%

19.2

Calculated using 1H NMR analysis.


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Antibacterial activity and mammalian cell toxicity. We tested the antibacterial activity of these amphiphilic polymers against E. coli (Gram-negative) and S. aureus (Gram-positive). We measured minimum inhibitory concentration (MIC) as the lowest concentration required to completely inhibit the bacterial growth. Also, we have examined the antibacterial activity of these derivatives against highly pathogenic MRSA and VRE. Next, the mammalian cell toxicity has been investigated by the release of hemoglobin and lactate dehydrogenase (LDH) as a measure of membrane permeabilization from human red blood cells (hRBCs) and human endothelial kidney (HEK) cells respectively. The concentration required for 50% release of hemoglobin and LDH was designated as HC50 and IC50 respectively. We investigated the effect of side chain structural variations on antibacterial activity and cytotoxicity. The non-quaternized polymer, PIBMI, in its protonated state (soluble in water) does not have potent antibacterial efficacy with MIC of 105 µg mL-1 and 91 µg mL-1 against E. coli and S. aureus respectively (data not shown) and hence quaternization was performed to improve the antibacterial activity. The higher hydrophobic polymer, Qn-penP (n-pentyl) was found to be highly antibacterial (MIC = 3-16 µg mL-1) towards all the four bacteria but was found to be highly toxic to hRBCs and HEK cells (Table 2). The amide polymer with n-propyl chain, QnprAP had potent antibacterial activity and less toxicity to mammalian cells. However, the ester and ether moiety containing polymers, Qn-prEP and Qn-BEGP were non-toxic to bacteria as well as mammalian cells. This resulted in the understanding that amide moiety is important in the side chain to obtain the optimized polymer. Keeping the amide moiety intact, the hydrophobic moieties appended to the amide were varied. The cyclobutyl side chain amide polymer, QCybuAP also showed potent antibacterial activity but was more toxic to mammalian cells than Qn-prAP. The cyclopropyl (QCyprAP), isomeric (Qi-prAP) and unsaturated (Qn-prenAP, Qn-


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Table 2. Antibacterial activity and mammalian cell toxicity of cationic-amphiphilic polymers.

Polymer E. coli

MICa (µg mL-1) S. aureus MRSA

VRE

HC50b (µg mL-1) hRBCs

IC50c (µg mL-1) HEK

Qn-PenP

14

3

16

4

343

40

QCybuAP

16

8

16

8

250

95

Qn-prAP

31

31

31

8

1000

100

Qn-prenAP

31

31

31

31

>1000

>100

Qi-prAP

31

62

125

62

>1000

>100

QCyprAP

31

16

125

125

>1000

>100

Qn-buenAP

31

31

62

16

>1000

>100

Qn-buynAP

31

31

125

62

>1000

>100

Qn-prEP

125

250

250

125

>1000

>100

Qn-BEGP

>1000

>1000

>1000

>1000

>1000

>100

a

Minimum inhibitory concentration (MIC) after 24 h; bConcentration required for 50% hemolysis of human red blood cells (hRBCs); cConcentration required for 50% of lactate dehydrogenase (LDH) release from human endothelial kidney (HEK) cells. MRSA-methicillin resistant Staphylococcus aureus; VRE - vancomycin resistant Enterococcus faecium.

buenAP and Qn-buynAP) amide side chain containing polymers though were non-toxic to mammalian cells but displayed good antibacterial activity against drug sensitive bacteria but were ineffective against drug-resistant bacteria (MRSA and VRE) compared to Qn-prAP and QCybuAP (Table 2). Over all, these results suggested that the nature of the side chain influences the antibacterial and mammalian toxicity profiles of these cationic-amphiphilic polymers. The plausible reason behind might be the culmination of hydrophobic and hydrogen bonding interactions with the membranes as the cationic charge density (for electrostatic interaction) is


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constant in these polymers. E.g. amide polymers are potent antibacterial agents than their ester counter parts may be due to strong hydrogen bonding interactions with the bacterial membranes.

Membrane-active properties. Mechanism of action was studied by investigating the membrane-active properties of the polymers by measuring the cytoplasmic membrane depolarization, permeabilization and ATP leakage against E. coli and S. aureus. All the experiments were performed at a concentration of 25 µg mL-1. Cytoplasmic membrane depolarization was measured using the dye, DiSC3(5) (3, 3’-dipropylthiadicarbocyanine iodide). DiSC3(5) intercalates into the cytoplasmic membrane of energized cells and initially the fluorescence is quenched. Upon disruption of the membrane potential, the release of the dye into the solution results in increase of fluorescence. The variation of side chain chemical structure in these amphiphilic polymers also had effect on the membrane-disruptive properties in bacteria. The ability to dissipate the membrane potential was high for the QCybuAP and Qn-prAP but QnbuenAP, Qn-buynAP and QCyprAP were less effective at causing greater extent of depolarization against E. coli (Fig. 2A). Against S. aureus, all these polymers had more or less similar effect on causing membrane depolarization (Fig. 2B). Kinetics of membrane permeabilization was studied by measuring the uptake of a fluorescent probe, propidium iodide (PI). This dye enters only membrane compromised cells and fluoresces upon binding to nucleic acids. Different extent of membrane permeabilization was observed for these polymers with different side chain structure against E. coli (Fig. 2C). Dissipation of membrane potential leads to limiting the energy required for the metabolic processes in bacteria. Leakage of ATP levels in E. coli (Fig. 2D) were measured after treatment with the polymers using the luciferin-luciferase bioluminescence assay to understand the energy limitation conditions. The levels of ATP


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Figure 2. Membrane-active properties of the polymers against E. coli and S. aureus. Membrane depolarization against E. coli (A) and S. aureus (B); Membrane permeabilization against E. coli (C); Leakage of ATP levels against E. coli (D). All the experiments were performed at 25 µg mL-1.

leakage varied depending on the side chain structure of the polymers against E. coli (Fig. 2D).

Hydrophobicity profiles. The differences in the antibacterial activity, mammalian cell toxicity and membrane-active profiles of all these polymers prompted us to determine the variations in their hydrophobicity profiles. The hydrophobicity profile of the polymers was investigated using reverse-phase high performance liquid chromatography (RP-HPLC) by observing the absorbance at 220 nm on a C-18 column (Fig. 3). The retention time (RT in min) of the polymers varied with respect to differences in hydrophobicity as well as side chain chemical structure. QCybuAP had 23 ACS Paragon Plus Environment

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Figure 3. Chromatograms (reverse phase-high performance liquid chromatography, RP-HPLC) of polymers having different side chains displaying different retention times (RT).

high RT of 21.2 min followed by 20.5 min for Qn-buenAP. Qn-prAP, Qn-buynAP and QCyprAP showed RT values of 19.8, 19.4 and 18.5 min. These results suggest that the variations in the side chain structure leads to tunable hydrophobicity of the polymers. The change in the hydrophobicity profiles of these polymers with different side chain structure to some extent explains the trends in antibacterial activity, cytotoxicity and membrane-active properties. The higher hydrophobic polymer, QCybuAP had potent antibacterial activity and high membranedisruptive properties but was more toxic to mammalian cells. Qn-prAP, probably with optimum amphiphilicity, showed good antibacterial activity and potent membrane-disruptive properties but was less toxic to mammalian cells. Qn-buenAP though had higher hydrophobicity showed low antibacterial activity and weak membrane-disruption but was also low toxic to mammalian cells. We believe that although molecular hydrophobicity is important but the interaction of the 24 ACS Paragon Plus Environment

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polymer with the bacterial cell membrane might different depending on the side chain structure. Qn-buynAP and QCyprAP with lower hydrophobicity had low antibacterial activity, low membrane-disruption and very low toxicity to mammalian cells. These results suggest that the variation in side chain structure leads to tunable hydrophobicity of the amphiphilic polymers thereby resulting in modulation of biological activity profiles like antibacterial activity and mammalian cell toxicity.

Anti-endotoxin properties. We isolated human peripheral blood mononuclear cells (hPBMCs) from fresh human blood using Ficoll-Hypaque density centrifugation technique. These are the monocytes circulating in the peripheral human blood that get stimulated by bacterial endotoxins and lead to inflammatory response. Before proceeding with the anti-endotoxin studies, the cytotoxicity of these polymers by measuring the release of LDH against hPBMCs was evaluated. Qn-PenP had 40% whereas all the other polymers had ~20% release of LDH even at 100 µg mL1, the highest concentration tested (Fig. 4). Next, hPBMCs were stimulated with LPS in the presence and absence of these amphiphilic polymers and the corresponding secretion of proinflammatory cytokines, TNF-α and IL-6 were measured using ELISA. LPS from E. coli O111:B4, a strain of considerable interest has been used because of its pathogenicity.64 This strain of E. coli was discovered in 1945 to be the main cause of nonspecific gastroenteritis of babies and is still frequently isolated from babies suffering from acute diarrheal episodes.64 Concentration dependent stimulation of LPS on hPBMCs was studied and it was found that 20 ng mL-1 is the minimum concentration that can produce high levels of cytokine response (Fig. S2). Amphiphilic polymers with different side chain structures were used to understand their effect towards anti-inflammatory properties. Polymers alone did not increase the production of


Biomacromolecules

100

% of LDH Release

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QCyprAP Qi-prAP Qn-prEP Qn-BEGP

Qn-PenP Qn-prAP QCybuAP Qn-buenAP Qn-buynAP

80 60

Qn-prenAP

40 20 0 -20 0

20

40

60

[Polymer] g mL

80

-1

100

)

Figure 4. Cytotoxicity of human PBMCs measured by the release of lactate dehydrogenase (LDH) due to membrane permeabilization. Percentage of LDH release after treatment with polymers containing cyclic and acyclic side chains of different hydrophobicity.

cytokines. LPS when co-incubated with 20 µg mL-1 of Qn-PenP, Qn-prAP and QCybuAP (more hydrophobic polymers) had more than 80% inhibition of cytokine secretion compared to LPS alone (Fig. 5A & 5B). Qn-BEGP, the highly hydrophilic polymer, co-treated with LPS had no effect on the production of both the cytokines. Qn-buenAP, QCyprAP, and Qi-prAP reduced the LPS induced production by 60-70% of TNF-α and 30-40% of IL-6 compared to LPS. Coincubation of LPS and Qn-buynAP, Qn-prenAP and Qn-prEP though reduced TNF-α secretion by 20-40% but did not inhibit IL-6 production compared to LPS. All the polymers also showed concentration dependent effect on the inhibition of cytokine secretion depending on their side


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Figure 5. Secretion of pro-inflammatory cytokines. Production of TNF-α (A & C) and IL-6 (B & D) after stimulation of hPBMCs with LPS either in the presence or absence of polymers with different side chain chemical structure. Polymers alone were also used as controls. Hank’s balance salt solution (HBSS) was used as negative control.

chain structure and hydrophobicity (Fig. 5C & 5D). These results suggested that the inhibition of LPS stimulated cytokine secretion is dependent of side chain chemical structure that in turn depends on the hydrophobicity of the polymers.

Mechanism of LPS neutralization. In a pursuit to determine the binding properties of these polymers to LPS, experiments were performed using BODIPY dye conjugate of LPS (BLPS) 27 ACS Paragon Plus Environment

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(Molecular Probes Inc.). BODIPY conjugate of LPS (BODIPY-LPS) has its fluorescence quenched due to aggregation of LPS.65 The fluorescence increases in presence of an amphiphilic molecule that can dissociate LPS aggregate structure. The fluorescence of BLPS (0.5 µg mL-1) increased in the presence of 2% sodium dodecyl sulphate (SDS) whereas it decreased in presence PMB (0.5 mg mL-1) compared to BLPS alone (Fig. 6A). These results indicated that SDS binds and dissociates LPS aggregates and hence increase the fluorescence to a great extent as known in literature.65 PMB was found to bind and promote the aggregation of LPS resulting in a decrease of fluorescence as reported in literature.24 On the other hand, fluorescence of BLPS increased in presence of polymers (0.5 mg mL-1) but to a lesser extent compared to SDS (Fig. 6B). The variations in the increase of fluorescence were found to be dependent on the side chain chemical structure. Concentration dependent effect of QCybuAP binding to BLPS was also studied. Either increasing or decreasing concentration of QCybuAP from 0.5 mg mL-1 decreased the fluorescence of BLPS. Polymers bind and cause changes in the aggregate structure of LPS which was evident through increase in fluorescence of BLPS. Thus, these findings led to the understanding that the polymers bind to LPS but whether they dissociated (like SDS) or promoted (like PMB, rBPI21 and ultrashort AMPs) LPS aggregate structure could not be explained with this experiment. Further, to probe polymer binding to LPS, we performed Dynamic light scattering (DLS) studies. As known in literature, we have observed polydisperse size distribution of LPS (12.5 µg mL-1) aggregates due to its spontaneous self-assembly in aqueous solutions (Fig. 6C). However, co-incubation of LPS and QCybuAP (12.5 µg mL-1 + 12.5 µg mL-1) resulted in a monodisperse size distribution compared to LPS alone (Fig. 6D & Fig. S3). LPS + Qn-PenP, LPS + Qn-prAP also had monodisperse size distribution compared to LPS alone. However, LPS + Qn-prEP and


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Figure 6. LPS binding and dissociation in presence of amphiphilic molecules. Fluorescence intensity of 0.5 µg mL-1 of BLPS measured in presence of sodium dodecyl sulphate (2% SDS), colistin (0.5 mg mL-1), QCybuAP (0.1 mg mL-1, 0.5 mg mL-1, 1 mg mL-1, & 2 mg mL-1) (A) and polymers with different side chain chemical structure (all at 0.5 mg mL-1) (B); Size distribution of LPS (12.5 µg mL-1) (C) and LPS + QCybuAP (12.5 µg mL-1 + 12.5 µg mL-1) (D). Light scattering intensity (E) and mean count rate (KCPS (kilo counts per second)) (F) of LPS alone and in the presence of polymers with different side chain chemical structure.

LPS + Qn-BEGP had similar polydisperse size distribution as LPS alone (Fig. S3). Interestingly, we found the effects of polymer binding to LPS with respect to light scattering intensity and mean count rate (KCPS (kilo counts per second)) in DLS measurements. These effects were studied for polymers with different side chain chemical structures. The intensity is naturally weighted according to the scattering intensity of each particle fraction which increases as the density or number of particles increase. Light scattering intensity (%) increased to 10% (LPS + Qn- PenP) and 12% (LPS + Qn-prAP and LPS + QCybuAP) whereas LPS + Qn-prEP and LPS + Qn-BEGP had same as LPS (8%) (Fig. 6E). This suggests that the particle density increased for LPS co-treated with Qn-PenP, Qn-prAP and QCybuAP but not those with Qn-prEP and QnBEGP compared to LPS alone. Mean count rate or KCPS in DLS is simply the number of 29 ACS Paragon Plus Environment

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photons detected and is usually stated in a “per second” basis. This is useful for monitoring stability as a function of time. We found that the count rates increased in the order of LPS, LPS + Qn-PenP, LPS + Qn-prAP and LPS + QCybuAP and decreased for LPS + Qn-prEP and LPS + Qn-BEGP (Fig. 6F). These results indicated that the stability of the LPS aggregates increase in presence of Qn-PenP, Qn-prAP and QCybuAP but not in presence of Qn- prEP and Qn-BEGP. Overall, the above mechanistic studies suggest that LPS binding by these cationic amphiphilic polymers is the mechanism of LPS neutralization. The variations in binding characteristics of different polymers with different side chain chemical structures also explained the ability of differential inhibition of cytokine secretion in hPBMCs.

Influence of side chain chemical structure in LPS neutralization. Since the cationic charge density of the cationic-amphiphilic polymers is nearly constant, the role of electrostatic interaction in LPS binding may be similar for these polymers with different side chain chemical structures. Hydrophobic/hydrophilic ratio. The highly hydrophilic polymer, Qn-BEGP showed no inhibition of cytokine production due to its inability to bind LPS. Whereas the higher hydrophobic polymers (Qn-PenP) and amide moiety containing (Qn-prAP, QCybuAP) polymers bind strongly to LPS and thereby inhibit LPS induced cytokine secretion to a great extent in hPBMCs. The weaker LPS binding polymers with ester moieties (Qn-prEP) and unsaturation (Qn-buynAP) in the side chains due to their reduced hydrophobicity had weaker inhibition of cytokine production. These results suggest that amphiphilicity (hydrophobic/hydrophilic ratio) of these polymers play an important role in LPS binding and neutralization. 30 ACS Paragon Plus Environment

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Role of hydrogen bonding. All the amide side chain containing polymers (Qn-prAP, QCybuAP, Qn-buenAP, QCyprAP and Qi-prAP) showed greater LPS neutralization compared to the corresponding ester polymer, QnprEP. This indicated the role of strong hydrogen bonding interactions in amide polymers for LPS neutralization compared to that of ester polymers. Overall, these results suggest that LPS binding by these cationic polymers might have been mediated through a concoction of electrostatic, hydrophobic and hydrogen bonding interactions that are tunable by varying the side chain chemical structure. More importantly, unlike the amphiphilic small molecules/AMPs or PMB/ rBPI21/ultrashort AMPs, these polymers bind but neither dissociates nor promotes LPS aggregation (Fig. 7). We believe that the binding of polymer to LPS leads to some sort of a pseudo-aggregate formation (Fig. 7) resulting in LPS neutralization/detoxification. This pseudo-aggregate formation got evident from the BODIPYLPS binding studies which showed that increase of fluorescence in presence of polymers is less than that of SDS and more than that of colistin. Thus, these polymers bind and transiently dissociate LPS aggregate structure and revert to a stable pseudo-aggregate containing both the oppositely charged macromolecules (anionic LPS and cationic polymer). These conclusions were also supported by DLS studies. The advantage of this mode of binding compared to the LPS disaggregation approach is that the pseudo-aggregate formation might also lead to phagocytosis by the immune cells (e.g. macrophages) thereby eliminating the effect of endotoxins.

CONCLUSIONS Taken together, the findings showed that these cationic amphiphilic polymers possess antibacterial and anti-endotoxin/anti-inflammatory properties. The high hydrophobic polymer, 31 ACS Paragon Plus Environment

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Figure 7. Binding of LPS to different molecules and their respective mechanisms of LPS neutralization.

Qn-PenP showed strong antibacterial and anti-inflammatory properties but was more toxic to mammalian cells. On the other hand, high hydrophilic polymer, Qn-BEGP showed very weak antibacterial and anti-inflammatory properties and was non-toxic to mammalian cells. Polymers containing ester (Qn-prEP), unsaturated (Qn-buenAP and Qn-buynAP) and lower hydrophobic amide side chain polymers (Qi-prAP, QCyprAP) were weak antibacterial and anti-inflammatory agents with reduced toxicity. Hence, Qn-prAP and QCybuAP with sufficient hydrophobicity and hydrogen bonding interactions were potent antibacterial and strong anti-inflammatory polymers with minimal toxicity to mammalian cells. In conclusion, we demonstrated cationic amphiphilic polymers that neutralize and detoxify LPS by inhibiting the LPS induced secretion of pro-inflammatory cytokines (TNF-α and Il-6) in human immune cells. We found that these polymers bind LPS aggregates but neither promote nor dissociate them and rather lead to a pseudo-aggregate formation. We believe this


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might prevent the recognition and binding of LPS to either the soluble or cell surface receptors that require the native LPS aggregate thereby resulting in LPS detoxification. This study provides novel synthetic cationic amphiphilic polymers for effective neutralization of bacterial endotoxins (e.g. LPS).

ASSOCIATED CONTENT Supporting information contains the synthesis, characterization and supporting figures. This material is available free of charge via the Internet at http://pubs.acs.org. AUTHOR INFORMATION Corresponding Author *Corresponding author. E-mail: [email protected] ACKNOWLEDGMENTS We thank Prof. C.N.R. Rao, FRS (JNCASR) for his constant support and encouragement. Uppu thanks UGC for Senior Research Fellowship (SRF). We thank Mr. Jiaul Hoque for helping with the GPC experiments.

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Lipopolysaccharide Neutralization by Cationic-Amphiphilic Polymers

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