Combination Therapy Using Synthetic : A Platform to

Combat Multidrug-Resistant

Rashin Namivandi Zangeneh

A thesis in fulfilment of the requirements for the degree of

Doctor of Philosophy

School of Chemical Engineering

Faculty of Engineering

April 2020

PLEASE TYPE THE UNIVERSITY OF NEW SOUTH WALES Thesis/Dissertation Sheet

Surname or Family name: Namivandi Zangeneh

First name: Rashin Other name/s:

Abbreviation for degree as given in the University calendar: PhD

School: School of Chemical Engineering Faculty: Faculty of Engineering

Title: Combination Therapy Using Synthetic Antimicrobial Polymers: A Platform to Combat Multidrug-Resistant Bacteria

Abstract 350 words maximum:

The widespread failure of antibiotics in the treatment of multidrug-resistant (MDR) or -associated infections is a critical global healthcare issue. Thus, there is an urgent need for the development of novel and effective antimicrobial agents or strategies. This dissertation explores the use of a potent synthetic antimicrobial that consists of biocompatible oligo(ethylene glycol), hydrophobic ethylhexyl and cationic primary functional groups as a potential alternative to currently available antibiotics. In particular, this work investigates the advantages of combination therapy involving synthetic antimicrobial polymers and other antimicrobial agents as a novel therapeutic approach against bacterial infections. Firstly, a potent antibiofilm agent was developed by incorporating NO donor moieties into the structure of the synthetic antimicrobial polymer. The NO-loaded polymer showed dual-action capability as it could release NO to disperse biofilm, while the polymer caused membrane disruption. A synergistic effect in biofilm dispersal, planktonic and biofilm killing activities was observed against the Gram-negative bacteria (P. aeruginosa). In a second approach, synergistic combinations containing the synthetic antimicrobial polymer and two antibiotics, namely doxycycline and colistin, were developed. Coadministration of these compounds significantly improved the bacteriostatic efficacy against wild-type and MDR P. aeruginosa strains. In addition, the combination involving doxycycline showed synergistic bactericidal activity, could hinder resistance development in P. aeruginosa and was capable of reviving susceptibility to treatment in the resistant strains. In a third approach, synthetic antimicrobial polymers in the form of core-shell micelles were used in combination with antimicrobial essential oils, namely carvacrol and eugenol, where the antimicrobial polymeric micelles played a secondary role as delivery vehicles for essential oils. Coadministration of these compounds led to significant biofilm inhibition and synergistic killing effects against wild-type and MDR P. aeruginosa strains. Finally, a structure-activity relationship study was conducted to investigate the effect of polymer topology on biological performance. Linear with different chain lengths, block and hyperbranched copolymers were synthesized and assessed for antimicrobial and hemolytic activities. The best performing polymer in terms of biological properties was determined to have hyperbranched architecture and was >156 times more selective toward P. aeruginosa over red blood cells.

Declaration relating to disposition of project thesis/dissertation

I hereby grant to the University of New South Wales or its agents the right to archive and to make available my thesis or dissertation in whole or in part in the University libraries in all forms of media, now or here after known, subject to the provisions of the Copyright Act 1968. I retain all property rights, such as patent rights. I also retain the right to use in future works (such as articles or books) all or part of this thesis or dissertation.

I also authorise University Microfilms to use the 350-word abstract of my thesis in Dissertation Abstracts International (this is applicable to doctoral theses only).

………………………………………… …………………………………….……… ……….……………………...…….… Signature Witness Signature Date

The University recognises that there may be exceptional circumstances requiring restrictions on copying or conditions on use. Requests for restriction for a period of up to 2 years must be made in writing. Requests for a longer period of restriction may be considered in exceptional circumstances and require the approval of the Dean of Graduate Research.

FOR OFFICE USE ONLY Date of completion of requirements for Award:

Page | ii

Originality statement ‘I hereby declare that this submission is my own work and to the best of my knowledge it contains no materials previously published or written by another person, or substantial proportions of material which have been accepted for the award of any other degree or diploma at UNSW or any other educational institution, except where due acknowledgement is made in the thesis. Any contribution made to the research by others, with whom I have worked at UNSW or elsewhere, is explicitly acknowledged in the thesis. I also declare that the intellectual content of this thesis is the product of my own work, except to the extent that assistance from others in the project's design and conception or in style, presentation and linguistic expression is acknowledged.’

Signed ……………………………………………......

Date ……………………………………………......

Page | iii

INCLUSION OF PUBLICATIONS STATEMENT

UNSW is supportive of candidates publishing their research results during their candidature as detailed in the UNSW Thesis Examination Procedure.

Publications can be used in their thesis in lieu of a Chapter if:  The student contributed greater than 50% of the content in the publication and is the “primary author”, ie. the student was responsible primarily for the planning, execution and preparation of the work for publication  The student has approval to include the publication in their thesis in lieu of a Chapter from their supervisor and Postgraduate Coordinator.  The publication is not subject to any obligations or contractual agreements with a third party that would constrain its inclusion in the thesis

Please indicate whether this thesis contains published material or not.

This thesis contains no publications, either published or submitted for publication ☐ (if this box is checked, you may delete all the material on page 2) Some of the work described in this thesis has been published and it has been documented in the relevant Chapters with acknowledgement (if this box is

☒ checked, you may delete all the material on page 2)

This thesis has publications (either published or submitted for publication) ☐ incorporated into it in lieu of a chapter and the details are presented below

CANDIDATE’S DECLARATION I declare that:  I have complied with the Thesis Examination Procedure  where I have used a publication in lieu of a Chapter, the listed publication(s) below meet(s) the requirements to be included in the thesis. Name Signature Date (dd/mm/yy)

Page | iv

Copyright statement ‘I hereby grant the University of New South Wales or its agents the right to archive and to make available my thesis or dissertation in whole or part in the University libraries in all forms of media, now or here after known, subject to the provisions of the Copyright Act 1968. I retain all proprietary rights, such as patent rights. I also retain the right to use in future works (such as articles or books) all or part of this thesis or dissertation. I also authorise University Microfilms to use the 350 word abstract of my thesis in Dissertation Abstract International (this is applicable to doctoral theses only). I have either used no substantial portions of copyright material in my thesis or I have obtained permission to use copyright material; where permission has not been granted I have applied/will apply for a partial restriction of the digital copy of my thesis or dissertation.

Signed……………………………

Date………………………………

Authenticity statement ‘I certify that the Library deposit digital copy is a direct equivalent of the final officially approved version of my thesis. No emendation of content has occurred and if there are any minor variations in formatting, they are the result of the conversion to digital format.

Signed……………………………

Date………………………………

Page | v

Acknowledgements

First and foremost, my most profound appreciation and gratitude go to my supervisor, Prof. Cyrille Boyer, for his invaluable support, patience and mentorship throughout my PhD. Also, my sincere thanks to my co-supervisor, Dr. Edgar Wong, for his tremendous guidance that has been instrumental in my academic development. Without them, none of this would have been possible.

I would like to acknowledge the University of New South Wales and the Australian Government for providing me with the scholarship that enabled me to perform my studies in one of the high-ranking universities in the world; this has been greatly effective on my personal and career development.

I would like to thank my masters and honours thesis students, Rebecca J Kwan, Maeva Sauvage-Nguyen, and Yiling Yang, for their contributions and help throughout the projects. Special thanks to Prof. Naresh Kumar, Dr. Kitty Ho, Prof. Mark Duncan Willcox and Dr. Debarun Dutta, for their helpful advice and contributions. My sincere thanks go to Eh Hau Pan and Camillo Taraborrelli for their invaluable efforts in managing the laboratories and for their help and support throughout my time in UNSW.

To all my PhD mates and friends: Zahra Sadrearhami, Parisa Moazzam, Peter Judzewitsch, Erna Wulandari, Yasemin Fadil, Gervase Ng, Sihao Xu, Chenyu Wu, Zack Huang, Rubby Noor, Dr. Khanh Nguyen, Dr. Susan Oliver, Dr, Ali Bagheri, Dr. Nathaniel

Corrigan, Dr. Kenward Jung, Dr. Awin Hadzir and Dr Siva Shanmugam; for their support, friendship and sharing all the happiness.

Finally, I wish to thank my family for the continuous support they have given me throughout my life; I will forever be grateful. My deepest thanks to my beloved husband Shakib for his understanding, patience, unconditional love, for giving me motivation and encouragement during my PhD and for being by my side through all the tough times.

Page | vi

Abstract

The widespread failure of antibiotics in the treatment of multidrug-resistant (MDR) or biofilm-associated infections is a critical global healthcare issue. Thus, there is an urgent need for the development of novel and effective antimicrobial agents or strategies. This dissertation explores the use of a synthetic antimicrobial polymer that consists of biocompatible oligo(ethylene glycol), hydrophobic ethylhexyl and cationic primary amine functional groups as a potential alternative to currently available antibiotics. In particular, this work investigates the advantages of combination therapy involving synthetic antimicrobial polymers and other antimicrobial agents as a novel therapeutic approach against bacterial infections. Firstly, a potent antibiofilm agent was developed by incorporating NO donor moieties into the structure of the synthetic antimicrobial polymer. The NO-loaded polymer showed dual-action capability as it could release NO to disperse biofilm, while the polymer caused membrane disruption. A synergistic effect in biofilm dispersal, planktonic and biofilm killing activities was observed against the Gram-negative bacteria Pseudomonas aeruginosa (P. aeruginosa). In a second approach, synergistic combinations containing the synthetic antimicrobial polymer and two antibiotics, namely doxycycline and colistin, were developed. Coadministration of these compounds significantly improved the bacteriostatic efficacy against wild-type and MDR P. aeruginosa strains. In addition, the combination involving doxycycline showed synergistic bactericidal activity, could hinder resistance development in P. aeruginosa and was capable of reviving susceptibility to treatment in the resistant strains. In a third approach, synthetic antimicrobial polymers in the form of core-shell micelles were used in combination with antimicrobial essential oils, namely carvacrol and eugenol, where the antimicrobial polymeric micelles played a secondary role as delivery vehicles for essential oils. Coadministration of these compounds led to significant biofilm inhibition and synergistic killing effects against wild-type and MDR P. aeruginosa strains. Finally, a structure-activity relationship study was conducted to investigate the effect of polymer topology on biological performance. Linear copolymers with different chain lengths, block and hyperbranched copolymers were synthesized and assessed for antimicrobial and hemolytic activities. The best performing polymer in terms of biological properties was determined to have hyperbranched architecture and was >156 times more selective toward P. aeruginosa over red blood cells.

Page | vii

List of publications

Publications contributing to this thesis

Rashin Namivandi-Zangeneh, Yiling Yang, Sihao Xu, Edgar H. H. Wong, and

Cyrille Boyer “Antibiofilm Platform based on the Combination of Antimicrobial

Polymers and Essential Oils” Biomacromolecules, 2020, 21 (1), pp 262–272.

Rashin Namivandi-Zangeneh, Zahra Sadrearhami, Debarun Dutta, Mark

D.P. Willcox, Edgar H. H. Wong, and Cyrille Boyer, “Synergy between Synthetic

Antimicrobial Polymer and Antibiotics: A Promising Platform to Combat

Multidrugresistant Bacteria” ACS Infectious Diseases, 2019, 5 (8), pp 1357–1365.

Rashin Namivandi-Zangeneh, Zahra Sadrearhami, Maeva Sauvage-Nguyen,

Kitty K. K. Ho, Naresh Kumar, Edgar H. H. Wong, and Cyrille Boyer “Nitric

Oxide-Loaded Antimicrobial Polymer for the Synergistic Eradication of Bacterial

Biofilm” ACS Macro Letters, 2018, 7 (5), pp 592–597.

Rashin Namivandi-Zangeneh, Rebecca J. Kwan, Thuy-Khanh Nguyen,

Jonathan Yeow, Frances L. Byrne, Stefan H. Oehlers, Edgar H. H. Wong, and

Cyrille Boyer, “The effects of polymer topology and chain length on the antimicrobial activity and hemocompatibility of amphiphilic ternary copolymers” Polymer Chemistry, 2018, 9 (13), pp 1735–1744.

Other publications

Zahra Sadrearhami, Rashin Namivandi-Zangeneh, Emily Price, Marta

Krasowska, Sameer A. Al-Bataineh, Jason Whittle, Edgar H. H. Wong, Anton

Blencowe and Cyrille Boyer, “S-Nitrosothiol Plasma-Modified Surfaces for the

Prevention of Bacterial Biofilm Formation” ACS Biomaterials Science &

Engineering, 2019, 5 (11), pp 5881–5887.

Page | viii

Francois‐Marie Allioux, Salma Merhebi, Jianbo Tang, Shuhada A. Idrus‐Saidi,

Roozbeh Abbasi, Maricruz G. Saborio, Mohammad B. Ghasemian, Jialuo Han,

Rashin Namivandi‐Zangeneh, Anthony P. O'Mullane, Pramod Koshy, Rahman

Daiyan, Rose Amal, Cyrille Boyer and Kourosh Kalantar‐Zadeh “Catalytic Metal

Foam by Chemical Melting and Sintering of Liquid Metal ”

Advanced Functional Materials, 2019, 30, 1-13.

Zahra Sadrearhami, Thuy-Khanh Nguyen, Rashin Namivandi-Zangeneh,

Kenward Jung, Edgar H. H. Wong and Cyrille Boyer “Recent advances in nitric oxide delivery using polymer-based systems” Journal of Materials Chemistry B,

2018, 6, 2945-2959.

Conference presentations

Rashin Namivandi Zangeneh, Edgar H. H. Wong and Cyrille Boyer “Nitric

Oxide-Loaded Antimicrobial Polymer for the Synergistic Eradication of Bacterial

Biofilm” presentation in 9th International Nanomedicine conference, June 2018,

Sydney, Australia.

Rashin Namivandi Zangeneh, Edgar H. H. Wong and Cyrille Boyer

“Synergy between Synthetic Antimicrobial Polymer and Antibiotics/Nitric

Oxide: A Promising Platform to Combat Multidrug-Resistant Bacteria” 37th

Australian Polymer Symposium, November 2019, Sunshine coast, Australia.

Page | ix

Table of contents

Acknowledgements ...... vi

Abstract ...... vii

List of publications ...... viii

Table of contents ...... x

List of figures ...... xiv

List of tables ...... xxi

List of schemes ...... xxi

List of abbreviations/symbols ...... xxii

1 Introduction ...... 2

1.1 Thesis motivation ...... 2

1.2 Thesis outline ...... 6

1.3 Reference...... 10

2 Literature Review ...... 14

2.1 Antibiotic resistance ...... 14

2.1.1 Mechanisms of antibiotic resistance ...... 14

2.1.2 Antibiotic resistance acquisition ...... 18

2.1.3 Adaptive Resistance ...... 19

2.2 Bacterial ...... 19

2.2.1 Biofilm life cycle...... 21

2.2.2 Recalcitrance of bacterial biofilms toward antibiotics ...... 22

2.2.3 Resistance acquisition in biofilms ...... 26

2.3 Novel therapeutic strategies to combat antibiotic resistance ...... 26

Page | x

2.4 Synthetic antimicrobial polymers ...... 30

2.4.1 Structure-activity relationships ...... 30

2.4.2 Mechanism of action of synthetic antimicrobial polymers ...... 52

2.4.3 Building well-defined polymers via reversible addition-fragmentation chain transfer (RAFT) polymerization ...... 54

2.5 Combination therapy ...... 57

2.5.1 Antimicrobial polymers in combination with nitric oxide ...... 60

2.5.2 Antimicrobial polymers in combination with antibiotics ...... 69

2.5.3 Antimicrobial polymers in combination with essential oils ...... 79

2.5.4 Antimicrobial polymers in combination with metal-based nanomaterials 82

2.5.5 Antimicrobial polymers in combination with carbon-based nanomaterials 103

2.6 Reference...... 111

3 Nitric Oxide-Loaded Antimicrobial Polymer for The Synergistic Eradication of Bacterial Biofilm ...... 128

3.1 Introduction ...... 128

3.2 Experimental Section ...... 130

3.3 Results and discussion ...... 137

3.3.1 Synthesis of amphiphilic ternary random ...... 137

3.3.2 Attachment of NONOate to ternary random copolymer ...... 138

3.3.3 Determination of the release and stability of nitric oxide ...... 140

3.3.4 Biofilm dispersal by NO-loaded polymer ...... 141

3.3.5 Planktonic and biofilm killing by NO-loaded polymer ...... 145

3.4 Summary ...... 148

3.5 References ...... 148

Page | xi

3.6 Appendix A ...... 152

4 Synergy between Synthetic Antimicrobial Polymer and Antibiotics: A Promising Platform to Combat Multidrug-resistant Bacteria ...... 157

4.1 Introduction ...... 157

4.2 Experimental Section ...... 160

4.3 Results and Discussion ...... 166

4.3.1 Assessment of antimicrobial polymer-antibiotic interactions ...... 167

4.3.2 Evaluation of bactericidal activity of combinations and individual agents 174

4.3.3 Evaluation of resistance development toward polymer-doxycycline combination ...... 177

4.4 Summary ...... 179

4.5 References ...... 180

4.6 Appendix B ...... 183

5 An Antibiofilm Platform based on the Combination of Antimicrobial Polymers and Essential Oils ...... 188

5.1 Introduction ...... 188

5.2 Experimental Section ...... 192

5.3 Results and Discussion ...... 201

5.3.1 Synthesis and characterization of antimicrobial and control polymers 201

5.3.2 Preparation and characterization of polymer-oil combinations ...... 205

5.3.3 Determination of the MIC values of synthesised polymers ...... 208

5.3.4 Biofilm inhibition activity of polymer-oil combinations and individual components ...... 209

5.3.5 Biofilm killing activity of polymer-oil combinations and individual components ...... 216

Page | xii

5.3.6 Hemocompatibility of polymer-oil combinations and individual components ...... 218

5.4 Summary ...... 220

5.5 Reference...... 221

5.6 Appendix C ...... 225

6 The Effects of Polymer Topology and Chain Length on the Antimicrobial Activity and Hemocompatibility of Amphiphilic Ternary Copolymers ...... 238

6.1 Introduction ...... 238

6.2 Experimental Section ...... 242

6.2.1 Synthesis of antimicrobial polymers ...... 244

6.3 Results and Discussion ...... 250

6.3.1 Effect of the polymer chain length on biological performance...... 252

6.3.2 Effect of block copolymer architecture on biological performance ...... 255

6.3.3 Effect of the hyperbranched architecture on biological performance ... 258

6.4 Summary ...... 261

6.5 Reference...... 262

6.6 Appendix D ...... 266

7 Concluding remarks and future perspectives ...... 273

7.1 Conclusions ...... 273

7.2 Challenges and future perspectives ...... 279

Page | xiii

List of figures

Figure 2.1. Intrinsic resistance mechanisms in a Gram-negative bacteria. Three different scenarios regarding the interaction of β-lactam antibiotics with Gram-negative bacteria to reach a penicillin-binding protein (PBP). Antibiotic A can cross the outer membrane via porins and bind to PBP. Antibiotic B can cross the outer membrane via porins but is expelled out via the efflux pumps. The penetration of antibiotic C is inhibited by the outer membrane.1...... 15

Figure 2.2. Antibiotic resistance by (a) antibiotic modification and (b) target modification.1 ...... 17

Figure 2.3. Resistance acquisition through (a) mutation and (b) horizontal gene transfer.24 ...... 19

Figure 2.4. Representative stages of the biofilm life cycle.34 ...... 22

Figure 2.5. The main mechanisms of antibiotic tolerance in bacterial biofilms.34 ...... 24

Figure 2.6. Illustration of the cross sections of mammalian and microbial cells’ membrane.130 ...... 32

Figure 2.7. Chemical structure of methacrylate-based antimicrobial copolymers containing primary or tertiary and QA groups as cationic moieties.137 ...... 33

Figure 2.8. Selective antimicrobial activity of tertiary amine-containing methacrylate polymers toward Mycobacterium tuberculosis over RBCs...... 34

Figure 2.9. Chemical structure of cationic amphiphilic polycarbonates bearing various QA functionalities (top). Antimicrobial and hemolytic activities of the polymer comprising N,N-dimethylbutylammonium QA group.146 ...... 37

Figure 2.10. Illustration of the self-assembly of the amphiphilic ternary random copolymers of different structures into single-chain polymeric nanoparticles in aqueous media (a). DLS volume distributions of PDab‑ F and PDab‑ Pyr and PC2 (PEG- 158 less version of PDab‑ F) in water and MHB...... 40

Figure 2.11. Synthesis of facially amphiphilic polymers via ring-opening metathesis polymerization of oxanorbornene-derived monomers (a). Antimicrobial (MIC90, µg −1 −1 -1 mL ) and hemolytic (HC50, µg mL ) activities of 3000 g mol series (b) and -1 −1 10,000 g mol series (c). Antimicrobial (MIC90, µg mL ) and hemolytic (HC50, µg mL−1) activities of the oligomers and polymers synthesised using the monomer with propyl group (d).161 ...... 42

Figure 2.12. Three spatial arrangements for cationic and hydrophobic moieties in the structure of synthetic antimicrobial polymers.110 ...... 43

Page | xiv

Figure 2.13. Chemical structure of random methacrylate copolymers bearing either primary amine or guanidine moieties as cationic functionalities with different RAFT R- and Z- groups.168 ...... 46

Figure 2.14. Schematic illustration of the structure, composition and segmentation of monomers within the polymer chains of the poly acrylamides synthesized via RAFT polymerization.174...... 48

Figure 2.15. Illustration of the chemical structure of cationic amphiphilic polycarbonate and its self-assembly into micelle structure in aqueous media.176 ...... 50

Figure 2.16. Schematic illustration of the chemical structure of glucosamine- functionalised star polymer.178 ...... 52

Figure 2.17. Proposed models for the membrane-disrupting action of antimicrobial peptides.180 ...... 53

Figure 2.18. Mechanism of RAFT polymerization.188 ...... 56

Figure 2.19. Proposed mechanisms for bactericidal action of nitric oxide through nitrosative and oxidative stress.197 ...... 60

Figure 2.20. Synthesis of dual-action dendrimers comprising quaternary moieties and N-diazeniumdiolate NO donors.204 ...... 64

Figure 2.21. Illustration of the chemical structure of dual-action nitric oxide-releasing alkyl-modified dendrimers (left). Confocal microscopy images showing the relative penetration of two representative dendrimers (G4 butyl and G4 hexyl) into S. aureus biofilm upon 1 h incubation at 50 µg mL−1(right).205 ...... 65

Figure 2.22. Schematic illustration of antifouling- releasing coatings prepared by covalent grafting of zwitterionic polymers to SNAP-doped CarboSil films.208 ..... 68

Figure 2.23. Synthesis of cationic polycarbonates comprising vitamin E and alkyl side chains (a). Chemical structure of doxycycline, streptomycin and penicillin-G (b).210 ...... 71

Figure 2.24. Chemical structure of cationic conjugated polymers (a) and polypeptide antibiotics polymyxin E (PLE) and polymyxin B (PLB) (b). Bactericidal activities of PLB and the combination of PLB and PBF as a function of concentration (c). Bactericidal activities of PBF, PLB and the combination of PLB and PBF as a function of time (d). SEM images of E. coli, 1: untreated, 2: treated with PBF, 3: treated with PLB, and 4: treated with the combination of PBF and PLB (e).211 ..... 73

Figure 2.25. Schematic illustration of the formation of β-lactam-metallopolymer bioconjugate via -pairing between antibiotic and metallopolymer (a). Release of β-lactam from β-lactam-metallopolymer bioconjugate in the presence of β- lactamases or lipoteichoic acid (b). Confocal microscopy images of HA-MRSA cells

Page | xv

before (control) and after incubation with penicillin-G, polymer and penicillin- G/polymer treatments (the control and treated bacteria were stained with BacLight live/dead kit) (scale bar = 50 μm) (c). SEM images of HA-MRSA before (control) and after incubation with penicillin-G, polymer and penicillin-G/ polymer treatments (scale bar =1 μm) (d).214 ...... 76

Figure 2.26. Chemical structure of multifunctional macromolecular system comprising cobaltocenium-containing metallopolymer, boronic acid, and penicillin (a). Schematic illustration of synergistic antimicrobial activity of multifunctional polymer against a Gram-negative bacteria (b).215 ...... 78

Figure 2.27. Chemical structure of the poly(oxanorborneneimide) copolymers (PONI- GAT), poly(maleic anhydride-alt-octadecene) crosslinking agent (p-MA-alt-OD) and carvacrol oil (a). Preparation of oil-in-water cross-linked polymeric nanocomposite (X-NCs) via emulsification of p-MA-alt-OD-loaded carvacrol into the aqueous solution of PONI-GAT (b). Bactericidal activities of 10 wt% X-NCs, carvacrol and PONI-GAT against 1-day-old P. aeruginosa after 3 h incubation as a function of emulsion concentration (c). Cytotoxicity and bactericidal activities of 10 wt% X-NCs against 3T3 fibroblast−P. aeruginosa biofilm coculture model after 3 h of treatment as a function of emulsion concentration (d).224 ...... 81

Figure 2.28. Synthesis of dual-action polymer-AgBr nanocomposites via on-site precipitation method.227...... 85

Figure 2.29. Synthetic scheme of qPDMAEMA-AgNP core-shell nanoparticles.229 .... 88

Figure 2.30. Schematic illustration of the synthesis of linear, slightly branched and highly branched PSAs and PSA-AgNP nanocomposites. Bactericidal activity of PSA- AgNPs against in disk-diffusion assay (bottom right corner).23191

Figure 2.31. Schematic representation of the synthesis of recyclable PMAG and qPDMAEMA-immobilised AuNPs.236 ...... 94

Figure 2.32. Schematic illustration of the synthesis of AuNPs grafted with metallopolymer-antibiotic bioconjugates.238 ...... 97

Figure 2.33. Synthesis of quaternized polycarbonate (PrBrT)-grafted MnFe2O4 nanoparticles.250 ...... 101

Figure 2.34. Chemical structure of penicillin, cationic metallopolymer (PCo) and cationic metallopolymer-penicillin bioconjugates (PCo-Peni) (top). Schematic illustration of the synthesis of recyclable magnetic nanoparticles grafted with cationic metallopolymer-penicillin bioconjugates (bottom).251 ...... 103

Figure 2.35. Schematic illustration of the synthesis of PEG and PHGC-modified graphene oxide sheets.258 ...... 105

Page | xvi

Figure 2.36. Schematic illustration of the proposed antibacterial mechanism of ZnO/GQD−PEI nanocomposite.263 ...... 109

Figure 3.1. General illustration depicting the dispersal and killing of bacteria biofilm using the NO-functionalized antimicrobial polymer in this study...... 130

Figure 3.2. FTIR spectra of P, fresh PNO, and PNO after 3 h of release study in PBS...... 139

Figure 3.3. XPS spectra of PNO and P...... 140

Figure 3.4. Cumulative release of NO from PNO in PBS buffer (pH 7.4, 37 oC) as measured using Griess assay...... 141

Figure 3.5. P. aeruginosa biofilm biomass reduction upon treatment with various concentrations (16-64 μgmL−1) of P, PNO, SperNO, and P + SperNO for 20 min (a) and 60 min (b). Data are representative of at least three independent experiments ± SD. Student’s t-test – asterisks indicate statistically significant difference of PNO treatments versus P treatment (**, p < 0.01; ****, p < 0.0001; ns, non-significant)...... 142

Figure 3.6. 2D and 3D tomographic microscopy images of the untreated, P, PNO, P + SperNO and SperNO treated P. aeruginosa biofilm samples. The compound concentration and treatment time was 64 μg mL−1 and 1 h, respectively. Images were taken at the center of the culture dish. Scale bar = 20 µm...... 144

Figure 3.7. 2D and 3D tomographic microscopy images of the untreated normal and thick P. aeruginosa biofilms compared to the PNO treated samples (64 μg mL−1 for 1 h). Note: two different spots of the culture dish were analyzed. Scale bar = 20 µm. 145

Figure 3.8. Bactericidal activity of P, PNO, SperNO and P + SperNO for 20 min (16 - 64 μg mL−1) on planktonic (a) and biofilm (b) P. aeruginosa as measured by colony- forming unit (CFU) analysis. Data are representative of at least three independent experiments ± SD. Student’s t-test – asterisks indicate statistically significant difference between PNO versus P treatments (****, p < 0.0001; ns, non-significant)...... 146

Figure 3.9. Bactericidal activity of P, PNO, SperNO and P + SperNO for 60 min (16 - 64 μg mL−1) on planktonic (a) and biofilm (b) P. aeruginosa as measured by colony- forming unit (CFU) analysis. Data are representative of at least three independent experiments ± SD. Student’s t-test – asterisks indicate statistically significant difference between PNO versus P treatments (****, p < 0.0001; ns, non-significant)...... 147

Figure 4.1. Synergistic antibacterial activity of synthetic antimicrobial polymer and doxycycline...... 159

Page | xvii

Figure 4.2. Chemical structure and reported mode of action of the antibiotics used in this work.44, 45 ...... 168

Figure 4.3. Checkerboard microdilution assay between P and selected antibiotics against P. aeruginosa PAO1. Bacterial growth, quantified by average OD600, is represented as a linear gradient from white to peach where darker colors represent less growth inhibition. Yellow and red bullets represent MIC values for P and antibiotics, respectively. Blue bullets represent concentrations exhibiting synergistic interaction. The data are representative of a minimum of two biological replicates...... 169

Figure 4.4. Checkerboard microdilution assay between P and selected antibiotics against E. coli K12. Bacterial growth, quantified by average OD600, is represented as a linear gradient from white to blue where darker colors represent less growth inhibition. Yellow and red bullets represent MIC values for P and antibiotics, respectively. Blue bullets represent concentrations exhibiting synergistic interaction. The data are representative of a minimum of two biological replicates...... 170

Figure 4.5. Checkerboard microdilution assay between P and D (a-e), and P and C (f-j) against P. aeruginosa PAO1, PA6294, PA ATCC 27853, PA32 and PA37. Bacterial growth, quantified by average OD600, is illustrated as a linear gradient from white to green and burgundy where darker colors represent less growth inhibition. Yellow and red bullets represent MIC values for P and antibiotic, respectively. Blue bullets indicate concentrations exhibiting synergistic interaction. The data are representative of a minimum of two biological replicates...... 172

Figure 4.6. Bactericidal activity of individual components and combinations. Bactericidal activity of P, D, and PD on planktonic (a) and biofilm (b) P. aeruginosa PAO1. Bactericidal activity of P, C, and PC on planktonic (c) and biofilm (d) P. aeruginosa PAO1. Bactericidal activity of P, D, and PD on planktonic (e) and biofilm (f) P. aeruginosa 6294. All bactericidal activities were determined by colony-forming unit (CFU) analysis upon 20 min incubation. Data are representative of at least three independent experiments ± SD. Two-way ANOVA test − asterisks indicate statistically significant difference of PD and PC vs P treatment (**p < 0.01; ***p < 0.001; ****p < 0.0001; ns, non-significant (p > 0.01))...... 175

Figure 4.7. Resistance development monitoring in P. aeruginosa PAO1 in the presence of sub-MIC levels of P, D and PD. The y-axis indicates the changes in MICs of the compounds over a period of 21 days as compared to the first day (0th passage). Note that the bacteria cultures from 20th passage were used in supplementary MIC test...... 177

Figure 5.1. Combination of synthetic antimicrobial block copolymer and essential oils as oil-in-water emulsion (left) and bacterial biofilm inhibition and eradication induced by this combination (right)...... 191

Figure 5.2. Chemical structure of the synthesized polymers and essential oils investigated in this study (top). Schematic illustrations of the emulsions containing antimicrobial polymers and essential oils in aqueous media (bottom)...... 204

Page | xviii

Figure 5.3. DLS normalized size distribution of RP/Car, RP/Eug, Car and Eug (a), BP, BP/Car and BP/Eug (b) and CP, CP/Car, CP/ Eug (c) in water...... 206

Figure 5.4. Inhibition of planktonic bacterial growth. P. aeruginosa PAO1 bacteria were grown in the presence of RP, Car, Eug, RP/Car and RP/Eug (a), and BP, CP, Car, Eug, BP/Car, BP/Eug, CP/Car and CP/Eug (b). Bacterial growth was quantified by measurement of the OD595 of the bacterial culture. Data are representative of at least two independent experiments ± SD. Two-way ANOVA test; black asterisks indicate a statistically significant difference of each combination vs individual polymer at the corresponding concentrations. Blue and red asterisks indicate a statistically significant difference of each combination vs individual Car or Eug, respectively. (**p < 0.01; ***p < 0.001; ****p < 0.0001; ns, nonsignificant (p > 0.01))...... 210

Figure 5.5. P. aeruginosa PAO1 biofilm formation inhibition. Biofilms were grown in the presence of RP, Car, Eug, RP/Car and RP/Eug (a), and BP, CP, Car, Eug, BP/Car, BP/Eug, CP/Car and CP/Eug (b). Biofilm biomass was quantified by measurement of the OD595 of the crystal violet stained biofilms. Data are representative of at least two independent experiments ± SD. Two-way ANOVA test; black asterisks indicate a statistically significant difference of each combination vs individual polymer at the corresponding concentrations. Blue and red asterisks indicate a statistically significant difference of each combination vs individual Car or Eug, respectively. (**p < 0.01; ***p < 0.001; ****p < 0.0001; ns, nonsignificant (p > 0.01))...... 211

Figure 5.6. Inhibition of planktonic bacterial growth induced. P. aeruginosa PA37 bacteria were grown in the presence of RP, Car, Eug, RP/Car and RP/Eug (a), and BP, CP, Car, Eug, BP/Car, BP/Eug, CP/Car and CP/Eug (b). Bacterial growth was quantified by measurement of the OD595 of the bacterial culture. Data are representative of at least two independent experiments ± SD. Two-way ANOVA test; black asterisks indicate a statistically significant difference of each combination vs individual polymer at the corresponding concentrations. Blue and red asterisks indicate a statistically significant difference of each combination vs individual Car or Eug, respectively. (**p < 0.01; ***p < 0.001; ****p < 0.0001; ns, nonsignificant (p > 0.01))...... 213

Figure 5.7. P. aeruginosa PA37 biofilm formation inhibition. Biofilms were grown in the presence of RP, Car, Eug, RP/Car and RP/Eug (a), and BP, CP, Car, Eug, BP/Car, BP/Eug, CP/Car and CP/Eug (b). Biofilm biomass was quantified by measurement of the OD595 of the crystal violet stained biofilms. Data are representative of at least two independent experiments ± SD. Two-way ANOVA test; black asterisks indicate a statistically significant difference of each combination vs individual polymer at the corresponding concentrations. Blue and red asterisks indicate a statistically significant difference of each combination vs individual Car or Eug, respectively. (**p < 0.01; ***p < 0.001; ****p < 0.0001; ns, nonsignificant (p > 0.01))...... 214

Figure 5.8. 2D and 3D tomographic microscopy images of P. aeruginosa PAO1 biofilm grown in the presence or absence of individual compounds and combinations for 6.5

Page | xix

h. Images were taken at the center of the culture dish and analyzed using 3D Nanolive Cell Explorer Steve digital staining software (scale bar = 20 μm). Note: Biofilm inhibition values based on this microscopy analysis are presented at the bottom right corner...... 215

Figure 5.9. Bactericidal activity of RP, Car, Eug, RP/Car and RP/Eug (a), and BP, CP, Car, Eug, BP/Car, BP/Eug, CP/Car and CP/Eug (b) against P. aeruginosa PAO1 biofilm. All bactericidal activities were determined by colony-forming unit (CFU) analysis upon 20 min of incubation. Data are representative of at least two independent experiments ± SD. Two-way ANOVA test; black asterisks indicate a statistically significant difference of each combination vs individual polymer at the corresponding concentrations. Blue and red asterisks indicate a statistically significant difference of each combination vs individual Car or Eug, respectively. (**p < 0.01; ***p < 0.001; ****p < 0.0001; ns, nonsignificant (p > 0.01))...... 216

Figure 5.10. Hemolytic activity of RP, BP, Car, Eug, RP/Car, BP/Car, RP/Eug and BP/Eug. Data are representative of at least two independent experiments ± SD. 219

Figure 6.1. The compositional structures and architectures of the amphiphilic ternary copolymers in this study...... 242

Figure 6.2. (a) GPC- differential refractive index (DRI) chromatograms of the Boc- protected polymers L1, L2 and L3. (b) DLS normalized volume distributions of Boc-deprotected L1, L2 and L3 polymers in water...... 253

Figure 6.3. (a) GPC DRI chromatograms of the macroRAFT agent and Boc-protected polymers B1 and B2. (b) DLS normalized volume distributions of Boc-deprotected B1 and B2 polymers in water. The inset illustrates the formation of micelles via the self-assembly of the block copolymers in water...... 256

Figure 6.4. (a) GPC DRI chromatograms of the Boc-protected hyperbranched polymers H1, H2, H3 and H4. (b) DLS normalized volume distributions of Boc-deprotected H1, H2, H3 and H4 polymers in water...... 259

Figure 6.5. 2D and 3D tomographic microscopy images of untreated control (a) and H2 treated samples (b-d). Scale bar = 20µm...... 261

Page | xx

List of tables

Table 2.1. Therapeutic strategies against antibiotic resistance and/or bacterial biofilms...... 27

Table 4.1. Checkerboard assay results indicating the synergistic activity of P with D and C and against five P. aeruginosa strains including PAO1, PA6294, PA ATCC 27853, PA32 and PA37. All the combinations showed moderate to high synergism (FICI ≤ 0.5)...... 173

Table 4.2. Supplementary antimicrobial activity test showing the susceptibility of modified strains of P. aeruginosa PAO1 (obtained from the 20th passage of resistant study, see Figure 4.7) to P, D and PD...... 178

Table 5.1. Hydrodynamic diameter (Dh) and zeta potential (ζ) values of individual compounds and combinations used in this study...... 207

Table 5.2. The antimicrobial activities of synthesized polymers against P. aeruginosa PAO1 and PA37...... 209

Table 6.1. Polymer Characterization by NMR, GPC, DLS and Zeta Potential Analysis...... 253

Table 6.2. Antimicrobial and Hemolytic Activities of Antimicrobial Ternary Copolymers...... 254

List of schemes

Scheme 3.1. Synthesis of NO-Modified Antimicrobial Polymer for Synergistic Eradication of Bacteria Biofilm ...... 138

Scheme 4.1. Synthesis of antimicrobial polymer, P, which is an amphiphilic random ternary copolymer, via reversible addition−fragmentation chain transfer (RAFT) polymerization...... 167

Page | xxi

List of abbreviations/symbols

Abs Absorbance AIBN 2,2’-azoisobutyronitrile AgNP Silver

Al2O3 Basic alumina AMPs Antimicrobial peptides ANOVA Analysis of variance AoSMC human aortic smooth muscle cells Attenuated Total Reflectance – Fourier Transform ATR-FTIR Infrared Spectroscopy ATRP Atom transfer radical polymerization gold AuNP Gold nanoparticles BPEI Branched polyethylenimine 2-methacryloyloxyethyl phosphorylcholine-co-butyl BPMPC methacrylate-co-benzophenone

CaCl2 Calcium chloride CCP Cationic conjugated polymer

CDCl3 Deuterated chloroform CFU Colony-forming unit CLSI Clinical and Laboratory Standards Institute CMC Critical micelle concentration CNT Carbon nanotube CV Crystal Violet Đ Polymer dispersity Dab tert-butyl (2-acrylamidoethyl) carbamate DB Degree of branching DCM Dichloromethane

Dh Hydrodynamic diameter DI Deionised DLS Dynamic light scattering DMAc N,N´-dimethylacetamide DMEM Dulbecco’s modified Eagle medium DMF N,N-dimethylformamide DMSO Dimethyl sulfoxide

Page | xxii

DPn Number-averaged degree of polymerization DTDS Dithiol−disulfide EGDMA Ethylene glycol dimethacrylate EPS Extracellular polymeric substance EOs Essential oils FBS Fetal bovine serum FDA Food and Drug Administration FeNP Iron oxide nanoparticle FICI Fractional bactericidal/inhibitory concentration index FRP Free radical polymerization FTIR Fourier transform infrared spectroscopy GO Graphene oxide GPC Gel permeation chromatography HCl Hydrochloric acid Concentration of a compound that cause 20% red blood HC20 cell lysis Concentration of a compound that cause 50% red blood HC50 cell lysis Concentration at which the cell viability is reduced by IC50 half IONPs Iron oxide nanoparticles

KH2PO4 Potassium phosphate monobasic LB Lysogeny broth MBC Minimum bactericical concentration MDR Multi-drug resistant

MgSO4 Magnesium sulfate MHB Mueller-Hinton broth MIC Minimal inhibitory concentration

Mn Number average molecular weight MNPs Magnetic nanoparticles MRSA Methicillin-resistant Mw Weight average molecular weight MWCNT Multiwalled carbon nanotube MWCO Molecular weight cut-off

Na2HPO4 Sodium phosphate dibasic NaCl Sodium chloride

Page | xxiii

NaHCO3 Sodium hydrogen carbonate NaOH Sodium hydroxide

Na2SO4 Sodium sulfate

NH4Cl Ammonium chloride NIH National Institutes of Health NMP Nitroxide-mediated polymerization NMR Nuclear magnetic resonance NO Nitric oxide NONOate N-diazeniumdiolates NPs Nanoparticles OD Optical density OEGA Oligoethylene glycol methyl acrylate PAGA Poly(2-(acrylamido) glucopyranose) PAMAM Poly(amidoamine) PAO1 Pseudomonas aeruginosa O1 PBF Polyfluorene derivative PBS Phosphate buffered saline PCo Cobaltocenium polymers PDI Particle dispersity index PDMAEMA poly [2-(dimethylamino)ethyl methacrylate] PEG Poly(ethylene glycol) PEI Polyethylenimine Photoinduced electron/energy transfer- Reversible PET-RAFT addition-fragmentation chain transfer PGSA Polyglucosamine PHEA Poly(N-hydroxyethyl acrylamide) PHGC Polyhexamethylene guanidine hydrochloride PLB Polymyxin B PLE Polymyxin E PLL Poly(L-lysine) PMAG Poly[(2-methacrylamide) glucopyranose] PMMA Poly(methyl methacrylate) PO Propylene oxide PPCs Polymer-protein complexes PPI Poly(propylene imine) PSAs Poly(sulfone amines)

Page | xxiv

PT Polythiophene derivative PVA Poly(vinylalcohol) QA Quaternary ammonium QS Quorum sensing RAFT Reversible addition-fragmentation chain transfer RBCs Red blood cells RDRP Reversible deactivation radical polymerization rGO Reduced GO nanosheets RNA Ribonucleic acid ROP Ring-opening polymerization ROS Reactive oxygen species RSNO S-nitrosothiols SCPNs Single-chain polymeric nanoparticles SEM Scanning electron microscope SNAP S-nitroso-N-acetylpenicillamine SNO S-nitrosothiols SO oxide SSD Silver sulfadiazine t1/2 Half-life TBAM 2-(tert-butylamino)ethyl methacrylate TEM Transmission electron microscopy TFA Trifluoroacetic acid TGA Thermogravimetric analysis THF Tetrahydrofuran TI Therapeutic index TiO2 Titanium(IV) oxide UA Usnic acid UV-Vis Ultraviolet-visible VRE Vancomycin-resistant Enterococcus WHO World Health Organization XPS X-ray photoelectron spectroscopy ZnO Zinc oxide nanoparticles ZnTPP, 5,10,15,20-tetraphenyl-21H,23H-porphinezinc λ Wavelength ζ Zeta potential

Page | xxv

CHAPTER ONE

Introduction

CHAPTER ONE

1 Introduction

1.1 Thesis motivation

The growing prevalence of infections caused by multidrug-resistant (MDR) bacteria is a serious threat to global health.1-3 The risks associated with antibiotic resistance are not limited to specific countries or continents and negatively impact human health worldwide. In particular, antibiotic resistance puts vulnerable patients (e.g., individuals with ongoing medical conditions or weakened immune systems) at risk. Without having access to effective antibiotics to prevent and treat infections we might no longer be able to perform medical procedures such as surgery, organ transplantation, caesarean section, chemotherapy, etc. due to the high risk of infections.4, 5 Besides causing significant problems in healthcare, antibiotic resistance poses a substantial economic burden on society.6, 7 It is estimated that if antibiotic resistance proceeds with the same rate as of today, by 2050, drug-resistant infections can lead to 10 million deaths per year and a total USD 100 trillion cost.8

Inappropriate use of antibiotics in the form of prescribing errors by practitioners (i.e., unnecessary prescription, wrong antibiotic type or incorrect treatment duration), access to antibiotics without prescription, and misuse and/or overuse of antibiotics in agricultural sectors are key drivers for the development of antibiotic resistance.9-11 The emergence and spread of antibiotic resistance have coincided with the slow progress in the development of new antibiotics, which further complicates the situation. Consequently, many

Page | 2 CHAPTER ONE

common and once easily-treated infectious diseases are becoming more challenging or even impossible to treat, heralding post-antibiotic era.12

According to the National Institutes of Health (NIH), the majority of clinical bacterial infections are biofilm-related.13, 14 In such infections, bacterial cells are shielded by a self-produced polymeric matrix which adheres cells to each other and to the surface.15, 16 Biofilm bacteria are much less susceptible to antibiotic therapies and host immune responses, which makes fighting them even more challenging, in particular, against multidrug-resistant (MDR) strains.17-19

Moreover, the biofilm mode of life stimulates the development and spread of antibiotic resistance. Due to the barrier effect of the extracellular polymeric matrix, the cells are exposed to reduced doses of antimicrobial agents, which induces mutation and resistance development.15 Also, horizontal gene transfer mechanisms are accelerated in the biofilm matrix due to the high cell density within the biofilm matrix, which facilitate the spread of resistance genes.20

The inability of many antibiotics to eradicate MDR or biofilm-related infections, along with the difficulty of producing new lay particular stress on the importance of the development of novel therapeutic approaches to counter these infections.21, 22 Antimicrobial approaches capable of combating MDR strains and bacterial biofilms with a low propensity for inducing resistance in bacterial cells are of particular interest. Mechanistic studies have revealed that antimicrobial agents or strategies with less specific targets such as those that physically disrupt bacterial cells are, in general, less probable to trigger resistance development in bacteria and are also able to target a wide range of

Page | 3 CHAPTER ONE

bacterial cells regardless of their metabolic activity or antibiotic susceptibility such as biofilm bacteria.23

Combining the science of natural antimicrobial peptides (AMPs) with polymerization techniques in the mid-2000s led to the emergence of synthetic antimicrobial polymers as a novel class of antimicrobial agents.24, 25 These polymers are simply synthetic mimics of AMP and exert their bactericidal activity through the same membrane disruption mechanism as AMP with a minimal risk of resistance development in bacteria.26-28 They can be synthesized from relatively inexpensive precursors using facile and well-controlled polymerization techniques. Furthermore, advanced polymerization techniques allow for the synthesis of well-defined polymers with tailorable biological properties for a wide range of antimicrobial applications.29, 30

Another promising strategy to tackle antibiotic resistance is combination therapy, where coadministration of two or more antimicrobial agents shows more potency compared to the individual agents.31-33 Mechanistic studies performed on a wide range of possible combinations have proven that membrane targeting compounds are potential coagents in many antimicrobial combinations.12 While the majority of reported antimicrobial combinations have been based on the use of conventional antibiotics, limited studies on the incorporation of synthetic antimicrobial polymers into combination therapy formulations have shown superior biological performance for these combinations.34

Given the potent biological activity of synthetic antimicrobial polymers individually and their potential positive contributory role in combination

Page | 4 CHAPTER ONE

therapies, coadministration of synthetic antimicrobial polymers and other

antimicrobial agents would offer a promising platform to combat antibiotic-

resistant and biofilm-related infections. This thesis aims to investigate the

potential use of a synthetic antimicrobial polymer, comprising poly(ethylene

glycol) (PEG), hydrophobic, and primary amine groups, in combination with

different antimicrobial agents as a novel therapeutic approach to combat

antibiotic resistance and improve the treatment efficiency against biofilm-

associated infections. The specific objectives of this thesis are:

1. To investigate the advantages of incorporating nitric oxide (NO) donor moieties

into the structure of the synthetic antimicrobial polymer as a novel therapeutic

approach against bacterial biofilms.

2. To study the interactions between the synthetic antimicrobial polymer and

various commercially available antibiotics against normal and MDR bacteria in

both planktonic and biofilm forms.

3. To evaluate the potential use of synthetic antimicrobial polymers in the form of

random and block copolymers in combination with antimicrobial essential oils

as a new antibiofilm platform.

4. To perform a structure-activity relationship study to investigate the effect of

polymer topology and chain length on the biological performance of the

synthesized polymers.

Page | 5 CHAPTER ONE

1.2 Thesis outline

This dissertation reports the development of a robust therapeutic approach based on the use of a synthetic antimicrobial polymer in three different combinations with either NO, antibiotics or essential oils as a potential alternative to current antibiotic therapies against MDR and biofilm-related infections.

With the subject matter of this thesis residing at combination therapy involving synthetic antimicrobial polymers as a potential treatment against MDR and biofilm-related infections, Chapter two provides a general overview of antibiotic resistance, bacterial biofilms and novel therapeutic strategies against antibiotic- resistant and biofilm-related infections. This chapter also includes the fundamental design aspects of synthetic antimicrobial polymers and related structure-activity relationship studies, followed by a brief section on the mechanism and advantages of the reversible addition-fragmentation chain transfer (RAFT) polymerisation technique. The chapter ends with an overview of combination therapy approach and a literature review on the studies that employed synthetic antimicrobial polymers in combination with other antimicrobial agents, including NO, antibiotics, essential oils, metal and metal oxide nanoparticles and finally carbon-based nanomaterials.

Chapter three describes the synthesis of a NO-releasing antimicrobial polymer as a novel antimicrobial/antibiofilm agent. NO donor moieties were incorporated into the structure of the polymer by reacting primary amine with NO gas. The

NO-loaded polymer demonstrated dual-action capability as the release of NO could induce biofilm dispersal, whereas the polymer exerted bactericidal action

Page | 6 CHAPTER ONE

via membrane wall disruption. Upon functionalizing the polymers with NO, a synergistic effect in biofilm dispersal, planktonic and biofilm killing activities against Pseudomonas aeruginosa (P. aeruginosa) was observed. The NO-loaded polymer resulted in 80% reduction in biofilm biomass and killed >99.999% of planktonic and biofilm P. aeruginosa cells within 1 h of treatment at a polymer concentration of 64 μgmL−1. The content of this chapter is based on the published work: Rashin Namivandi-Zangeneh et al., “Nitric Oxide-Loaded Antimicrobial

Polymer for the Synergistic Eradication of Bacterial Biofilm” ACS Macro Letters,

2018,7, 592-597.

Chapter four presents a potent antimicrobial platform based on the physical mixture of commercially available antibiotics and a synthetic antimicrobial polymer. The interactions between the antimicrobial polymer and selected antibiotics from ten different antibiotic classes were assessed using checkerboard assays against P. aeruginosa and (E. coli). Among tested combinations, two synergistic combinations containing doxycycline and colistin were discovered. Results of checkerboard assays showed that the coadministration of the antimicrobial polymer and either of these antibiotics could improve the bacteriostatic efficacy, especially against MDR P. aeruginosa strains PA32 and PA37, where the minimal inhibitory concentrations (MICs) of the polymer and antibiotics were reduced by at least 4-fold. We also observed the synergism in the killing activity, when the antimicrobial polymer was used in combination with doxycycline. This combination could kill >99.999% of planktonic and biofilm P. aeruginosa PAO1 after 20 min of incubation at a polymer concentration of 128 μg mL−1 and doxycycline concentration of 64 μg mL−1. Furthermore, this synergistic combination could efficiently reduce the rate

Page | 7 CHAPTER ONE

of resistance development in P. aeruginosa in comparison with the individual compounds and was able to revive the susceptibility to treatment in the resistant strains. The content of this chapter is based on the published work: Rashin

Namivandi-Zangeneh et al., “Synergy between Synthetic Antimicrobial Polymer and Antibiotics: A Promising Platform To Combat Multidrug-Resistant Bacteria”

ACS Infectious Diseases, 2019,5, 1357-1365.

Chapter five describes an effective antibiofilm platform based on the use of synthetic antimicrobial polymers in combination with essential oils in the form of an oil-in-water emulsion. Apart from being an antimicrobial agent, the antimicrobial polymer played a secondary role as a delivery vehicle for essential oils in these combinations. Given the hydrophobicity of essential oils and their low distribution in aqueous media, we synthesized an antimicrobial polymer in the form of a diblock copolymer to ensure that the antimicrobial polymer can sufficiently stabilize oil droplets and act as a carrier for them. The block copolymer was synthesized by incorporating a second hydrophobic homopolymer block into the structure of our random linear copolymer that consisted of cationic primary amines, low-fouling oligo(ethylene glycol)and hydrophobic ethylhexyl groups. The combinations of the synthesized copolymers and essential oils, namely carvacrol and eugenol, were prepared as oil-in-water emulsions through emulsifying essential oils into an aqueous solution of each polymer. The diblock copolymer underwent self-assembly into micelle morphology in aqueous media and encapsulated oil droplets inside their hydrophobic cavities. In contrast, linear random copolymer could not encapsulate the oil droplets and just merely stabilized them. Combination of antimicrobial polymers, either random or block, and essential oils showed higher

Page | 8 CHAPTER ONE

potency in a biofilm inhibition test against P. aeruginosa biofilms compared to the individual compounds. A 60−75% and 70−85% biofilm inhibition effect was observed for all tested combinations against wild-type P. aeruginosa PAO1 and

MDR strain PA37, respectively, upon 6.5 h of incubation time. In contrast, only the block copolymer acted synergistically with essential oils in killing assay against biofilm bacteria. Treatment of PAO1 biofilm for 20 min with the block copolymer−oil combinations resulted in the killing of >99.99% of biofilm bacteria, while the combinations containing random copolymer did not show any significant improvement relative to their individual components in killing assay.

We attributed the observed synergism in bactericidal activity to the targeted delivery of essential oils to the biofilm site, which was driven by the electrostatic interaction between positively charged delivery vehicles, in the form of polymeric micelles, and negatively charged bacteria. The content of this chapter is based on the published work: Rashin Namivandi-Zangeneh et al., “Antibiofilm

Platform based on the Combination of Antimicrobial Polymers and Essential

Oils” Biomacromolecules, 2020,21, 262-272.

Chapter six presents a detailed investigation on the effects of structural features, including the polymer chain length and architecture on biological performance

(i.e., the antimicrobial activity and hemocompatibility) of antimicrobial ternary copolymers that consisted of low-fouling oligo(ethylene glycol), cationic and hydrophobic side chains. Assessment of the antimicrobial activity against P. aeruginosa and E. coli showed that the polymer with longer chains (DP = 100) was slightly more bacteriostatic that polymers with shorter chain lengths (DP = 20-

50). In terms of hemocompatibility, polymers with shorter chains were more prone to induce hemagglutination. Interestingly, when the hydrophilic and

Page | 9 CHAPTER ONE

hydrophobic segments were separated into distinct blocks, the antimicrobial activity was lost in the resulting diblock copolymer. This was probably due the stable core–shell morphology of the resulting micelles, which were unable to interact with the bacterial membrane. Finally, polymers with the branched architecture were evaluated in terms of antimicrobial and hemolytic activities.

The hyperbranched polymer, which consisted of 2-ethylhexyl groups as hydrophobic side-chains had similar antimicrobial activity to the linear random copolymers (MIC = 64 μg mL−1). However, this polymer showed more than 4-fold increase in HC50 (defined as the polymer concentration that caused 50% red blood cell lysis) compared to the linear random copolymers (HC50 >10,000 μg mL−1) with no hemagglutination. The hyperbranched polymers also exhibited bactericidal activity and killed ≥99% and 90% of planktonic and biofilm P. aeruginosa, respectively, upon a 1 h treatment at the concentration of 64 μg mL−1. The content of this chapter is based on the published work: Rashin Namivandi-Zangeneh et al., “The effects of polymer topology and chain length on the antimicrobial activity and hemocompatibility of amphiphilic ternary copolymers” Polymer

Chemistry, 2018, 9, 1735-1744.

Finally, Chapter seven provides a brief summary of the main outcomes of this thesis and provides some recommendations for future works.

1.3 Reference

1. Blair, J. M. A.; Webber, M. A.; Baylay, A. J.; Ogbolu, D. O.; Piddock, L. J. V., Molecular mechanisms of antibiotic resistance. Nat. Rev. Microbiol. 2015, 13 (1), 42-51. 2. Talebi Bezmin Abadi, A.; Rizvanov, A. A.; Haertlé, T.; Blatt, N. L., World Health Organization Report: Current Crisis of Antibiotic Resistance. BioNanoScience 2019, 9 (4), 778-788.

Page | 10 CHAPTER ONE

3. Schillaci, D.; Spanò, V.; Parrino, B.; Carbone, A.; Montalbano, A.; Barraja, P.; Diana, P.; Cirrincione, G.; Cascioferro, S., Pharmaceutical Approaches to Target Antibiotic Resistance Mechanisms. J. Med. Chem. 2017, 60 (20), 8268-8297. 4. Brown, E. D.; Wright, G. D., Antibacterial drug discovery in the resistance era. Nature 2016, 529 (7586), 336-343. 5. Sommer, M. O. A.; Munck, C.; Toft-Kehler, R. V.; Andersson, D. I., Prediction of antibiotic resistance: time for a new preclinical paradigm? Nat. Rev. Microbiol. 2017, 15 (11), 689-696. 6. Roberts, R. R.; Hota, B.; Ahmad, I.; Scott, R. D., II; Foster, S. D.; Abbasi, F.; Schabowski, S.; Kampe, L. M.; Ciavarella, G. G.; Supino, M.; Naples, J.; Cordell, R.; Levy, S. B.; Weinstein, R. A., Hospital and Societal Costs of Antimicrobial-Resistant Infections in a Chicago Teaching Hospital: Implications for Antibiotic Stewardship. Clin. Infect. Dis. 2009, 49 (8), 1175-1184. 7. Gandra, S.; Barter, D. M.; Laxminarayan, R., Economic burden of antibiotic resistance: how much do we really know? Clin. Microbiol. Infect. 2014, 20 (10), 973- 980. 8. O'Neill, J. Tackling antimicrobial resistance, http://www.rsc.org/news- events/features/2015/may/tackling-antimicrobial-resistance/. 9. Ventola, C. L., The antibiotic resistance crisis: part 1: causes and threats. P T 2015, 40 (4), 277-283. 10. Berendonk, T. U.; Manaia, C. M.; Merlin, C.; Fatta-Kassinos, D.; Cytryn, E.; Walsh, F.; Bürgmann, H.; Sørum, H.; Norström, M.; Pons, M.-N.; Kreuzinger, N.; Huovinen, P.; Stefani, S.; Schwartz, T.; Kisand, V.; Baquero, F.; Martinez, J. L., Tackling antibiotic resistance: the environmental framework. Nat. Rev. Microbiol. 2015, 13 (5), 310-317. 11. Hay, S. I.; Rao, P. C.; Dolecek, C.; Day, N. P. J.; Stergachis, A.; Lopez, A. D.; Murray, C. J. L., Measuring and mapping the global burden of antimicrobial resistance. BMC Med. 2018, 16 (1), 78-78. 12. Brochado, A. R.; Telzerow, A.; Bobonis, J.; Banzhaf, M.; Mateus, A.; Selkrig, J.; Huth, E.; Bassler, S.; Zamarreño Beas, J.; Zietek, M.; Ng, N.; Foerster, S.; Ezraty, B.; Py, B.; Barras, F.; Savitski, M. M.; Bork, P.; Göttig, S.; Typas, A., Species-specific activity of antibacterial drug combinations. Nature 2018, 559 (7713), 259-263. 13. Jamal, M.; Ahmad, W.; Andleeb, S.; Jalil, F.; Imran, M.; Nawaz, M. A.; Hussain, T.; Ali, M.; Rafiq, M.; Kamil, M. A., Bacterial biofilm and associated infections. J. Chin. Med. Assoc. 2018, 81 (1), 7-11. 14. Miquel, S.; Lagrafeuille, R.; Souweine, B.; Forestier, C., Anti-biofilm Activity as a Health Issue. Front. Microbiol. 2016, 7, 592-592. 15. Lebeaux, D.; Ghigo, J.-M.; Beloin, C., Biofilm-related infections: bridging the gap between clinical management and fundamental aspects of recalcitrance toward antibiotics. Microbiol. Mol. Biol. Rev. 2014, 78 (3), 510-543. 16. Sadrearhami, Z.; Nguyen, T.-K.; Namivandi-Zangeneh, R.; Jung, K.; Wong, E. H. H.; Boyer, C., Recent advances in nitric oxide delivery for antimicrobial applications using polymer-based systems. J. Mater. Chem. B 2018, 6 (19), 2945-2959. 17. Mah, T.-F. C.; O'Toole, G. A., Mechanisms of biofilm resistance to antimicrobial agents. Trends Microbiol. 2001, 9 (1), 34-39. 18. Römling, U.; Balsalobre, C., Biofilm infections, their resilience to therapy and innovative treatment strategies. J. Intern. Med. 2012, 272 (6), 541-561. 19. Wolfmeier, H.; Pletzer, D.; Mansour, S. C.; Hancock, R. E. W., New Perspectives in Biofilm Eradication. ACS Infect. Dis 2018, 4 (2), 93-106.

Page | 11 CHAPTER ONE

20. Madsen, J. S.; Burmølle, M.; Hansen, L. H.; Sørensen, S. J., The interconnection between biofilm formation and horizontal gene transfer. FEMS IMMUNOL MED MIC 2012, 65 (2), 183-195. 21. Costerton, J. W.; Stewart, P. S.; Greenberg, E. P., Bacterial Biofilms: A Common Cause of Persistent Infections. Science 1999, 284 (5418), 1318-1322. 22. Stewart, P. S.; William Costerton, J., Antibiotic resistance of bacteria in biofilms. Lancet 2001, 358 (9276), 135-138. 23. Zasloff, M., Antimicrobial peptides of multicellular organisms. Nature 2002, 415 (6870), 389-395. 24. Ergene, C.; Yasuhara, K.; Palermo, E. F., Biomimetic antimicrobial polymers: recent advances in molecular design. Polym. Chem. 2018, 9 (18), 2407-2427. 25. Ganewatta, M. S.; Tang, C., Controlling macromolecular structures towards effective antimicrobial polymers. Polymer 2015, 63, A1-A29. 26. Jain, A.; Duvvuri, L. S.; Farah, S.; Beyth, N.; Domb, A. J.; Khan, W., Antimicrobial Polymers. Adv. Healthc. Mater. 2014, 3 (12), 1969-1985. 27. Konai, M. M.; Bhattacharjee, B.; Ghosh, S.; Haldar, J., Recent Progress in Polymer Research to Tackle Infections and Antimicrobial Resistance. Biomacromolecules 2018, 19 (6), 1888-1917. 28. Timofeeva, L.; Kleshcheva, N., Antimicrobial polymers: mechanism of action, factors of activity, and applications. Appl. Microbiol. Biotechnol. 2011, 89 (3), 475-492. 29. Boyer, C.; Bulmus, V.; Davis, T. P.; Ladmiral, V.; Liu, J.; Perrier, S., Bioapplications of RAFT Polymerization. Chem. Rev. 2009, 109 (11), 5402-5436. 30. Kenawy, E.-R.; Worley, S. D.; Broughton, R., The Chemistry and Applications of Antimicrobial Polymers: A State-of-the-Art Review. Biomacromolecules 2007, 8 (5), 1359-1384. 31. Fischbach, M. A., Combination therapies for combating antimicrobial resistance. Curr. Opin. Microbiol. 2011, 14 (5), 519-523. 32. Tamma, P. D.; Cosgrove, S. E.; Maragakis, L. L., Combination Therapy for Treatment of Infections with Gram-Negative Bacteria. Clin. Microbiol. Rev. 2012, 25 (3), 450-470. 33. Karaiskos, I.; Antoniadou, A.; Giamarellou, H., Combination therapy for extensively-drug resistant gram-negative bacteria. Expert Rev. Anti Infect. Ther. 2017, 15 (12), 1123-1140. 34. Yañez-Macías, R.; Muñoz-Bonilla, A.; De Jesús-Tellez, M. A.; Maldonado- Textle, H.; Guerrero-Sánchez, C.; Schubert, U. S.; Guerrero-Santos, R., Combinations of Antimicrobial Polymers with Nanomaterials and Bioactives to Improve Biocidal Therapies. Polymers 2019, 11 (11), 1789.

Page | 12

CHAPTER TWO

Literature review

CHAPTER TWO

2 Literature Review

2.1 Antibiotic resistance

The rising number of infectious diseases caused by multidrug-resistant (MDR) bacteria is a critical global health issue1, 2 that can lead to prolonged hospital stay periods, and increased medical costs and mortality. Although resistance development in bacteria is a natural phenomenon,3, 4 the overuse and misuse of antibiotics have accelerated the rate of resistance development over the past few decades. For instance, resistance acquisition has been observed within short periods (few months) in clinical settings.5, 6 Antibiotic resistance is the evolution of mechanisms in bacterial cells to diminish the effect of antibiotics.7 While there exists different classes of antibiotics that act on specific intracellular targets to interfere with cell wall biosynthesis, protein synthesis, DNA replication or folate biosynthesis,8, 9 bacteria have successfully developed several resistance mechanisms to reduce the effectiveness of almost all available antibiotics.

2.1.1 Mechanisms of antibiotic resistance

Three main categories of antibiotic resistance, namely intrinsic, acquired and adaptive, have been identified in bacterial cells. 10-13 Antibiotic resistance might be an innate ability of the bacteria to reduce the susceptibility to a specific antibiotic regardless of the previous antibiotic exposure (intrinsic resistance).

Intrinsic resistance is the result of the unique structural or functional characteristic of the bacterial species such as the reduced membrane permeability in Gram-negative bacteria and the activity of efflux pumps (Figure 2.1).14

Page | 14 CHAPTER TWO

Antibiotic resistance can also be obtained through mutation or horizontal gene transfer mechanisms (acquired resistance). Finally, bacteria can develop resistance as an adaptive transient response to the unfavourable surrounding environment (adaptive resistance). The four major antibiotic resistance mechanisms in bacteria are: (1) reduced membrane permeability (exclusive to

Gram-negative species), (2) efflux pump, (3) antibiotic modification, and (4) target modification.1, 15, 16

Figure 2.1. Intrinsic resistance mechanisms in a Gram-negative bacteria. Three different scenarios regarding the interaction of β-lactam antibiotics with Gram-negative bacteria to reach a penicillin-binding protein (PBP). Antibiotic A can cross the outer membrane via porins and bind to PBP. Antibiotic B can cross the outer membrane via porins but is expelled out via the efflux pumps. The penetration of antibiotic C is inhibited by the outer membrane.1

2.1.1.1 Reduced permeability of outer membrane

The unique membrane structure in Gram-negative bacteria plays a key role in intrinsic resistance by limiting the influx of antibiotics.14 The outer membrane, composed of phospholipids, lipopolysaccharides (LPS) and proteins in the form of porins, provides a formidable barrier against the influx of a wide range of

Page | 15 CHAPTER TWO

substances and antibiotics. Moreover, changes in the nature and relative composition of the membrane components due to mutation or horizontal gene transfer can significantly increase the resistance toward certain antibiotics.

Downregulation of porins or reduction of their activity is a major mechanism of resistance towards antibiotics that need to pass through porins to access their intracellular targets (e.g., β-lactams, , and fluoroquinolones).17

Alterations to the structure of LPS can also lead to the reduced permeability of the outer membrane to hydrophobic and cationic antibiotics such as aminoglycosides and polymyxins.18

2.1.1.2 Antibiotic efflux

Expulsion of antibiotics out of bacterial cells is another important mechanism of inherent resistance in many bacterial species. Efflux pumps are embedded proteins in the structure of the membrane in both Gram-negative and Gram- positive bacteria.11 These proteins can pump out antibiotics and therefore keep the intracellular antibiotic concentration low enough to minimize and even eliminate damage.1, 15 Efflux pumps can be highly selective to pump out a wide range of molecules (i.e., multidrug resistance efflux pumps).1 As well as being an intrinsic resistance mechanism, overexpression of the efflux pumps due to mutation or as a result of environmental conditions is also common.

2.1.1.3 Antibiotic modification

Changing the chemical structure of the antibiotics is an effective approach used by bacterial cells to diminish the efficacy of antibiotics. These changes can be done through of the antibiotic or by altering the chemical functionalities of the antibiotic (Figure 2.2 a).1, 19 Hydrolysis is mediated by Page | 16 CHAPTER TWO

which are produced by bacteria and can destroy the structure of antibiotic at a very high rate. β-lactamase enzymes are probably the best-known example in this category which hydrolyse the β-lactam ring of antibiotics such as penicillin and cephalosporin.20 Antibiotic modification can also occur through changing the active sites of the antibiotic molecule (warheads) which would eventually decrease the affinity of antibiotic and its target. This is especially effective against antibiotics with many active sites such as members of aminoglycoside family.21, 22

Figure 2.2. Antibiotic resistance by (a) antibiotic modification and (b) target modification.1

2.1.1.4 Target modification

While all three aforementioned resistance mechanisms aim to reduce the effective intracellular concentration of antibiotics, bacteria can still evade the action of antibiotics even in the presence of sufficient intact antibiotic by camouflaging or reprogramming the antibiotic binding sites (Figure 2.2 b).

Changing the antibiotic target site will negatively affect the affinity between antibiotic and its target, binding efficiency and therefore, antibiotic efficacy.1, 16

Page | 17 CHAPTER TWO

Alteration of the target sites might happen through mutation, enzymatic alternation, overproduction of target site or replacement of the target.16, 23

Resistance to macrolides by methylation of ribosomes or resistance to rifampicin by mutation of the RNA polymerase are well-known examples in this category.

8, 23

2.1.2 Antibiotic resistance acquisition

In addition to intrinsic resistance, bacterial cells are also capable of developing resistance through mutational changes and horizontal gene transfer (Figure 2.3).

In the former, resistance genes emerge and pass to progeny during division while in the latter, resistance genes are transferred from one bacteria to another bacteria.24 Horizontal gene transfer is probable between bacterial cells from different strains or even different species. Three major mechanisms contributing to horizontal gene transfer are transformation, transduction and conjugation. In transformation, bacteria influx the DNA segments which are released by dead bacterial cells in the environment. Transduction refers to the transfer of resistance genes between two bacteria by bacteriophages. Finally, conjugation is mediated by direct physical contact between bacterial cells and subsequent transfer of conjugative plasmids via pilus.25 Horizontal evolution mechanisms are known to be the main cause for the development and global distribution of antibiotic resistance for clinically important bacteria.26

Page | 18 CHAPTER TWO

Figure 2.3. Resistance acquisition through (a) mutation and (b) horizontal gene transfer.24

2.1.3 Adaptive Resistance

In response to hostile environmental conditions, bacteria can develop temporary resistance as an adaptive reaction to ensure their survival.27 This transient resistance usually involves the formation of new bacterial phenotypes and changes in genes and proteins expression. Two common adaptive resistance approaches are biofilm development and formation of persister cells,12, 28 which will be pointed out in the following part after a short introduction to bacterial biofilms.

2.2 Bacterial biofilms

Biofilm-associated infections contribute to the majority of chronic and recurrent infections and constitute a threat to public health and economy worldwide.29-32 These infections are highly recalcitrance toward host immune

Page | 19 CHAPTER TWO

responses and antibiotic treatments, even in patients with decent general health condition.33, 34 The unique characteristics of bacterial biofilms make the diagnosis and treatment of related infections more challenging compared to the infections caused by their free-floating planktonic counterparts.35, 36 As there has never been an antimicrobial agent that is exclusively effective toward bacterial biofilm, current approaches against biofilm-related infections are based on available antibiotics. These treatments involve long-term antibiotic therapies at high dosages along with the removal of the infected tissues (wound debridement) or implants, in the case of implant-related infections.37, 38 Thus, novel antimicrobial agents or therapeutic approaches that are specially designed against bacterial biofilm are highly demanded.

Biofilm bacteria are correlated with antibiotic resistance and tolerance in many ways. Biofilms are inherently tolerant toward antimicrobial therapy as a result of their extraordinary features, including the presence of extracellular polymeric substance (EPS) as a protecting shield, heterogeneity within the structure of biofilm and the presence of persister cells.34 The biofilm mode of existence can also accelerate the resistance development rate in bacteria by facilitating the horizontal gene transfer between the bacterial cells within the biofilm community.34, 39 Moreover, biofilm-related infections are frequently caused by antibiotic resistance strains, which makes them even harder to be eradicated by common antibiotic therapies. Gram-negative Pseudomonas aeruginosa and Gram- positive Staphylococcus aureus are major contributors to biofilm infections, and both belong to the high-risk group of antibiotic resistance announced by World Health Organisation (WHO).2

Page | 20 CHAPTER TWO

Biofilms are multicellular/multispecies communities of bacterial cells which are embedded in a self-produced three-dimensional matrix of EPS.31 These saturated non-mobile aggregates of bacteria are prevalent in many aqueous environments and inhabit on many biological surfaces, a wide range of medical devices, and inert surfaces.35, 40, 41 Biofilm formation is known to happen as a response to unfavourable environmental conditions.42 This unique lifestyle, however, is the dominant form of life in almost all humid or aqueous habitats and constitutes a major part of the bacterial biomass.43, 44

2.2.1 Biofilm life cycle

The switch in the lifestyle from free-floating cells to the sessile community involves a multi-stage process and causes major changes in the physiology and gene expression of the bacterial cells (Figure 2.4).38, 45 The first step in biofilm development is the transient attachment of bacterial cells to a surface, which is mediated by hydrophobic and electrostatic interactions and also van der Waals forces. Afterward, irreversible attachments are formed through surface appendages, including pili and fimbriae.46, 47 Attached cells then grow into microcolonies which establish the foundation of biofilm structure. Driven by altered gene expression, attached cells start a process of proliferation and secretion of a wide range of and biomolecules including proteins, polysaccharides, extra-cellular DNA and lipids to build the biofilm matrix.

During this process, the matrix grows into an elaborate three-dimensional structure with built-in water channels to deliver oxygen and nutrients to the cells which are embedded in the inner layers of biofilm, and also to dispose any metabolic by-products.46, 48 This stage of biofilm development is known as

Page | 21 CHAPTER TWO

maturation. The last step in the bacterial biofilm life cycle is dispersal, which can happen due to external physical forces (passive dispersal) or as a response to the environmental or biological cues (active dispersal).49, 50 Environmental cues for biofilm dispersal include changes in the physio-chemical properties of biofilm habitat such as temperature, pH, oxygen or nutrient levels while, biological cues are signals that are produced by the cells within the biofilm matrix. Active dispersal is mainly controlled through quorum sensing and often coordinate with regulated cell death and autolysis.50, 51 Upon sensing the dispersal cue, biofilm matrix undergoes degradation and bacterial cells are released from the biofilm matrix. These planktonic bacteria are differentiated versions of bacterial cells which are capable of colonizing new habitats and causing acute and recurrent infections.49

Figure 2.4. Representative stages of the biofilm life cycle.34

2.2.2 Recalcitrance of bacterial biofilms toward antibiotics

A chief characteristic of bacterial biofilms which differentiate them from planktonic bacteria is their recalcitrance toward antimicrobial agents, even if the biofilm is grown from susceptible strains of bacteria.31, 52, 53 Although the higher Page | 22 CHAPTER TWO

cell density in biofilm matrix (108-11 colony forming units (CFU) cm−2) than planktonic bacteria (103-4 CFU mL−1) might account for their resilience, it has been observed that biofilms are still less susceptible to antimicrobial agents even at the equivalent cell density.54 Besides, the fact that bacterial cells revive their susceptibility to treatment upon biofilm dispersal confirms that other factors, apart from causes of intrinsic and acquired resistance, should be involved in biofilm recalcitrance.15, 55

Biofilm recalcitrance emerges as the combined result of resistance and tolerance to antibiotics. Antibiotic resistance is the ability of bacteria to proliferate in the presence of antibiotics and usually involves mechanisms that prevent the antibiotic from binding to its target. This ability is genetic-based and can be transferred during cell division or through horizontal gene transfer.34, 39, 56

Tolerance, on the other hand, refers to the survival of bacteria in the presence of antibiotics. Tolerance can be either genotypic, caused by genetic alterations, or phenotypic, caused by environmental conditions. In both cases, there is a reduction in the ability of antibiotics to kill the bacteria. While the genotypic type of tolerance is permanent and heritable, changes in the environmental conditions might revert phenotypic tolerance.34, 39, 57 In the following section, we will briefly explain the main mechanisms that are involved in bacterial biofilm recalcitrance.

It is worthwhile noting that in the majority of the cases, a combination of these mechanisms contributes to the overall resilience of the biofilm.

2.2.2.1 The barrier effect of EPS

In biofilms, bacteria are enclosed in a self-produced three-dimensional scaffold consist of different biopolymers. This polymeric matrix supports the

Page | 23 CHAPTER TWO

biofilm integrity and adhesion to the surface.58 EPS constitute more than 90% of the dry mass of bacterial biofilms and is known to play an important role in biofilm tolerance.39, 58 Biofilm matrix can affect the access of antibiotics to the cells within the biofilm through interactions with the antibiotic molecules (Figure 2.5 a). Absence of these interactions, however, leads to the complete diffusion of antibiotic through biofilm matrix.59 It is now well accepted that the biofilm matrix is not impermeable, yet, it can reduce the penetration or activity of the antimicrobial agents.60, 61 This reduction might be due to the adsorption on or reaction with the biofilm matrix, where the latter usually causes antibiotic deactivation.30, 62 The diffusion of antibiotics through the biofilm matrix is directly related to the chemical structure of both antibiotic and biofilm matrix.53 While the diffusion is greatly retarded for a specific antibiotic, another antibiotic might easily pass through the same biofilm matrix. It is worthwhile noting that the complete penetration of an antibiotic into the biofilm matrix does not guarantee eradication of the biofilm, as the antibiotic efficiency might be reduced due to the simultaneous effect of other factors.34

Figure 2.5. The main mechanisms of antibiotic tolerance in bacterial biofilms.34

Page | 24 CHAPTER TWO

2.2.2.2 Metabolic heterogeneity

Metabolic activity and growth rate are correlated with antibiotic susceptibility in both planktonic and biofilm bacteria, although to a greater extent for the biofilm bacteria. The reduction of the growth rate from exponential to stationary phase would increase the tolerance toward antibiotics, as the majority of antibiotics are only effective against metabolically active cells.31, 63, 64 Studies have shown that metabolic activity of the bacterial cells dramatically decreases when moving from the outer layers toward the centre of biofilm matrix (Figure 2.5 b).

This reduction in the metabolic activity is associated with the gradual changes in the physicochemical parameters of the microenvironment of EPS matrix such as pH, nutrients, oxygen, and waste levels.65 Additionally, an increased tolerance has been observed in aged biofilms compared to young ones, as the population of cells with low metabolic activity increase once the biofilm is aged.39

2.2.2.3 Persisters subpopulation

The presence of persister cells as an isogenic sub-population in biofilms can elucidate the biofilm recalcitrant in cases where the antibiotic can completely diffuse into the biofilm matrix and efficiently kill the non-dividing cells (e.g., fluoroquinolone).66 Persisters are not genetic mutants, but transient phenotypes that constitute less than 1% of bacterial population in both planktonic and biofilms. They can inevitably survive in the presence of antibiotics, escape the immune responses of the host and eventually resume proliferation once the antibiotic treatment is ended (Figure 2.5 c).67

Page | 25 CHAPTER TWO

2.2.3 Resistance acquisition in biofilms

It has been proven that the biofilm cycle can significantly promote resistance development in bacterial cells.67, 68 Biofilm matrix can facilitate horizontal gene transfer between bacterial cells within the biofilm community. In particular, plasmid conjugation is accelerated due to the high cell density in the biofilm matrix which favour cell-cell physical contact.68 Moreover, the inhibitory effect of

EPS on the antibiotic diffusion can trigger resistance development by exposing the bacterial cells to subinhibitory concentrations of antibiotics for a long period which in turn would increase the chance of mutation.34

2.3 Novel therapeutic strategies to combat antibiotic resistance

Antibiotic resistance is the main cause of treatment failure using conventional antibiotic therapies. There is an urgent need to design novel antimicrobial agents or strategies to address this global crisis. While pharmaceutical companies are slow to invest in new antibiotic discovery,69, 70 various novel compounds and strategies have been developed by researchers over the past years. An ideal alternative to traditional antibiotics should exhibit a lower risk of resistance development in bacteria and at the same time, be able to overcome the resistant strains. Antibiofilm agents and approaches, although aiming for different strategies, are also contributing to the battle against antibiotic resistance by targeting adaptive resistance and inhibiting resistance acquisition in biofilms.

Table 2.1 outlines the ongoing research on therapeutic strategies against antibiotic resistance and bacterial biofilms.

Page | 26 CHAPTER TWO

Table 2.1. Therapeutic strategies against antibiotic resistance and/or bacterial biofilms.

Approach Description Mode of action Ref.

  Co-administration of two or more Synergistic interactions between compounds lead to Combination therapy 69, 71-74 antimicrobial agents at the same time higher treatment efficiency  Attachment and replication of bacteriophages inside  75-78 Phage therapy Using bacteriophages as bactericidal the bacterial cells cause cell lysis agents   Using visible light and Generation of reactive oxygen species (ROS) which Photodynamic therapy photosensitiser molecules to treat cause oxidative damage to the and 79-81 localized infections DNA  Applying a low/ high voltage current  2 2 to a conductive surface to eradicate Generation of H O and ROS which induce oxidative Electrochemical biofilm damage to cell membrane and DNA 38, 82-84 therapy   Using antibiotics in combination with Increasing the susceptibility of biofilms to antibiotics electrical current (Bioelectric effect)

 Using low-frequency ultrasound to eradicate bacterial biofilm (both alone  Inducing cavitation in or near the bacteria and and in combination with antibiotics) generation of peroxides Ultrasound-mediated  Using acoustically activated  Increase the efficacy of antibiotics by enhancing the 85-87 therapy microbubbles for localized delivery of permeability of cell membrane and biofilm matrix therapeutic agents and physical eradication of biofilms

Page | 27 CHAPTER TWO

 Limiting the interactions between bacteria and surfaces to prevent bacterial adhesion subsequent  Modification of surfaces with low- formation of biofilm Antimicrobial surfaces  88-91 fouling or antimicrobial compounds Killing the bacterial cells upon physical contact of cells with the surface or due to the release of active biocidal agents from the surface

  Modifying the chemical structure of Extending the life of existing antibiotics by Analog development existing families of antibiotics in an circumventing resistance mechanisms 8, 92, 93  effort to achieve novel antibiotics Generating new classes of antibiotics

 Co-administration of conventional  Blocking the main resistance mechanisms in bacteria Antibiotic adjuvants antibiotics along with antibiotic and prolonging the efficient lifetime of antibiotics 94-96 adjuvants (molecules with no or low antimicrobial activity)

 Exploiting the potential antibacterial  A wide range of antimicrobial compounds in the Natural compounds activity of natural antimicrobial form of essential oils, proteins, glycoproteins, 97-99 agents originated from plants and polysaccharides with diverse modes of action animal  Natural peptides produced as the first  Demonstrating antimicrobial activity by cell wall Antimicrobial peptides defence line in all multicellular and membrane disruption 100-104 organisms  Synthetic mimics of natural Antimicrobial antimicrobial peptides  Demonstrating antimicrobial activity by cell wall 105-111 polymers and membrane disruption

Page | 28 CHAPTER TWO

 Antibacterial and antibiofilm activities are exerted  Using polymeric, carbon-based, metal by various mechanisms, including membrane and metal oxide nanomaterials with disruption, oxidative stress through ROS generation, 112-115 Nanomaterials inherent antimicrobial activity DNA damage and etc.  Designing nanoparticle-based  Targeted and responsive antibiotic delivery to the delivery platforms site of infection

 At low concentrations, NO works as a biological signalling molecule to induce biofilm dispersal  Incorporation of NO as a potent  Nitric oxide (NO) At high concentrations, NO demonstrates 37, 116-118 antibacterial and antibiofilm agent in bactericidal activity through causing nitrosative and the therapeutic regimen oxidative stress

 Using pro-drug antibiotics, compounds that exhibit bactericidal activity only once entered the bacterial  Targeting persister population to cell  fully eradicate chronic and recurrent Combination therapy based on antibiotics and other Anti-persister therapy 67, 119-121 infections which are usually caused antimicrobial agents or adjuvants  by bacterial biofilm Invading bacterial cells regardless of their metabolic activity (physical or chemical disruption of bacterial cells)

 Using natural or synthetic bioactive  Preventing or decreasing the biosynthesis of QS molecules to quench or inhibit the QS Quorum sensing (QS) signal molecules system, interfere with bacteria cell-  122-126 inhibition/quenching Using quorum quenching enzymes to inactivate the cell communication and finally QS signal molecules decrease biofilm formation

Page | 29 CHAPTER TWO

In line with the general scope of this thesis, which is on the use of synthetic antimicrobial polymers in combination with other antimicrobial agents as potential alternative to traditional antibiotics, the focus of the remaining parts of this chapter will be on the fundamental characteristic of synthetic antimicrobial polymers, followed by a brief introduction to the principle and advantages of the reversible addition-fragmentation chain transfer (RAFT) polymerization method. Finally, a literature review of reported studies on combination therapy based on synthetic antimicrobial polymers and other antimicrobial agents will be presented.

2.4 Synthetic antimicrobial polymers

Inspired by the chemical structure of natural antimicrobial peptides (AMPs) and driven by the advances of controlled polymerization techniques, synthetic antimicrobial polymers have been developed as potential alternatives to conventional antibiotics.105, 106, 127, 128 These polymers offer many advantages over their natural counterparts and also small-molecule antibiotics. For example, they can be synthesized in a facile manner at low cost, while their biological activity can be easily tailored through modification of their chemical structure, and finally, they are active against a wide range of pathogens with minimal risk of inducing resistance development in bacteria.129

2.4.1 Structure-activity relationships

Antimicrobial polymers in the form we know today have been through an evolutionary transformation which started as biocidal polymers and gradually

CHAPTER TWO

evolved to imitate the unique characteristics of AMPs.110, 130 Continuous modification and optimisation of these polymers, performed through systematic structure-activity relationship studies, have significantly improved their biological performance. Recently, such studies are conducted using high- throughput screening processes which allow for the fast and accurate production and evaluation of novel antimicrobial polymers.131, 132 In this section, we aim to summarise the most critical design criteria for antimicrobial polymers based on promising structure-activity relationship studies conducted over the past two decades. The reader is referred to comprehensive discussions on the design, synthesis and properties of antimicrobial polymers in several recent publications for more information in this regard.105, 108, 110, 130, 133

2.4.1.1 Cationic monomers

To achieve the same antimicrobial potency as AMPs, the most critical characteristic of AMPs to mimic in synthetic polymers is the chemical composition. Natural AMPs are composed of a variety of amino acid residues bearing cationic, hydrophobic or hydrophilic functionalities.134, 135 Cationic residues (e.g., lysine and arginine) are vital components in the structure of AMPs, as they trigger the bactericidal action by inducing electrostatic interactions with the anionic of bacterial membrane.100

A wide range of cationic groups has been used in the structure of antimicrobial polymers, where the source of positive charge might be moieties like amines, ammonium salts, guanidium, phosphonium, sulfonium, etc.105, 130 Considering the dissimilarities in the outer membrane layer of microbial species, namely, Gram-

Negative, Gram-positive, and fungal cells (Figure 2.6), species-specific activities

Page | 31 CHAPTER TWO

of cationic moieties are expected. Therefore, several investigations have been performed to explore the effect of cationic groups on the antimicrobial activity of synthetic antimicrobial polymers.

Figure 2.6. Illustration of the cross sections of mammalian and microbial cells’ membrane.130

In general, a higher antimicrobial performance has been observed for the primary amine groups against Gram-negative bacteria in comparison with other cationic functionalities such as tertiary amines or quaternary ammonium (QA) groups.136-138 In a study reported by Kuroda and Palermo, series of methacrylate- based antimicrobial polymers were synthesized using primary or tertiary amines and also QA as the cationic functionality, and methyl or butyl methacrylate as the hydrophobic monomer (Figure 2.7).137 When tested against Escherichia coli

DH5, primary or tertiary amine group containing copolymers demonstrated higher antimicrobial activity than QA containing polymers regardless of the type of hydrophobic monomer. For instance, in the series of polymers synthesized using methyl methacrylate as hydrophobic monomer, the best antimicrobial activity was observed for a primary amine containing polymer with the minimum inhibitory concentration (MIC) of 8.1 µg mL−1, while the corresponding

Page | 32 CHAPTER TWO

tertiary amine and QA bearing polymers (with the same mole fraction of methyl side chains of 0.65) had the MIC values of 840 and >2000 µg mL−1, respectively.

Moreover, when assessed for hemolytic activity, as an initial indicator of mammalian cell biocompatibility, polymers based on primary or tertiary amines showed generally higher selectivity (defined as the ratio of polymer concentration that caused 50% red blood cell (RBC) lysis, HC50, to MIC) compared to QA containing polymers.

Figure 2.7. Chemical structure of methacrylate-based antimicrobial copolymers containing primary or tertiary amines and QA groups as cationic moieties.137

Primary amines show high affinity to phospholipid component of the outer membrane leaflet in Gram-negative bacteria and tend to interact with the membrane through a combination of hydrogen bonding and electrostatic interactions. At the same time, this cationic group demonstrates lower toxicity towards mammalian cells in comparison with QA groups.110 In contrast, primary amines might show low or even no activity against Gram-positive or mycobacteria. In a recent work by Judzewitsch et al., a remarkable increase in the activity against Mycobacterium smegmatis (M. smegmatis) was observed when the cationic group was changed from primary to tertiary and quaternary ammonium.132 In a similar work by Phillips et al., tertiary amine bearing Page | 33 CHAPTER TWO

methacrylate polymers showed higher potency against Mycobacterium tuberculosis (M. tuberculosis) in comparison with primary amine bearing polymers

(Figure 2.8).139 Fluorescent microscopy and transmission electron microscopy

(TEM) images showed that the membrane of M. tuberculosis was not disrupted after the treatment with tertiary amine containing polymer, suggesting that a mechanism distinct from membrane disruption might be involved in the antimicrobial activity of this polymer against M. tuberculosis. Moreover, tertiary amine containing polymer were less hemolytic compared to primary amine containing polymers over a wide range of concentrations and caused less than

5% hemolysis at 5 mg mL−1.

Figure 2.8. Selective antimicrobial activity of tertiary amine-containing methacrylate polymers toward Mycobacterium tuberculosis over RBCs.

Another potent cationic moiety which is also present in the structure of AMPs is guanidium. Antimicrobial polymers based on guanidium as their source of positive charge have shown promising biological performance in comparison with primary or tertiary amines as demonstrated by the groups of Tew and

Page | 34 CHAPTER TWO

Locock.140, 141 In both studies, the highest antimicrobial activity and selectivity was observed for guanidinium containing polymers, where analogous polymers with primary amines either were inactive or represented high levels of hemolytic activity toward RBCs. In a recent work by Yang and co-workers, series of biodegradable polycarbonates bearing guanidinium functionalities were synthesized using metal-free organocatalytic ring-opening polymerization

(ROP).142 These polymers showed broad-spectrum activity against both Gram- negative and Gram-positive bacteria including MDR Acinetobacter baumannii (A. baumannii), Klebsiella pneumoniae (K. pneumonia), P. aeruginosa, E. coli, and methicillin-resistant S. aureus (MRSA) in both in vitro and in vivo studies.

Moreover, no evidence of resistance development was observed after 30 passages in the presence of sub-MIC levels of guanidinium-functionalized polycarbonates

(with ethylene spacer) against A. baumannii.

It is worth noting that the effect of cationic functionality might be tuned by other structural determinants such as molecular weight and amphiphilic balance, therefore, species-specific antimicrobial and hemolytic activities of cationic moieties should not be overgeneralized. As an example, the difference in the hemolytic activity of the same primary amine-containing polymers in Figure 2.7 and Figure 2.8 is due to the significant difference in the molecular weight of the polymers.

2.4.1.2 Amphiphilic balance

Amphiphilic balance is probably the linchpin of cell-type selectivity in the design of antimicrobial polymers. AMPs represent the optimal amphiphilic balance by containing the minimum level of hydrophobic segments required to

Page | 35 CHAPTER TWO

exhibit an antimicrobial effect (≥30%).100, 134 Differences in the composition of cell membrane between mammalian and bacterial cells are the basis of the selectivity of AMPs and their synthetic mimics (Figure 2.6).143, 144 While the outer leaflet of the mammalian cell membrane is mainly composed of neutral zwitterionic lipids, bacterial cells possess a negatively charged outer leaflet mainly due to the presence of acidic lipids. As a result, the interactions between positively charged

AMPs or their synthetic mimics are much weaker and less favourable with mammalian cells in comparison with bacterial species. Nonetheless, in the presence of excessive hydrophobic components, membrane disruption would happen even in the case of mammalian cells. With regard to the design of antimicrobial polymers, increasing the hydrophobicity can improve the antimicrobial activity, however, overuse of hydrophobic groups in the structure can significantly increase the non-specific membrane disruption. On the other hand, low hydrophobicity can lead to a loss in bactericidal activity. Amphiphilic balance is controlled by various physiochemical parameters; however, the most common approaches to optimise it are tuning the relative contribution of hydrophobic and cationic moieties and tailoring the structure of hydrophobic moieties (i.e., hydrophobicity).143, 145, 146

The effect of amphiphilic balance on the biological properties of biodegradable polycarbonate was investigated by Chin et al.146 A series of polycarbonates were prepared by organocatalytic ROP of functionalized cyclic carbonate monomers, followed by a quaternization step to yield QA moieties with different pendant groups (Figure 2.9). A thorough investigation into the biological performance of the synthesized polymers clearly showed that the hydrophobic character of the polymers can profoundly affect the antimicrobial and hemolytic activities. The

Page | 36 CHAPTER TWO

homopolymer with single QA center (N,N-dimethylbutylammonium) demonstrated potent antimicrobial activity against MRSA, vancomycin-resistant

Enterococcus (VRE), carbapenem-resistant A. baumannii and fluconazole-resistant

Cryptococcus neoformans with MIC values of 7.8, 3.9, 62.5 and 31.3 µg mL−1, respectively. This polymer also demonstrated high hemocompatibility with the

HC50 value of >4000 µg mL−1, suggesting high selectivity toward bacteria over

RBCs. Interestingly, polymers containing more hydrophobic pendant groups such as hexyl, octyl, cyclohexyl, etc. demonstrated higher toxicity toward RBCs and did not necessarily show higher antimicrobial activity.

Figure 2.9. Chemical structure of cationic amphiphilic polycarbonates bearing various QA functionalities (top). Antimicrobial and hemolytic activities of the polymer comprising N,N- dimethylbutylammonium QA group.146

Although control over amphiphilicity is necessary in the design of antimicrobial polymers, the presence of hydrophobic side chains is not

Page | 37 CHAPTER TWO

mandatory.147 Many recent reports suggest that optimal amphiphilic balance is achievable even in systems with no distinct hydrophobic pendant groups.148-151

Polymer backbone can be an important contributor to the overall hydrophobicity and thus can be tailored to adjust this pivotal requirement.

2.4.1.3 Hydrophilic Monomers

Until recently, only cationic and hydrophobic functionalities were used in the structure of antimicrobial polymers; however, it is now well understood that hydrophilic moieties are also a crucial part of antimicrobial polymer structure.

Such moieties are also present in the structure of AMPs (e.g., amino acids such as serine, threonine, tryptophan and glycine).110 The incorporation of hydrophilic moieties into the structure of polymer is an effective way to control the amphiphilic balance and reduce toxicity toward mammalian cells.152-156 These moieties are in essence neutral hydrophilic groups such as hydroxyl groups, zwitterionic moieties, sugar molecules or short polyethylene glycol (PEG) chains.

Grafted PEG oligomers, when used as hydrophilic moiety, can preserve the efficacy of antimicrobial polymers in biological medium by inhibiting undesirable protein complexation.157, 158 In 2017, Nguyen et al. reported the development of ternary antimicrobial polymers which formed single-chain polymeric nanoparticles (SCPNs) in aqueous media due to the self-folding property of the chains (Figure 2.10 a).158 These linear random copolymers were highly effective against various Gram-negative bacteria including E. coli, P. aeruginosa and Vibrio cholerae and at the same time demonstrated excellent biocompatibility when tested against RBCs and rat H4IIE liver cells. The impact of the incorporation of hydrophilic side chains on the biological performance of

Page | 38 CHAPTER TWO

the synthesized polymers was assessed by comparing the efficacy of a representative ternary copolymer and its PEG-less counterpart. While, the ternary copolymer synthesized from tert-butyl (2-acrylamidoethyl) carbamate

(Dab), 2-phenylethyl acrylate (F) and oligoethylene glycol methyl ether acrylate

(OEGA) monomers, denoted as PDab-F, demonstrated strong antimicrobial activity and high mammalian cell biocompatibility, its PEG-less counterpart, denoted as

PC2, was inactive against P. aeruginosa and E. coli and showed high toxicity toward

H4IIE cells. The authors attributed this to the formation of polymer-protein complexes (PPCs) in the bacterial cell culture media which was used in the experiments (i.e., MHB). This was further confirmed by dynamic light scattering

(DLS) analysis, where the peak associated with PPCs formation was only observed for the PEG-less polymer in MHB media (Figure 2.10 b). Ternary copolymers were also tested for PPC formation in Dulbecco’s modified Eagle medium (DMEM) supplemented by 10% fetal bovine serum (FBS), which is a frequently used cell culture medium. Similarly, no evidence of PPC formation was observed in this medium.

Page | 39 CHAPTER TWO

Figure 2.10. Illustration of the self-assembly of the amphiphilic ternary random copolymers of different structures into single-chain polymeric nanoparticles in aqueous media (a). DLS volume distributions of PDab‐F and PDab‐Pyr and PC2 (PEG-less version of PDab‐F) in water and

MHB.158

2.4.1.4 Molecular weight

Considering the mode of action of antimicrobial polymers which requires physical contact with bacterial membrane, changing the molecular weight could be an effective way to modulate the antimicrobial activity. Although it seems reasonable to speculate that increasing the molecular weight might potentiate the electrostatic interactions and membrane disruption, decline in antimicrobial activity is also probable due to adverse effects like reduced solubility or membrane diffusion issues.130, 159 Besides, high molecular weight polymers usually show an increased hemolytic activity compared to their low molecular weight counterparts. In general, low molecular weight polymers, inspired by short chain AMPs with 10-50 amino acid residues,134, 160 are more favourable in terms of antimicrobial activity and selectivity.161, 162 Series of facially amphiphilic oxanorbornene polymers were synthesized and systematically studied for the

Page | 40 CHAPTER TWO

effect of structural parameters on the biological performance by Tew and co- workers.161 The authors developed a versatile way for the synthesis of oxanorbornene-derived monomers and subsequently used ring-opening metathesis polymerization to prepare antimicrobial polymers (Figure 2.11 a). To investigate the impact of molecular weight on the antimicrobial and hemolytic activities, two series of polymers were synthesized with molecular weights of

3000 and 10000 g mol−1. The best antimicrobial performance (i.e., highest selectivity) was observed for the polymer with ethyl hydrophobic group and molecular weight of 3000 g mol-1, denoted as Ethyl_3K, which was 28 times more selective toward E. coli and S. aureus over RBCs (Figure 2.11 b and c). In general, increasing the molecular weight of synthesized polymers to 10,000 g mol-1 led to the reduction and loss of activity against E. coli and S. aureus, respectively. The loss of activity against S. aureus was attributed to the specific structure of the cell membrane in Gram-positive bacteria, where the thick murein layer inhibits the high molecular weight polymers from reaching the plasma membrane. To further investigate the effect of molecular weight on the antimicrobial performance, various oligomers were prepared using propyl monomers and tested. The individual monomer was also tested to see if a minimum chain length is required for antimicrobial activity. Interestingly, the individual monomer was inactive toward both tested bacteria, which further confirms the pivotal role of amphiphilic polymeric structure for antimicrobial activity. As expected, short oligomers were nonhemolytic, however, by further increasing the molecular weight to 3000 and 10000 g mol-1, an approximately 10-fold reduction in HC50 values was observed (Figure 2.11 d). Increasing the molecular weight of oligomers clearly enhanced the antimicrobial activity against E. coli, while an

Page | 41 CHAPTER TWO

opposite effect was observed against S. aureus. This observation was in line with the assumption on the barrier effect of the membrane in Gram-positive bacteria for high molecular weight polymers.

Figure 2.11. Synthesis of facially amphiphilic polymers via ring-opening metathesis polymerization of oxanorbornene-derived monomers (a). Antimicrobial (MIC90, µg mL−1) and hemolytic (HC50, µg mL−1 ) activities of 3000 g mol-1 series (b) and 10,000 g mol-1 series

(c). Antimicrobial (MIC90, µg mL−1) and hemolytic (HC50, µg mL−1) activities of the oligomers and polymers synthesised using the monomer with propyl alkyl group (d).161

2.4.1.5 Spatial arrangement of cationic and hydrophobic moieties

The position of cationic and hydrophobic functionalities in the polymer chain is another key factor in the design of antimicrobial polymers. In AMPs, cationic and hydrophobic moieties reside in separate residues (different center), whereas for synthetic mimics, other arrangements are also possible (Figure 2.12).110

Bifunctional monomers have been widely used in the synthesis of antimicrobial polymers. In these monomers, cationic and hydrophobic moieties can either be

Page | 42 CHAPTER TWO

connected to each other directly (same center) or be spatially separated (facially amphiphilic). The arrangement of active functionalities can profoundly affect the final conformation of antimicrobial polymers in aqueous media and subsequently mediate their interactions with the bacterial or mammalian cells membrane. The impact of different spatial arrangement of functionalities in antimicrobial polymers on the biological performance has been thoroughly studied by Tew, Kuroda, and other researchers.163-165 Although promising biological performance has been observed for each of the above mentioned arrangements, the overall findings suggest that the same center and facially amphiphilic arrangements might lead to higher selectivity, albeit at the expense of compromising the antimicrobial activity. In the case of different center design, the probable segregation of hydrophobic segments of the polymer chain can increase unfavourable non-specific membrane disruption.

Figure 2.12. Three spatial arrangements for cationic and hydrophobic moieties in the structure of synthetic antimicrobial polymers.110

2.4.1.6 Alkyl spacer/tail/side chain length

Another design parameter which modulate the antimicrobial and hemolytic activities of synthetic antimicrobial polymers is the length of alkyl spacer/tail/side chain. These small hydrophobic segments either connect the cationic moiety to the polymer backbone (cationic spacer arm), exist as pendent groups which are attached to the positive charge center (alkyl tail) or constitute Page | 43 CHAPTER TWO

the hydrophobic monomer (side chain). Variations in the length or structure of these groups can directly affect polymer conformation, hydrophobicity and amphiphilic balance.110, 130, 133 The optimum alkyl length might vary based on other characteristics of the system, however, C4 – C6 seem to be the most reported optimum length in the literature.146, 163, 166, 167 A thorough investigation on the impact of cationic spacer arms on the antimicrobial activity was reported by

Palermo et al.166 To this end, various methacrylate random copolymers were synthesized either as homopolymers using cationic monomers with different alkyl spacers or in the form copolymers along with ethyl or butyl methacrylate as hydrophobic monomers. The preliminary results showed a stark difference between the performance of tested polymers, with selectivity indexes in the range of 0.23 to 62. Based on the reported results, polymers containing 6- aminobutyl methacrylate had the lowest MIC value against both E. coli and S. aureus, however, they were highly hemolytic. Meanwhile, polymers based on 2- aminobutyl methacrylate showed the opposite behaviour by having the highest

MIC and HC50 values. The best performing polymer consisted of 4-aminobutyl methacrylate and ethyl methacrylate (mole fraction of hydrophobic repeating units = 0.2-0.3), which demonstrated high antimicrobial activity while maintaining its hemocompatibility. This polymer demonstrated MIC values of

21 and 63 µg mL−1 against E. coli and S. aureus, respectively, and the HC50 value of 1300 µg mL−1. Molecular dynamics simulations revealed that alkyl spacer length can affect polymer conformation upon contact with bacterial membrane.

While the polymers with longer spacer lengths (butyl and hexyl) could effectively diffuse into membrane bilayer, the polymer containing ethyl spacer only interact with the surface of the bilayer.

Page | 44 CHAPTER TWO

2.4.1.7 End groups

The effect of end groups on the biological performance of antimicrobial polymers is known to be mostly associated with the amphiphilic balance.

Therefore, this effect is more sensible for low molecular weight polymers or polymer with a large number of end groups such as dendrimers or star polymers.168, 169 In these cases, incorporation of hydrophobic end groups into the structure can increase antimicrobial activity and probably hemolytic activity.

Griesser et al. studied the impact of RAFT end-groups on the overall performance of a series of well-defined cationic methacrylate polymers.168 They systematically changed the R- and Z-RAFT end-groups in relatively short polymers (i.e., DP =

24) containing either primary amine or guanidine groups (Figure 2.13). The polymers were then assessed in terms of antimicrobial and hemolytic activities against a wide range of clinically relevant pathogens and RBCs, respectively.

Results from these tests demonstrated that the RAFT R-groups was mostly involved in controlling the hemolytic level of the polymers, while, the Z-groups appeared to mediate the antimicrobial activity of the polymers.

Page | 45 CHAPTER TWO

Figure 2.13. Chemical structure of random methacrylate copolymers bearing either primary amine or guanidine moieties as cationic functionalities with different RAFT R- and Z- groups.168

The end group effect is also significant for telechelic polymers, where the -α and -ω end-groups are responsible for the antimicrobial activity of a .170-172

2.4.1.8 Macromolecular architectures and nanostructures

Although the majority of research in the field of antimicrobial polymers have focused upon discovery and development of polymers in the form of linear random copolymers, polymers with complex architectures have gained considerable attention in recent years.110, 111, 130 Facile synthesis of polymers with precise control over architecture and composition, using advanced polymerization techniques, have enabled the production of antimicrobial polymers with various kinds of architectural designs.127, 173 Major changes in the biological activity of antimicrobial polymers have been observed upon changing the polymer architecture.

Page | 46 CHAPTER TWO

The effect of segmentation of cationic and hydrophobic functionalities on the biological activity of antimicrobial polymers was studied by Perrier and co- workers in 2017.174 A small library of statistical, diblock, and multiblock copolymers was prepared via RAFT polymerization using N- isopropylacrylamide and 2-aminoethyl acrylamide as the hydrophobic and cationic monomer, respectively (Figure 2.14). The resulting polymers were thoroughly studied for antimicrobial activity and biocompatibility and their overall performance was assessed based on the selectivity. In general, diblock and multiblock copolymers showed better performance in comparison with statistical copolymers. Moreover, at low content of cationic comonomer, the multiblock outperformed its statistical and diblock counterparts especially against P. aeruginosa and Staphylococcus epidermidis (S. epidermidis). These observations were attributed to the increased hydrophobicity of diblock and multiblock copolymers, as assessed via HPLC, and also to the improved interactions between cationic moieties and bacterial membrane upon charge segregation in the block copolymers.

Page | 47 CHAPTER TWO

Figure 2.14. Schematic illustration of the structure, composition and segmentation of monomers within the polymer chains of the poly acrylamides synthesized via RAFT polymerization.174

In another study, Judzewitsch et al. investigated the effects of monomer distribution within linear antimicrobial polymers comprising aminoethyl, phenylethyl, and hydroxyethyl acrylamides.175 A library of well-defined copolymers was synthesized via photoinduced electron transfer–reversible addition–fragmentation chain transfer polymerization (PET-RAFT) and screened for biological performance. It was observed that antimicrobial and hemolytic activities were greatly influenced by the sequence and composition of blocks within the polymer chain. While separating hydrophobic monomers into a distinct block led to the loss of antimicrobial activity, polymers with localized cationic segments and amphiphilic segments containing both hydrophobic and hydrophilic monomers demonstrated potent antimicrobial activity.

Additionally, genus specificity was observed for some of the designed polymers, which further confirms the potential of block copolymer design for possible clinical applications.

Page | 48 CHAPTER TWO

Changing the monomer distribution from random to segregated blocks might induce the self-assembly of antimicrobial polymer into a core-shell micelle morphology.130 Micellization has shown to impact the antimicrobial activity of the polymers in two opposite ways.111 Micellization can pose a detrimental effect on the biological activity of block copolymers by limiting the mobility of polymer chains and minimising the integration of hydrophobic segments into the membrane lipid bilayer. On the other hand, some studies have shown the positive impact of micellization on antimicrobial performance due to the increased local concentration of effective functionalities. This positive role of micellization on antimicrobial performance is most likely to happen when the micelle structure can undergo disassembly upon contact with the bacterial membrane.

Nederberg et al. reported the synthesis of biodegradable antimicrobial micelles based on cationic polycarbonates.176 Amphiphilic triblock polycarbonates with a well-defined structure and narrow molecular weight distribution were prepared via metal-free organocatalytic ROP, followed by a quaternization step using trimethylamine. The resulting polymers simultaneously underwent self- assembly in aqueous media to form cationic micellar nanoparticles (Figure 2.15).

These cationic micelles demonstrated remarkable efficacy against the growth of selected Gram-positive bacteria including MRSA strain and also against

Cryptococcus neoformans. It is worthwhile noting that the MICs of the synthesised polymers were above their CMC (critical micelle concentration) values in the buffer used for growing the bacteria, indicating that antimicrobial action was induced by polymeric micelles and not individual polymers. The authors postulated that micellization could potentiate the antimicrobial activity by

Page | 49 CHAPTER TWO

increasing the local concentration of polymers and therefore positive charges, which led to strong electrostatic interactions between nanoparticles and microbial membrane. Meanwhile, the micelles exhibited low hemolytic activity

(<15% hemolysis at 500 µg mL−1) and did not cause significant toxicity even at concentrations higher than MIC when tested in vivo using a mice model.

Figure 2.15. Illustration of the chemical structure of cationic amphiphilic polycarbonate and its self-assembly into micelle structure in aqueous media.176

Branched antimicrobial polymers in the forms of dendrimer, star, hyperbranched or brush polymers have also gained increased attention recently.144 The higher density of active functionalities and end groups endows these polymers with inherent multivalent property which can facilitate the interactions between the polymer and anionic microbial membrane.130 Moreover, branched antimicrobial polymers usually exhibit less hemolytic activity

Page | 50 CHAPTER TWO

compared with their linear counterparts due to the reduction in non-specific interactions. The other attractive feature of branched antimicrobial polymers is the possibility of loading cargo molecules or therapeutic agents into the nanostructures for drug delivery purposes.111, 144

Allcock and co-workers reported the synthesis of star- and comb-shaped molecular brushes comprising poly[2-(dimethylamino)ethyl methacrylate] via atom transfer radical polymerization (ATRP).177 The resulting polymers were subsequently quaternized using different alkyl iodide compounds to investigate the effect of side chain hydrophobicity on the antimicrobial activity. When assessed for their antimicrobial activity against E. coli, star-shaped brushes quaternized with iodoheptane had the lowest MIC value (250 µg mL−1) compared to other polymers tested. Antimicrobial activity was also observed for the quaternized star-shaped brush in the form of fibres prepared by electrospinning.

The lower antimicrobial activity of comb-shaped brushes was attributed to the limited interactions of these nanoparticle and bacterial membrane due to excessive and unfavourable positive charge density.

Using an arm-first approach, Wong et al. developed a series of functionalized star polymers with promising antimicrobial activity against Gram-positive bacteria, including S. aureus, MRSA, Enterococcus faecalis, and VRE.178 These star polymers were composed of a mixture of poly(L-lysine) (PLL) and polyglucosamine (PGSA) arms (Figure 2.16). While PLL was selected as the bactericidal component, PGSA was incorporated to endow the polymer with biocompatibility. Moreover, PGSA was expected to increase the penetration of the star polymers through the bacterial cell membrane due to its similarity to the

Page | 51 CHAPTER TWO

structure of peptidoglycan component of the bacterial cell membrane. Star polymers with different molar ratios of arms were synthesized and compared to individual arms and also AMPs such as melittin and polymyxin B in terms of their biological performance. While all synthesized star polymers were active against Gram-positive bacteria, the star polymer with 25% GSA and 75% PLL content (S-GSA 25) exhibited the best activity. In terms of biocompatibility, all the stars were non-hemolytic up to 1 mg mL-1 and demonstrated low toxicity toward human aortic smooth muscle cells (AoSMC). The authors attributed the excellent performance of S-GSA 25 to the star architecture of the polymer, as the physical mixture of PLL and PGSA arms did not show the same level of biocompatibility.

Figure 2.16. Schematic illustration of the chemical structure of glucosamine-functionalised star polymer.178

2.4.2 Mechanism of action of synthetic antimicrobial polymers

The mechanism behind the antimicrobial activity of synthetic antimicrobial polymers resemble the mechanism of their natural counterparts (i.e., AMPs).

Membrane disruption through physical contact with the bacterial cell is the most recognized mode of action for AMPs and antimicrobial polymers; however, other mechanisms might be involved simultaneously.100, 130, 179 This bactericidal action

Page | 52 CHAPTER TWO

is triggered by the electrostatic interactions between cationic moieties of AMPs or polymer and anionic lipid heads of the outer leaflet of the bacterial membrane, which allows them to attach to the bacterial membrane. Upon physical contact, the membrane integrity is disrupted through the diffusion of the hydrophobic segments into the phospholipid bilayers. Several models have been proposed for this step of mechanism including toroidal, barrel-stave and carpet models

(Figure 2.17).180 Membrane permeabilization is followed by the leakage of cytoplasmic components, and finally, cell death. The major differences in the design of membrane in mammalian and bacterial cells form the basis of the selectivity of AMPs and their synthetic mimics toward the bacterial cells (Figure

2.6).130 The presence of negatively charged components in the structure of the bacterial membrane strengthen the electrostatic interactions with polycationic peptide or polymer. In contrast to small molecule antibiotics which act on specific intracellular targets, the bactericidal effect of AMPs and their polymeric mimics is mainly mediated by nonspecific actions on the cell membrane. Thus, the risk of resistance development against these compounds is significantly reduced.100

Figure 2.17. Proposed models for the membrane-disrupting action of antimicrobial peptides.180

Page | 53 CHAPTER TWO

2.4.3 Building well-defined polymers via reversible addition-fragmentation

chain transfer (RAFT) polymerization

Synthetic polymers have proven useful in the biomedical fields over the past few decades.127, 181 They have been rationally implemented in applications such as targeted drug/gene delivery, tissue engineering, coatings and recently in antimicrobial applications. While each specific application requires a unique polymer design in macroscopic, microscopic and nanoscale levels, another key factor to consider is the uniformity in the structural properties of the synthesized polymer, both within a polymer chain and between individual chains.182 The superior performance of natural biopolymers such as polypeptides and polynucleotides is partially related to the high degree of uniformity in their structure even at nanoscale levels. Thus, along with tailoring the polymer structure, special attention should be given to the consistency of structural properties such as molecular weight, chain architecture, composition and site- specific functionality of the synthesized polymers. This would also enable the systematic optimisation of the polymeric therapeutic platforms as the effect of each structural parameter can be solely detected and customized.

Reversible deactivation radical polymerization (RDRP) is a powerful technique in polymer synthesis, which offers great control over the structural features of the polymers as oppose to free radical polymerization (FRP) technique.183 The main difference in the mechanism of polymerization between these two techniques is associated with the propagation step. In FRP, propagating radicals are irreversibly terminated via either disproportionation reactions or radical coupling, whereas in RDRP, propagating radical species are reversibly activated and deactivated. The reversible termination or chain transfer

Page | 54 CHAPTER TWO

steps in RDRP minimises the undesirable termination of the propagating radicals and regulates the concentration of propagating radical species throughout the polymerization. Furthermore, polymers synthesized via RDRP retain their living characteristic, which allows them to undergo further chain propagation upon addition of extra monomers. Among several prominent RDRP techniques, nitroxide-mediated polymerization (NMP), ATRP, and RAFT polymerization have been widely studied and used in literature.

RAFT polymerization was first introduced by Moad, Rizzardo and Thang in

1998 and since then have attracted increasing attention.184 RAFT polymerization is probably the most versatile RDRP technique due to the following advantages:

(1) tolerant of a wide range of monomers and functionalities, (2) compatible with a broad range of solvents (aqueous, polar/nonpolar organic solvents and ionic), reaction temperatures, initiation processes (thermal, redox and photoinitiated) and polymerization methods (solution, suspension, emulsion and dispersion) (3) requires comparatively inexpensive and simple experimental setup, (4) offers great control over the molecular weight and the molecular weight distribution,

(5) capable of synthesising various architectures including block, gradient, hyperbranched, stars, brushes and graft polymers, and (6) feasible to be performed from various surfaces.

To switch the polymerization process from FRP to RAFT, a suitable chain transfer agent which is typically a thiocarbonylthio compound is added to the system.185, 186 This chain transfer agent with the general formula “RSC(Z)=S” is referred to as RAFT agent. The selection of appropriate RAFT agent is crucial to achieve well-controlled polymerization and consequently well-defined

Page | 55 CHAPTER TWO

polymers.187 In the same manner as FRP, decomposition of a radical initiator is the first step in the RAFT process (Figure 2.18). The resulting radicals subsequently react with monomers to generate propagating radicals (Pn •). In the next step, a Pn • adds to the RAFT agent (1) to form an intermediate species (2).

Upon fragmentation of an intermediate radical, a dormant macroRAFT agent (3) and a R-group derived radical (4) are generated, which the latter resumes monomer propagation and forms a new propagating radical (Pm •). Repeated cycles of formation and fragmentation of the intermediate species lead to an equilibrium between the dormant macroRAFTs (3 and 6) and propagating radicals (Pn •and Pm •). RAFT exchange equilibrium facilitates the uniform chain growth and therefore leads to a narrow molecular weight distribution. Although less likely to occur in comparison with FRP, termination might happen through the reaction between propagating radicals or intermediate species.

Figure 2.18. Mechanism of RAFT polymerization.188

Page | 56 CHAPTER TWO

RAFT polymerization is particularly very practical for the synthesis of antimicrobial polymers, as it allows for the synthesis of well-defined polymers with predetermined molecular weights and narrow polydispersities (to imitate

AMPs). Furthermore, polymers with antimicrobial pendant moieties or end- groups are readily synthesized via RAFT process, since the technique is highly compatible with variety of monomers and functionalities.132, 158, 189 Polymers with diverse and complex architectures that are of interest for antimicrobial applications, such as block copolymers, hyperbranched, dendritic and star structures can be achieved by RAFT polymerization.175, 190 Finally, this technique is highly appropriate for the preparation of antimicrobial surfaces as in situ polymerization can be performed on various substrates.191-193 Considering the mentioned advantages of RAFT polymerization, this technique was used for the synthesis of all the polymers that were used in this thesis.

2.5 Combination therapy

Considering the slow progress in new antibiotic discovery and the emerging threat of antibiotic resistance worldwide, the development of novel antimicrobial strategies involving the use of currently available antibiotics via combination therapy could fast track these developments into clinical settings.73 In addition, the application of such combination therapy may enable the resensitization and potentiation of conventional antibiotics that are no longer effective against MDR strains.72, 73, 194

Combination therapy can be simply defined as the co-administration of an antibiotic along with other antimicrobial or adjuvant agents. Although achieving

Page | 57 CHAPTER TWO

the right and effective combinations against MDR strains requires a great deal of effort, the potential advantages of combination therapy over monotherapy are significant.74, 195, 196 These advantages include: (1) broader spectrum antimicrobial activity via multiple cellular targets (empirical therapy), (2) greater potential for narrow-spectrum activity via species-specific and strain-specific interactions, (3) lower resistance development, (4) revival of susceptibility to treatment in MDR bacteria, and (5) improved treatment efficiency with lower treatment dose and subsequent side effects.

Depending on the nature of involved compounds, combinations can be classified into three major categories: (1) congruous combinations, where each compound has independent antimicrobial activity, (2) syncretic combinations, where one of the compounds has no antimicrobial activity (i.e., adjuvant), but it can potentiate the activity of the other component, and (3) coalism, where components demonstrate antimicrobial activity only in the presence of each other.73 Congruous combinations based on two or more antibiotics are the most widely studied and used combinations. With respect to the modes of action, contributing compounds in combination therapy might have different pathways, same pathway but different targets or the same pathway and target.72

Dictated by the modes of action and therefore, the interactions between compounds, combination therapy might lead to synergistic, addition or antagonistic results. In synergistic interactions, the combination exhibits stronger antimicrobial activity than the sum of individual components. An additive interaction (also known as indifference) makes no changes in the efficiency of the treatment. Finally, an antagonistic interaction leads to a total antimicrobial

Page | 58 CHAPTER TWO

activity which is less than the sum of the individual agents. A recent comprehensive study by Typas and co-workers suggests that additive and antagonistic interactions are more prevalent than synergistic interactions.194 Their results showed that synergism is more likely to happen between antibiotics which belong to the same class or those which target the same cellular process in bacteria. An interesting finding in this work was that membrane-targeting compounds were involved in a significant number of synergistic combinations.194

A possible explanation for this observation might be the increase in the intracellular concentration of the coagent in the presence of a membrane- targeting compound. They observed the highest synergistic interactions between colistin, a member of polymyxin class of antibiotics with a membrane disruption mechanism, and members of the macrolides class.

Combination therapy is not only an effective way to extend the efficient lifetime of approved antibiotics, but also a fast and cheap platform to develop novel therapeutic regimens. Moreover, considering the poor activity of available antibiotics against bacterial biofilm, co-administration of antibiotics with an antibiofilm agent might lead to robust antibiofilm activity. To develop more potent antimicrobial platforms based on combination therapy, we might need to think outside the box of antibiotics. Given the promising role of membrane- targeting compounds in synergistic combinations,194 synthetic antimicrobial polymers could be potential coagents in combination therapy. The remainder of this review provides an overview of antimicrobial strategies that entail the use of synthetic antimicrobial polymers in combination with other antimicrobial agents including nitric oxide, antibiotics, natural antimicrobial compounds and nanomaterials to combat antibiotic-resistant and biofilm-related infections.

Page | 59 CHAPTER TWO

2.5.1 Antimicrobial polymers in combination with nitric oxide

Nitric oxide (NO) is a gaseous molecule with potent bactericidal and antibiofilm properties in a dose-dependent manner.37 At high concentrations, NO can demonstrate bactericidal activity by inducing extracellular nitrosative stress and intracellular oxidative stress (Figure 2.19).116, 197 At low concentrations, whether produced endogenously or as an exogenous agent, NO can induce the dispersal of bacterial biofilms.37, 198 In both modes of action, there is a very low risk of resistance development in bacteria upon treatment with NO.116, 117

Figure 2.19. Proposed mechanisms for bactericidal action of nitric oxide through nitrosative and oxidative stress.197

Despite its promising antibacterial activity, clinical applications of NO have been greatly affected by its very short half-life in biological environments.

Therefore, several small-molecule NO donors such as N-diazeniumdiolates

(NONOates) and S-nitrosothiols (RSNO) have been developed.199 However, these NO donors still lack the localized and controlled release profile which is

Page | 60 CHAPTER TWO

favourable for biological applications. To address this issue, a wide range of delivery and encapsulation approaches based on polymeric and non-polymeric scaffolds have been developed.200, 201 Polymer-based systems in the form of micelles, liposomes, hydrogels and branched polymers have shown great potential in increasing the antimicrobial efficacy of NO.37 Incorporation of synthetic antimicrobial polymers into NO encapsulation or delivery platforms could further increase the potency of these systems due to the probable synergistic or additive interactions.

Schoenfisch and co-workers have focused on using hyperbranched polymers and dendrimers as scaffolds for NO delivery and release. In the majority of these studies, the structure of polymeric scaffold is modified by alkyl chains, QA moieties, PEG chains, etc., prior to the incorporation of NONOates moieties using high pressure NO gas at alkaline condition. Although the chemical structure of the used polymeric scaffolds in some of these studies might not reflect the ideal characteristics of antimicrobial polymers, the inherent cationic nature of these polymers (due to the high density of amine/amide functionalities) along with the modification of the polymeric scaffolds with alkyl or QA groups endow these polymers with antimicrobial activity to some extent.

In 2012, Sun et al. reported the synthesis of NO-releasing poly(propylene imine) (PPI) dendrimers as potential antimicrobial agents.202 They post-modified the primary amine-functionalized dendrimers with propylene oxide (PO), styrene oxide (SO) or poly(ethylene glycol) methyl ether acrylate (PEG) to obtain secondary amine-functionalized dendrimers and subsequently used high pressure NO gas to incorporate NONOate moieties to the structure of

Page | 61 CHAPTER TWO

dendrimers. Antimicrobial activity was observed to be influenced by dendrimer size (generation) and exterior functionalities. When tested against P. aeruginosa and S. aureus, dendrimers of higher generation or those bearing hydrophobic exterior functionality (e.g., SO) demonstrated higher potency compared to other dendrimers. Larger dendrimers possessed a high capacity for NO loading and showed significantly greater NO flux and NO release in comparison to their smaller counterparts. TheNO-releasing dendrimers showed superior bactericidal activity against Gram-negative and Gram-positive bacteria compared dendrimers without NO. Moreover, incorporation of NONOate moieties reduced the toxicity of dendrimers toward L929 mouse fibroblast cells. As a result, NO-releasing dendrimers offered higher therapeutic efficiency by showing improved antimicrobial activity and biocompatibility in comparison to the individual components, which might be due to the synergistic or additive effects of NO and antimicrobial scaffold.

In another study, Kim and co-workers conjugated Pluronic® F68, a thermosensitive polymer with the chemical formula of (PEO)78-(PPO)30-(PEO)78, to a branched polyethylenimine (BPEI) and subsequently incorporated

NONOate moieties to the structure.203 F68 is a water-soluble, biocompatible polymer which has been approved by Food and Drug Administration (FDA) and is mostly used as a non-ionic surfactant in cell culture. In that study, however,

F68 was used as a polymeric scaffold for NO-releasing BPEI (F68-BPEI-

NONOate). The NO release profile, obtained using a chemiluminescence NO analyser, revealed an initial fast burst release with the half-life of 29.75 min for

F68-BPEI-NONOate. Evaluation of the bactericidal activities of NO-releasing polymer and its non-NO-releasing analogue showed that the incorporation of

Page | 62 CHAPTER TWO

NO could significantly improve the efficacy of F68-BPEI against E. coli. However,

F68-BPEI and F68-BPEI-NONOate exhibited similar bactericidal activities against S. aureus. The authors attributed this observation to the differences in the structure of the cell membrane in Gram-negative and Gram-positive bacteria.

While the F68-BPEI could efficiently eradicate S. aureus through membrane disruption, it was less effective against the lipid bilayer of P. aeruginosa. Notably,

F68-BPEI-NONOate had the same bactericidal efficiency against MRSA as observed against wild type S. aureus with minimum bactericidal concentration

(MBC) value of 3.32 mg mL−1. In addition, F68-BPEI-NONOate exhibited lower toxicity toward NIH/3T3 cells compared to F68-BPEI probably due to the negative surface charge of F68-BPEI-NONOate which reduced the interactions with mammalian cells.203

Worley et al. developed a dual-action antimicrobial agent by incorporating both NONOate and QA functionalities into the structure of poly(amidoamine)

(PAMAM) dendrimers (Figure 2.20).204 They investigated the effect of dendrimer generation and QA alkyl chain length on bactericidal activity of NO-releasing and control dendrimers. Regardless of the dendrimer generation, increase in the length of the alkyl tail led to an increase in bactericidal activity against both P. aeruginosa and S. aureus. Moreover, a generation dependency in antimicrobial activity was observed for all synthesized dendrimers, where G4 dendrimers represented higher activity in comparison to their G1 counterparts. Comparison of the bactericidal activities of NO-releasing and control dendrimers showed that the positive effect of adding NO is only limited to dendrimers with shorter alkyl tails. The authors postulated that the intracellular concentration of NO could not reach the required level to trigger bactericidal action due to the extensive

Page | 63 CHAPTER TWO

membrane damage caused by dendrimers with long alkyl tails (octyl and dodecyl).204 All dual-action dendrimers showed high biocompatibility when incubated with L929 mouse fibroblast cells at concentrations equal to their MBC values for 4 h. Notably, incorporation of NO not only increased the antimicrobial activity of QA-modified dendrimers (with short alkyl chains) but also improved their biocompatibility in comparison with unmodified and QA-modified dendrimers.

Figure 2.20. Synthesis of dual-action dendrimers comprising quaternary ammonium moieties and N-diazeniumdiolate NO donors.204

In a follow-up study, the same group introduced another version of dual- action NO-releasing dendrimers. The primary amine-containing PAMAM dendrimers were modified with short to medium alkyl chains via ring-opening reaction with epoxides (Figure 2.21).205 Subsequently, NONOate moieties were incorporated via the secondary amine functionalities. NO-loaded dendrimers demonstrated an initial burst release in PBS which was followed by a steady reduction in the NO release for up to 10 h. The bactericidal activity of control and

NO-releasing dendrimers of different generations (G1 to G4) were then assessed against planktonic and biofilm bacteria. Both control and NO-releasing dendrimers represented higher bactericidal activity against P. aeruginosa compared to S. aureus and MRSA planktonic or biofilm cells. Similar to their previous study,204 the addition of NO donor moieties only improved the

Page | 64 CHAPTER TWO

bactericidal action of the dendrimers that were modified with short alkyl chains.

This was confirmed by confocal microscopy where hexyl-modified dendrimers notably damaged the membrane of bacteria in P. aeruginosa biofilm within 1 h treatment, which led to low intracellular NO content. In contrast, a significant amount of intracellular NO was observed upon 1 h incubation with butyl- modified dendrimers. In terms of antibiofilm activity, hexyl-modified dendrimers demonstrated higher potency against all tested strains, possibly due to the ability of these polymers to effectively penetrate into biofilm matrix as supported by confocal microscopy images. In general, G3 hexyl-modified dendrimers (with or without NO) were the best performing compounds by exhibiting potent activity against both planktonic and biofilm at concentrations

~2-4 times lower than their IC50 values, defined as the concentration at which the cell viability is reduced by half, towards L929 mouse fibroblasts cells.

Figure 2.21. Illustration of the chemical structure of dual-action nitric oxide-releasing alkyl- modified dendrimers (left). Confocal microscopy images showing the relative penetration of two representative dendrimers (G4 butyl and G4 hexyl) into S. aureus biofilm upon 1 h incubation at 50 µg mL−1(right).205

Using a different polymer architecture, Yang et al. reported the synthesis of

NO-releasing hyperbranched polyamidoamines (h-PAMAM) as potential Page | 65 CHAPTER TWO

antimicrobial agents against oral infections.206 In comparison with the defect-free structure of dendrimers, hyperbranched polymers have irregular structure with dendritic, terminal and linear units. The presence of linear units in the structure of h-PAMAM induced structural defects and increased the number of secondary amine groups. Synthesized hyperbranched polymers were initially modified with PO to convert the primary amine groups to secondary amines and subsequently were loaded with NO. The NO payload and release were found to be greatly controlled by PO modification. While modification with one molar equivalent of PO with respect to primary amines enhanced the NO payload of h-

PAMAM, excessive PO modification led to a significant reduction in NO payload as the secondary amines were also consumed. The bactericidal activities of resulting polymers were then assessed against Gram-negative periodontal and

Gram-positive cariogenic bacteria. A decline in bactericidal activity was observed for PO-modified polymers in comparison with the control polymers, probably due to the reduction in the number of primary amine moieties and consequently lower membrane disruption. Similarly, the introduction of NO to the structure of polymeric scaffolds reduced the antimicrobial activity. This observation was attributed to the negative charge of NONOate moieties which reduced the interactions of polymeric scaffold and bacterial membrane. In general, the best bactericidal activity was observed for control polymer with no

PO and NO modifications. However, excessive PO modification (at all tested concentrations, i.e., 0.1-16 mg mL−1) and NO modifications (at polymer concentrations ≤4 mg mL−1 and incubation time of 2 h) improved the biocompatibility of h-PAMAM polymers towards human gingival fibroblasts

(HGF-1). Longer incubation time of 24 h resulted in an overall reduction in cell

Page | 66 CHAPTER TWO

viability for all tested polymers, especially for NO-releasing polymers at high polymer concentrations probably due to the toxicity of NO for mammalian cells at high doses. Considering both antimicrobial activity and biocompatibility, the combination of PO and NO modifications improved the therapeutic potential of the cationic h-PAMAM. The NO-releasing h-PAMAM modified by excess amounts of PO demonstrated low toxicity (>75% cell viability) toward HGF-1 after 2 h incubation at 4 mg mL−1 which is the highest MBC value against all bacteria tested. Finally, to investigate the effect of polymer architecture on antimicrobial activity, the efficacy of NO-releasing PO-modified h-PAMAM polymer was compared to analogues of dendritic scaffold with identical unit structure, molecular weight and exterior functionalities. Despite the lower degree of regularity in the structure, NO-releasing PO-modified h-PAMAM polymer exhibited comparable antimicrobial activity and biocompatibility to the analogous dendric scaffold.

Considering the promising antimicrobial potency of NO-releasing hyperbranched polyamidoamines, the authors conducted an in vivo study against Porphyromonas gingivalis, which is a Gram-negative bacteria associated with periodontitis.207 In accord with the in vitro results, NO-releasing hyperbranched polymer demonstrated higher activity compared to control polymer in vivo. Treatment with NO-releasing PO-modified h-PAMAM polymer at 37 mg kg−1 once a day for three days resulted in full eradication of bacteria, >6 log10 reductions in the number of viable bacteria, in subcutaneous chamber model.

Page | 67 CHAPTER TWO

Extending from this concept, Locklin and co-workers developed a dual-action coating composed of zwitterionic terpolymers and NO-releasing CarboSil

(silicone-polycarbonate-urethane thermoplastic) (Figure 2.22).208 Zwitterionic polymers are mostly known and used for their antifouling properties; however, they also exhibit modest antimicrobial activity.130 In this work, zwitterionic polymer, 2-methacryloyloxyethyl phosphorylcholine-co-butyl methacrylate-co- benzophenone (BPMPC), was covalently grafted to S-nitroso-N- acetylpenicillamine (SNAP) doped CarboSil films via UV-irradiation. The presence of zwitterionic polymer in the coating reduced the SNAP leaching in physiological conditions, increase NO release flux and decrease the proteins adsorption and accumulation on the surface. These antifouling-biocide releasing coatings showed high potency against S. aureus as a result of the combined effects of NO (bactericidal effect) and zwitterionic polymer (protein and bacteria repellent effect). Dual-action coatings could effectively reduce >99.9 % of the viable bacteria which was higher than the effect of coatings which were prepared with individual BPMPC or NO-doped Carbosil.

Figure 2.22. Schematic illustration of antifouling-biocide releasing coatings prepared by covalent grafting of zwitterionic polymers to SNAP-doped CarboSil films.208

Page | 68 CHAPTER TWO

2.5.2 Antimicrobial polymers in combination with antibiotics

Synthetic antimicrobial polymers have emerged as potential candidates to be used in combination therapy formulations along with traditional antibiotics given the positive impact of membrane-targeting antimicrobial agents in antibacterial drug combinations.194. In these combinations, antibiotics might be either used in a physical mixture with the synthetic polymer, be encapsulated into the polymeric nanostructure or be conjugated to the polymer chains. The synergistic interaction between synthetic antimicrobial polymer and antibiotic not only improves the treatment efficiency but also provides an effective way to extend the lifetime of antibiotic by reviving the susceptibility of the MDR bacteria to antibiotics.

In 2008, Wang and co-workers studied the potential use of polyethylenimine

(PEI) in combination with 16 antibiotics from 10 different classes against a resistant P. aeruginosa strain.209 Considering the polycationic nature and membrane permeabilization effect of PEI, synergistic interactions with selected antibiotics were expected. Authors investigated the interactions between PEI and antibiotics using checkerboard and time killing assays. Among the tested combinations, synergistic interactions were observed for five antibiotics, including novobiocin, ceftazidime, cefotaxime, chloramphenicol, and rifampicin in checkerboard assay. Rest of the antibiotics exhibited antagonism or addition interactions with PEI. PEI at subinhibitory concentration was able to reduce the

MIC value of these antibiotics up to 40-fold. Synergistic combinations were then tested for bactericidal activity at subinhibitory concentrations of PEI and antibiotics (25% of the MICs values) using time-kill assay. Except for cefotaxime,

Page | 69 CHAPTER TWO

all antibiotics showed synergistic killing effects in combination with PEI with 5- to 8-log10 reductions in CFU compared to untreated bacteria. The observed antagonism interactions between PEI and three antibiotics, namely tobramycin, polymyxin B and vancomycin were attributed to the reduction in the uptake of the antibiotics in the presence of PEI. The strong affinity between PEI and bacterial LPS, could reduce the available cation-binding sites of LSP for these antibiotics.

In an interesting work by Yang and co-workers, a small library of cationic polycarbonates comprising vitamin E moiety and propyl or benzyl side chains was synthesized using ROP, and then quaternized with trimethylamine (Figure

2.23 a).210 Structure-activity studies showed that the incorporation of biologically active vitamin E into the structure of polymers can significantly increase the antimicrobial activity against E. coli, S. aureus and . The authors postulated that vitamin E could modulate the amphiphilic balance of the synthesized polymers, which in turn led to an improved antimicrobial activity.

In addition, the resulting polymers demonstrated relatively low hemolytic activities with the HC20 values (defined as the concentration of polymer that caused 20% RBC lysis) in the range of 500-1000 µg mL−1. However, all of the synthesized polymers exhibited poor antimicrobial activity against P. aeruginosa.

To enhance the activity against P. aeruginosa, the lead polymer of the study was used in combination with doxycycline, streptomycin and penicillin (Figure 2.23 b). The potency of these combinations was then evaluated using checkerboard assay. Although all three combinations were synergistic, the best outcome was observed for the combination based on doxycycline with FICI (fractional bactericidal/inhibitory concentration index) values of 0.11 and 0.16, which

Page | 70 CHAPTER TWO

correlated to 8- and 20-fold reductions in the MBC value of doxycycline, respectively. Preliminary mechanistic studies using confocal microscope and scanning electron microscope (SEM) images suggested that the increased membrane permeability due to the presence of cationic polymer could potentially facilitate the influx of doxycycline and led to a potent bactericidal activity.

Figure 2.23. Synthesis of cationic polycarbonates comprising vitamin E and alkyl side chains

(a). Chemical structure of doxycycline, streptomycin and penicillin-G (b).210

Synergistic combinations of a cationic conjugated polymer (CCP) and membrane targeting polypeptide antibiotics (polymyxin E and polymyxin B) were reported by Cheng and co-workers in 2017 (Figure 2.24 a and b).211

Although CCPs show higher antimicrobial activity under UV irradiation, all the experiments were conducted under dark condition in that study. Two CCPs, a polyfluorene derivative (PBF) and a polythiophene derivative (PT), were synthesized and used in combination with ampicillin, kanamycin, sulfamethizol, gentamicin, polymyxin B (PLB) and polymyxin E (PLE). Evaluation of the MIC values of individual agents and combinations revealed two synergistic

Page | 71 CHAPTER TWO

combinations involving polypeptide antibiotics and PBF. The MIC values of PLE and PLB were reduced from 15.1 and 3.7 µg mL−1 to 2.5 and 2.3 µg mL−1, respectively, in the presence of PBF at subinhibitory concentration (10 μM).

Significantly higher bactericidal efficacy of PBF-PLB combination relative to individual compounds also confirmed the synergistic interaction between PBF and PLB (Figure 2.24 c and d). At subinhibitory concentrations, both PBF and

PLE caused minimal damage to bacterial cell membrane with no obvious change in cell morphology. However, when used in combination, they exhibited strong membrane disruption activity which led to clear morphological damage as evidenced by SEM images (Figure 2.24 e).

Page | 72 CHAPTER TWO

Figure 2.24. Chemical structure of cationic conjugated polymers (a) and polypeptide antibiotics polymyxin E (PLE) and polymyxin B (PLB) (b). Bactericidal activities of PLB and the combination of PLB and PBF as a function of concentration (c). Bactericidal activities of

PBF, PLB and the combination of PLB and PBF as a function of time (d). SEM images of E. coli, 1: untreated, 2: treated with PBF, 3: treated with PLB, and 4: treated with the combination of PBF and PLB (e).211

Incorporation of antibiotics into the structure of antimicrobial polymers can also increase the antimicrobial performance. In this regard, Lu and co-workers developed a series of water-soluble amphiphilic copolymers containing cationic,

Page | 73 CHAPTER TWO

hydrophobic and ciprofloxacin pendant groups.212, 213 The presence of ciprofloxacin significantly reduced the MIC value of the ternary copolymers compared to the control polymer, which only contained cationic and hydrophobic monomers. In ternary antimicrobial polymers containing QA moieties as the source of positive charge, the best antimicrobial activity was observed for the polymer with 56.4, 4.3 and 39.3% QA, ciprofloxacin and hydrophobic content, respectively.212 This polymer showed the MIC of 4 µg mL−1 against E. coli, which was ~10-fold lower than the MIC value of the control polymer with no ciprofloxacin moieties. Moreover, strong bactericidal activity against E. coli was observed for polymer-modified fibers in disk- diffusion assay. However, the applicability of these polymers in biomedical applications remains inconclusive as no information regarding the cytotoxicity of the ternary copolymers toward mammalian cells was presented in that study.

Tang and co-workers reported the synergistic interaction between an antimicrobial polymer with a relatively different chemical structure and β-lactam antibiotics against various MRSA strains.214 They introduced novel cationic cobaltocenium-containing polymers with membrane disruption activity and the ability to reduce the activity of β-lactamase enzymes. These metallopolymers showed high affinity to members of the β-lactam family and formed β-lactam- metallopolymer bioconjugates via ion-pairing between their cationic cobaltocenium moieties and carboxylate anions of the antibiotics (Figure 2.25 a).

However, in the presence of β-lactamase or upon contact with bacterial membrane, antibiotic molecules could be released from the β-lactam- metallopolymer bioconjugates due to the ion-exchange with carboxylate anions of the β-lactamase or negatively charged components of the bacterial cell

Page | 74 CHAPTER TWO

membrane (Figure 2.25 b). β-Lactam-metallopolymer bioconjugates were able to inhibit the activity of β-lactamase (obtained from HA-MRSA strain) by more than

80% at the subinhibitory concentration of 5 μM. The bactericidal activities of individual metallopolymer, β-lactam antibiotics (penicillin-G, cefazolin, amoxicillin, and ampicillin) and β-lactam-metallopolymer bioconjugates were then assessed using disk-diffusion assay against three strains of MRSA. It was found that β-lactam-metallopolymer bioconjugates demonstrated higher activity in comparison with the individual agents, in particular against HA-MRSA strain, where the diameter of inhibition zone was increased from less than 8 mm for both metallopolymer and penicillin-G to more than 16 mm for the corresponding bioconjugate. The synergistic killing effect of β-lactam-metallopolymer bioconjugates was further confirmed by confocal microscopy and SEM images, using penicillin-G as a representative antibiotic (Figure 2.25 c and d),where the highest bactericidal effect was observed for the β-lactam-metallopolymer bioconjugates. SEM images clearly show membrane damage and cell lysis in the presence of penicillin-G-metallopolymer bioconjugates (Figure 2.25 d). While the majority of common β-lactamase inhibitors are inactive against bacteria, the introduced metallopolymers exhibited potent antimicrobial activity against tested bacteria with MIC values in the range of 15-100 µg mL−1. Moreover, negligible hemolytic activity and low in vivo toxicity were observed for this antimicrobial polymer.

Page | 75 CHAPTER TWO

Figure 2.25. Schematic illustration of the formation of β-lactam-metallopolymer bioconjugate via ion-pairing between antibiotic and metallopolymer (a). Release of β-lactam from β- lactam-metallopolymer bioconjugate in the presence of β-lactamases or lipoteichoic acid (b).

Confocal microscopy images of HA-MRSA cells before (control) and after incubation with penicillin-G, polymer and penicillin-G/polymer treatments (the control and treated bacteria were stained with BacLight live/dead kit) (scale bar = 50 μm) (c). SEM images of HA-MRSA before (control) and after incubation with penicillin-G, polymer and penicillin-G/ polymer treatments (scale bar =1 μm) (d).214

Page | 76 CHAPTER TWO

In a follow-up study by the same group, a multifunctional macromolecular system was designed by incorporating phenylboronic acid and β-lactam antibiotics into the structure of cationic cobaltocenium-containing polymers

(Figure 2.26 a).215 The addition of boronolectins to the structure of the polymer enhanced the interactions between the polymer and bacterial membrane through the reaction of phenylboronic acid moieties with polyols components of lipopolysaccharide or peptidoglycan in the membrane of Gram-negative or

Gram-positive bacteria, respectively. This additional binding strategy further improved the efficacy of the metallopolymers against Gram-negative bacteria in particular (Figure 2.26 b). Synergistic interactions between three active components in this macromolecular system led to an improved antimicrobial activity compared to individual antibiotic or antibiotic-metallopolymer bioconjugate against both Gram-negative and Gram-positive bacteria. As an example, when assessed against E. coli, the MIC value of penicillin was reduced from 12.6 µg mL−1 to 7.1 and 3.7 µg mL−1 when it was used in combination with metallopolymer and boronolectin-containing metallopolymer, respectively. The phenylboronic acid-containing penicillin-complexed cobaltocenium metallopolymers exhibited minimal cytotoxicity against RBCs (<10% hemolysis at 500 µg mL−1) and also against immune cells in both in vitro and in vivo experiments.

Page | 77 CHAPTER TWO

Figure 2.26. Chemical structure of multifunctional macromolecular system comprising cobaltocenium-containing metallopolymer, boronic acid, and penicillin (a). Schematic illustration of synergistic antimicrobial activity of multifunctional polymer against a Gram- negative bacteria (b).215

Antimicrobial dendrimers also showed potential as coagents in combination with antibiotics. Besides having antimicrobial activity, the branched structure of these polymers allows for the efficient antibiotic loading or conjugation.

Combination therapy systems involving dendrimers (mostly PPI or PAMAM) and antibiotics have shown potent antimicrobial activity against a wide range of pathogens.190, 216-218

Lisowka and co-workers investigated the possible enhancement of antimicrobial activity against Gram-negative bacteria through co-administration of nadifloxacin (i.e., a fluoroquinolone antibiotic) with PPI or maltose-modified

PPI dendrimers. Antimicrobial activity of individual agents and combinations were tested using microdilution assay against P. aeruginosa, E. coli and Proteus hauseri. Results showed that the addition of PPI and maltose-modified PPI

Page | 78 CHAPTER TWO

dendrimers at sub-inhibitory levels (1-5 µM) could significantly increase the inhibition effect of nadifloxacin on the growth of P. aeruginosa and E. coli. Against

P. hauseri, however, the increased antimicrobial efficiency was only observed for the combinations at the highest concentrations of nadifloxacin and dendrimers.

Cytotoxicity studies against various mammalian cells showed that PPI dendrimers were toxic at concentrations above 1 µM, while nadifloxacin showed no toxicity up to 6 µg mL−1 (the highest concentration used in antimicrobial test).

In general, the combination of nadifloxacin and PPI dendrimers (at 1 µM) inhibited the growth of Gram-negative bacteria and displayed minimal toxicity toward mammalian cells.

2.5.3 Antimicrobial polymers in combination with essential oils

Plant-derived antimicrobial compounds have been known and used in medicine, food and packaging industries for some time.99 These natural antimicrobial agents are produced as secondary metabolites by many plants and represent high diversity in their chemical compositions and antimicrobial activities. Essential oils (EOs) can be extracted from different parts of the plants and commonly contain low molecular weight phenolic compounds as their active antimicrobial components.219, 220 Despite the promising antimicrobial/antibiofilm activity against a wide range of pathogens, potential clinical application of EOs has been limited by their poor solubility and stability in aqueous media.

Therefore, various encapsulation and delivery platforms have been reported to address these drawbacks.221-223

Rotello and co-workers synthesized a series of oil-in-water cross-linked polymeric nanocomposites with strong antibiofilm activity.224-226 In 2017, the

Page | 79 CHAPTER TWO

development of carvacrol loaded cross-linked polymer nanocomposite (X-NC) was reported by this group.224 Poly(oxanorborneneimide) copolymers (PONI-

GAT) bearing primary amine, guanidinium, and tetraethylene glycol monomethyl ether groups were synthesized as both stabilising agent and delivery vehicle for carvacrol (Figure 2.27 a). Nanocomposites were prepared by emulsifying carvacrol into PONI-GAT aqueous solution at pH 10. To obtain cross-linked nanocomposites, poly(maleic anhydride-alt-octadecene) (p-MA-alt-

OD) was loaded into the carvacrol as the cross-linking agent (Figure 2.27 b).

Upon emulsification, PONI-GAT deposited on the oil−water interface, followed by the fast cross-linking reaction between amine groups of PONI-GAT and p-

MA-alt-OD to yield highly stable X-NCs. The X-NCs prepared using optimal formulation had a net negative charge based on zeta (ζ) potential measurements and were around 250 nm in size, as confirmed by TEM and DLS. These nanocomposites were highly stable in serum-containing media (probably due to the overall net negative charge of nanocomposites) and could effectively diffuse into the biofilm matrix. The antibiofilm activities of X-NCs and individual components, PONI-GAT and carvacrol, were then assessed against 1-day-old E. coli (CD-2), S. aureus (CD-489), P. aeruginosa (CD-1006), and Enterobacter cloacae complex (CD-1412) biofilms. Against all four tested bacteria and at all tested concentrations, X-NCs showed the best bactericidal activity (Figure 2.27 c).

Finally, the therapeutic potential of X-NCs was evaluated against fibroblast−biofilm coculture wound model. X-NCs could efficiently eradicate P. aeruginosa biofilms (∼99.5%) after 3 h treatment at 15 v/v% of X-NC emulsion, without any significant toxicity toward 3T3 fibroblast cells (Figure 2.27 d).

Page | 80 CHAPTER TWO

Figure 2.27. Chemical structure of the poly(oxanorborneneimide) copolymers (PONI-GAT), poly(maleic anhydride-alt-octadecene) crosslinking agent (p-MA-alt-OD) and carvacrol oil

(a). Preparation of oil-in-water cross-linked polymeric nanocomposite (X-NCs) via emulsification of p-MA-alt-OD-loaded carvacrol into the aqueous solution of PONI-GAT (b).

Bactericidal activities of 10 wt% X-NCs, carvacrol and PONI-GAT against 1-day-old P. aeruginosa after 3 h incubation as a function of emulsion concentration (c). Cytotoxicity and bactericidal activities of 10 wt% X-NCs against 3T3 fibroblast−P. aeruginosa biofilm coculture model after 3 h of treatment as a function of emulsion concentration (d).224

In a subsequent study conducted by the same group, the chemical composition of cross-linked nanocomposite was modified so that they could be easily degraded in the presence of glutathione or esterase enzymes.225 Thus, a different cross-linking strategy was implemented using dithiol−disulfide (DTDS) cross-

Page | 81 CHAPTER TWO

linker and ester-linked maleimide groups. The resulting biodegradable cross- linked nanocomposites (X-BNCs) were stable in serum-containing media; however, they were quickly degraded in the presence of glutathione or esterase as confirmed by an increase in the size of X-BNCs in the DLS analysis. Similar to their previous study,224 the synthesized nanocomposites could readily penetrate into the biofilm matrix and effectively eradicate the biofilm of various Gram- negative and Gram-positive bacteria (~100% reduction in viability) after 3 h incubation at 5 v/v% of the emulsion. X-BNCs also demonstrated therapeutic potential against wound biofilm by causing a 4-fold reduction in the CFU of P. aeruginosa with minimal toxicity toward 3T3 fibroblast cells in a coculture wound model. Markedly, no resistance toward X-BNCs was observed in E. coli even after

20 serial passages in the presence of sub-MIC levels of X-BNCs.

2.5.4 Antimicrobial polymers in combination with metal-based

nanomaterials

Nanoparticle-based strategies have recently emerged as potential alternatives to traditional therapeutic approaches using conventional antibiotics.113, 114 By acting through different mechanisms, antimicrobial nanoparticles can impede the resistance acquisition in bacteria. Moreover, due to the facile synthesis procedures and relatively high colloidal stability, antimicrobial nanoparticles are promising candidates for potential clinical applications. Although still under debate, the most widely accepted mechanisms for the antimicrobial activity of metallic nanomaterials is known to be a combination of bacterial cell membrane

(physical damage) disruption, oxidative stress through ROS generation and interference of essential cellular processes such as DNA synthesis.115 The

Page | 82 CHAPTER TWO

mechanism of antimicrobial activity of nanoparticles is mediated by their physiochemical properties, including composition, surface charge and functionalities, size and shape.114 Moreover, the antimicrobial activity of nanomaterials seems to be cell-type-dependent and varies based on the characteristics of the bacterial cells such as cell wall composition.113 Using nanomaterials in combination with synthetic antimicrobial polymers can be a potent therapeutic approach to combat antibiotic resistance by further increasing the non-specific actions on bacterial cells and broadening the spectrum of antimicrobial activity. Therefore, various types of metal-based nanomaterials have been used in combination with antimicrobial polymers in an effort to achieve synergistic interactions and consequently, higher therapeutic potential.

Silver, either in the form of metal, ion or nanoparticle has shown great antimicrobial properties against a wide range of pathogens. Ag-containing systems exert their antimicrobial activity through membrane disruption and continuous release of Ag+ . Ag+ ions can induce oxidative stress, inactivate cellular enzymes and damage DNA in bacterial cells.

One of the earliest studies on the co-administration of cationic polymers and silver-containing nanoparticles was published by Sen and co-workers in 2006.227

The cationic polymer-silver bromide nanocomposites were synthesized via a simple one-pot precipitation technique. The synthesis procedure started by partial N-alkylation of the pyridine groups of a commercially available poly(4- vinylpyridine), followed by the on-site precipitation of the bromide counterions through the gradual addition of silver para-toluene sulfonate solution (Figure

2.28). By changing the degree of N-alkylation of the polymers (21 and 42%) and

Page | 83 CHAPTER TWO

varying the molar ratios between silver ions and polymer bromide ions (1:1 and

1:2), four cationic polymer-stabilized silver bromide nanoparticles with different sizes and Ag contents were obtained. TEM images showed that a lower degree of N-alkylation and lower Ag+ to Br− ratio would lead to a reduction in the size of synthesized AgBr nanoparticles. Comparison of the antimicrobial activities of polymer-AgBr nanocomposites and individual components in Luria-Bertani (LB) broth proved the superior performance of nanocomposites against E. coli and

Bacillus cereus. All four tested nanocomposites had the MIC value of 50 µg mL−1 against both tested bacteria and were bactericidal at this concentration as confirmed by CFU analysis. However, when tested via the disk-diffusion method, polymer-AgBr nanocomposite with the smaller particle size exhibited higher bactericidal activity (i.e., larger inhibition zone). The authors attributed this observation to the higher rate of Ag+ release from the smaller nanoparticles.

This was confirmed by the results of atomic emission spectrometry analysis which showed that Ag+ release was inversely proportional to the nanoparticle size. This offered an efficient way of tuning the antimicrobial activity of the nanocomposites by tailoring the structure of nanoparticles. The combined results from the antimicrobial tests performed on the surface and in the broth suggested that the antibacterial activity of the nanocomposites was induced by a combination of membrane disruption and release of Ag+ ions from AgBr nanoparticles. The polymer-AgBr nanocomposites also showed great efficiency against P. aeruginosa biofilm formation for 24 to 72 h, when used as coating for glass substrates. In contrast, the control glass substrate which was coated with corresponding alkyl-modified poly(4-vinylpyridine) could only partially inhibit biofilm formation for the first 24 h and lost its efficacy as the dead bacteria

Page | 84 CHAPTER TWO

covered the surface of the coating. Synergistic interactions between the cationic polymer and AgBr nanoparticles resulted in a significant improvement in bacteriostatic, bactericidal and antibiofilm activities against both Gram-negative and Gram-positive bacteria, compared to individual components. Furthermore, due to the sustained release of Ag+, these nanocomposites showed extended bactericidal activity for up to 17 days in LB broth, while individual AgBr nanoparticles and polymers exhibited poor activities in the same condition.

Figure 2.28. Synthesis of dual-action polymer-AgBr nanocomposites via on-site precipitation method.227

In 2012, Jang and co-workers reported the development of silver nanoparticle- embedded cationic polymer nanofibers using one-pot aqueous dispersion polymerizations in the presence of the poly(vinylalcohol) (PVA).228 In the synthesis procedure, 2,2-azobis(isobutyronitrile) (AIBN) played two roles as the reducing agent for Ag+ and radical initiator for 2-(tert-butylamino)ethyl methacrylate (TBAM) polymerization. Nanofibers were then formed by the

Page | 85 CHAPTER TWO

assembly of the synthesized silver nanoparticles (AgNPs) and PTBAM on PVA chains due to the high stirring rate. Fabricated PTBAM-AgNP nanofibers demonstrated higher antimicrobial performance against E. coli and S. aureus compared with the analogous nanofibers that were prepared using poly(methyl methacrylate)(PMMA), (PMMA)-AgNP nanofibers, and silver sulfadiazine

(SSD) at the same total silver concentration. For example, when tested against E. coli, the MIC value of PTBAM-AgNP nanofibers was 2 and 16 times lower than the MIC values for PMMA-AgNP nanofibers and SSD, respectively. PTBAM-

AgNP nanofibers also demonstrated promising bactericidal activity as evaluated via disk-diffusion assay (modified Kirby-Bauer test) against S. aureus. The diameter of inhibitory zone was 50.2 and 17.6 mm for PTBAM-AgNP nanofibers and SSD, respectively. The promising antimicrobial activity of PTBAM-AgNP nanofibers was attributed to sustained release of Ag+ and AgNPs from the fibers as well as the antimicrobial activity of PTBAM substrate.

In an interesting study conducted by Mei et al., core-shell nanoparticles composed of AgNP core and quaternized poly [2-(dimethylamino)ethyl methacrylate] (qPDMAEMA) shell were synthesized and systematically investigated in terms of biological performance.229 qPDMAEMA was synthesized via RAFT polymerization of DMAEMA monomers followed by a quaternization step. Subsequently, core-shell nanoparticles (qPDMAEMA-AgNPs) were formed via the reduction of AgNO3 with NaBH4 in the presence of qPDMAEMA (Figure

2.29). The antimicrobial activity and cytotoxicity of core-shell nanoparticles were optimised by changing the length of alkyl tail, molar ratio of qPDMAEMA to silver, NaBH4 volume and molecular weight of qPDMAEMA. The optimum

AgNP-qPDMAEMA core-shell nanoparticle was found to have pendant butyl

Page | 86 CHAPTER TWO

groups, a molecular weight of 8427 Da and was synthesized with a 1:1 molar ratio of qPDMAEMA to silver. The core-shell nanoparticles showed higher antimicrobial activity against both P. aeruginosa and S. aureus compared to individual qPDMAEMA and AgNPs as evidenced by larger diameter of the zone of inhibition in the disk-diffusion assay. Authors correlated the observed potency of the core-shell nanoparticles to the polyvalent and synergistic antimicrobial activity of the components. This postulation was further confirmed by the results of the mechanistic study, which suggested that the antimicrobial activity of nanoparticles involved the inhibition of intracellular enzymatic activity and membrane disruption. qPDMAEMA-AgNPs demonstrated low toxicity toward

NIH3T3 fibroblast cells with more than 80% cell viability after one day incubation at ≤8.0 µg mL−1 which is well above the MIC values of 0.2 and 0.4 µg mL−1 against

P. aeruginosa and S. aureus ,respectively. The potency of these nanoparticles was also tested in vivo using P. aeruginosa and S. aureus-induced wound infection models in healthy and diabetic rats. The wound healing was significantly improved in the treatment groups, which had received qPDMAEMA-AgNP once a day for 24 days.

Page | 87 CHAPTER TWO

Figure 2.29. Synthetic scheme of qPDMAEMA-AgNP core-shell nanoparticles.229

In a follow-up study, the same group incorporated a biocompatible carbohydrate polymer into the structure of qPDMAEMA-AgNP nanocomposites to further improve their biological performance.230 The nanocomposite comprising qPDMAEMA and poly(2-(acrylamido) glucopyranose) (PAGA) shell and AgNPs core, qPDMAEMA/PAGA-AgNPs, were simply synthesized by the reduction of Ag+ in the presence of both polymers. The combined results from hemolysis and cytotoxicity assays clearly showed that the presence of carbohydrate polymers on the surface of nanoparticles could significantly improve the biocompatibility of the nanocomposites in comparison with qPDMAEMA-AgNPs across all concentration tested. Both PAGA-AgNPs and qPDMAEMA/PAGA-AgNPs caused less than 10% hemolysis over a wide range of concentrations (0.15-10 µmol L−1). When tested against NIH3T3 cells using

MTT assay, minimal toxicity (>80% cell viability) was observed for PAGA-

AgNPs and qPDMAEMA/PAGA-AgNPs at concentrations up to 10 and 2.5 µmol

Page | 88 CHAPTER TWO

L−1, respectively. The qPDMAEMA/PAGA-AgNPs also showed enhanced antimicrobial activity due to the synergistic interactions between the three components. Besides the antimicrobial activity of AgNPs and qPDMAEMA,

PAGA also contributed to the antimicrobial activity via glucosamine-mediated penetration into the bacterial membrane. The antimicrobial activities of these nanocomposites against P. aeruginosa, E. coli, S. aureus, and Bacillus amyloliquefaciens (B. amyloliquefaciens) biofilms were subsequently studied via confocal microscopy 3D imaging, biofilm dispersal and colony counting assays.

Results from biofilm dispersal assay clearly showed the synergistic interactions between AgNPs and PAGA and also between AgNPs and qPDMAEMA. The biofilm dispersal caused by either of PAGA-AgNPs or qPDMAEMA-AgNPs was higher than the effect of their individual components. However, the biofilm dispersal induced by qPDMAEMA/PAGA-AgNPs was just slightly higher than the effect of qPDMAEMA-AgNPs especially at concentrations higher than 5

µmol L−1. Colony counting assay showed the ability of qPDMAEMA/PAGA-

AgNPs to kill the bacteria in both planktonic and biofilm forms for all tested bacteria. However, higher potency was observed against two tested Gram- positive bacteria, namely S. aureus and B. amyloliquefaciens, with more than 3 and

4 log10 reductions in CFU compared with the untreated samples for planktonic and biofilm, respectively. In vivo study on mouse implant infection model showed that qPDMAEMA/PAGA-AgNPs could effectively eradicate bacterial biofilm and reduce the number of viable bacteria on the implant after only one treatment at the concentration of 2.5 µmol L−1.

Apart from linear polymers, cationic polymers with branched architectures have also been used for functionalisation of AgNPs. The high density of

Page | 89 CHAPTER TWO

functionalities in the polymeric nanostructures such as hyperbranched or star polymers not only improves the biological performance of the polymer but also favours the formation and stabilization of Ag nanoparticles.

A series of cationic branched polymer-AgNP nanocomposites was developed by Yan and co-workers.231 Branched poly(sulfone amines) (PSAs) were synthesized by the reaction of divinylsulfone and 1-(2-aminoethyl)piperazine, where the degree of branching (DB) was dictated by the volumetric ratio between

DMF and water (Figure 2.30). The PSA-coated AgNPs (PSA-AgNPs) were formed through the complexation of Ag+ with the PSAs and subsequent in situ reduction of Ag+ in the presence of extensive amine functionalities of PSAs. TEM images showed a reduction in the size of AgNPs upon increasing the DB of PSAs.

Antimicrobial activities of PSAs and PSA-AgNPs against Aspergillus niger and E. coli were evaluated via standard disk-diffusion assay and LB broth method, respectively. PSA-AgNPs were found to be more effective than pure PSAs in both assays regardless of the DB. Surprisingly, increasing the DB affected the antimicrobial activities of PSAs and PSA-AgNPs in opposite ways. The maximum antimicrobial activity was observed at the lowest DP (0.04) and highest DB (0.41) for PSAs and PSA-AgNPs, respectively. The PSA-AgNP nanocomposite comprising the PSA polymer with the DB of 0.41, produced an inhibitory zone with the diameter of 2.3 mm against tested fungi species in disk- diffusion assay and could effectively inhibit the growth of E. coli by more than

80% at the concentration of 0.003 mmol mL-1. The authors attributed this to the effect of DB on the size of AgNPs. Based on the TEM images, increasing the DB of PSAs resulted in a reduction in the size of AgNPs, which in turn improved the surface interactions and penetration ability of the nanocomposite. The toxicity of

Page | 90 CHAPTER TWO

the pure PSA was examined against COS-7 cells (i.e., kidney fibroblast cell line) over a wide range of concentrations and DBs. PSA were biocompatible across all tested concentrations with the lowest toxicity at the highest DB (about 80% cell viability at DB = 041). However, no information on the cytotoxicity of PSA-

AgNPs nanocomposite was provided in the study.

Figure 2.30. Schematic illustration of the synthesis of linear, slightly branched and highly branched PSAs and PSA-AgNP nanocomposites. Bactericidal activity of PSA-AgNPs against

Aspergillus niger in disk-diffusion assay (bottom right corner).231

In a similar study by Dai et al., the impact of the structure of cationic ligands

(linear or star polymers) on the antimicrobial activity of AgNP nanocomposites was evaluated against resistant bacteria in both planktonic and biofilm forms.232

The structure of cationic ligand was optimised via structure-activity relationship studies and eventually the star polymer with 8 arms and butyl pendant groups

Page | 91 CHAPTER TWO

(8-arm-PEG-b-DMAEMA-C4) was selected as the best ligand for AgNPs. The resulting nanocomposite demonstrated high hemocompatibility and very low toxicity towards NIH3T3 cells due to the presence of biocompatible PEG block and star architecture. Evaluation of the antimicrobial activity against Gram- negative and Gram-positive bacteria in both planktonic and biofilm forms confirmed the improved efficacy of the nanocomposites formed using 8-arm-

PEG-b-DMAEMA-C4 compared to analogous nanocomposites with linear or 4- arm cationic ligands and also the individual component (i.e., AgNPs and 8-arm-

PEG-b-DMAEMA-C4). The 8-arm-PEG-b-DMAEMA-C4-AgNP nanocomposite was able to effectively inhibit the bacterial growth and eradicate biofilms without inducing resistance development in bacterial cells as a result of the synergistic interactions between membrane targeting ligand and AgNPs.

In comparison with the remarkable inherent antimicrobial activity of AgNPs, gold nanoparticles (AuNPs) are considered to be biologically inert.233 However, they are potential candidates to be involved in antimicrobial therapy and many other biomedical applications due to the high biocompatibility, facile and fast preparation, straightforward surface modification and unique optical properties.

Functionalisation of AuNPs with cationic antimicrobial polymers provides an effective platform to improve the antimicrobial activity of components as well as modulating the physiochemical properties of nanoparticles in biological medium.234

The synthesis of multivalent polymer-AuNP nanocomposites thorough in situ reduction of chloroauric acid in the presence qPDMAEMA was reported by Li and co-workers.235 Presentation of qPDMAEMA on the surface of AuNPs

Page | 92 CHAPTER TWO

stabilized the AuNPs in biological medium and increased the interactions between nanoparticles and bacterial membrane due to the high density of positive charges on the surface of nanoparticles. The growth of planktonic P. aeruginosa and S. aureus was inhibited by more than 90% in the presence of qPDMAEMA-AuNPs at a concentration as low as 1.5 µg mL−1. In disk-diffusion assay, qPDMAEMA-AuNPs could produce larger inhibition zone than that of pure qPDMAEMA against both P. aeruginosa and S. aureus. The authors related the potent bactericidal activity of the polymer-coated AuNPs to the polyvalent effect of polymer on the surface of AuNPs. These nanocomposites had relatively low toxicity when tested against NIH3T3 fibroblast cells with more than 80% cell viability at the concentration of 12 µg mL−1.

In 2016, Yuan and co-workers developed novel recyclable polymer-AuNP nanocomposites by immobilizing qPDMAEMA and poly[(2-methacrylamide) glucopyranose] (PMAG) on the surface of AuNPs.236 Both polymers were synthesized by RAFT polymerization using 4-cyano-4-

(phenylcarbonothioylthio)-pentanoic acid as the chain transfer agent. The synthesized polymers were then reacted with excess amounts of 2-aminoethanol to afford terminal thiol groups. Finally, polymer immobilized-AuNPs were prepared by grafting the terminally thiolated PMAG and qPDMAEMA onto the preformed citrate-protected AuNPs (Figure 2.31). The resulting nanocomposites demonstrated antimicrobial activity due to the presence of qPDMAEMA, and more importantly, were selective toward E. coli as a result of the PMAG components. PMAG showed high affinity toward E. coli pili and therefore could endow the designed nanoparticles with target specificity. Expectedly, poor bactericidal activity was observed against S. aureus due to the absence of glycol-

Page | 93 CHAPTER TWO

sensitive fimbriae. Notably, these nanocomposites could be efficiently recycled after each treatment via the addition of excess amounts of mannose as the lectin on E. coli pili has higher affinity to mannose over glucose moieties of PMAG.

Recycled nanocomposites could retain their efficacy after three cycles. Evaluation of the antibacterial activity using colony counting assay clearly showed the synergistic interactions between qPDMAEMA and PMAG. AuNPs bearing both polymers exhibited higher bactericidal activity in comparison with AuNPs grafted with PMAG or qPDMAEMA. Although PMAG is inactive against bacteria, it could potentiate the overall antimicrobial activity by increasing the interactions between nanoparticles and bacterial cell membrane and lowering the required treatment does. Moreover, the introduction of PMAG onto the surface of AuNPs reduced the toxic effect of qPDMAEMA towards L929 fibroblast cells, as evidenced by MTT assay. The observed improvement in the biocompatibility of composite nanoparticles was attributed to the reduction in the required treatment dose and also the cytocompatibility of PMAG.

Figure 2.31. Schematic representation of the synthesis of recyclable PMAG and qPDMAEMA-immobilised AuNPs.236

Similarly, Fullam and co-workers reported the synthesis of PDMAEMA and poly(N-hydroxyethyl acrylamide) (PHEA) coated AuNPs using a direct

Page | 94 CHAPTER TWO

reduction method.237 Although antimicrobial activity was mostly controlled by

PDMAEMA, the presence of PHEA was essential for the stability of nanoparticles in biological media. Preliminary assessment of the antimicrobial activity showed that the MIC values of the nanocomposite bearing PDMAEMA was two and eight times lower than the MIC of the bulk polymer against M. smegmatis and E. coli, respectively. This observation confirmed that the multivalent presentation of antimicrobial polymers on the surface of AuNPs could improve the efficacy of antimicrobial polymers, especially against E. coli. However, introduction of the

PHEA onto the surface of the nanoparticles led to a reduction in antimicrobial activity in comparison with the nanoparticles containing PDMAEMA only.

Examination of the bactericidal activities revealed that all tested nanocomposites were bactericidal against E. coli (MBC corresponded to MIC) but bacteriostatic against M. smegmatis (MBC significantly higher than MIC). Fluorescence microscopy results suggested that membrane disruption might not be involved in antimicrobial mechanism of these nanoparticles against M. smegmatis since no evidence of membrane permeabilization was observed even after treatment at 2

× MIC. In terms of hemocompatibility, all the tested nanocomposites were non- hemolytic (<5% hemolysis) even at 1600 µg mL−1, however, hemagglutination effect was observed even below the MIC, which might be due to the multivalent presentation of cationic polymers on nanoparticles.

In a recent study, Tang and co-workers reported the development of an antimicrobial agent by incorporating their previously reported metallopolymer- antibiotic bioconjugates on the surface of gold nanoparticles.238 The cationic cobaltocenium polymers (PCo) with three different molecular weights (6000,

15000 and 30000 g mol-1) were synthesized by RAFT polymerization and then

Page | 95 CHAPTER TWO

subjected to ligand exchange using tetrabutylammonium chloride to yield water- soluble PCo with Cl− as the counterion. Subsequently, the polymers were reacted with NaBH4 to reduce the dithioester end groups of the RAFT agent to thiol groups. The resulting polymers were then grafted onto AuNPs to afford PCo-

AuNPs (Figure 2.32). Based on the combined results from disk-diffusion and microdilution assays, PCo-AuNPs demonstrated higher antimicrobial activity compared to individual AuNPs and PCo, against S. aureus and K. pneumonia.

Between all tested samples, the PCo-AuNPs formed using the PCo with the molecular weight of 15000 g mol−1 (PCo-15K) produced the largest zone of inhibition against both bacteria tested and had the lowest MIC values of 54 and

49 µg mL−1 against S. aureus and K. pneumonia, respectively. Bactericidal and bacteriostatic activities were further improved upon the conjugation of penicillin-G to the nanocomposite via counterion-exchange (PCo/Peni-AuNPs)

(Figure 2.32). For instance, when tested via microdilution assay against S. aureus,

PCo/Peni-AuNPs had higher antimicrobial activity with the MIC value of 2.6 µg mL−1, compared to individual PCo-Peni and penicillin-G with the MIC values of

2.6 and 15.8 µg mL−1, respectively. The superior antimicrobial activity of

PCo/Peni-AuNPs was attributed to synergistic interactions of the individual components.

Page | 96 CHAPTER TWO

Figure 2.32. Schematic illustration of the synthesis of AuNPs grafted with metallopolymer- antibiotic bioconjugates.238

Titanium oxide (TiO2) is a broad-spectrum antimicrobial agent that induce antimicrobial activity through membrane disruption and ROS generation.115 TiO2 also shows a unique photocatalytic antibacterial activity, where ROS generation is enhanced under irradiation with visible light, near-UV light or UV-A.

Encouraged by unique bactericidal activity of TiO2 nanoparticles, Jang and co- workers developed polymer-TiO2 nanocomposite in the form of core-shell nanoparticles by photopolymerization of TBAM and ethylene glycol dimethacrylate (EGDMA) on the surface of TiO2 nanoparticles.239 The resulting nanoparticles (PTBAM-co-PEGDMA-TiO2) showed higher antibacterial activity compared to individual components regardless of the presence of UV irradiation.

While bare TiO2 nanoparticles only exhibited antimicrobial activity under UV irradiation (76% killing), PTBAM-co-PEGDMA-TiO2 nanoparticles could effectively kill S. aureus in dark (95.7%) and under UV irradiation (99.8%). The superior antimicrobial performance of PTBAM-co-PEGDMA-TiO2 nanoparticles

Page | 97 CHAPTER TWO

was attributed to the synergistic interactions between TiO2 nanoparticles and biocidal polymer and the increased local concentration of antimicrobial polymers which improved the interactions with bacterial membrane compared to bulk poly(TBAM-co-EGDMA).

Iron oxide nanoparticles and especially superparamagnetic iron oxide nanoparticles have gained special attention in the design of antimicrobial agents recently. Besides exhibiting some levels of antimicrobial activity through ROS generation,240, 241 unique magnetic properties of these nanoparticles make them highly appealing for antimicrobial applications such as hyperthermia therapy,242,

243 bacterial detection systems244, 245 and design of recyclable antibacterial agents.246, 247 However, surface modification is usually an imperative step before they can be incorporated into potential therapeutic or diagnostic platforms.248

Surface modification of magnetic nanoparticles with antimicrobial polymers would assist with their biocompatibility and colloidal stability for biomedical applications and might potentially lead to improved antimicrobial performance due to synergistic interactions between the components.

In a study conducted by Taresco et al., polymer-coated magnetic nanoparticles were synthesized and loaded with usnic acid (UA), a natural antimicrobial compound, as potential antimicrobial agents for the treatment of localized infections.249 Magnetic nanoparticles (MNPs) in the form of manganese iron oxide were synthesized via co-precipitation of metal ions in a microemulsion system and subsequently functionalized with either a hydrophobic branched poly-ε- caprolactone or a cationic polyacrylamide with inherent antimicrobial activity.

UA loading happened through hydrophobic interactions and acid-base

Page | 98 CHAPTER TWO

interactions for poly-ε-caprolactone and polyacrylamide coated MNPs, respectively. While UA loading was relatively low for pristine MNPs, both polymer-coated MNPs had significant UA payloads, with a higher UA loading for polyacrylamide-coated MNPs. When tested against S. epidermidis, UA-loaded polyacrylamide-coated MNPs demonstrated activity in growth inhibition assay with MIC value of 100 µg mL−1, which is significantly lower than values for UA- loaded MNPs and UA-loaded poly-ε-caprolactone-coated MNPs, >2 and 5 mg mL−1, respectively. Authors attributed this enhancement in antimicrobial activity to the synergistic interactions of antimicrobial polymer and UA. However, the contribution of MNPs in the antimicrobial activity of these UA-loaded hybrid nanoparticles is unclear. Based on the presented data, the MNPs served as scaffold for the co-delivery of antimicrobial polymer and UA.

In 2016, Duan and co-workers reported the synthesis of hybrid antimicrobial nanoparticles composed of superparamagnetic MnFe2O4 core and cationic polycarbonate shell.250 Quaternized polycarbonates (PrBrT) were synthesized via

ROP of 5-methyl-5-(3-bromopropyl) oxycarbonyl-1,3- dioxan-2-one (MTC-

(CH2)3Br) monomers followed by deprotection of phosphonate groups and finally a quaternization step using excess amounts of N,N,N′,N′- tetramethylethylenediamine. PrBrT were then grafted onto the surface of

MnFe2O4 nanoparticles via ligand exchange (Figure 2.33). Comparison of the antimicrobial activities of bulk PrBrT and PrBrT-coated MnFe2O4 nanoparticles

(PrBrT-MnFe2O4) against E. coli and S. aureus showed a significant difference in the bactericidal and bacteriostatic activities of tested compounds. PrBrT-MnFe2O4 exhibited the same bactericidal and bacteriostatic activities as bulk PrBrT at significantly lower concentrations (approximately 3 orders of magnitude) even

Page | 99 CHAPTER TWO

without magnetic hyperthermia treatment. While the individual polymer only showed bactericidal activities at 100 mg mL−1, PrBrT-MnFe2O4 could effectively eradicate 97 and 98% of E. coli and S. aureus, respectively, at 120 µg mL−1. This was correlated to the increased local concentration of positive charges for PrBrT-

MnFe2O4 as the analogous nanoparticles with lower polymer grafting density

(i.e., lower positive charge density) had lower antimicrobial activity. Notably, a significant enhancement in bactericidal efficiency of the hybrid nanoparticles was observed under magnetic heating, especially against E. coli. The killing efficiency of PrBrT-MnFe2O4 was increased from almost zero to 100% at the concentration of 60 µg mL−1 when a magnetic field was applied. This improvement was attributed to the synergistic effects of two bactericidal mechanisms including, membrane disruption by cationic polymer and magnetic hyperthermia function of MnFe2O4 core, as the control core-shell nanoparticles decorated with PEG were inactive against bacteria even when exposed to magnetic heating. Moreover, PrBrT-MnFe2O4 showed minimal hemolytic effect

(<3% hemolysis) at a high concentration of 1 mg mL−1.

Page | 100 CHAPTER TWO

Figure 2.33. Synthesis of quaternized polycarbonate (PrBrT)-grafted MnFe2O4 nanoparticles.250

In a study by Pageni et al., charged cobaltocenium-containing metallopolymers were grafted from magnetic iron oxide nanoparticles via surface-initiated RAFT polymerization.251 Iron oxide nanoparticles (FeNPs) were synthesized by coprecipitation method and subsequently functionalized with 4- cyanopentanoic acid dithiobenzoate as RAFT agent. Controlled radical polymerization of 2- cobaltocenium amidoethyl methacrylate hexaflurophosphate monomer was then conducted using AIBN as initiator and

DMF as solvent. The resulting polymer-coated nanoparticles were then subjected to ion exchange in two steps, to yield hydrophilic chloride-paired nanoparticles

(FeNP-Cl) and finally penicillin-G conjugated nanoparticles (FeNP-Peni) (Figure

2.34). The final penicillin-loaded nanoparticles had a Dh of 33 nm as evidenced by DLS analysis with 65 wt% cobaltocenium and approximately 30 wt% penicillin based on TGA measurements. To investigate the impact of the introduction of penicillin-conjugated metallopolymers (PCo-Peni) onto the surface of iron oxide nanoparticles on bactericidal activity, disk-diffusion assay

Page | 101 CHAPTER TWO

was performed at constant penicillin content against two Gram-positive and three Gram-negative bacterial strains. Similar outcome was observed against all tested bacteria where FeNP-Peni exhibited the highest bactericidal activity, corresponding to the largest zone of inhibition, followed by PCo-Peni, penicillin and FeNP-Cl, respectively. The poor performance of FeNP-Cl was attributed to the low concentration of cobaltocenium in tested sample. When tested against S. aureus, the diameter of the inhibition zone was increased from 8 mm for penicillin to 11 and 18 mm for PCo-Peni and FeNP-Peni, respectively. FeNP-Peni also showed significant improvement in bacteriostatic activity compared to penicillin as confirmed by the lower MIC values against both Gram-negative and Gram-

Positive bacteria. For instance, the MIC value was reduced from 13.5 µg mL−1 for penicillin to 3.4 µg mL−1 for FeNP-Peni against S. aureus. The authors postulated that the superior antibacterial efficacy of FeNP-Peni was due to the synergistic interactions between cobaltocenium and penicillin along with the increased local concentration of effective components afforded by the nanoparticle scaffold.

These nanoparticles were shown to be easily recycled in response to an external magnetic field. However, to ensure a constant penicillin concentration, an additional batch of penicillin was added before each cycle. Furthermore, resistance study conducted against E. coli and S. aureus showed that the FeNP-

Peni retained almost the same MIC values after 15 serial passages. Notably,

FeNP-Peni exhibited very low hemolytic activity (<5%) at high concentrations as

500 mg mL-1.

Page | 102 CHAPTER TWO

Figure 2.34. Chemical structure of penicillin, cationic metallopolymer (PCo) and cationic metallopolymer-penicillin bioconjugates (PCo-Peni) (top). Schematic illustration of the synthesis of recyclable magnetic nanoparticles grafted with cationic metallopolymer- penicillin bioconjugates (bottom).251

2.5.5 Antimicrobial polymers in combination with carbon-based

nanomaterials

Carbon nanomaterials, namely fullerene, carbon nanotube (CNT), graphene and carbon dot, are well known for their unique physiochemical properties which are caused by the quantum confinement effect. Owing to their small size,

Page | 103 CHAPTER TWO

large surface area, unique optical/electrical properties, and superior mechanical strength, these low-dimensional materials are potential candidates for many biomedical applications such as drug/gene delivery, imaging, photothermal therapy, biosensor design, and tissue engineering.252 Additionally, moderate antimicrobial activity has also been observed for some members of the carbon nanomaterial family, namely CNTs, graphene-based nanomaterials and carbon dots.253, 254 Although the mechanism behind the antimicrobial activity of these nanomaterials still remains indefinitive, physical damage to the bacterial membrane, oxidative stress through ROS generation, and entrapment of bacterial cells within the aggregated nanoparticles are considered to be involved in their antimicrobial action.255 However, surface modification is required to improve the applicability of these nanomaterials for antimicrobial applications. Although limited studies have been conducted thus far, surface modification of GO nanosheets,256-259 CNT260-262 and carbon quantum dots263 with cationic polymers has improved the overall antimicrobial performance of these nanomaterials.

In this regard, the synthesis of polymer-functionalized graphene oxide (GO) nano sheets was reported by Sun and co-workers.258 In that study, PEG and polyhexamethylene guanidine hydrochloride (PHGC) were covalently conjugated to the surface of GO sheets which were prepared based on modified

Hummers method (Figure 2.35). Antimicrobial activity of PEG- and PHGC- modified GO (GO-PEG-PHGC) was studied by time-kill assay; GO, GO-PEG and

GO-PHGC were also included for comparison. Expectedly, GO-PEG-PHGC exhibited the highest bactericidal activity against both E. coli and S. aureus compared to the rest of the compounds (although at relatively high concentration), where both bacteria were fully eradicated after 60 min incubation

Page | 104 CHAPTER TWO

with GO-PEG-PHGC at 4 mg mL−1. This enhancement in antibacterial activity was correlated to the bactericidal effect of PHGC along with the improved dispersity of GO-PEG-PHGC due to the PEG modification. While PEGylation could significantly enhance the colloidal stability of GO in saline solution and facilitate the interactions between GO-PEG-PHGC nanosheets and bacterial cells,

GO-PEG had the lowest bactericidal activity in time-kill assay against both bacteria tested.

Figure 2.35. Schematic illustration of the synthesis of PEG and PHGC-modified graphene oxide sheets.258

Besides acting as antimicrobial agent, polymer-modified GO could potentially be used in the development of antimicrobial surfaces. Wang and co-workers synthesized GO-qPDMAEMA nanosheets and subsequently deposited them onto PEI-coated glass coverslips to obtain GO-qPDMAEMA films.259 Prepared

Page | 105 CHAPTER TWO

films showed high resistance to adsorption of proteins, bacterial and mammalian cells. Although control films prepared using GO-PDMAEMA and GO-COOH showed similar protein-repelling capabilities as GO-qPDMAEMA, their anti- adhesion activities were much lower when tested against bacterial and mammalian cells. The authors postulated that the presence of QA moieties on the surface of GO-qPDMAEMA films induced toxicity toward bacterial and mammalian cells.

Very recently, Zhou et al. have developed novel hybrid nanomaterials composed of polymer-modified reduced graphene oxide nanosheets and AgNPs with promising antimicrobial activity.264 Catechol-terminated hydrophilic quaternized polymer (qCP) was prepared via RAFT polymerization of

DMAEMA and subsequent quaternization of tertiary amines using benzyl chloride. The resulting polymer was then grafted onto the reduced GO nanosheets (rGO) via mussel-inspired modification strategy to obtain qCP- modified rGO (rGO-qCP). Finally, AgNPs were synthesized by in situ reduction of Ag+ ions in the presence of rGO-qCP nanosheets (rGO-qCP-Ag). Antimicrobial activity of rGO-qCP-Ag was assessed via growth inhibition and killing assays against E. coli and S. aureus, and compared to the activity of individual GO, qCP, rGO-qCP and rGO-Ag. Against both bacteria and at all concentrations tested, rGO-qCP-Ag showed the best antimicrobial activity compared with the other compounds. rGO-qCP-Ag showed higher potency against E. coli than S. aureus with MIC and MBC values of 16 and 32 μgmL−1, respectively. This superior activity was attributed to the synergistic interactions between cationic polymer,

AgNPs and rGO nanosheets. Cationic polymer has proven to be a key component in this synergistic hybrid nanocomposite as it could simultaneously promote the

Page | 106 CHAPTER TWO

physical contact with bacterial cells, stabilize the rGO nanosheets, increase the silver loading content, reduce the size of AgNPs on the surface of rGO and enhance the intracellular ROS levels. Mechanistic studies revealed that a combination of membrane disruption and ROS production was involved in the antibacterial action of the rGO-QCP-Ag.

In similar studies, hybrid nanoparticles composed of amphiphilic polymers,

AgNPs and CNTs have been synthesized and assessed for their antimicrobial performance.260, 265 In a study conducted by Murugan et al., polymer- functionalized AgNPs-deposited multiwalled carbon nanotubes (MWCNTs) were synthesized via a multistep synthesis procedure.260 PPI dendrimers were covalently grafted onto carboxyl functionalized multi-walled CNTs (MWCNTs-

COOH) and then subjected to quaternization to form MWCNTs-qPPI. This was followed by in situ reduction and deposition of AgNPs onto the functionalised

MWCNTs (MWCNTs-qPPI-AgNPs). Compared to MWCNTs-COOH and

MWCNTs-qPPI, MWCNTs-qPPI-AgNPs exhibited superior bactericidal activity against E. coli, S. aureus and . For instance, when tested against E. coli, MWCNTs-qPPI-AgNPs displayed the strongest killing efficiency by eradicating 93.1 ± 0.5% of the bacterial cells after 2 h treatment at 25 µg mL−1, compared to 40 ± 1.5 and 87 ± 0.5% killing efficiencies for MWCNTs-COOH and

MWCNTs-qPPI, respectively. The increased efficacy of the MWCNTs-qPPI-

AgNPs hybrid nanoparticles was attributed to the simultaneous antimicrobial activity of MWCNTs, qPPI and AgNPs.

In 2019, Chen and co-workers reported the synthesis of water-soluble nanocomposites based on PEI-functionalized graphene quantum dots

Page | 107 CHAPTER TWO

(GQD−PEI) and zinc oxide nanoparticles (ZnO) using a sol-gel method.263 To investigate the effect of PEI functionalization on the antimicrobial activity, hybrid nanoparticles composed of GQD−PEI and ZnO (ZnO/GQD-PEI) and control nanocomposite with no PEI capping layer (ZnO/GQD) were tested against E. coli using microdilution assay. Although both nanocomposites showed modest antimicrobial activities, ZnO/GQD-PEI nanocomposites with different

PEI weight ratios (MIC values in the range of 2-2.8 mg mL−1) outperformed

ZnO/GQD (MIC value of 4 mg mL−1). This higher antibacterial efficacy was attributed to the enhanced interactions of the nanocomposites with the bacterial membrane due to the presence of the cationic PEI capping layer. Considering the photochemical activity of ZnO and GQD, the bactericidal activities of nanocomposites were then assessed in the presence of UV irradiation. Notably,

ZnO/GQD-PEI showed higher bactericidal activity than ZnO/GQD and killed

96.9% of the bacterial cell under UV irradiation for 5 min at the concentration of

2 mg mL−1. Interestingly, electron spin resonance measurements showed that the

ROS generation was higher for ZnO/GQD-PEI in comparison with ZnO/GQD, suggesting that the PEI might assist in the generation of ROS by ZnO and GQD.

The authors postulated that the synergistic interactions between three constituents have led to the improved bactericidal activity of ZnO/GQD-PEI.

Cationic PEI, in particular, played a key role in the overall antimicrobial activity of the nanocomposites by reducing the ZnO/GQD-PEI agglomeration, promoting the electrostatic interactions between nanocomposite and bacterial membrane, and increasing the generation of ROS (Figure 2.36).

Page | 108 CHAPTER TWO

Figure 2.36. Schematic illustration of the proposed antibacterial mechanism of

ZnO/GQD−PEI nanocomposite.263

2.6 Summary

In line with the urgent need to design novel antimicrobial agents or strategies to address antibiotic resistance crisis, combination therapy approaches have been developed by researchers over the past decades. The majority of the proposed and tested combinations have been based on the use of traditional antibiotics.

Although such combinations have shown remarkable biological performances, the high risk associated with the development of resistance to individual antibiotics and also the inability of the antibiotics to fight biofilm-related infections limit the clinical application of these combinations. Novel antimicrobial combinations based on the inclusion of membrane targeting compounds, such as antimicrobial polymers, in the therapeutic regimens have Page | 109 CHAPTER TWO

also demonstrated promising antimicrobial potency. Synthetic antimicrobial polymers offer many advantages over their natural counterparts (i.e., AMP) and also traditional antibiotics. There is a very low risk regarding the development of resistance towards antimicrobial polymers in bacteria, and they have shown strong antibiofilm activity as opposed to small molecules antibiotics. Moreover, advances in polymer science provide the ability to readily manipulate the properties of antimicrobial polymers according to the characteristics of their coagent(s) and also the requirements of each specific antimicrobial application.

For instance, the structure of the antimicrobial polymer can be easily modified to enable the polymer to act as a delivery vehicle to improve the localized delivery and also to enhance the stability of the antimicrobial coagents.

Combination therapy platforms based on synthetic antimicrobial polymers and other antimicrobial agents such as NO, antibiotics, metal and metal oxide nanoparticles, etc. are potential alternatives for current antibiotic therapies.

Although combinations involving antimicrobial polymers have been reported with demonstrated activity against planktonic bacteria, the assessment of the antibiofilm activity of such combinations is missing in many of these reports.

Considering the significant role of bacterial biofilm in chronic and recurrent infections and also their contribution to the development and spread of resistance, assessing the proposed antimicrobial combinations against bacterial biofilm is necessary. This work describes the development of novel combinations based on synthetic antimicrobial polymers and NO, antibiotics or essential oils and investigates the potency of these combinations against both planktonic and biofilm bacteria.

Page | 110 CHAPTER TWO

2.7 Reference

1. Blair, J. M. A.; Webber, M. A.; Baylay, A. J.; Ogbolu, D. O.; Piddock, L. J. V., Molecular mechanisms of antibiotic resistance. Nat. Rev. Microbiol. 2015, 13 (1), 42-51. 2. Talebi Bezmin Abadi, A.; Rizvanov, A. A.; Haertlé, T.; Blatt, N. L., World Health Organization Report: Current Crisis of Antibiotic Resistance. BioNanoScience 2019, 9 (4), 778-788. 3. Crofts, T. S.; Gasparrini, A. J.; Dantas, G., Next-generation approaches to understand and combat the antibiotic resistome. Nat. Rev. Microbiol. 2017, 15 (7), 422- 434. 4. D’Costa, V. M.; King, C. E.; Kalan, L.; Morar, M.; Sung, W. W. L.; Schwarz, C.; Froese, D.; Zazula, G.; Calmels, F.; Debruyne, R.; Golding, G. B.; Poinar, H. N.; Wright, G. D., Antibiotic resistance is ancient. Nature 2011, 477 (7365), 457-461. 5. Davies, J., Bacteria on the rampage. Nature 1996, 383 (6597), 219-220. 6. English, B. K.; Gaur, A. H., The Use and Abuse of Antibiotics and the Development of Antibiotic Resistance. In Hot Topics in Infection and Immunity in Children VI, Finn, A.; Curtis, N.; Pollard, A. J., Eds. Springer New York: New York, NY, 2010; pp 73-82. 7. Wright, G. D., Solving the Antibiotic Crisis. ACS Infect. Dis. 2015, 1 (2), 80-84. 8. Rossiter, S. E.; Fletcher, M. H.; Wuest, W. M., Natural Products as Platforms To Overcome Antibiotic Resistance. Chem. Rev. 2017, 117 (19), 12415-12474. 9. Kohanski, M. A.; Dwyer, D. J.; Collins, J. J., How antibiotics kill bacteria: from targets to networks. Nat. Rev. Microbiol. 2010, 8 (6), 423-435. 10. Culyba, M. J.; Mo, C. Y.; Kohli, R. M., Targets for Combating the Evolution of Acquired Antibiotic Resistance. Biochemistry 2015, 54 (23), 3573-3582. 11. Schillaci, D.; Spanò, V.; Parrino, B.; Carbone, A.; Montalbano, A.; Barraja, P.; Diana, P.; Cirrincione, G.; Cascioferro, S., Pharmaceutical Approaches to Target Antibiotic Resistance Mechanisms. J. Med. Chem. 2017, 60 (20), 8268-8297. 12. Pang, Z.; Raudonis, R.; Glick, B. R.; Lin, T.-J.; Cheng, Z., Antibiotic resistance in Pseudomonas aeruginosa: mechanisms and alternative therapeutic strategies. Biotechnol. Adv. 2019, 37 (1), 177-192. 13. Peterson, E.; Kaur, P., Antibiotic Resistance Mechanisms in Bacteria: Relationships Between Resistance Determinants of Antibiotic Producers, Environmental Bacteria, and Clinical Pathogens. Front. Microbiol. 2018, 9, 1-21. 14. Delcour, A. H., Outer membrane permeability and antibiotic resistance. Biochim. Biophys. Acta 2009, 1794 (5), 808-816. 15. Walsh, C., Molecular mechanisms that confer antibacterial drug resistance. Nature 2000, 406 (6797), 775-781. 16. Munita, J. M.; Arias, C. A., Mechanisms of Antibiotic Resistance. In Virulence Mechanisms of Bacterial Pathogens, Fifth Edition, American Society of Microbiology: 2016. 17. Nikaido, H., Molecular Basis of Bacterial Outer Membrane Permeability Revisited. Microbiol. Mol. Biol. Rev. 2003, 67 (4), 593-656. 18. Vaara, M., Agents that increase the permeability of the outer membrane. Microbiol. Rev. 1992, 56 (3), 395-411. 19. Wright, G. D., Bacterial resistance to antibiotics: Enzymatic degradation and modification. Adv. Drug Del. Rev. 2005, 57 (10), 1451-1470. 20. Wilke, M. S.; Lovering, A. L.; Strynadka, N. C. J., β-Lactam antibiotic resistance: a current structural perspective. Curr. Opin. Microbiol. 2005, 8 (5), 525-533.

Page | 111 CHAPTER TWO

21. Shaw, K. J.; Rather, P. N.; Hare, R. S.; Miller, G. H., Molecular genetics of aminoglycoside resistance genes and familial relationships of the aminoglycoside- modifying enzymes. Microbiol. Rev. 1993, 57 (1), 138-163. 22. Mingeot-Leclercq, M. P.; Glupczynski, Y.; Tulkens, P. M., Aminoglycosides: activity and resistance. Antimicrob Agents Chemother 1999, 43 (4), 727-737. 23. Petchiappan, A.; Chatterji, D., Antibiotic Resistance: Current Perspectives. ACS omega 2017, 2 (10), 7400-7409. 24. Sommer, M. O. A.; Munck, C.; Toft-Kehler, R. V.; Andersson, D. I., Prediction of antibiotic resistance: time for a new preclinical paradigm? Nat. Rev. Microbiol. 2017, 15 (11), 689-696. 25. Thomas, C. M.; Nielsen, K. M., Mechanisms of, and Barriers to, Horizontal Gene Transfer between Bacteria. Nat. Rev. Microbiol. 2005, 3 (9), 711-721. 26. Gilmore, M. S.; Lebreton, F.; van Schaik, W., Genomic transition of enterococci from gut commensals to leading causes of multidrug-resistant hospital infection in the antibiotic era. Curr. Opin. Microbiol. 2013, 16 (1), 10-16. 27. Sandoval-Motta, S.; Aldana, M., Adaptive resistance to antibiotics in bacteria: a systems biology perspective. Wiley Interdiscip. Rev. Syst. Biol. Med. 2016, 8 (3), 253- 267. 28. Sánchez-Romero, M. A.; Casadesús, J., Contribution of phenotypic heterogeneity to adaptive antibiotic resistance. Proc. Natl. Acad. Sci. U. S. A. 2014, 111 (1), 355-360. 29. Römling, U.; Balsalobre, C., Biofilm infections, their resilience to therapy and innovative treatment strategies. J. Intern. Med. 2012, 272 (6), 541-561. 30. Stewart, P. S.; William Costerton, J., Antibiotic resistance of bacteria in biofilms. Lancet 2001, 358 (9276), 135-138. 31. Mah, T.-F. C.; O'Toole, G. A., Mechanisms of biofilm resistance to antimicrobial agents. Trends Microbiol. 2001, 9 (1), 34-39. 32. Bjarnsholt, T., The role of bacterial biofilms in chronic infections. APMIS 2013, 121 (s136), 1-58. 33. Koo, H.; Allan, R. N.; Howlin, R. P.; Stoodley, P.; Hall-Stoodley, L., Targeting microbial biofilms: current and prospective therapeutic strategies. Nat. Rev. Microbiol. 2017, 15 (12), 740-755. 34. Lebeaux, D.; Ghigo, J.-M.; Beloin, C., Biofilm-related infections: bridging the gap between clinical management and fundamental aspects of recalcitrance toward antibiotics. Microbiol Mol Biol Rev 2014, 78 (3), 510-543. 35. Arciola, C. R.; Campoccia, D.; Montanaro, L., Implant infections: adhesion, biofilm formation and immune evasion. Nat. Rev. Microbiol. 2018, 16 (7), 397-409. 36. Wolfmeier, H.; Pletzer, D.; Mansour, S. C.; Hancock, R. E. W., New Perspectives in Biofilm Eradication. ACS Infect. Dis. 2018, 4 (2), 93-106. 37. Sadrearhami, Z.; Nguyen, T.-K.; Namivandi-Zangeneh, R.; Jung, K.; Wong, E. H. H.; Boyer, C., Recent advances in nitric oxide delivery for antimicrobial applications using polymer-based systems. J. Mater. Chem. B 2018, 6 (19), 2945-2959. 38. Wolfmeier, H.; Pletzer, D.; Mansour, S. C.; Hancock, R. E. W., New Perspectives in Biofilm Eradication. ACS Infectious Diseases 2018, 4 (2), 93-106. 39. Flemming, H.-C.; Wingender, J.; Szewzyk, U.; Steinberg, P.; Rice, S. A.; Kjelleberg, S., Biofilms: an emergent form of bacterial life. Nat. Rev. Microbiol. 2016, 14 (9), 563-575. 40. del Pozo, J. L.; Patel, R., The Challenge of Treating Biofilm-associated Bacterial Infections. Clin. Pharmacol. Ther. 2007, 82 (2), 204-209.

Page | 112 CHAPTER TWO

41. Arciola, C. R.; Campoccia, D.; Ehrlich, G. D.; Montanaro, L., Biofilm-Based Implant Infections in Orthopaedics. In Biofilm-based Healthcare-associated Infections: Volume I, Donelli, G., Ed. Springer International Publishing: Cham, 2015; pp 29-46. 42. Costerton, J. W.; Stewart, P. S.; Greenberg, E. P., Bacterial Biofilms: A Common Cause of Persistent Infections. Science 1999, 284 (5418), 1318-1322. 43. Henrici, A. T., Studies of Freshwater Bacteria: I. A Direct Microscopic Technique. J. Bacteriol. 1933, 25 (3), 277-287. 44. Geesey, G. G.; Mutch, R.; Costerton, J. W.; Green, R. B., Sessile bacteria: An important component of the microbial population in small mountain streams 1. Limnol. Oceanogr. 1978, 23 (6), 1214-1223. 45. Singer, S. W.; Erickson, B. K.; VerBerkmoes, N. C.; Hwang, M.; Shah, M. B.; Hettich, R. L.; Banfield, J. F.; Thelen, M. P., Posttranslational modification and sequence variation of redox-active proteins correlate with biofilm life cycle in natural microbial communities. ISME J 2010, 4 (11), 1398-1409. 46. van Wolferen, M.; Orell, A.; Albers, S.-V., Archaeal biofilm formation. Nat. Rev. Microbiol. 2018, 16 (11), 699-713. 47. Garrett, T. R.; Bhakoo, M.; Zhang, Z., Bacterial adhesion and biofilms on surfaces. PROG NAT SCI-MATER 2008, 18 (9), 1049-1056. 48. Klausen, M.; Gjermansen, M.; Kreft, J.-U.; Tolker-Nielsen, T., Dynamics of development and dispersal in sessile microbial communities: examples from Pseudomonas aeruginosa and Pseudomonas putida model biofilms. FEMS Microbiol. Lett. 2006, 261 (1), 1-11. 49. McDougald, D.; Rice, S. A.; Barraud, N.; Steinberg, P. D.; Kjelleberg, S., Should we stay or should we go: mechanisms and ecological consequences for biofilm dispersal. Nat. Rev. Microbiol. 2012, 10 (1), 39-50. 50. Kaplan, J. B., Biofilm dispersal: mechanisms, clinical implications, and potential therapeutic uses. J Dent Res 2010, 89 (3), 205-218. 51. Guilhen, C.; Forestier, C.; Balestrino, D., Biofilm dispersal: multiple elaborate strategies for dissemination of bacteria with unique properties. Mol. Microbiol. 2017, 105 (2), 188-210. 52. Anderl, J. N.; Franklin, M. J.; Stewart, P. S., Role of antibiotic penetration limitation in Klebsiella pneumoniae biofilm resistance to ampicillin and ciprofloxacin. Antimicrob Agents Chemother 2000, 44 (7), 1818-1824. 53. Davies, D., Understanding biofilm resistance to antibacterial agents. Nat. Rev. Drug Discov. 2003, 2 (2), 114-122. 54. Larsen, T., Susceptibility of Porphyromonas gingivalis in biofilms to amoxicillin, doxycycline and metronidazole. Oral Microbiol Immunol 2002, 17 (5), 267-271. 55. Williams, I.; Venables, W. A.; Lloyd, D.; Paul, F.; Critchley, I., The effects of adherence to silicone surfaces on antibiotic susceptibility in Staphylococcus aureus. Microbiology 1997, 143 (7), 2407-2413. 56. Brauner, A.; Fridman, O.; Gefen, O.; Balaban, N. Q., Distinguishing between resistance, tolerance and persistence to antibiotic treatment. Nat. Rev. Microbiol. 2016, 14 (5), 320-330. 57. Olsen, I., Biofilm-specific antibiotic tolerance and resistance. Eur. J. Clin. Microbiol. Infect. Dis. 2015, 34 (5), 877-886. 58. Flemming, H.-C.; Wingender, J., The biofilm matrix. Nat. Rev. Microbiol. 2010, 8 (9), 623-633. 59. Van Acker, H.; Van Dijck, P.; Coenye, T., Molecular mechanisms of antimicrobial tolerance and resistance in bacterial and fungal biofilms. Trends Microbiol. 2014, 22 (6), 326-333.

Page | 113 CHAPTER TWO

60. Suci, P. A.; Mittelman, M. W.; Yu, F. P.; Geesey, G. G., Investigation of ciprofloxacin penetration into Pseudomonas aeruginosa biofilms. Antimicrob Agents Chemother 1994, 38 (9), 2125-2133. 61. Vrany, J. D.; Stewart, P. S.; Suci, P. A., Comparison of recalcitrance to ciprofloxacin and levofloxacin exhibited by Pseudomonas aeruginosa bofilms displaying rapid-transport characteristics. Antimicrob. Agents Chemother 1997, 41 (6), 1352-1358. 62. Daddi Oubekka, S.; Briandet, R.; Fontaine-Aupart, M. P.; Steenkeste, K., Correlative time-resolved fluorescence microscopy to assess antibiotic diffusion-reaction in biofilms. Antimicrob. Agents Chemother 2012, 56 (6), 3349-3358. 63. Amato, S. M.; Fazen, C. H.; Henry, T. C.; Mok, W. W. K.; Orman, M. A.; Sandvik, E. L.; Volzing, K. G.; Brynildsen, M. P., The role of metabolism in bacterial persistence. Front. Microbiol. 2014, 5, 70-70. 64. Evans, D. J.; Allison, D. G.; Brown, M. R. W.; Gilbert, P., Susceptibility of Pseudomonas aeruginosa and Escherichia coli biofilms towards ciprofloxacin: effect of specific growth rate. J. Antimicrob. Chemother. 1991, 27 (2), 177-184. 65. Stewart, P. S.; Franklin, M. J., Physiological heterogeneity in biofilms. Nat. Rev. Microbiol. 2008, 6 (3), 199-210. 66. Lewis, K., Multidrug Tolerance of Biofilms and Persister Cells. In Bacterial Biofilms, Romeo, T., Ed. Springer Berlin Heidelberg: Berlin, Heidelberg, 2008; pp 107- 131. 67. Lewis, K., Persister cells, dormancy and infectious disease. Nat. Rev. Microbiol. 2007, 5 (1), 48-56. 68. Madsen, J. S.; Burmølle, M.; Hansen, L. H.; Sørensen, S. J., The interconnection between biofilm formation and horizontal gene transfer. FEMS Immunology & Medical Microbiology 2012, 65 (2), 183-195. 69. Cottarel, G.; Wierzbowski, J., Combination drugs, an emerging option for antibacterial therapy. Trends Biotechnol. 2007, 25 (12), 547-555. 70. Projan, S. J., Why is big Pharma getting out of antibacterial drug discovery? Curr. Opin. Microbiol. 2003, 6 (5), 427-430. 71. Worthington, R. J.; Melander, C., Combination approaches to combat multidrug- resistant bacteria. Trends Biotechnol. 2013, 31 (3), 177-184. 72. Fischbach, M. A., Combination therapies for combating antimicrobial resistance. Curr. Opin. Microbiol. 2011, 14 (5), 519-523. 73. Tyers, M.; Wright, G. D., Drug combinations: a strategy to extend the life of antibiotics in the 21st century. Nat. Rev. Microbiol. 2019, 17 (3), 141-155. 74. Tamma, P. D.; Cosgrove, S. E.; Maragakis, L. L., Combination Therapy for Treatment of Infections with Gram-Negative Bacteria. Clin. Microbiol. Rev. 2012, 25 (3), 450-470. 75. Kortright, K. E.; Chan, B. K.; Koff, J. L.; Turner, P. E., Phage Therapy: A Renewed Approach to Combat Antibiotic-Resistant Bacteria. Cell Host Microbe 2019, 25 (2), 219-232. 76. Levin, B. R.; Bull, J. J., Population and evolutionary dynamics of phage therapy. Nat. Rev. Microbiol. 2004, 2 (2), 166-173. 77. Hua, Y.; Luo, T.; Yang, Y.; Dong, D.; Wang, R.; Wang, Y.; Xu, M.; Guo, X.; Hu, F.; He, P., Phage Therapy as a Promising New Treatment for Lung Infection Caused by Carbapenem-Resistant Acinetobacter baumannii in Mice. Front. Microbiol. 2018, 8 (2659). 78. Leppänen, M.; Maasilta, I. J.; Sundberg, L.-R., Antibacterial Efficiency of Surface-Immobilized Flavobacterium-Infecting Bacteriophage. ACS Appl. Bio Mater. 2019, 2 (11), 4720-4727.

Page | 114 CHAPTER TWO

79. Hamblin, M. R.; Hasan, T., Photodynamic therapy: a new antimicrobial approach to infectious disease? Photochem. Photobiol. Sci 2004, 3 (5), 436-450. 80. Jori, G.; Fabris, C.; Soncin, M.; Ferro, S.; Coppellotti, O.; Dei, D.; Fantetti, L.; Chiti, G.; Roncucci, G., Photodynamic therapy in the treatment of microbial infections: Basic principles and perspective applications. Lasers Surg Med 2006, 38 (5), 468-481. 81. Cieplik, F.; Deng, D.; Crielaard, W.; Buchalla, W.; Hellwig, E.; Al-Ahmad, A.; Maisch, T., Antimicrobial photodynamic therapy – what we know and what we don’t. Crit. Rev. Microbiol. 2018, 44 (5), 571-589. 82. Istanbullu, O.; Babauta, J.; Duc Nguyen, H.; Beyenal, H., Electrochemical biofilm control: mechanism of action. Biofouling 2012, 28 (8), 769-778. 83. Freebairn, D.; Linton, D.; Harkin-Jones, E.; Jones, D. S.; Gilmore, B. F.; Gorman, S. P., Electrical methods of controlling bacterial adhesion and biofilm on device surfaces. Expert Rev. Med. Devices 2013, 10 (1), 85-103. 84. Del Pozo, J. L.; Rouse, M. S.; Patel, R., Bioelectric Effect and Bacterial Biofilms. a Systematic Review. NT J ARTIF ORGANS 2008, 31 (9), 786-795. 85. LuTheryn, G.; Glynne-Jones, P.; Webb, J. S.; Carugo, D., Ultrasound-mediated therapies for the treatment of biofilms in chronic wounds: a review of present knowledge. Microb. Biotechnol. 2019, 13 (3), 613-628. 86. Bartley, J.; Young, D., Ultrasound as a treatment for chronic rhinosinusitis. Med. Hypotheses 2009, 73 (1), 15-17. 87. Dong, Y.; Li, J.; Li, P.; Yu, J., Ultrasound Microbubbles Enhance the Activity of Vancomycin Against Staphylococcus epidermidis Biofilms In Vivo. J. Ultrasound Med. 2018, 37 (6), 1379-1387. 88. Hetrick, E. M.; Schoenfisch, M. H., Reducing implant-related infections: active release strategies. Chem. Soc. Rev. 2006, 35 (9), 780-789. 89. Cheng, G.; Zhang, Z.; Chen, S.; Bryers, J. D.; Jiang, S., Inhibition of bacterial adhesion and biofilm formation on zwitterionic surfaces. Biomaterials 2007, 28 (29), 4192-4199. 90. Campoccia, D.; Montanaro, L.; Arciola, C. R., A review of the biomaterials technologies for infection-resistant surfaces. Biomaterials 2013, 34 (34), 8533-8554. 91. Teughels, W.; Van Assche, N.; Sliepen, I.; Quirynen, M., Effect of material characteristics and/or surface topography on biofilm development. Clin. Oral Implants Res. 2006, 17 (S2), 68-81. 92. Pawlowski, A. C.; Johnson, J. W.; Wright, G. D., Evolving medicinal chemistry strategies in antibiotic discovery. Curr. Opin. Biotechnol. 2016, 42, 108-117. 93. Coates, A. R. M.; Halls, G.; Hu, Y., Novel classes of antibiotics or more of the same? Br. J. Pharmacol. 2011, 163 (1), 184-194. 94. González-Bello, C., Antibiotic adjuvants – A strategy to unlock bacterial resistance to antibiotics. Bioorg. Med. Chem. Lett. 2017, 27 (18), 4221-4228. 95. Wright, G. D., Antibiotic Adjuvants: Rescuing Antibiotics from Resistance. Trends Microbiol. 2016, 24 (11), 862-871. 96. Gill, E. E.; Franco, O. L.; Hancock, R. E. W., Antibiotic Adjuvants: Diverse Strategies for Controlling Drug-Resistant Pathogens. Chem. Biol. Drug Des. 2015, 85 (1), 56-78. 97. Silva, L. N.; Zimmer, K. R.; Macedo, A. J.; Trentin, D. S., Plant Natural Products Targeting Bacterial Virulence Factors. Chem. Rev. 2016, 116 (16), 9162-9236. 98. Martelli, G.; Giacomini, D., Antibacterial and antioxidant activities for natural and synthetic dual-active compounds. Eur. J. Med. Chem. 2018, 158, 91-105.

Page | 115 CHAPTER TWO

99. Gyawali, R.; Ibrahim, S. A., Natural products as antimicrobial agents. Food Control 2014, 46, 412-429. 100. Zasloff, M., Antimicrobial peptides of multicellular organisms. Nature 2002, 415 (6870), 389-395. 101. Ganz, T., Defensins: antimicrobial peptides of innate immunity. Nat. Rev. Immunol. 2003, 3 (9), 710-720. 102. Brown, K. L.; Hancock, R. E. W., Cationic host defense (antimicrobial) peptides. Curr. Opin. Immunol. 2006, 18 (1), 24-30. 103. Kumar, P.; Kizhakkedathu, N. J.; Straus, K. S., Antimicrobial Peptides: Diversity, Mechanism of Action and Strategies to Improve the Activity and Biocompatibility In Vivo. Biomolecules 2018, 8 (1), 1-24. 104. Xie, S.-X.; Song, L.; Yuca, E.; Boone, K.; Sarikaya, R.; VanOosten, S. K.; Misra, A.; Ye, Q.; Spencer, P.; Tamerler, C., Antimicrobial Peptide–Polymer Conjugates for Dentistry. ACS Appl. Polym. Mater. 2020, 2 (3), 1134-1144. 105. Jain, A.; Duvvuri, L. S.; Farah, S.; Beyth, N.; Domb, A. J.; Khan, W., Antimicrobial Polymers. Adv. Healthc. Mater. 2014, 3 (12), 1969-1985. 106. Konai, M. M.; Bhattacharjee, B.; Ghosh, S.; Haldar, J., Recent Progress in Polymer Research to Tackle Infections and Antimicrobial Resistance. Biomacromolecules 2018, 19 (6), 1888-1917. 107. Kuroda, K.; Caputo, G. A., Antimicrobial polymers as synthetic mimics of host- defense peptides. WIRES Nanomed Nanobiotechnol 2013, 5 (1), 49-66. 108. Timofeeva, L.; Kleshcheva, N., Antimicrobial polymers: mechanism of action, factors of activity, and applications. Appl. Microbiol. Biotechnol. 2011, 89 (3), 475-492. 109. Takahashi, H.; Palermo, E. F.; Yasuhara, K.; Caputo, G. A.; Kuroda, K., Molecular Design, Structures, and Activity of Antimicrobial Peptide-Mimetic Polymers. Macromol. Biosci. 2013, 13 (10), 1285-1299. 110. Ergene, C.; Yasuhara, K.; Palermo, E. F., Biomimetic antimicrobial polymers: recent advances in molecular design. Polym. Chem. 2018, 9 (18), 2407-2427. 111. Lam, S. J.; Wong, E. H. H.; Boyer, C.; Qiao, G. G., Antimicrobial polymeric nanoparticles. Prog. Polym. Sci. 2018, 76, 40-64. 112. Gao, W.; Thamphiwatana, S.; Angsantikul, P.; Zhang, L., Nanoparticle approaches against bacterial infections. WIREs Nanomed Nanobiotechnol 2014, 6 (6), 532-547. 113. Hajipour, M. J.; Fromm, K. M.; Akbar Ashkarran, A.; Jimenez de Aberasturi, D.; Larramendi, I. R. d.; Rojo, T.; Serpooshan, V.; Parak, W. J.; Mahmoudi, M., Antibacterial properties of nanoparticles. Trends Biotechnol. 2012, 30 (10), 499-511. 114. Gupta, A.; Mumtaz, S.; Li, C.-H.; Hussain, I.; Rotello, V. M., Combatting antibiotic-resistant bacteria using nanomaterials. Chem. Soc. Rev. 2019, 48 (2), 415-427. 115. Huh, A. J.; Kwon, Y. J., “Nanoantibiotics”: A new paradigm for treating infectious diseases using nanomaterials in the antibiotics resistant era. J. Control. Release 2011, 156 (2), 128-145. 116. Carpenter, A. W.; Schoenfisch, M. H., Nitric oxide release: Part II. Therapeutic applications. Chem. Soc. Rev. 2012, 41 (10), 3742-3752. 117. Wo, Y.; Brisbois, E. J.; Bartlett, R. H.; Meyerhoff, M. E., Recent advances in thromboresistant and antimicrobial polymers for biomedical applications: just say yes to nitric oxide (NO). Biomaterials Science 2016, 4 (8), 1161-1183. 118. Fleming, D.; Rumbaugh, K. P., Approaches to Dispersing Medical Biofilms. 2017, 5 (2), 15. 119. Allison, K. R.; Brynildsen, M. P.; Collins, J. J., Metabolite-enabled eradication of bacterial persisters by aminoglycosides. Nature 2011, 473 (7346), 216-220.

Page | 116 CHAPTER TWO

120. Zhang, Y.; Yew, W. W.; Barer, M. R., Targeting Persisters for Tuberculosis Control. Antimicrob. Agents Chemother 2012, 56 (5), 2223. 121. Conlon, B. P.; Rowe, S. E.; Lewis, K., Persister Cells in Biofilm Associated Infections. In Biofilm-based Healthcare-associated Infections: Volume II, Donelli, G., Ed. Springer International Publishing: Cham, 2015; pp 1-9. 122. Galloway, W. R. J. D.; Hodgkinson, J. T.; Bowden, S. D.; Welch, M.; Spring, D. R., Quorum Sensing in Gram-Negative Bacteria: Small-Molecule Modulation of AHL and AI-2 Quorum Sensing Pathways. Chem. Rev. 2011, 111 (1), 28-67. 123. Kalia, V. C., Quorum sensing inhibitors: An overview. Biotechnol. Adv. 2013, 31 (2), 224-245. 124. Rasmussen, T. B.; Givskov, M., Quorum sensing inhibitors: a bargain of effects. Microbiology 2006, 152 (4), 895-904. 125. Fong, J.; Zhang, C.; Yang, R.; Boo, Z. Z.; Tan, S. K.; Nielsen, T. E.; Givskov, M.; Liu, X.-W.; Bin, W.; Su, H.; Yang, L., Combination Therapy Strategy of Quorum Quenching and Quorum Sensing Inhibitor in Suppressing Multiple Quorum Sensing Pathways of P. aeruginosa. Sci. Rep. 2018, 8 (1), 1155. 126. Amara, N.; Mashiach, R.; Amar, D.; Krief, P.; Spieser, S. A. H.; Bottomley, M. J.; Aharoni, A.; Meijler, M. M., Covalent Inhibition of Bacterial Quorum Sensing. J. Am. Chem. Soc. 2009, 131 (30), 10610-10619. 127. Boyer, C.; Bulmus, V.; Davis, T. P.; Ladmiral, V.; Liu, J.; Perrier, S., Bioapplications of RAFT Polymerization. Chem. Rev. 2009, 109 (11), 5402-5436. 128. Matyjaszewski, K.; Xia, J., Atom Transfer Radical Polymerization. Chem. Rev. 2001, 101 (9), 2921-2990. 129. Sovadinova, I.; Palermo, E. F.; Urban, M.; Mpiga, P.; Caputo, G. A.; Kuroda, K., Activity and Mechanism of Antimicrobial Peptide-Mimetic Amphiphilic Polymethacrylate Derivatives. Polymers 2011, 3 (3), 1512-1532. 130. Ganewatta, M. S.; Tang, C., Controlling macromolecular structures towards effective antimicrobial polymers. Polymer 2015, 63, A1-A29. 131. Judzewitsch, P. R.; Corrigan, N.; Trujillo, F.; Xu, J.; Moad, G.; Hawker, C. J.; Wong, E. H. H.; Boyer, C., High-Throughput Process for the Discovery of Antimicrobial Polymers and Their Upscaled Production via Flow Polymerization. 2020, 53 (2), 631-639. 132. Judzewitsch, P. R.; Zhao, L.; Wong, E. H. H.; Boyer, C., High-Throughput Synthesis of Antimicrobial Copolymers and Rapid Evaluation of Their Bioactivity. Macromolecules 2019, 52 (11), 3975-3986. 133. Kenawy, E.-R.; Worley, S. D.; Broughton, R., The Chemistry and Applications of Antimicrobial Polymers: A State-of-the-Art Review. Biomacromolecules 2007, 8 (5), 1359-1384. 134. Hancock, R. E. W.; Sahl, H.-G., Antimicrobial and host-defense peptides as new anti-infective therapeutic strategies. Nat. Biotechnol. 2006, 24 (12), 1551-1557. 135. Brogden, K. A.; Ackermann, M.; McCray, P. B.; Tack, B. F., Antimicrobial peptides in animals and their role in host defences. Int. J. Antimicrob. Agents 2003, 22 (5), 465-478. 136. Palermo, E. F.; Lee, D.-K.; Ramamoorthy, A.; Kuroda, K., Role of Cationic Group Structure in Membrane Binding and Disruption by Amphiphilic Copolymers. J. Phys. Chem. B 2011, 115 (2), 366-375. 137. Palermo, E. F.; Kuroda, K., Chemical Structure of Cationic Groups in Amphiphilic Polymethacrylates Modulates the Antimicrobial and Hemolytic Activities. Biomacromolecules 2009, 10 (6), 1416-1428.

Page | 117 CHAPTER TWO

138. Paslay, L. C.; Abel, B. A.; Brown, T. D.; Koul, V.; Choudhary, V.; McCormick, C. L.; Morgan, S. E., Antimicrobial Poly(methacrylamide) Derivatives Prepared via Aqueous RAFT Polymerization Exhibit Biocidal Efficiency Dependent upon Cation Structure. Biomacromolecules 2012, 13 (8), 2472-2482. 139. Phillips, D. J.; Harrison, J.; Richards, S.-J.; Mitchell, D. E.; Tichauer, E.; Hubbard, A. T. M.; Guy, C.; Hands-Portman, I.; Fullam, E.; Gibson, M. I., Evaluation of the Antimicrobial Activity of Cationic Polymers against Mycobacteria: Toward Antitubercular Macromolecules. Biomacromolecules 2017, 18 (5), 1592-1599. 140. Locock, K. E. S.; Michl, T. D.; Valentin, J. D. P.; Vasilev, K.; Hayball, J. D.; Qu, Y.; Traven, A.; Griesser, H. J.; Meagher, L.; Haeussler, M., Guanylated Polymethacrylates: A Class of Potent Antimicrobial Polymers with Low Hemolytic Activity. Biomacromolecules 2013, 14 (11), 4021-4031. 141. Gabriel, G. J.; Madkour, A. E.; Dabkowski, J. M.; Nelson, C. F.; Nüsslein, K.; Tew, G. N., Synthetic Mimic of Antimicrobial Peptide with Nonmembrane-Disrupting Antibacterial Properties. Biomacromolecules 2008, 9 (11), 2980-2983. 142. Chin, W.; Zhong, G.; Pu, Q.; Yang, C.; Lou, W.; De Sessions, P. F.; Periaswamy, B.; Lee, A.; Liang, Z. C.; Ding, X.; Gao, S.; Chu, C. W.; Bianco, S.; Bao, C.; Tong, Y. W.; Fan, W.; Wu, M.; Hedrick, J. L.; Yang, Y. Y., A macromolecular approach to eradicate multidrug resistant bacterial infections while mitigating drug resistance onset. Nat. Commun. 2018, 9 (1), 1-4. 143. Kuroda, K.; Caputo, G. A.; DeGrado, W. F., The Role of Hydrophobicity in the Antimicrobial and Hemolytic Activities of Polymethacrylate Derivatives. Chem. Eur. J. 2009, 15 (5), 1123-1133. 144. Engler, A. C.; Wiradharma, N.; Ong, Z. Y.; Coady, D. J.; Hedrick, J. L.; Yang, Y.-Y., Emerging trends in macromolecular antimicrobials to fight multi-drug-resistant infections. Nano Today 2012, 7 (3), 201-222. 145. Venkataraman, S.; Zhang, Y.; Liu, L.; Yang, Y.-Y., Design, syntheses and evaluation of hemocompatible pegylated-antimicrobial polymers with well-controlled molecular structures. Biomaterials 2010, 31 (7), 1751-1756. 146. Chin, W.; Yang, C.; Ng, V. W. L.; Huang, Y.; Cheng, J.; Tong, Y. W.; Coady, D. J.; Fan, W.; Hedrick, J. L.; Yang, Y. Y., Biodegradable Broad-Spectrum Antimicrobial Polycarbonates: Investigating the Role of Chemical Structure on Activity and Selectivity. Macromolecules 2013, 46 (22), 8797-8807. 147. Palermo, E. F.; Lienkamp, K.; Gillies, E. R.; Ragogna, P. J., Antibacterial Activity of Polymers: Discussions on the Nature of Amphiphilic Balance. Angew. Chem. Int. Ed. 2019, 58 (12), 3690-3693. 148. Cuthbert, T. J.; Hisey, B.; Harrison, T. D.; Trant, J. F.; Gillies, E. R.; Ragogna, P. J., Surprising Antibacterial Activity and Selectivity of Hydrophilic Polyphosphoniums Featuring Sugar and Hydroxy Substituents. Angew. Chem. Int. Ed. 2018, 57 (39), 12707- 12710. 149. Kurowska, M.; Eickenscheidt, A.; Guevara-Solarte, D.-L.; Widyaya, V. T.; Marx, F.; Al-Ahmad, A.; Lienkamp, K., A Simultaneously Antimicrobial, Protein- Repellent, and Cell-Compatible Polyzwitterion Network. Biomacromolecules 2017, 18 (4), 1373-1386. 150. Ergene, C.; Palermo, E. F., Self-immolative polymers with potent and selective antibacterial activity by hydrophilic side chain grafting. J. Mater. Chem. B 2018, 6 (44), 7217-7229. 151. Ergene, C.; Palermo, E. F., Cationic Poly(benzyl ether)s as Self-Immolative Antimicrobial Polymers. Biomacromolecules 2017, 18 (10), 3400-3409.

Page | 118 CHAPTER TWO

152. Allison, B. C.; Applegate, B. M.; Youngblood, J. P., Hemocompatibility of Hydrophilic Antimicrobial Copolymers of Alkylated 4-Vinylpyridine. Biomacromolecules 2007, 8 (10), 2995-2999. 153. Mortazavian, H.; Foster, L. L.; Bhat, R.; Patel, S.; Kuroda, K., Decoupling the Functional Roles of Cationic and Hydrophobic Groups in the Antimicrobial and Hemolytic Activities of Methacrylate Random Copolymers. Biomacromolecules 2018, 19 (11), 4370-4378. 154. Sellenet, P. H.; Allison, B.; Applegate, B. M.; Youngblood, J. P., Synergistic Activity of Hydrophilic Modification in Antibiotic Polymers. Biomacromolecules 2007, 8 (1), 19-23. 155. Punia, A.; Mancuso, A.; Banerjee, P.; Yang, N.-L., Nonhemolytic and Antibacterial Acrylic Copolymers with Hexamethyleneamine and Poly(ethylene glycol) Side Chains. ACS Macro Lett. 2015, 4 (4), 426-430. 156. Yang, X.; Hu, K.; Hu, G.; Shi, D.; Jiang, Y.; Hui, L.; Zhu, R.; Xie, Y.; Yang, L., Long Hydrophilic-and-Cationic Polymers: A Different Pathway toward Preferential Activity against Bacterial over Mammalian Membranes. Biomacromolecules 2014, 15 (9), 3267-3277. 157. Lam, S. J.; Wong, E. H. H.; O’Brien-Simpson, N. M.; Pantarat, N.; Blencowe, A.; Reynolds, E. C.; Qiao, G. G., Bionano Interaction Study on Antimicrobial Star- Shaped Peptide Polymer Nanoparticles. ACS Applied Materials & Interfaces 2016, 8 (49), 33446-33456. 158. Nguyen, T.-K.; Lam, S. J.; Ho, K. K. K.; Kumar, N.; Qiao, G. G.; Egan, S.; Boyer, C.; Wong, E. H. H., Rational Design of Single-Chain Polymeric Nanoparticles That Kill Planktonic and Biofilm Bacteria. ACS Infectious Diseases 2017, 3 (3), 237-248. 159. Lienkamp, K.; Kumar, K.-N.; Som, A.; Nüsslein, K.; Tew, G. N., “Doubly Selective” Antimicrobial Polymers: How Do They Differentiate between Bacteria? Chem. Eur. J. 2009, 15 (43), 11710-11714. 160. Hancock, R. E. W.; Lehrer, R., Cationic peptides: a new source of antibiotics. Trends Biotechnol. 1998, 16 (2), 82-88. 161. Lienkamp, K.; Madkour, A. E.; Musante, A.; Nelson, C. F.; Nüsslein, K.; Tew, G. N., Antimicrobial Polymers Prepared by ROMP with Unprecedented Selectivity: A Molecular Construction Kit Approach. J. Am. Chem. Soc. 2008, 130 (30), 9836-9843. 162. Mowery, B. P.; Lindner, A. H.; Weisblum, B.; Stahl, S. S.; Gellman, S. H., Structure−activity Relationships among Random Nylon-3 Copolymers That Mimic Antibacterial Host-Defense Peptides. J. Am. Chem. Soc. 2009, 131 (28), 9735-9745. 163. Sambhy, V.; Peterson, B. R.; Sen, A., Antibacterial and Hemolytic Activities of Pyridinium Polymers as a Function of the Spatial Relationship between the Positive Charge and the Pendant Alkyl Tail. Angew. Chem. Int. Ed. 2008, 47 (7), 1250-1254. 164. Gabriel, G. J.; Maegerlein, J. A.; Nelson, C. F.; Dabkowski, J. M.; Eren, T.; Nüsslein, K.; Tew, G. N., Comparison of Facially Amphiphilic versus Segregated Monomers in the Design of Antibacterial Copolymers. Chem. Eur. J. 2009, 15 (2), 433- 439. 165. Takahashi, H.; Caputo, G. A.; Vemparala, S.; Kuroda, K., Synthetic Random Copolymers as a Molecular Platform To Mimic Host-Defense Antimicrobial Peptides. Bioconjugate Chem. 2017, 28 (5), 1340-1350. 166. Palermo, E. F.; Vemparala, S.; Kuroda, K., Cationic Spacer Arm Design Strategy for Control of Antimicrobial Activity and Conformation of Amphiphilic Methacrylate Random Copolymers. Biomacromolecules 2012, 13 (5), 1632-1641.

Page | 119 CHAPTER TWO

167. Punia, A.; He, E.; Lee, K.; Banerjee, P.; Yang, N.-L., Cationic amphiphilic non- hemolytic polyacrylates with superior antibacterial activity. Chem. Commun. 2014, 50 (53), 7071-7074. 168. Michl, T. D.; Locock, K. E. S.; Stevens, N. E.; Hayball, J. D.; Vasilev, K.; Postma, A.; Qu, Y.; Traven, A.; Haeussler, M.; Meagher, L.; Griesser, H. J., RAFT- derived antimicrobial polymethacrylates: elucidating the impact of end-groups on activity and cytotoxicity. Polymer Chemistry 2014, 5 (19), 5813-5822. 169. Chen, C. Z.; Cooper, S. L., Interactions between dendrimer and bacterial membranes. Biomaterials 2002, 23 (16), 3359-3368. 170. Waschinski, C. J.; Tiller, J. C., Poly(oxazoline)s with Telechelic Antimicrobial Functions. Biomacromolecules 2005, 6 (1), 235-243. 171. Benski, L.; Tiller, J. C., Telechelic biocidal poly(2-oxazoline)s and polycations. Eur. Polym. J. 2019, 120, 109233. 172. Waschinski, C. J.; Herdes, V.; Schueler, F.; Tiller, J. C., Influence of Satellite Groups on Telechelic Antimicrobial Functions of Polyoxazolines. Macromol. Biosci. 2005, 5 (2), 149-156. 173. Braunecker, W. A.; Matyjaszewski, K., Controlled/living radical polymerization: Features, developments, and perspectives. Prog. Polym. Sci. 2007, 32 (1), 93-146. 174. Kuroki, A.; Sangwan, P.; Qu, Y.; Peltier, R.; Sanchez-Cano, C.; Moat, J.; Dowson, C. G.; Williams, E. G. L.; Locock, K. E. S.; Hartlieb, M.; Perrier, S., Sequence Control as a Powerful Tool for Improving the Selectivity of Antimicrobial Polymers. ACS Appl. Mater. Interfaces 2017, 9 (46), 40117-40126. 175. Judzewitsch, P. R.; Nguyen, T.-K.; Shanmugam, S.; Wong, E. H. H.; Boyer, C., Towards Sequence-Controlled Antimicrobial Polymers: Effect of Polymer Block Order on Antimicrobial Activity. Angew. Chem. Int. Ed. 2018, 57 (17), 4559-4564. 176. Nederberg, F.; Zhang, Y.; Tan, J. P. K.; Xu, K.; Wang, H.; Yang, C.; Gao, S.; Guo, X. D.; Fukushima, K.; Li, L.; Hedrick, J. L.; Yang, Y.-Y., Biodegradable nanostructures with selective lysis of microbial membranes. Nat. Chem. 2011, 3 (5), 409- 414. 177. Liu, X.; Zhang, H.; Tian, Z.; Sen, A.; Allcock, H. R., Preparation of quaternized organic–inorganic hybrid brush polyphosphazene-co-poly[2-(dimethylamino)ethyl methacrylate] electrospun fibers and their antibacterial properties. Polym. Chem. 2012, 3 (8), 2082-2091. 178. Wong, E. H. H.; Khin, M. M.; Ravikumar, V.; Si, Z.; Rice, S. A.; Chan-Park, M. B., Modulating Antimicrobial Activity and Mammalian Cell Biocompatibility with Glucosamine-Functionalized Star Polymers. Biomacromolecules 2016, 17 (3), 1170- 1178. 179. Brogden, K. A., Antimicrobial peptides: pore formers or metabolic inhibitors in bacteria? Nat. Rev. Microbiol. 2005, 3 (3), 238-250. 180. Corrêa, J. A. F.; Evangelista, A. G.; Nazareth, T. d. M.; Luciano, F. B., Fundamentals on the molecular mechanism of action of antimicrobial peptides. Materialia 2019, 8, 100494. 181. Fairbanks, B. D.; Gunatillake, P. A.; Meagher, L., Biomedical applications of polymers derived by reversible addition – fragmentation chain-transfer (RAFT). Adv. Drug Del. Rev. 2015, 91, 141-152. 182. Ali, M.; Brocchini, S., Synthetic approaches to uniform polymers. Adv. Drug Del. Rev. 2006, 58 (15), 1671-1687. 183. Moad, G.; Rizzardo, E.; Thang, S. H., Living Radical Polymerization by the RAFT Process – A Third Update. Aust. J. Chem. 2012, 65 (8), 985-1076.

Page | 120 CHAPTER TWO

184. Chiefari, J.; Chong, Y. K.; Ercole, F.; Krstina, J.; Jeffery, J.; Le, T. P. T.; Mayadunne, R. T. A.; Meijs, G. F.; Moad, C. L.; Moad, G.; Rizzardo, E.; Thang, S. H., Living Free-Radical Polymerization by Reversible Addition−Fragmentation Chain Transfer: The RAFT Process. Macromolecules 1998, 31 (16), 5559-5562. 185. Moad, G.; Thang, S. H., RAFT Polymerization: Materials of The Future, Science of Today: Radical Polymerization–The Next Stage. Aust. J. Chem. 2009, 62 (11), 1379- 1381. 186. Moad, G.; Chiefari, J.; Chong, Y. K.; Krstina, J.; Mayadunne, R. T. A.; Postma, A.; Rizzardo, E.; Thang, S. H., Living free radical polymerization with reversible addition – fragmentation chain transfer (the life of RAFT). Polym. Int. 2000, 49 (9), 993- 1001. 187. Moad, G.; Rizzardo, E.; Thang, S. H., Living Radical Polymerization by the RAFT ProcessA First Update. Aust. J. Chem. 2006, 59 (10), 669-692. 188. Hill, M. R.; Carmean, R. N.; Sumerlin, B. S., Expanding the Scope of RAFT Polymerization: Recent Advances and New Horizons. Macromolecules 2015, 48 (16), 5459-5469. 189. Himmelsbach, A.; Schneider-Chaabane, A.; Lienkamp, K., Asymmetrically Substituted Poly(diitaconates) Obtained by Reversible Addition-Fragmentation Chain Transfer (RAFT) Polymerization: Synthesis, Copolymerization Parameters, and Antimicrobial Activity. Macromol. Chem. Phys. 2019, 220 (24), 1-10. 190. Wrońska, N.; Felczak, A.; Zawadzka, K.; Poszepczyńska, M.; Różalska, S.; Bryszewska, M.; Appelhans, D.; Lisowska, K., Poly(Propylene Imine) Dendrimers and Amoxicillin as Dual-Action Antibacterial Agents. Molecules 2015, 20 (10), 19330- 19342. 191. Roy, D.; Knapp, J. S.; Guthrie, J. T.; Perrier, S., Antibacterial Cellulose Fiber via RAFT Surface Graft Polymerization. Biomacromolecules 2008, 9 (1), 91-99. 192. Wang, B.; Xu, Q.; Ye, Z.; Liu, H.; Lin, Q.; Nan, K.; Li, Y.; Wang, Y.; Qi, L.; Chen, H., Copolymer Brushes with Temperature-Triggered, Reversibly Switchable Bactericidal and Antifouling Properties for Biomaterial Surfaces. ACS Appl. Mater. Interfaces 2016, 8 (40), 27207-27217. 193. Wang, B.; Ye, Z.; Tang, Y.; Han, Y.; Lin, Q.; Liu, H.; Chen, H.; Nan, K., Fabrication of nonfouling, bactericidal, and bacteria corpse release multifunctional surface through surface-initiated RAFT polymerization. Int J Nanomedicine 2016, 12, 111-125. 194. Brochado, A. R.; Telzerow, A.; Bobonis, J.; Banzhaf, M.; Mateus, A.; Selkrig, J.; Huth, E.; Bassler, S.; Zamarreño Beas, J.; Zietek, M.; Ng, N.; Foerster, S.; Ezraty, B.; Py, B.; Barras, F.; Savitski, M. M.; Bork, P.; Göttig, S.; Typas, A., Species-specific activity of antibacterial drug combinations. Nature 2018, 559 (7713), 259-263. 195. Walvekar, P.; Gannimani, R.; Govender, T., Combination drug therapy via nanocarriers against infectious diseases. Eur. J. Pharm. Sci. 2019, 127, 121-141. 196. Karaiskos, I.; Antoniadou, A.; Giamarellou, H., Combination therapy for extensively-drug resistant gram-negative bacteria. Expert Rev. Anti Infect. Ther. 2017, 15 (12), 1123-1140. 197. Hetrick, E. M.; Shin, J. H.; Stasko, N. A.; Johnson, C. B.; Wespe, D. A.; Holmuhamedov, E.; Schoenfisch, M. H., Bactericidal Efficacy of Nitric Oxide-Releasing Silica Nanoparticles. ACS Nano 2008, 2 (2), 235-246. 198. Arora, D. P.; Hossain, S.; Xu, Y.; Boon, E. M., Nitric Oxide Regulation of Bacterial Biofilms. Biochemistry 2015, 54 (24), 3717-3728. 199. Riccio, D. A.; Schoenfisch, M. H., Nitric oxide release: Part I. Macromolecular scaffolds. Chem. Soc. Rev. 2012, 41 (10), 3731-3741.

Page | 121 CHAPTER TWO

200. Quinn, J. F.; Whittaker, M. R.; Davis, T. P., Delivering nitric oxide with nanoparticles. J. Control. Release 2015, 205, 190-205. 201. Kim, J.; Saravanakumar, G.; Choi, H. W.; Park, D.; Kim, W. J., A platform for nitric oxide delivery. J. Mater. Chem. B 2014, 2 (4), 341-356. 202. Sun, B.; Slomberg, D. L.; Chudasama, S. L.; Lu, Y.; Schoenfisch, M. H., Nitric Oxide-Releasing Dendrimers as Antibacterial Agents. Biomacromolecules 2012, 13 (10), 3343-3354. 203. Park, J.; Kim, J.; Singha, K.; Han, D.-K.; Park, H.; Kim, W. J., Nitric oxide integrated polyethylenimine-based tri-block copolymer for efficient antibacterial activity. Biomaterials 2013, 34 (34), 8766-8775. 204. Worley, B. V.; Slomberg, D. L.; Schoenfisch, M. H., Nitric Oxide-Releasing Quaternary Ammonium-Modified Poly(amidoamine) Dendrimers as Dual Action Antibacterial Agents. Bioconjugate Chem. 2014, 25 (5), 918-927. 205. Worley, B. V.; Schilly, K. M.; Schoenfisch, M. H., Anti-Biofilm Efficacy of Dual-Action Nitric Oxide-Releasing Alkyl Chain Modified Poly(amidoamine) Dendrimers. Mol. Pharm. 2015, 12 (5), 1573-1583. 206. Yang, L.; Wang, X.; Suchyta, D. J.; Schoenfisch, M. H., Antibacterial Activity of Nitric Oxide-Releasing Hyperbranched Polyamidoamines. Bioconjugate Chem. 2018, 29 (1), 35-43. 207. Yang, L.; Jing, L.; Jiao, Y.; Wang, L.; Marchesan, J. T.; Offenbacher, S.; Schoenfisch, M. H., In Vivo Antibacterial Efficacy of Nitric Oxide-Releasing Hyperbranched Polymers against Porphyromonas gingivalis. Mol. Pharm. 2019, 16 (9), 4017-4023. 208. Liu, Q.; Singha, P.; Handa, H.; Locklin, J., Covalent Grafting of Antifouling Phosphorylcholine-Based Copolymers with Antimicrobial Nitric Oxide Releasing Polymers to Enhance Infection-Resistant Properties of Medical Device Coatings. Langmuir 2017, 33 (45), 13105-13113. 209. Khalil, H.; Chen, T.; Riffon, R.; Wang, R.; Wang, Z., Synergy between polyethylenimine and different families of antibiotics against a resistant clinical isolate of Pseudomonas aeruginosa. Antimicrob Agents Chemother 2008, 52 (5), 1635-1641. 210. Ng, V. W. L.; Ke, X.; Lee, A. L. Z.; Hedrick, J. L.; Yang, Y. Y., Synergistic Co-Delivery of Membrane-Disrupting Polymers with Commercial Antibiotics against Highly Opportunistic Bacteria. Adv. Mater. 2013, 25 (46), 6730-6736. 211. Tian, J.; Zhang, J.; Yang, J.; Du, L.; Geng, H.; Cheng, Y., Conjugated Polymers Act Synergistically with Antibiotics to Combat Bacterial Drug Resistance. ACS Appl. Mater. Interfaces 2017, 9 (22), 18512-18520. 212. He, M.; Xiao, H.; Zhou, Y.; Lu, P., Synthesis, characterization and antimicrobial activities of water-soluble amphiphilic copolymers containing ciprofloxacin and quaternary ammonium salts. J. Mater. Chem. B 2015, 3 (18), 3704-3713. 213. He, M.; Zhou, Y.; Xiao, H.; Lu, P., Amphiphilic cationic copolymers with ciprofloxacin: preparation and antimicrobial activities. New J. Chem. 2016, 40 (2), 1354- 1364. 214. Zhang, J.; Chen, Y. P.; Miller, K. P.; Ganewatta, M. S.; Bam, M.; Yan, Y.; Nagarkatti, M.; Decho, A. W.; Tang, C., Antimicrobial Metallopolymers and Their Bioconjugates with Conventional Antibiotics against Multidrug-Resistant Bacteria. J. Am. Chem. Soc. 2014, 136 (13), 4873-4876. 215. Yang, P.; Bam, M.; Pageni, P.; Zhu, T.; Chen, Y. P.; Nagarkatti, M.; Decho, A. W.; Tang, C., Trio Act of Boronolectin with Antibiotic-Metal Complexed Macromolecules toward Broad-Spectrum Antimicrobial Efficacy. ACS Infect. Dis. 2017, 3 (11), 845-853.

Page | 122 CHAPTER TWO

216. Felczak, A.; Zawadzka, K.; Wrońska, N.; Janaszewska, A.; Klajnert, B.; Bryszewska, M.; Appelhans, D.; Voit, B.; Lisowska, K., Enhancement of antimicrobial activity by co-administration of poly(propylene imine) dendrimers and nadifloxacin. New J. Chem. 2013, 37 (12), 4156-4162. 217. Mishra, M. K.; Kotta, K.; Hali, M.; Wykes, S.; Gerard, H. C.; Hudson, A. P.; Whittum-Hudson, J. A.; Kannan, R. M., PAMAM dendrimer-azithromycin conjugate nanodevices for the treatment of Chlamydia trachomatis infections. Nanomed. Nanotechnol. Biol. Med. 2011, 7 (6), 935-944. 218. Cheng, Y.; Qu, H.; Ma, M.; Xu, Z.; Xu, P.; Fang, Y.; Xu, T., Polyamidoamine (PAMAM) dendrimers as biocompatible carriers of quinolone antimicrobials: An in vitro study. Eur. J. Med. Chem. 2007, 42 (7), 1032-1038. 219. Dorman, H. J. D.; Deans, S. G., Antimicrobial agents from plants: antibacterial activity of plant volatile oils. J. Appl. Microbiol. 2000, 88 (2), 308-316. 220. Helander, I. M.; Alakomi, H.-L.; Latva-Kala, K.; Mattila-Sandholm, T.; Pol, I.; Smid, E. J.; Gorris, L. G. M.; von Wright, A., Characterization of the Action of Selected Essential Oil Components on Gram-Negative Bacteria. J. Agric. Food Chem. 1998, 46 (9), 3590-3595. 221. Chong, Y.-B.; Zhang, H.; Yue, C. Y.; Yang, J., Fabrication and Release Behavior of Microcapsules with Double-Layered Shell Containing Clove Oil for Antibacterial Applications. ACS Appl. Mater. Interfaces 2018, 10 (18), 15532-15541. 222. Jobdeedamrong, A.; Jenjob, R.; Crespy, D., Encapsulation and Release of Essential Oils in Functional Silica Nanocontainers. Langmuir 2018, 34 (44), 13235- 13243. 223. Amato, D. N.; Amato, D. V.; Mavrodi, O. V.; Martin, W. B.; Swilley, S. N.; Parsons, K. H.; Mavrodi, D. V.; Patton, D. L., Pro-Antimicrobial Networks via Degradable Acetals (PANDAs) Using Thiol–Ene Photopolymerization. ACS Macro Lett. 2017, 6 (2), 171-175. 224. Landis, R. F.; Gupta, A.; Lee, Y.-W.; Wang, L.-S.; Golba, B.; Couillaud, B.; Ridolfo, R.; Das, R.; Rotello, V. M., Cross-Linked Polymer-Stabilized Nanocomposites for the Treatment of Bacterial Biofilms. ACS Nano 2017, 11 (1), 946-952. 225. Landis, R. F.; Li, C.-H.; Gupta, A.; Lee, Y.-W.; Yazdani, M.; Ngernyuang, N.; Altinbasak, I.; Mansoor, S.; Khichi, M. A. S.; Sanyal, A.; Rotello, V. M., Biodegradable Nanocomposite Antimicrobials for the Eradication of Multidrug-Resistant Bacterial Biofilms without Accumulated Resistance. J. Am. Chem. Soc. 2018, 140 (19), 6176-6182. 226. Li, C.-H.; Chen, X.; Landis, R. F.; Geng, Y.; Makabenta, J. M.; Lemnios, W.; Gupta, A.; Rotello, V. M., Phytochemical-Based Nanocomposites for the Treatment of Bacterial Biofilms. ACS Infect. Dis. 2019, 5 (9), 1590-1596. 227. Sambhy, V.; MacBride, M. M.; Peterson, B. R.; Sen, A., Silver Bromide Nanoparticle/Polymer Composites: Dual Action Tunable Antimicrobial Materials. J. Am. Chem. Soc. 2006, 128 (30), 9798-9808. 228. Song, J.; Kang, H.; Lee, C.; Hwang, S. H.; Jang, J., Aqueous Synthesis of Silver Nanoparticle Embedded Cationic Polymer Nanofibers and Their Antibacterial Activity. ACS Appl. Mater. Interfaces 2012, 4 (1), 460-465. 229. Mei, L.; Lu, Z.; Zhang, X.; Li, C.; Jia, Y., Polymer-Ag Nanocomposites with Enhanced Antimicrobial Activity against Bacterial Infection. ACS Appl. Mater. Interfaces 2014, 6 (18), 15813-15821. 230. Guo, Q.; Zhao, Y.; Dai, X.; Zhang, T.; Yu, Y.; Zhang, X.; Li, C., Functional Silver Nanocomposites as Broad-Spectrum Antimicrobial and Biofilm-Disrupting Agents. ACS Appl. Mater. Interfaces 2017, 9 (20), 16834-16847.

Page | 123 CHAPTER TWO

231. Wang, R.; Wang, L.; Zhou, L.; Su, Y.; Qiu, F.; Wang, D.; Wu, J.; Zhu, X.; Yan, D., The effect of a branched architecture on the antimicrobial activity of poly(sulfone amines) and poly(sulfone amine)/silver nanocomposites. J. Mater. Chem. 2012, 22 (30), 15227-15234. 232. Dai, X.; Chen, X.; Zhao, J.; Zhao, Y.; Guo, Q.; Zhang, T.; Chu, C.; Zhang, X.; Li, C., Structure–Activity Relationship of Membrane-Targeting Cationic Ligands on a Silver Nanoparticle Surface in an Antibiotic-Resistant Antibacterial and Antibiofilm Activity Assay. ACS Appl. Mater. Interfaces 2017, 9 (16), 13837-13848. 233. Amin, R. M.; Mohamed, M. B.; Ramadan, M. A.; Verwanger, T.; Krammer, B., Rapid and sensitive microplate assay for screening the effect of silver and gold nanoparticles on bacteria. Nanomedicine 2009, 4 (6), 637-643. 234. Allahverdiyev, A. M.; Kon, K. V.; Abamor, E. S.; Bagirova, M.; Rafailovich, M., Coping with antibiotic resistance: combining nanoparticles with antibiotics and other antimicrobial agents. Expert Rev. Anti Infect. Ther. 2011, 9 (11), 1035-1052. 235. Mei, L.; Zhang, X.; Wang, Y.; Zhang, W.; Lu, Z.; Luo, Y.; Zhao, Y.; Li, C., Multivalent polymer–Au nanocomposites with cationic surfaces displaying enhanced antimicrobial activity. Polym. Chem. 2014, 5 (8), 3038-3044. 236. Yuan, Y.; Liu, F.; Xue, L.; Wang, H.; Pan, J.; Cui, Y.; Chen, H.; Yuan, L., Recyclable Escherichia coli-Specific-Killing AuNP–Polymer (ESKAP) Nanocomposites. ACS Appl. Mater. Interfaces 2016, 8 (18), 11309-11317. 237. Richards, S.-J.; Isufi, K.; Wilkins, L. E.; Lipecki, J.; Fullam, E.; Gibson, M. I., Multivalent Antimicrobial Polymer Nanoparticles Target Mycobacteria and Gram- Negative Bacteria by Distinct Mechanisms. Biomacromolecules 2018, 19 (1), 256-264. 238. Yang, P.; Pageni, P.; Rahman, M. A.; Bam, M.; Zhu, T.; Chen, Y. P.; Nagarkatti, M.; Decho, A. W.; Tang, C., Gold Nanoparticles with Antibiotic- Metallopolymers toward Broad-Spectrum Antibacterial Effects. Adv. Healthc. Mater. 2019, 8 (6), 1800854. 239. Kong, H.; Song, J.; Jang, J., Photocatalytic Antibacterial Capabilities of TiO2−Biocidal Polymer Nanocomposites Synthesized by a Surface-Initiated Photopolymerization. Environ. Sci. Technol. 2010, 44 (14), 5672-5676. 240. Taylor, E.; Webster, T. J., Reducing infections through nanotechnology and nanoparticles. Int J Nanomedicine 2011, 6, 1463-1473. 241. Behera, S. S.; Patra, J. K.; Pramanik, K.; Panda, N.; Thatoi, H., Characterization and evaluation of antibacterial activities of chemically synthesized iron oxide nanoparticles. 2012. 242. Thomas, L. A.; Dekker, L.; Kallumadil, M.; Southern, P.; Wilson, M.; Nair, S. P.; Pankhurst, Q. A.; Parkin, I. P., -stabilised iron oxide nanoparticles for use in magnetic hyperthermia. J. Mater. Chem. 2009, 19 (36), 6529-6535. 243. Nguyen, T.-K.; Duong, H. T. T.; Selvanayagam, R.; Boyer, C.; Barraud, N., Iron oxide nanoparticle-mediated hyperthermia stimulates dispersal in bacterial biofilms and enhances antibiotic efficacy. Sci. Rep. 2015, 5 (1), 18385. 244. Kaittanis, C.; Naser, S. A.; Perez, J. M., One-Step, Nanoparticle-Mediated Bacterial Detection with Magnetic Relaxation. Nano Lett. 2007, 7 (2), 380-383. 245. Kaittanis, C.; Santra, S.; Perez, J. M., Emerging nanotechnology-based strategies for the identification of microbial pathogenesis. Adv. Drug Del. Rev. 2010, 62 (4), 408- 423. 246. Dong, H.; Huang, J.; Koepsel, R. R.; Ye, P.; Russell, A. J.; Matyjaszewski, K., Recyclable Antibacterial Magnetic Nanoparticles Grafted with Quaternized Poly(2- (dimethylamino)ethyl methacrylate) Brushes. Biomacromolecules 2011, 12 (4), 1305- 1311.

Page | 124 CHAPTER TWO

247. Jeong, C. J.; Sharker, S. M.; In, I.; Park, S. Y., Iron Oxide@PEDOT-Based Recyclable Photothermal Nanoparticles with Poly(vinylpyrrolidone) Sulfobetaines for Rapid and Effective Antibacterial Activity. ACS Appl. Mater. Interfaces 2015, 7 (18), 9469-9478. 248. Gupta, A. K.; Gupta, M., Synthesis and surface engineering of iron oxide nanoparticles for biomedical applications. Biomaterials 2005, 26 (18), 3995-4021. 249. Taresco, V.; Francolini, I.; Padella, F.; Bellusci, M.; Boni, A.; Innocenti, C.; Martinelli, A.; D'Ilario, L.; Piozzi, A., Design and characterization of antimicrobial usnic acid loaded-core/shell magnetic nanoparticles. Mater. Sci. Eng. C 2015, 52, 72-81. 250. Pu, L.; Xu, J.; Sun, Y.; Fang, Z.; Chan-Park, M. B.; Duan, H., Cationic polycarbonate-grafted superparamagnetic nanoparticles with synergistic dual-modality antimicrobial activity. Biomater. Sci. 2016, 4 (5), 871-879. 251. Pageni, P.; Yang, P.; Bam, M.; Zhu, T.; Chen, Y. P.; Decho, A. W.; Nagarkatti, M.; Tang, C., Recyclable magnetic nanoparticles grafted with antimicrobial metallopolymer-antibiotic bioconjugates. Biomaterials 2018, 178, 363-372. 252. Hong, G.; Diao, S.; Antaris, A. L.; Dai, H., Carbon Nanomaterials for Biological Imaging and Nanomedicinal Therapy. Chem. Rev. 2015, 115 (19), 10816-10906. 253. Liu, S.; Zeng, T. H.; Hofmann, M.; Burcombe, E.; Wei, J.; Jiang, R.; Kong, J.; Chen, Y., Antibacterial Activity of Graphite, Graphite Oxide, Graphene Oxide, and Reduced Graphene Oxide: Membrane and Oxidative Stress. ACS Nano 2011, 5 (9), 6971- 6980. 254. Kang, S.; Herzberg, M.; Rodrigues, D. F.; Elimelech, M., Antibacterial Effects of Carbon Nanotubes: Size Does Matter! Langmuir 2008, 24 (13), 6409-6413. 255. Zou, X.; Zhang, L.; Wang, Z.; Luo, Y., Mechanisms of the Antimicrobial Activities of Graphene Materials. J. Am. Chem. Soc. 2016, 138 (7), 2064-2077. 256. Pan, N.; Liu, Y.; Fan, X.; Jiang, Z.; Ren, X.; Liang, J., Preparation and characterization of antibacterial graphene oxide functionalized with polymeric N- halamine. J. Mater. Sci. 2017, 52 (4), 1996-2006. 257. Liu, T.; Liu, Y.; Liu, M.; Wang, Y.; He, W.; Shi, G.; Hu, X.; Zhan, R.; Luo, G.; Xing, M.; Wu, J., Synthesis of graphene oxide-quaternary ammonium nanocomposite with synergistic antibacterial activity to promote infected wound healing. Burns Trauma 2018, 6 (1). 258. Li, P.; Sun, S.; Dong, A.; Hao, Y.; Shi, S.; Sun, Z.; Gao, G.; Chen, Y., Developing of a novel antibacterial agent by functionalization of graphene oxide with guanidine polymer with enhanced antibacterial activity. Appl. Surf. Sci. 2015, 355, 446- 452. 259. Tu, Q.; Tian, C.; Ma, T.; Pang, L.; Wang, J., Click synthesis of quaternized poly(dimethylaminoethyl methacrylate) functionalized graphene oxide with improved antibacterial and antifouling ability. Colloids Surf. B. Biointerfaces 2016, 141, 196-205. 260. Murugan, E.; Vimala, G., Effective functionalization of multiwalled carbon nanotube with amphiphilic poly(propyleneimine) dendrimer carrying silver nanoparticles for better dispersability and antimicrobial activity. J. Colloid Interface Sci. 2011, 357 (2), 354-365. 261. Joo, Y. T.; Jung, K. H.; Kim, M. J.; Kim, Y., Preparation of antibacterial PDMAEMA-functionalized multiwalled carbon nanotube via atom transfer radical polymerization. J. Appl. Polym. Sci. 2013, 127 (3), 1508-1518. 262. Koromilas, N. D.; Lainioti, G. C.; Gialeli, C.; Barbouri, D.; Kouravelou, K. B.; Karamanos, N. K.; Voyiatzis, G. A.; Kallitsis, J. K., Preparation and Toxicological Assessment of Functionalized Carbon Nanotube-Polymer Hybrids. PLoS One 2014, 9 (9), 1-15.

Page | 125 CHAPTER TWO

263. Liu, J.; Shao, J.; Wang, Y.; Li, J.; Liu, H.; Wang, A.; Hui, A.; Chen, S., Antimicrobial Activity of Zinc Oxide–Graphene Quantum Dot Nanocomposites: Enhanced Adsorption on Bacterial Cells by Cationic Capping Polymers. ACS Sustain. Chem. Eng. 2019, 7 (19), 16264-16273. 264. Zhou, S.; Ji, H.; Fu, Y.; Yang, Y.; Lü, C., Mussel-inspired fabrication of cationic polymer modified rGO supported silver nanoparticles hybrid with robust antibacterial and catalytic reduction performance. Appl. Surf. Sci. 2020, 506, 144655. 265. Yuan, W.; Jiang, G.; Che, J.; Qi, X.; Xu, R.; Chang, M. W.; Chen, Y.; Lim, S. Y.; Dai, J.; Chan-Park, M. B., Deposition of Silver Nanoparticles on Multiwalled Carbon Nanotubes Grafted with Hyperbranched Poly(amidoamine) and Their Antimicrobial Effects. J. Phys. Chem. C. 2008, 112 (48), 18754-18759.

Page | 126

CHAPTER THREE

Nitric Oxide-Loaded Antimicrobial Polymer for The Synergistic Eradication of Bacterial Biofilm

CHAPTER THREE

3 Nitric Oxide-Loaded Antimicrobial Polymer for The

Synergistic Eradication of Bacterial Biofilm

This chapter is based on the work published in ACS Macro Letters,

2018, 7 (5),592–597.

3.1 Introduction

The formation of bacterial biofilms on the surfaces of body tissues and medical devices often leads to health complications such as chronic and recurrent infections.1-3 Treatment of biofilm-related infections is challenging4-7 because the bacteria cells are embedded in a polymeric self- secreted matrix which makes them highly resistant to hostile environmental conditions and antibiotic treatments.8-10 The failure of many commercial antibiotics, natural antimicrobial peptides, and synthetic antimicrobial agents in eradicating biofilms illustrates the urgent need for designing novel antimicrobial/antibiofilm agents.

Page | 128 CHAPTER THREE

Nitric oxide (NO), a diatomic gas molecule, has recently been proposed as a potential antibiofilm agent.11, 12 NO demonstrates excellent antimicrobial activity against both planktonic and biofilm bacteria with a low chance of resistance development.9, 13 In addition, NO has a dose-dependent antimicrobial activity where it disperses bacteria biofilms at low concentrations and kills at higher doses.11, 14-16 Although NO is a promising antimicrobial agent,10, 13 the potential clinical applications of NO (gas) are hindered by its instability in biological media and lack of controlled and localized delivery.17 To address these issues, the use of polymeric and non- polymeric vehicles have been considered.9, 18, 19 Various small molecule NO donors such as S-nitrosothiols (RSNOs)20, 21 and N- diazeniumdiolates (NONOates)22-26 have been encapsulated into or conjugated to polymerics scaffolds, resulting in improved stability and NO delivery compared to small molecule NO donors.

Previous studies have suggested that the co-administration of two or more mechanistically different antimicrobial agents can effectively combat bacterial infections by lowering the required treatment dose, decreasing the risk of resistance development and increasing the overall treatment efficicacy.23, 27-30 For example, the co-administration of NO with other antimicrobial agents such as synthetic antimicrobial polymers,31-33 silver-releasing materials,34, 35 and antibiotics23, 36, 37 has resulted in better antimicrobial performance than the individual compounds.

Encouraged by these early studies on NO co-administration systems, and on the basis of our recent experiences with NO delivery platforms22, 23 and antimicrobial polymer synthesis,38-41 we herein describe the development of a

Page | 129 CHAPTER THREE

novel NO- releasing antimicrobial/antibiofilm polymer for the synergistic eradication bacterial biofilm (Figure 3.1). In contrast to previous studies22, 23 where the polymer was only utilized to encapsulate NO donors, in this study, the polymer has an active role as antimicrobial agent as well as NO carrier. However, given that bacterial biofilm are generally more challenging to combat, it is our aim in this study to improve the antibiofilm activity by endowing the polymers with NO-releasing functional groups (i.e., NONOate).

Figure 3.1. General illustration depicting the dispersal and killing of bacteria biofilm using the NO-functionalized antimicrobial polymer in this study.

3.2 Experimental Section

Materials

Ethylenediamine (Sigma-Aldrich, ≥ 99%), di-tert-butyl dicarbonate (Aldrich,

99%), triethylamine (Scharlau, 99%), acryloyl chloride (Merck, ≥ 96%), oligoethylene glycol methyl ether acrylate (OEG acrylate) Mn 480 g mol–1

(Aldrich), 2-ethylhexyl acrylate (Aldrich, 98%), trifluoroacetic acid (TFA) (Sigma-

Aldrich, 99%), hexane (Merck), diethyl ether (Merck), 1,4-dioxane (Merck) and basic alumina (Al2O3) (LabChem) were used as received. 2,2’-azobis (2

Page | 130 CHAPTER THREE

methylpropionitrile) (AIBN) (Acros, 98%) was purified by recrystallization from methanol. Sodium sulfate (Na2SO4), magnesium sulfate (MgSO4), sodium hydrogen carbonate (NaHCO3), tetrahydrofuran, and acetone were obtained from Chem-Supply and used as received. Milli-Q water with a resistivity of > 18

MΩ⋅cm was obtained from an in-line Millipore RiOs/Origin water purification system.

Analytical instruments

1H Nuclear magnetic resonance (NMR) spectra were attained using a Bruker

AC300F spectrometer. Deuterated solvents D2O or CDCl3 (obtained from

Cambridge Isotope Laboratories) were used as reference solvents and samples with concentration of ca. 10-20 mg mL–1 were prepared.

Gel permeation chromatography (GPC) analysis was performed using a

Shimadzu liquid chromatography system equipped with a Shimadzu refractive index detector and two MIX C columns (Polymer Lab) operating at 40 °C.

Tetrahydrofuran was used as the eluent at a flow rate of 1 mL min–1. The system was calibrated with poly(methyl methacrylate) standards with molecular weights of 200 to 106 g mol–1.

Dynamic light scattering (DLS) and zeta-potential measurements were conducted using a Malvern Zetasizer Nano ZS apparatus equipped with a He-

Ne laser operated at λ = 633 nm and at a scattering angle of 173°. All samples were prepared at a concentration of ca. 2 mg mL–1 where filtered Milli-Q water

(using 0.45 μm pore size filter) was used as the solvent to solubilize the polymers.

Page | 131 CHAPTER THREE

Attenuated total reflectance-Fourier transform infrared spectroscopy (ATR-

FTIR) measurement of samples was performed using a Bruker IFS66/S Fourier transform spectrometer by averaging 128 scans with a resolution of 4 cm–1.

Polymer samples were pre-dried as thin films for ATR-FTIR analysis.

UV-Visible spectra were recorded in a quartz cuvette using a CARY 3000 spectrometer from Bruker at 25 °C.

X-ray photoelectron spectroscopy (XPS) measurements were performed on a

ESCALAB250Xi instrument (Thermo Scientific, UK) with a monochromated Al

Kα source (energy 1486 eV) at 120 W over 500 μm at a 90° angle. Background vacuum pressure was 2 × 10–9 mbar.

Transmission electron microscopy (TEM) was performed using a TEM

(CM200) at an accelerating voltage of 200 kV. Samples were prepared by placing a droplet of a 4 mg mL–1 polymer solution in water on a carbon-coated copper grid. Uranyl acetate staining was applied to increase contrast.

Synthesis of NO-loaded antimicrobial polymer

Synthesis of cationic monomer

Cationic monomer tert-butyl (4-acrylamidobutyl) carbamate was prepared in the same manner as reported previously.38 Briefly, ethylenediamine (0.3 mol) was dissolved in chloroform (400 mL), followed by the dropwise addition of di-tert- butyl dicarbonate (0.03 mol in 100 mL) over 2 h at 0−5 °C. The reaction mixture was stirred overnight at 25 °C. White precipitates were filtered, and the organic phase was washed exhaustively with water using a separation funnel to remove

Page | 132 CHAPTER THREE

excess diamines. The organic layer was then dehydrated over MgSO4, filtered, and dried using a rotary evaporation unit to yield a pale-yellow oil.

Tetrahydrofuran (100 mL) was added to dissolve the intermediate product.

Triethylamine (36 mmol) and acryloyl chloride (31.5 mmol) were added dropwise to the solution at 0−5 °C with N2 bubbling. The contents were stirred at

25 °C for 1 h. The byproducts were filtered, and the solvent was removed in vacuo. The crude product was dissolved in chloroform (150 mL) and washed against brine (1 × 75 mL). The organic phase was stirred with MgSO4 and basic

Al2O3 for 10 min, filtered, and concentrated in vacuo. The product was further purified by repeated precipitation steps in hexane to yield the tert- butyloxycarbonyl Boc- protected monomer as a fine white powder, which was dried in vacuo.

tert-Butyl (2-acrylamidoethyl) carbamate: 1H NMR (300MHz, CDCl3,25 °C), δH

6.56 (br s, 1H, NH), 6.28 (dd, J = 17.1 Hz, 1.5 Hz, 1H, CHH=CH), 6.12 (dd, J = 17.1

Hz, 10.2Hz, 1H, CHH=CH), 5.65 (dd, J = 10.2 Hz, 1.5 Hz, 1H, CHH=CH), 5.05 (br s, 1H, NH), 3.49−3.41 (m, 2H, CH2), 3.34−3.28 (m, 2H, CH2), 1.45 (s, 9H, CH3); 13C

NMR (300 MHz, CDCl3,25 °C), δC 166.23, 157.50, 130.88, 126.30, 79.85, 41.05, 40.09,

28.35.

Synthesis of amphiphilic ternary random copolymer

The synthesis of amphiphilic ternary statistical copolymer proceeded in the same manner as previously described.38 Briefly, The RAFT agent benzyl dodecyl carbonotrithioate (11.6 μmol), AIBN (4.6 μmol), OEGA (350 μmol), cationic monomer (580 μmol), and hydrophobic monomer (230 μmol) were dissolved in

Page | 133 CHAPTER THREE

1,4-dioxane (such that the total monomer concentration in solvent is 1 M). The solution was purged with N2 for 20 min in an ice bath. The polymerization was conducted for 20 h at 70 °C and then quenched in an ice bath with exposing to air. The polymer was purified by precipitation in a hexane/diethyl ether (7:3) mixture thrice and subsequently dried in vacuo. The monomer composition of the polymers was calculated using the following equation ʃa,b/6 : ʃc/9 : ʃd/2 where

ʃa,b, ʃc, and ʃd correspond to the integrals of the characteristic protons of 2- ethylhexyl acrylate (methyl -CH3- groups, δH 0.80-0.98 ppm), cationic monomer

(tert-butyl -CH3- groups, δH 1.38-1.52 ppm) and OEGA (ester -CH2O- groups, 4.10-

4.30 ppm), respectively (Figure A 1).

The Boc protecting groups were removed using TFA in the same manner as reported previously.38 In general, the polymer solution in dichloromethane (ca.

10 wt % polymer) was treated with TFA (20 mol equiv with respect to the Boc group) for 1.5 h at 25 °C. Boc- deprotected polymer was subsequently precipitated into diethyl ether/hexane (4:1). The precipitate was isolated by centrifugation, dissolved in methanol, and reprecipitated two more times. The polymer was then dried in vacuo and further purified by dialysis against water

(Cellu-Sep 3500 MWCO). The aqueous solution was lyophilized to yield the Boc- deprotected polymer (P).

Attachment of NONOate to ternary random copolymer

A solution of the Boc-deprotected polymer P in acetonitrile (30 mg mL–1) was placed in a Parr apparatus and clamped. The apparatus was sealed and purged with N2 three times, followed by purging with NO gas (25 °C, 5 atm, 24 h).

Page | 134 CHAPTER THREE

Afterwards, the solution was purged with N2 to remove the excess NO. The resultant NO-loaded cationic polymer (PNO) was stored at 4 °C.

In vitro NO release study

The NO released profile of PNO was determined using a standard Griess reagent kit.22, 23 Briefly, a solution of PNO in acetonitrile (2 mL of a 2 mg mL–1) was transferred to a dialysis membrane (Cellu-Sep 3500 MWCO). The dialysis membrane was immersed in 8 mL of PBS buffer (pH 7.4) and placed in an incubator at 37 °C. A 100 µL aliquot of the surrounding PBS solution was taken at different time points and mixed with 100 µL of Griess reagent and 1.8 mL of

PBS buffer. After 15 min incubation at room temperature, the UV-Vis absorbance of the resulting solutions was measured at 540 nm. NO concentration was subsequently calculated using a standard calibration curve.

Biofilm dispersal study

The laboratory strain P. aeruginosa PAO1 was used to grow biofilm. In all assays, a single colony of PAO1 was inoculated in 10 mL of Luria Bertani medium

(LB 10) at 37 oC with shaking at 200 rpm overnight. The overnight culture was diluted 1:200 in freshly prepared M9 minimal medium containing 48 mM

Na2HPO4, 22 mM KH2PO4, 9 mM NaCl, 19 mM NH4Cl, pH 7.0, supplemented with 2 mM MgSO4, 100 μM CaCl2 and 20 mM glucose. The bacterial suspension was then aliquoted 1 mL per well of tissue-culture treated 24-well plates (Costar,

Corning®). The plates were incubated at 37 °C with shaking at 180 rpm in an orbital shaker that does not stop agitation when the door is opened (model

OM11, Ratek, Boronia, Australia) and the biofilm cultures were allowed to grow

Page | 135 CHAPTER THREE

for 6.5 h without any disruption. To characterize the effect on biofilm dispersal, preformed PAO1 biofilms were treated with different compounds at various concentrations and incubation times. Biofilm biomass was quantified using the crystal violet (CV) staining method. Briefly, after treatment, the culture supernatant was removed and the biofilm on the well surfaces was washed once with 1 mL of PBS, followed by the addition of 1 mL 0.03% CV stain made from a

1:10 dilution of Gram crystal violet (BD) in PBS. The plates were incubated on the bench for 20 min before the wells were washed twice with PBS. The CV stained biofilms were mixed with 1 mL 100% ethanol and quantified by measuring the

OD595 of the homogenized suspension using a microtiter plate reader (FLUOstar

Omega, BMG Labtech). All assays included two replicates and were repeated in at least three independent experiments.

Killing study

To characterize the bactericidal activity, PAO1 biofilms were grown in the same manner as above. The compounds were then added to the wells and the plates were incubated for either 20 or 60 min. After treatment, the planktonic and biofilm viability analysis were determined by a drop plate method. For planktonic analysis, free-floating cells in the biofilm supernatant were serially diluted in sterile PBS and plated onto LB agar. For biofilm analysis, cells attached on the interior surfaces of the well (surface area 4.5 cm2) were washed twice with sterile PBS to remove loosely attached bacteria, before being resuspended and homogenized in PBS by incubating in an ultrasonication bath (150 W, 40 kHz;

Unisonics, Australia) for 20 min. Resuspended biofilm cells were then serially diluted and plated onto LB agar. Planktonic and biofilm colonies were counted

Page | 136 CHAPTER THREE

and CFU was calculated after 24 h incubation at 37 °C. All assays included two replicates and were repeated in at least three independent experiments.

Biofilm imaging

To visualize the effect of polymer on biofilm, PAO1 biofilms were grown on

35 mm tissue culture dishes (FluoroDish, World Precision Instruments Inc.,

Sarasota, FL, USA) in the same way as above. The compound was then added to the well and incubated for 1 h. After treatment, the supernatant was removed and the biofilm on the well surface was washed twice with 2 mL of PBS, followed by the addition of 1 mL PBS. The wells were analyzed with a 3D tomographic microscope (3D Cell Explorer, NanoLive, Lausanne, Switzerland) equipped with a digital staining software. All assays were repeated in at least two independent experiments.

3.3 Results and discussion

3.3.1 Synthesis of amphiphilic ternary random copolymer

For this, a linear amphiphilic statistical ternary copolymer (P) that contained

30 repeat units of biocompatible oligo- ethylene glycol, 20 repeat units of hydrophobic ethylhexyl, and 50 repeat units of primary amino groups was first prepared via reversible addition-fragmentation chain transfer (RAFT) polymerization,42, 43 followed by the removal of tert-butyloxycarbonyl (Boc) groups as reported previously (Scheme 3.1). This polymer composition is necessary to achieve optimal antimicrobial activity and biocompatibility.38, 39 1H

Page | 137 CHAPTER THREE

NMR analysis confirmed that the chemical composition of the synthesized polymer P was similar to the molar feed ratio. The number-averaged molecular weight (Mn) and dispersity (Đ) of the Boc-protected polymer were estimated to be 15000 g mol–1 and 1.3, respectively, based on gel permeation chromatography

(GPC) analysis (Figure A 2). After Boc-deprotection, zeta potential measurement revealed that polymer P has a net positive charge of +18.4 mV as expected due to the presence of cationic primary amino groups.

Scheme 3.1. Synthesis of NO-Modified Antimicrobial Polymer for Synergistic Eradication of

Bacteria Biofilm

3.3.2 Attachment of NONOate to ternary random copolymer

Functionalization of P with NONOate was achieved by sparging the polymer solution with NO gas, resulting in the NO-loaded polymer, PNO (Scheme 3.1).

The successful loading of NO was verified by FTIR analysis by the presence of two new peaks at 1510 and 1290 cm–1, which are attributed to the presence of

Page | 138 CHAPTER THREE

NONOate functional groups (NO donor molecules)22, 23 (Figure 3.2).

Furthermore, XPS analysis was employed to confirm the presence of NONOate moieties (Figure 3.3). While the primary amine and amide groups of P were observed at 399.7 and 401.5 eV, a new peak at 406 eV was observed in the PNO spectra which corresponds to the NONOate functionality, in accord with previous report (Figure 3.3).44 The size and morphology of P and PNO were also analysed by transmission electron microscopy (TEM). The TEM measurements which were performed in the dry state revealed that P has an average diameter of 12 ± 4 nm, while PNO formed larger aggregates with an average diameter of

130 ± 25 nm (Figure A 3). These aggregates may have been caused by the drying process.

Figure 3.2. FTIR spectra of P, fresh PNO, and PNO after 3 h of release study in PBS.

Page | 139 CHAPTER THREE

Figure 3.3. XPS spectra of PNO and P.

3.3.3 Determination of the release and stability of nitric oxide

Once the loading of NO was confirmed, the NO release profile of PNO was subsequently determined using a Griess assay.22, 23 Cumulative NO release was measured in phosphate buffered saline (PBS, pH 7.4) at 37 °C. The polymer PNO showed steady NO release over 60 min upon contact with aqueous media before reaching a plateau during which all stored NO was assumed to be released

(Figure 3.4), as supported by the absence of peaks corresponding to the

NONOate groups in FTIR and XPS spectra after 3 h (Figure A 4). The half-life of

NO release was about 12.3 min. Based on this assay, there are approximately seven NONOate groups per polymer chain, which corresponds to around 14% of primary amine converted into NONOate groups. The instability of NO is a well- known limit regarding the long-term storage and potential clinical application of

NO-containing systems. However, the use of small molecule NO donors in combination with polymeric systems can improve the stability and localised

Page | 140 CHAPTER THREE

delivery of NO. A stability study was also performed on PNO during a six-month storage period in acetonitrile at 4 °C to determine the stability of NONOates in the polymer. The total NO content decreased by 54 and 89% after three and six months of storage, respectively (Figure A 5). The storage of the NO-loaded polymer at –20 or –80 °C might improve the stability of NONOate groups. It is noteworthy that fresh PNO was used in all other experiments.

Figure 3.4. Cumulative release of NO from PNO in PBS buffer (pH 7.4, 37 oC) as measured using Griess assay.

3.3.4 Biofilm dispersal by NO-loaded polymer

To investigate the synergistic effect of PNO in combating bacterial biofilm, biofilm dispersal experiments were initially conducted. For this, Pseudomonas aeruginosa PAO1 biofilms were grown in cell culture media M9 for 6.5 h prior to incubation with selected compounds at different concentrations (16, 32 and 64 µg mL–1) for either 20 or 60 min. Specifically, the biofilm dispersal activity of PNO and other compounds which served as controls, i.e. (i) P, (ii) commercially available NO donor, spermine NONOate (SperNO), and (iii) cocktail mixture of

P and SperNO (P + SperNO), were compared. At both incubation times and in

Page | 141 CHAPTER THREE

all concentrations tested, PNO induced the best biofilm dispersal compared to the rest (Figure 3.5 a, b and Table A 1).

a) b)

Figure 3.5. P. aeruginosa biofilm biomass reduction upon treatment with various concentrations (16-64 μgmL−1) of P, PNO, SperNO, and P + SperNO for 20 min (a) and 60 min (b). Data are representative of at least three independent experiments ± SD. Student’s t- test – asterisks indicate statistically significant difference of PNO treatments versus P treatment (**, p < 0.01; ****, p < 0.0001; ns, non-significant).

Treatment for 20 min with PNO resulted in 70.8 ± 7.3, 70.0 ± 5.7 and 65.7 ± 7.4% reduction of biofilm biomass at polymer concentrations of 16, 32 and 64 µg mL–1, respectively, as determined by crystal violet (CV) staining relative to the untreated control biofilms (Figure 3.5). By increasing the incubation time to 60 min, a slight increase in biofilm biomass reduction was observed (Figure 3.5). For example, at 64 µgmL–1, the biomass reduction increased from 65.7 ± 7.4 to 80 ±

4.3%. In contrast, P exhibited a lower biofilm dispersal capability, for instance at

64 µgmL–1 only 54% biofilm dispersal was observed after 60 min. This value is consistent with our previous report.38 Whereas the biofilm dispersal ability of

PNO was just as effective across the concentrations tested, interestingly, P

Page | 142 CHAPTER THREE

showed a dose-dependent activity. Considering that the concentrations of NO molecules (7-30 µM) found at these polymer concentrations are usually insubstantial in dispersing biofilms alone, as proven by the lack of activity of

SperNO found here and in our previous studies.22, 23 The biofilm dispersal activity observed for PNO can be attributed to a synergistic effect between the released

NO molecules and the reformed antimicrobial polymer P. To confirm this assumption, we have made a physical mixture of P + SperNO which has the same polymer and NO concentration as PNO, and investigated the biofilm dispersal properties. In the case of this cocktail mixture (P + SperNO), the biofilm biomass reduction was similar to P alone under all conditions tested, which indicated the absence of synergy between P and SperNO. This strongly suggests that the conjugation of NO on antimicrobial polymer, to yield a single chemical entity

PNO, is important toward affording a synergistic antimicrobial effect on bacteria biofilms. This synergestic effect might be attributed to localized delivery of NO to the bacteria biofilm promoted by the electrostatic interaction between the cationic charges present on PNO and the anionic bacteria surface. This electrostatic interaction most likely resulted in the increase of localized NO concentration in the proximity of biofilm.

The biofilm biomass following 60 min of treatment at 64 µgmL–1 was also visualized using a 3D tomographic microscope (Nanolive) equipped with a digital staining software (Figure 3.6). The obtained 2D and 3D tomographic images revealed remaining biofilm biomass values that were consistent to those obtained from CV staining experiments.

Page | 143 CHAPTER THREE

Figure 3.6. 2D and 3D tomographic microscopy images of the untreated, P, PNO, P +

SperNO and SperNO treated P. aeruginosa biofilm samples. The compound concentration and treatment time was 64 μg mL−1 and 1 h, respectively. Images were taken at the center of the culture dish. Scale bar = 20 µm.

Encouraged by these promising biofilm dispersal results, we then tested the ability of PNO to disperse thicker biofilm (Figure 3.7). Thicker biofilms (i.e., 1.8 × thicker than the above-mentioned regular biofilm model) were grown by doubling the volume of cultured bacteria. Interestingly, PNO maintained the same efficacy against the thicker biofilm, which are usually more resistant to antimicrobial treatments (Figure 3.7).5, 8

Page | 144 CHAPTER THREE

Figure 3.7. 2D and 3D tomographic microscopy images of the untreated normal and thick P. aeruginosa biofilms compared to the PNO treated samples (64 μg mL−1 for 1 h). Note: two different spots of the culture dish were analyzed. Scale bar = 20 µm.

3.3.5 Planktonic and biofilm killing by NO-loaded polymer

Subsequently, the bactericidal activity of PNO was investigated and compared to the same set of control samples (vide supra). Colony-forming unit (CFU) analysis was used to assess the bactericidal activity of the compounds against both planktonic and biofilm bacteria. P. aeruginosa biofilms were grown in the same manner as in the biofilm dispersal study and treated for either 20 or 60 min at compound concentrations of 16 and 64 µgmL–1. Likewise in the biofilm dispersal study, PNO showed the best bactericidal activity compared with the

Page | 145 CHAPTER THREE

other compounds against both planktonic and biofilm bacteria (Figure 3.8 and

Figure 3.9 and Table A 2).

a) b)

Figure 3.8. Bactericidal activity of P, PNO, SperNO and P + SperNO for 20 min (16 - 64 μg mL−1) on planktonic (a) and biofilm (b) P. aeruginosa as measured by colony-forming unit

(CFU) analysis. Data are representative of at least three independent experiments ± SD.

Student’s t-test – asterisks indicate statistically significant difference between PNO versus P treatments (****, p < 0.0001; ns, non-significant).

For the planktonic cells, treatment with PNO for 20 min at polymer concentrations of 16 and 64 µgmL–1 resulted in 1.4 and 5.2 log10 reductions in CFU, respectively, compared to the untreated sample (Figure 3.8 a). P exhibited bactericidal activity toward planktonic cells at 64 µgmL–1 with a 2.2 log10 reduction in CFU. Interestingly, in the case of both SperNO and P + SperNO physical mixture samples, we did not observe any significant reduction in bacterial cell viability. Treatment for 60 min showed a similar trend (Figure 3.9 a). These observations confirm the synergistic effect of PNO in killing planktonic bacteria cells. Although NO is known for its ability to disperse and kill bacteria cells at low (pM-nM) and high (100 µM-mM) concentrations,14-16 respectively, it

Page | 146 CHAPTER THREE

is interesting to note that a relatively low amount of released NO molecules (7-

30 µM) was sufficient in inducing cell death when it is combined with P. We are not entirely certain on the actual synergistic mechanism which led to this enhanced bactericidal activity but we can only affirm that the synergistic effect is only present when both NO donors and antimicrobial polymer exist as a single compound.

Figure 3.9. Bactericidal activity of P, PNO, SperNO and P + SperNO for 60 min (16 - 64 μg mL−1) on planktonic (a) and biofilm (b) P. aeruginosa as measured by colony-forming unit

(CFU) analysis. Data are representative of at least three independent experiments ± SD.

Student’s t-test – asterisks indicate statistically significant difference between PNO versus P treatments (****, p < 0.0001; ns, non-significant).

PNO also exhibited excellent bactericidal property against bacteria biofilm with a dose-dependent trend (Figure 3.8 b and Figure 3.9 b). Treatment with this polymer for 20 min at concentrations of 16 and 64 µgmL–1 led to 2.8 and 4.0 log10 reductions in CFU, respectively, compared with the untreated sample. Extending the incubation time to 60 min at 64 µgmL–1 resulted in a further decrease of biofilm viability (5.2 log10 reduction), while the change in the bactericidal activity

Page | 147 CHAPTER THREE

at 16 µgmL–1 was not significant based on ANOVA analysis (Figure 3.9 b). As expected, both SperNO and P + SperNO did not result in any significant reduction in biofilm cell viability, which further confirms the synergistic nature of PNO.

3.4 Summary

In summary, we have developed a robust antibiofilm agent by incorporating

NO donor functional groups into the structure of a synthetic antimicrobial ternary copolymer wherein the NO- loaded antimicrobial polymer was designed to act potentially as both the active agent as well as a delivery vehicle for the localized and targeted delivery of NO to bacteria cells. A synergistic effect in biofilm dispersal, planktonic and biofilm killing activities was achieved against the Gram-negative bacteria P. aeruginosa. Crucially, we observed that this synergistic effect was only achievable when both NO donors and antimicrobial polymer exist as a single chemical entity, as the cocktail mixture of individual compounds did not suggest any synergy in antimicrobial activity. The overall enhanced antimicrobial performance of the NO-loaded polymer compared to the native antimicrobial polymer further suggests that combination therapy may be the way forward in combating bacteria biofilms.

3.5 References

1. Høiby, N.; Ciofu, O.; Johansen, H. K.; Song, Z. j.; Moser, C.; Jensen, P. Ø.; Molin, S.; Givskov, M.; Tolker‑ Nielsen, T.; Bjarnsholt, T., The clinical impact of bacterial biofilms. Int J Oral Sci 2011, 3, 55-65. 2. Chen, M.; Yu, Q.; Sun, H., Novel Strategies for the Prevention and Treatment of Biofilm Related Infections. Int. J. Mol. Sci. 2013, 14 (9), 18488-18501.

Page | 148 CHAPTER THREE

3. Kaplan, J. B., Biofilm Dispersal: Mechanisms, Clinical Implications, and Potential Therapeutic Uses. J. Dent. Res. 2010, 89 (3), 205-218. 4. Costerton, J. W.; Stewart, P. S.; Greenberg, E. P., Bacterial Biofilms: A Common Cause of Persistent Infections. Science 1999, 284 (5418), 1318-1322. 5. Stewart, P. S.; William Costerton, J., Antibiotic resistance of bacteria in biofilms. Lancet 2001, 358 (9276), 135-138. 6. Mah, T.-F.; Pitts, B.; Pellock, B.; Walker, G. C.; Stewart, P. S.; O'Toole, G. A., A genetic basis for Pseudomonas aeruginosa biofilm antibiotic resistance. Nature 2003, 426, 306-310. 7. Hoyle, B. D.; Costerton, J. W., Bacterial resistance to antibiotics: The role of biofilms. In Progress in Drug Research / Fortschritte der Arzneimittelforschung / Progrès des recherches pharmaceutiques, Jucker, E., Ed. Birkhäuser Basel: 1991; Vol. 37, pp 91-105. 8. Mah, T.-F. C.; O'Toole, G. A., Mechanisms of biofilm resistance to antimicrobial agents. Trends Microbiol 2001, 9 (1), 34-39. 9. Wo, Y.; Brisbois, E. J.; Bartlett, R. H.; Meyerhoff, M. E., Recent advances in thromboresistant and antimicrobial polymers for biomedical applications: just say yes to nitric oxide (NO). Biomater. Sci. 2016, 4 (8), 1161-1183. 10. McDougald, D.; Rice, S. A.; Barraud, N.; Steinberg, P. D.; Kjelleberg, S., Should we stay or should we go: mechanisms and ecological consequences for biofilm dispersal. Nat. Rev. Microbiol. 2011, 10, 39-50. 11. Liu, N.; Xu, Y.; Hossain, S.; Huang, N.; Coursolle, D.; Gralnick, J. A.; Boon, E. M., Nitric Oxide Regulation of Cyclic di-GMP Synthesis and Hydrolysis in Shewanella woodyi. Biochem. 2012, 51 (10), 2087-2099. 12. Lundberg, J. O.; Gladwin, M. T.; Weitzberg, E., Strategies to increase nitric oxide signalling in cardiovascular disease. Nat. Rev. Drug Discov. 2015, 14, 623-641. 13. Carpenter, A. W.; Schoenfisch, M. H., Nitric oxide release: Part II. Therapeutic applications. Chem. Soc. Rev. 2012, 41 (10), 3742-3752. 14. Schairer, D. O.; Chouake, J. S.; Nosanchuk, J. D.; Friedman, A. J., The potential of nitric oxide releasing therapies as antimicrobial agents. Virulence 2012, 3 (3), 271-279. 15. Barraud, N.; Schleheck, D.; Klebensberger, J.; Webb, J. S.; Hassett, D. J.; Rice, S. A.; Kjelleberg, S., Nitric Oxide Signaling in Pseudomonas aeruginosa Biofilms Mediates Phosphodiesterase Activity, Decreased Cyclic Di-GMP Levels, and Enhanced Dispersal. J. Bacteriol. 2009, 191 (23), 7333-7342. 16. Barraud, N.; J. Kelso, M.; A. Rice, S.; Kjelleberg, S., Nitric Oxide: A Key Mediator of Biofilm Dispersal with Applications in Infectious Diseases. Curr. Pharm. Des. 2015, 21 (1), 31-42. 17. Riccio, D. A.; Schoenfisch, M. H., Nitric oxide release: Part I. Macromolecular scaffolds. Chem. Soc. Rev. 2012, 41 (10), 3731-3741. 18. Nichols, S. P.; Storm, W. L.; Koh, A.; Schoenfisch, M. H., Local delivery of nitric oxide: Targeted delivery of therapeutics to bone and connective tissues. Adv. Drug Deliv. Rev. 2012, 64 (12), 1177-1188. 19. Jen, M. C.; Serrano, M. C.; van Lith, R.; Ameer, G. A., Polymer-Based Nitric Oxide Therapies: Recent Insights for Biomedical Applications. Adv. Funct. Mater. 2012, 22 (2), 239-260. 20. Liu, T.; Zhang, W.; Yang, X.; Li, C., Hollow polymer nanoparticles with S- nitrosothiols as scaffolds for nitric oxide release. J. Colloid Interface Sci. 2015, 459, 115- 122.

Page | 149 CHAPTER THREE

21. Sundaram, J.; Pant, J.; Goudie, M. J.; Mani, S.; Handa, H., Antimicrobial and Physicochemical Characterization of Biodegradable, Nitric Oxide-Releasing Nanocellulose– Packaging Membranes. J. Agric. Food Chem. 2016, 64 (25), 5260-5266. 22. Sadrearhami, Z.; Yeow, J.; Nguyen, T.-K.; Ho, K. K. K.; Kumar, N.; Boyer, C., Biofilm dispersal using nitric oxide loaded nanoparticles fabricated by photo-PISA: influence of morphology. ChemComm 2017, 53 (96), 12894-12897. 23. Nguyen, T.-K.; Selvanayagam, R.; Ho, K. K. K.; Chen, R.; Kutty, S. K.; Rice, S. A.; Kumar, N.; Barraud, N.; Duong, H. T. T.; Boyer, C., Co-delivery of nitric oxide and antibiotic using polymeric nanoparticles. Chem. Sci. 2016, 7 (2), 1016-1027. 24. Parzuchowski, P. G.; Frost, M. C.; Meyerhoff, M. E., Synthesis and Characterization of Polymethacrylate-Based Nitric Oxide Donors. J. Am. Chem. Soc. 2002, 124 (41), 12182-12191. 25. Lu, Y.; Slomberg, D. L.; Shah, A.; Schoenfisch, M. H., Nitric Oxide-Releasing Amphiphilic Poly(amidoamine) (PAMAM) Dendrimers as Antibacterial Agents. Biomacromolecules 2013, 14 (10), 3589-3598. 26. Reighard, K. P.; Ehre, C.; Rushton, Z. L.; Ahonen, M. J. R.; Hill, D. B.; Schoenfisch, M. H., Role of Nitric Oxide-Releasing Chitosan Oligosaccharides on Mucus Viscoelasticity. ACS Biomater Sci Eng 2017, 3 (6), 1017-1026. 27. Cottarel, G.; Wierzbowski, J., Combination drugs, an emerging option for antibacterial therapy. Trends Biotechnol 2007, 25 (12), 547-555. 28. Fischbach, M. A., Combination therapies for combating antimicrobial resistance. Curr. Opin. Microbiol. 2011, 14 (5), 519-523. 29. Ng, V. W. L.; Ke, X.; Lee, A. L. Z.; Hedrick, J. L.; Yang, Y. Y., Synergistic Co- Delivery of Membrane-Disrupting Polymers with Commercial Antibiotics against Highly Opportunistic Bacteria. Adv. Mater. 2013, 25 (46), 6730-6736. 30. Zhao, Y.; Chen, Z.; Chen, Y.; Xu, J.; Li, J.; Jiang, X., Synergy of Non-antibiotic Drugs and Pyrimidinethiol on Gold Nanoparticles against Superbugs. J. Am. Chem. Soc. 2013, 135 (35), 12940-12943. 31. Carpenter, A. W.; Worley, B. V.; Slomberg, D. L.; Schoenfisch, M. H., Dual Action Antimicrobials: Nitric Oxide Release from Quaternary Ammonium- Functionalized Silica Nanoparticles. Biomacromolecules 2012, 13 (10), 3334-3342. 32. Worley, B. V.; Slomberg, D. L.; Schoenfisch, M. H., Nitric Oxide-Releasing Quaternary Ammonium-Modified Poly(amidoamine) Dendrimers as Dual Action Antibacterial Agents. Bioconjugate Chem. 2014, 25 (5), 918-927. 33. Yang, L.; Wang, X.; Suchyta, D. J.; Schoenfisch, M. H., Antibacterial Activity of Nitric Oxide-Releasing Hyperbranched Polyamidoamines. Bioconjugate Chem. 2018, 29 (1), 35-43. 34. Storm, W. L.; Johnson, J. A.; Worley, B. V.; Slomberg, D. L.; Schoenfisch, M. H., Dual action antimicrobial surfaces via combined nitric oxide and silver release. J Biomed Mater Res A 2015, 103 (6), 1974-1984. 35. Privett, B. J.; Deupree, S. M.; Backlund, C. J.; Rao, K. S.; Johnson, C. B.; Coneski, P. N.; Schoenfisch, M. H., Synergy of Nitric Oxide and Silver Sulfadiazine against Gram-Negative, Gram-Positive, and Antibiotic-Resistant Pathogens. Mol. Pharm. 2010, 7 (6), 2289-2296. 36. Ren, H.; Wu, J.; Colletta, A.; Meyerhoff, M. E.; Xi, C., Efficient Eradication of Mature Pseudomonas aeruginosa Biofilm via Controlled Delivery of Nitric Oxide Combined with Antimicrobial Peptide and Antibiotics. Front Microbiol 2016, 7 (1260).

Page | 150 CHAPTER THREE

37. Ravikumar, G.; Chakrapani, H., Synergistic Activities of Nitric Oxide and Various Drugs In Nitric Oxide Donors, Academic Press: 2017; pp 293-312. 38. Nguyen, T.-K.; Lam, S. J.; Ho, K. K. K.; Kumar, N.; Qiao, G. G.; Egan, S.; Boyer, C.; Wong, E. H. H., Rational Design of Single-Chain Polymeric Nanoparticles That Kill Planktonic and Biofilm Bacteria. ACS Infect Dis 2017, 3 (3), 237-248. 39. Namivandi-Zangeneh, R.; Kwan, R. J.; Nguyen, T.-K.; Yeow, J.; Byrne, F. L.; Oehlers, S. H.; Wong, E. H. H.; Boyer, C., The effects of polymer topology and chain length on the antimicrobial activity and hemocompatibility of amphiphilic ternary copolymers. Polym. Chem. 2018, DOI: 10.1039/C7PY01069A. 40. Lam, S. J.; Wong, E. H. H.; Boyer, C.; Qiao, G. G., Antimicrobial polymeric nanoparticles. Prog. Polym. Sci. 2018, 76, 40-64. 41. Judzewitsch, P. R.; Nguyen, T.-K.; Shanmugam, S.; Wong, E. H. H.; Boyer, C. A. J. M., Towards Sequence-Controlled Antimicrobial Polymers: Effect of Polymer Block Order on Antimicrobial Activity. Angew. Chem. Int. Ed. 2018, DOI: 10.1002/anie.201713036. 42. Moad, G.; Rizzardo, E.; Thang, S. H., RAFT polymerization and some of its applications. Chem. Asian J. 2013, 8 (8), 1634-44. 43. Boyer, C.; Bulmus, V.; Davis, T. P.; Ladmiral, V.; Liu, J.; Perrier, S., Bioapplications of RAFT Polymerization. Chem. Rev. 2009, 109 (11), 5402-5436. 44. Ho, K. K. K.; Ozcelik, B.; Willcox, M. D. P.; Thissen, H.; Kumar, N., Facile solvent-free fabrication of nitric oxide (NO)-releasing coatings for prevention of biofilm formation. ChemComm 2017, 53 (48), 6488-6491.

Page | 151 CHAPTER THREE

3.6 Appendix A

Figure A 1. 1H NMR spectra of the Boc-protected version of P in CDCl3 (a) and the corresponding Boc-deprotected polymer in D2O (b).

Figure A 2. GPC-differential refractive index (DRI) chromatogram of the Boc-protected version of P.

Page | 152 CHAPTER THREE

Figure A 3. TEM images of P (a) and PNO (b) in water. Insets show enlarged images of the polymers.

Figure A 4. PS spectra of PNO after 3 h of release study in PBS.

Page | 153 CHAPTER THREE

Figure A 5. NO content of PNO as a function of storage time.

Table A 1. Biofilm biomass upon treatment with various concentrations (16-64 μg mL−1) of

P, PNO, SperNO, and P + SperNO for 20 and 60 min, as measured using the CV staining method at OD595.

Biofilm Biomassa

20 min 60 min

Entry 16 μg mL−1 32 μg mL−1 64 μg mL−1 16 μg mL−1 32 μg mL−1 64 μg mL−1

Control 0.93± 0.08 1.01±0.09

P 1.16±0.10 0.96±0.02 0.70±0.11 1.07±0.21 0.97±0.10 046±0.14

PNO 0.27±0.07 0.28±0.06 0.33±0.07 0.23±0.01 0.24±0.02 0.20±0.04

SperNO 1.01±0.01 1.00±0.12 0.89±0.03 1.10±0.08 1.18±0.15 0.94±0.11

P + SperNO 1.25±0.20 0.95±0.07 0.55±0.11 1.31±0.03 0.75±0.18 0.47±0.05 aValues are the mean of three independent experiments ± standard deviation.

Page | 154 CHAPTER THREE

Table A 2. Log10 reduction of viable P. aeruginosa bacterial colonies upon treatment with various concentrations (16-64 μg mL−1) of P, PNO, SperNO and P + SperNO for 20 and 60 min, as determined using CFU analysis.

Log10 reductiona 20 min 60 min Planktonic Biofilm Planktonic Biofilm 16 16 64 16 64 64 16 64 Entry μg μg mL−1 μg mL−1 μg mL−1 μg mL−1 μg mL−1 μg mL−1 μg mL−1 mL−1

P 0.72±0.43 2.17±1.22 0.19±0.50 1.39±0.47 0.82±0.61 2.03±0.77 0.52±0.41 1.42±0.52

PNO 1.40±0.35 5.23±1.25 2.77±0.72 4.01±0.97 1.89±1.37 5.85±1.37 2.12±1.31 5.15±1.49

- - SperNO - 0.16±0.57 - 0.24±0.32 - - 0.26±0.43 0.15±0.30

P+ SperNO - 1.08±0.45 - 0.76±0.69 - 1.38±0.40 - 0.98±0.56 aValues are the mean of three independent experiments ± standard deviation.

Page | 155

CHAPTER FOUR

Synergy between Synthetic Antimicrobial

Polymer and Antibiotics: A Promising

Platform to Combat Multidrug-resistant

Bacteria

CHAPTER FOUR

4 Synergy between Synthetic Antimicrobial Polymer and

Antibiotics: A Promising Platform to Combat

Multidrug-resistant Bacteria

This chapter is based on the work published in ACS Infectious Diseases, 2019, 5,

1357–1365.

4.1 Introduction

The rising number of infections caused by multidrug-resistant (MDR) bacteria is a critical global healthcare concern.1-5 Although resistance development is a natural phenomenon, the extensive overuse of antibiotics has accelerated the process in bacteria over the past few decades.6-9 As a result, the efficiency of many antibiotics has diminished, causing longer hospital stays, higher medical costs and increased mortality.10, 11 To address this global issue, more effective therapeutic approaches are required. These include improving the efficiency of current antibiotics through chemical modification or using adjuvants,12-15

Page | 157 CHAPTER FOUR

employing antimicrobial agents with novel cellular targets,16-21 and applying combination therapy over monotherapy22, 23.

In combination therapy, the coadministration of traditional antibiotics along with other antibiotics24-26 or non-antibiotic drugs27, 28 has been reported to improve treatment efficiency compared to using individual compounds. Furthermore, combination therapy not only hinders resistance development but also revives

MDR bacteria’s susceptibility to treatment.11, 29 Additionally, synergistic combinations can effectively reduce the required treatment dose and possible side effects.30 However, to achieve synergistic activity, compound pairings need to be selected rationally. A recent report on combination therapy shows that indifference or antagonism is more likely to occur than synergism.31 Although many reports have been published on combination therapy using common antibiotics and drugs, only few studies have focused upon using synthetic antimicrobial polymers in combination with antibiotics.32-34

By mimicking the fundamental composition of natural antimicrobial peptides, synthetic antimicrobial polymers have recently emerged as potential antimicrobial agents that target the bacteria cell membrane.16, 17 Generally, electrostatic interactions between the cationic residues of antimicrobial polymers and the anionic components of bacteria cell walls or outer membranes are the first step in bactericidal activity, followed by the insertion of the polymer hydrophobic sub-units into the cytoplasmic membrane to induce membrane disruption/permeabilization leading to cell death. Such a bactericidal mechanism hinders resistance development in bacteria, thereby making synthetic

Page | 158 CHAPTER FOUR

antimicrobial polymers a promising co-agent in combination therapy to create more potent antimicrobial strategies.

On the basis of our experience in the development of antimicrobial polymers35-

37 and the potential benefits of combination therapy,38-41 we herein investigate the efficacy of an antimicrobial platform based on our lead synthetic antimicrobial polymer35, 39 in combination with different classes of commercially available antibiotics (e.g., doxycycline from the family, as depicted in Figure

4.1). The potency of these combinations is investigated via the checkerboard assay, and the synergistic combinations are further studied in terms of bactericidal activity and resistance development in bacteria.

Figure 4.1. Synergistic antibacterial activity of synthetic antimicrobial polymer and doxycycline.

Page | 159 CHAPTER FOUR

4.2 Experimental Section

Materials

Materials. Ethylenediamine (Sigma-Aldrich, ≥ 99%), di-tert-butyl dicarbonate

(Aldrich, 99%), triethylamine (Scharlau, 99%), acryloyl chloride (Merck, ≥ 96%), oligoethylene glycol methyl ether acrylate (OEGA) Mn 480 g mol–1 (Aldrich), 2- ethylhexyl acrylate (Aldrich, 98%), trifluoroacetic acid (TFA) (Sigma-Aldrich,

99%), hexane (Merck), diethyl ether (Merck), 1,4-dioxane (Merck) and basic alumina (Al2O3) (LabChem) were used as received. 2,2’-azobis (2 methylpropionitrile) (AIBN) (Acros, 98%) was purified by recrystallization from methanol. Sodium sulfate (Na2SO4), magnesium sulfate (MgSO4), sodium hydrogen carbonate (NaHCO3), tetrahydrofuran, and acetone were obtained from Chem-Supply and used as received. Milli-Q water with a resistivity of > 18

MΩ⋅cm was obtained from an in-line Millipore RiOs/Origin water purification system. Antibiotics (amoxicillin, ampicillin, azithromycin dihydrate, ceftriaxone disodium salt hemi (hepatahydrate), ciprofloxacin hydrochloride, clarithromycin, colistin sodium methanesulfonate and doxycycline hydrochloride) were purchased from Sigma-Aldrich. Gentamicin sulfate was purchased from Enzo life sciences and tobramycin was purchased from Biogal.

Analytical instruments

Characterizations. 1H Nuclear magnetic resonance (NMR) spectra were obtained using a Bruker AC300F spectrometer. Deuterated solvents D2O or

CDCl3 (obtained from Cambridge Isotope Laboratories) were used as reference solvents and samples with concentration of ca. 10-20 mg mL–1 were prepared. The

Page | 160 CHAPTER FOUR

monomer composition of the polymers was calculated using the following equation ʃa,b/6 : ʃc/9 : ʃd/2 where ʃa,b, ʃc, and ʃd correspond to the integrals of the characteristic protons of 2-ethylhexyl acrylate (methyl -CH3- groups, δH 0.80-0.98 ppm), cationic monomer (tert-butyl -CH3- groups, δH 1.38-1.52 ppm) and OEGA

(ester -CH2O- groups, 4.10-4.30 ppm), respectively (Figure B 1).

Gel permeation chromatography (GPC) analysis was performed using a

Shimadzu liquid chromatography system equipped with a Shimadzu refractive index detector and two MIX C columns (Polymer Lab) operating at 40 °C.

Tetrahydrofuran was used as the eluent at a flow rate of 1 mL min–1. The system was calibrated with poly(methyl methacrylate) standards with molecular weights of 200 to 106 g mol–1.

Zeta-potential measurements were conducted using a Malvern Zetasizer

Nano ZS apparatus equipped with a He-Ne laser operated at λ = 633 nm and at a scattering angle of 173o. Samples were prepared at a concentration of ca. 2 mg mL–1 where filtered Milli-Q water (using a 0.45 μm pore size filter) was used as the solvent to solubilize the polymer.

Synthesis of antimicrobial polymer

Synthesis of cationic monomer

Cationic monomer tert-butyl (4-acrylamidobutyl) carbamate was prepared in the same manner as reported previously.35 Briefly, ethylenediamine (0.3 mol) was dissolved in chloroform (400 mL), followed by the dropwise addition of di-tert- butyl dicarbonate (0.03 mol in 100 mL) over 2 h at 0−5 °C. The reaction mixture was stirred overnight at 25 °C. White precipitates were filtered, and the organic

Page | 161 CHAPTER FOUR

phase was washed exhaustively with water using a separation funnel to remove excess diamines. The organic layer was then dehydrated over MgSO4, filtered, and dried using a rotary evaporation unit to yield a pale-yellow oil.

Tetrahydrofuran (100 mL) was added to dissolve the intermediate product.

Triethylamine (36 mmol) and acryloyl chloride (31.5 mmol) were added dropwise to the solution at 0−5 °C with N2 bubbling. The contents were stirred at

25 °C for 1 h. The byproducts were filtered, and the solvent was removed in vacuo. The crude product was dissolved in chloroform (150 mL) and washed against brine (1 × 75 mL). The organic phase was stirred with MgSO4 and basic

Al2O3 for 10 min, filtered, and concentrated in vacuo. The product was further purified by repeated precipitation steps in hexane to yield the tert- butyloxycarbonyl (Boc)-protected monomer as a fine white powder, which was dried in vacuo.

tert-Butyl (2-acrylamidoethyl) carbamate: 1H NMR (300MHz, CDCl3,25 °C), δH

6.56 (br s, 1H, NH), 6.28 (dd, J = 17.1 Hz, 1.5 Hz, 1H, CHH=CH), 6.12 (dd, J = 17.1

Hz, 10.2Hz, 1H, CHH=CH), 5.65 (dd, J = 10.2 Hz, 1.5 Hz, 1H, CHH=CH), 5.05 (br s, 1H, NH), 3.49−3.41 (m, 2H, CH2), 3.34−3.28 (m, 2H, CH2), 1.45 (s, 9H, CH3); 13C

NMR (300 MHz, CDCl3,25 °C), δC 166.23, 157.50, 130.88, 126.30, 79.85, 41.05, 40.09,

28.35.

Synthesis of amphiphilic ternary random copolymer

The synthesis of amphiphilic ternary statistical copolymer proceeded in the same manner as reported previously.35 Briefly, the reversible addition−fragmentation chain transfer (RAFT) agent benzyl dodecyl carbonotrithioate (11.6 μmol), AIBN (4.6 μmol), OEGA (350 μmol), tert-butyl (4-

Page | 162 CHAPTER FOUR

acrylamidobutyl) carbamate (580 μmol), and 2-ethylhexyl acrylate (230 μmol) were dissolved in 1,4-dioxane (such that the total monomer concentration in solvent is 1 M). The solution was purged with N2 for 20 min in an ice bath. The polymerization was conducted for 20 h at 70 °C and then quenched in an ice bath with exposure to air. The polymer was purified by precipitation in a hexane/diethyl ether (7:3) mixture thrice and subsequently dried in vacuo. The

Boc- protected polymer produced monomodal molecular weight distributions with dispersity (Đ) value of 1.3 and the number-averaged molecular weight (Mn) of 17 500 g mol-1 as evidenced by GPC analysis (Figure B 2)

The Boc protecting groups were removed using TFA in the same manner as reported previously.35 In general, the polymer solution in dichloromethane (ca.

10 wt % polymer) was treated with TFA (20 mol equivalent with respect to the

Boc group) for 3 h at 25 °C. Boc- deprotected polymer was subsequently precipitated into diethyl ether/hexane (4:1). The precipitate was isolated by centrifugation, dissolved in methanol, and reprecipitated two more times. The polymer was then dried in vacuo and further purified by dialysis against water

(Cellu-Sep 3500 MWCO). The aqueous solution was lyophilized to yield the Boc- deprotected polymer P (P, amphiphilic random ternary copolymer).

Minimum inhibitory concentration (MIC) determination

The MIC of polymer and selected antibiotics were determined via broth microdilution method according to Clinical and Laboratory Standards Institute

(CLSI) guidelines. Briefly, a single colony was cultured in 10 mL of Mueller-

Hinton broth (MHB) at 37 °C with shaking at 200 rpm overnight. Subsequently, a subculture was prepared from the overnight culture by diluting 1:100 in 10 mL

Page | 163 CHAPTER FOUR

MHB and allowed to grow to mid log phase; then, it was diluted to the appropriate concentration for the MIC test. A 2-fold dilution series of 100 μL of polymers solution in MHB was added into 96-well microplates followed by the addition of 100 μL of the subculture suspension. The final concentration of bacteria in each well was ca. 5 × 105 cells mL–1. The plates were incubated at 37 °C for 20 h, and the absorbance at 600 nm was measured with a microtiter plate reader (FLUOstar Omega, BMG Labtech). MIC values were defined as the lowest concentration of sample that showed no visible growth and inhibited cell growth by more than 90%. Positive controls without polymer and negative controls without bacteria were included. All assays included two replicates and were repeated in at least three independent experiments.

Checkerboard microdilution assay

The checkerboard assay was performed in 96-well cell culture plates (Costar,

Corning®) containing polymer and antibiotic in MHB. Concentration gradients of polymer and selected antibiotics were prepared in the horizontal and vertical direction in a 10 × 8 layout, respectively. Bacterial suspensions were prepared in the same manner as above-mentioned for the MIC test and added to the plates.

Positive and negative controls, without antimicrobial agent and bacteria, respectively, were also included. The plates were incubated at 37 °C for 20 h, and the absorbance at 600 nm was recorded subsequently. The MICs of individual polymer and antibiotics were computed from the corresponding column or row which contains only one component, while the remaining columns and rows were screened for fractional inhibitory concentration index (FICI). The FICI values were calculated as follows:

Page | 164 CHAPTER FOUR

MIC in combination MIC in combination FICI = A + B MICA MICB

The FICI data was interpreted as follows: ≤0.5, synergistic effect; 0.5 < FICI < 1, indifference; ≥1, antagonistic. All experiments were repeated in at least two independent experiments.

Killing study

To evaluate the bactericidal activity of selected compounds and combinations,

Pseudomonas aeruginosa (P. aeruginosa) PAO1 and 6294 biofilms were grown. A single colony of PAO1 or PA6294 was cultured overnight in 10 mL of Luria-

Bertani medium (LB 10) at 37 oC with shaking at 200 rpm. The overnight culture was diluted 1:200 in freshly prepared M9 minimal medium containing 48 mM

Na2HPO4, 22 mM KH2PO4, 9 mM NaCl, and 19 mM NH4Cl, pH 7.0, supplemented with 2 mM MgSO4, 100 μM CaCl2 and 20 mM glucose. The bacterial suspension was then aliquoted (1 mL per well) into tissue-culture treated 24-well plates

(Costar, Corning®). The plates were incubated at 37 °C with shaking at 180 rpm in an orbital shaker that does not stop agitation when the door is opened (model

OM11, Ratek, Boronia, Australia), and the biofilm cultures were allowed to grow for 6.5 h without any disruption. The compounds were then added to the wells, and the plates were incubated for 20 min. After treatment, the planktonic and biofilm viability analyses were determined by a drop plate method. For planktonic analysis, free-floating cells in the biofilm supernatant were serially diluted in sterile PBS and plated onto LB agar. For biofilm analysis, cells attached on the interior surfaces of the well (surface area 4.5 cm2) were washed twice with sterile PBS to remove loosely attached bacteria, before being resuspended and homogenized in PBS by incubating in an ultrasonication bath (150 W, 40 kHz; Page | 165 CHAPTER FOUR

Unisonics, Australia) for 20 min. Resuspended biofilm cells were then serially diluted and plated onto LB agar. Planktonic and biofilm colonies were counted, and CFU was calculated after 24 h of incubation at 37 °C. All assays included two replicates and were repeated in at least three independent experiments.

Antimicrobial resistance study

To investigate the resistance development in Pseudomonas aeruginosa PAO1, bacterial suspensions at exponential phase (~107 cells mL–1) were subjected to sequential passaging in the presence of selected compounds at sub-inhibitory concentrations (i.e., 1/4 × MIC, 1/2 × MIC, 1 × MIC and, 2 × MIC) for 21 days. Cells were incubated at 37 °C and passaged at 24 h intervals. After incubation, the cultures were checked for growth. Cultures from the second highest concentrations that allow growth (OD600 ≥ 2.00) were diluted to an OD600 of 0.01 per milliliter in fresh MHB containing 1/4 × MIC, 1/2 × MIC, 1 × MIC and, 2 × MIC of selected compound. Assays were performed with two independent experiments.

4.3 Results and Discussion

The ability of synthetic antimicrobial polymers to combat MDR bacteria provides the key motivation for us to investigate their efficacy in combination therapy along with traditional antibiotics. For this, we used an antimicrobial polymer in the form of an amphiphilic random ternary copolymer (P) that was developed by our group.35 Large quantities of polymer can be made in a facile manner via a controlled radical polymerization technique, termed reversible addition−fragmentation chain transfer (RAFT) polymerization (Scheme 4.1).42, 43 Page | 166 CHAPTER FOUR

This polymer was rationally designed to have optimal antimicrobial activity and biocompatibility based on the three selected monomers. It contains 30 repeat units of biocompatible oligoethylene glycol to impart low-fouling properties, 20 repeat units of hydrophobic ethylhexyl groups to induce membrane disruption, and 50 repeat units of primary amino groups to establish electrostatic interactions with the bacterial membrane.35, 36

Scheme 4.1. Synthesis of antimicrobial polymer, P, which is an amphiphilic random ternary copolymer, via reversible addition−fragmentation chain transfer (RAFT) polymerization.

4.3.1 Assessment of antimicrobial polymer-antibiotic interactions

To assess the potential synergistic effect between P and commercially available antibiotics, their interactions were evaluated using the checkerboard assay.

Figure 4.2 shows the chemical structure and mode of action of ten common antibiotics used in this study. These antibiotics were selected on the basis of their ability to act on different mechanisms, such as targeting protein synthesis

(doxycycline, clarithromycin, azithromycin, gentamicin, and tobramycin), cell wall biosynthesis (ampicillin, amoxicillin, ceftriaxone), cell wall integrity

(colistin), and DNA synthesis (ciprofloxacin) in bacteria.

Page | 167 CHAPTER FOUR

Figure 4.2. Chemical structure and reported mode of action of the antibiotics used in this work.44, 45

Checkerboard plots of tested combinations against two Gram-negative bacteria Pseudomonas aeruginosa PAO1 and Escherichia coli K12 are represented in

Figure 4.3 and Figure 4.4; Gram-positive bacteria were not included in this study as the antimicrobial activity of P is better against Gram-negative bacteria.37 This

Page | 168 CHAPTER FOUR

assay is conducted to evaluate the bacteriostatic activity of proposed combinations. Minimum inhibitory concentrations (MICs) of the used antimicrobial agents alone and in combination, and fractional inhibitory concentration indexes (FICIs) of the tested combinations against P. aeruginosa

PAO1 and E. coli K12 are shown in Table B 1 and Table B 2 It is worthwhile to note that FICI is a parameter commonly used to determine the synergism/antagonism of compounds in combination therapy. Although both tested bacteria were susceptible to all selected antibiotics, only doxycycline (D) and colistin methanesulfonate (C) showed synergy (FICI ≤ 0.5) with P. Notably, indifference (0.5 < FICI < 1) was the most common interaction between P and antibiotics, but none of the combinations demonstrated antagonism interaction

(FICI ≥1) against the tested bacteria strains. It is noteworthy that clarithromycin showed synergy with P exclusively against P. aeruginosa PAO1.

Figure 4.3. Checkerboard microdilution assay between P and selected antibiotics against P. aeruginosa PAO1. Bacterial growth, quantified by average OD600, is represented as a linear gradient from white to peach where darker colors represent less growth inhibition. Yellow and red bullets represent MIC values for P and antibiotics, respectively. Blue bullets

Page | 169 CHAPTER FOUR

represent concentrations exhibiting synergistic interaction. The data are representative of a minimum of two biological replicates.

Figure 4.4. Checkerboard microdilution assay between P and selected antibiotics against E. coli K12. Bacterial growth, quantified by average OD600, is represented as a linear gradient from white to blue where darker colors represent less growth inhibition. Yellow and red bullets represent MIC values for P and antibiotics, respectively. Blue bullets represent concentrations exhibiting synergistic interaction. The data are representative of a minimum of two biological replicates.

The combination of P and D (PD) yielded synergistic activity against P. aeruginosa PAO1 and E. coli K12 with FICI values of 0.38–0.50 and 0.50, respectively. Co-administration of these antimicrobial agents resulted in at least a 4-fold decrease in MIC values of P and D against P. aeruginosa PAO1 (Table B

1 and Table B 2). Meanwhile, the P and C combination (PC) demonstrated stronger synergy against P. aeruginosa PAO1 (FICI = 0.38) compared to E. coli K12

(FICI = 0.50). On the basis of these results, both combinations demonstrated slight species-specific activity, where slightly greater synergy was observed against P. aeruginosa PAO1.

Page | 170 CHAPTER FOUR

The mechanism behind the synergistic interactions is complicated; however, the mode of action of individual components might explain the observed synergy. When used in combination with other compounds that act on intracellular targets, membrane-targeting compounds are expected to modulate the intracellular drug concentration. Synergistic interaction will lead to increased drug influx while antagonistic interaction will do the opposite.31 We postulate that, in the synergistic combination PD, P can enhance D uptake through membrane wall disruption. In spite of this, we are unsure as to why the other antibiotics, which act on intracellular targets, mainly displayed neutral interactions. Synergistic interactions are also probable when drugs that act on a similar mechanism are used in combination.22, 31 Given that both P and C predominantly act on the same mechanism of membrane wall disruption, this might be the reason behind the observed synergistic activity in PC.

Following these results, the potency of the synergistic combinations (PD and

PC) was evaluated against four more P. aeruginosa strains including PA ATCC

27853, an invasive strain isolated from microbial keratitis (PA 6294),46 and two

MDR strains (PA32 and PA37) also isolated from cases of microbial keratitis47, 48 via the checkerboard method (Figure 4.5 and Table 4.1).

Page | 171 CHAPTER FOUR

Figure 4.5. Checkerboard microdilution assay between P and D (a-e), and P and C (f-j) against P. aeruginosa PAO1, PA6294, PA ATCC 27853, PA32 and PA37. Bacterial growth, quantified by average OD600, is illustrated as a linear gradient from white to green and burgundy where darker colors represent less growth inhibition. Yellow and red bullets represent MIC values for P and antibiotic, respectively. Blue bullets indicate concentrations exhibiting synergistic interaction. The data are representative of a minimum of two biological replicates.

Page | 172 CHAPTER FOUR

Table 4.1. Checkerboard assay results indicating the synergistic activity of P with D and C and against five P. aeruginosa strains including PAO1, PA6294, PA ATCC 27853, PA32 and

PA37. All the combinations showed moderate to high synergism (FICI ≤ 0.5).

-1 -1 MIC (µg mL ) MIC (µg mL )

FICI FICI Alone In Combination Alone In Combination P D P D P C P C

8 1 0.38 4 4 0.38 PAO1 32 8 32 16 8 2 0.50 8 2 0.38

16 1 0.38 8 2 0.38 PA6294 64 8 64 8 16 2 0.50 16 1 0.38

PA 32 16 8 4 0.50 64 32 16 8 0.50 ATCC

32 4 0.28 16 2 0.38

PA32 128 128 32 8 0.31 64 16 16 4 0.50

16 32 0.38 - - -

16 32 0.25 32 1 0.31

PA37 128 256 32 16 0.31 128 16 32 2 0.38

- - - 16 4 0.38

PD and PC exhibited synergism against all tested P. aeruginosa strains. The greatest synergistic effect was found against MDR strains PA32 and PA37 for both PD and PC. The strongest synergistic activities with FICI values of 0.28 and

0.38 against PA32, and 0.25 and 0.31 against PA37 were observed for PD and PC, respectively (Table 4.1). As an example, although PA32 showed resistance toward both P and D, we observed a 4- to 8-fold and 4- to 32-fold decrease in the

MIC values of P and D, respectively, when PD was used. P at a sub-MIC level as low as 16 μg mL−1 (MIC = 128 μgmL−1) was able to reduce the MIC of D from 128 to 4 μg mL−1 (Table 4.1) when used in combination.

Page | 173 CHAPTER FOUR

4.3.2 Evaluation of bactericidal activity of combinations and individual

agents

Subsequently, bactericidal and antibiofilm activity of PD and PC synergistic combinations were investigated. Colony-forming unit (CFU) analysis was used to assess the bactericidal activity of the combinations against both planktonic and biofilm bacteria. P. aeruginosa PAO1 biofilms were grown in cell culture media

M9 for 6.5 h prior to incubation with selected compounds at different concentrations for 20 min.

First, the bactericidal activities of PD and PC at fixed P concentration of 128

μg mL−1 and three antibiotic concentrations of 16, 32, and 64 μg mL−1, denoted as

128/16, 128/32 and 128/64 μg mL−1, were examined against PAO1. For comparison, individual P and antibiotics (D and C) were also tested (Figure 4.6 a-d).

Page | 174 CHAPTER FOUR

Figure 4.6. Bactericidal activity of individual components and combinations. Bactericidal activity of P, D, and PD on planktonic (a) and biofilm (b) P. aeruginosa PAO1. Bactericidal activity of P, C, and PC on planktonic (c) and biofilm (d) P. aeruginosa PAO1. Bactericidal activity of P, D, and PD on planktonic (e) and biofilm (f) P. aeruginosa 6294. All bactericidal activities were determined by colony-forming unit (CFU) analysis upon 20 min incubation.

Data are representative of at least three independent experiments ± SD. Two-way ANOVA test − asterisks indicate statistically significant difference of PD and PC vs P treatment (**p <

0.01; ***p < 0.001; ****p < 0.0001; ns, non-significant (p > 0.01)).

For the planktonic cells, treatment with P at 128 μg mL−1 for 20 min resulted in an average of 3.3 ± 0.24 log10 reduction in CFU compared to the untreated sample, while D and C did not cause any significant reduction in bacterial cell viability

(Figure 4.6 a, c and Table B 3). Both PD and PC exhibited a bactericidal property against planktonic bacteria in a dose-dependent manner. Treatment with PD at concentrations of 128/64, 128/32, and 128/16 μg mL−1 led to 5.4 ± 0.29, 5.3 ± 0.61 and 4.8 ± 0.48 log10 reductions in CFU, respectively, compared to the untreated sample. These results confirm the synergistic effect of PD in killing planktonic Page | 175 CHAPTER FOUR

bacteria cells at the tested concentrations. PC, on the other hand, showed significant synergy only at 128/64 μg mL−1 causing 5.1 ± 0.59 log10 reduction in

CFU compared to the untreated sample.

Synergistic bactericidal activity was also observed against the bacterial biofilm upon treatment with PD in the same dose-dependent manner (Figure 4.6 b and

Table B 3). We observed synergistic effect at the highest D concentration (128/64

μg mL−1), whereas it was non-significant at lower D concentrations (128/32 and

128/16 μg mL−1). Treatment with PD for 20 min at 128/64 μg mL−1 resulted in 5.5

± 0.43 log10 reductions in CFU, compared to the untreated biofilm. However, the differences between the bactericidal activities of P and PC at all tested concentrations were non-significant based on ANOVA analysis (Figure 4.6 d and

Table B 3).

We then tested the bactericidal activity of PD against P. aeruginosa 6294. We observed synergy against planktonic PA6294 with 6.2 ± 0.10, 6.0 ± 0.39 and 5.9 ±

0.87 log10 reductions in CFU compared to the untreated sample at treatment concentrations of 128/64, 128/32 and 128/16 μg mL−1 respectively (Figure 4.6 e and

Table B 3). Interestingly, PD demonstrated a reverse dose-dependent trend against bacterial biofilm, where the lowest D concentration caused the highest reduction in CFU with a 4.7±0.14 log10 reduction in CFU at 128/16 μg mL−1 compared to the untreated sample (Figure 4.6 f and Table B 3).

The observed synergy between P and D in the killing assays might be caused by the bacteriostatic characteristic of D, where it can potentiate the bactericidal activity of P by restricting bacterial growth.

Page | 176 CHAPTER FOUR

4.3.3 Evaluation of resistance development toward polymer-doxycycline

combination

Next, an antimicrobial resistance study was conducted to investigate if PD can induce resistance in P. aeruginosa PAO1. P and D were also included as controls.

For this, 20 serial passages of bacterial cells were done over a period of 21 days in the presence of sub-MIC levels of P, D, and PD (Figure 4.7). We demonstrated in our previous work P. aeruginosa PAO1 could not develop resistance toward C over a period of 22 days;35 therefore, PC and C were not included in the resistance study.

Figure 4.7. Resistance development monitoring in P. aeruginosa PAO1 in the presence of sub-

MIC levels of P, D and PD. The y-axis indicates the changes in MICs of the compounds over a period of 21 days as compared to the first day (0th passage). Note that the bacteria cultures from 20th passage were used in supplementary MIC test.

P. aeruginosa PAO1 developed resistance toward D rapidly, where the MIC increased 32 times the original value and plateaued after six days. P, on the other hand, triggered minor resistance in P. aeruginosa which is in accordance with previously published data.35 The MIC value changed between 1 × MIC and 2 ×

Page | 177 CHAPTER FOUR

MIC over the test period and finally reached 4× MIC. This behaviour is attributed to the membrane disruption mechanism of P, which minimizes the likelihood of resistance development in bacteria. The MIC for PD was less volatile, fluctuating between 1× MIC and 2× MIC but not exceeding 2× MIC during the test period, which shows the potency of PD in suppressing resistance development in bacteria.

A supplementary antimicrobial activity test was subsequently performed against the obtained modified strains from the resistance study to evaluate their susceptibility to further treatments. For this, bacteria cells were derived from the

20th passage of all three treatments (PAO1-20-D, PAO1-20-P and PAO1-20-PD) and subjected to P, D and, PD. Table 4.2 shows the MIC values of P, D and, PD against unmodified and modified strains of P. aeruginosa PAO1.

Table 4.2. Supplementary antimicrobial activity test showing the susceptibility of modified strains of P. aeruginosa PAO1 (obtained from the 20th passage of resistant study, see Figure

4.7) to P, D and PD.

MIC (µg mL-1)

Treatment PAO1 PAO1-20-D PAO1-20-P PAO1-20-PD

D 8 256 8-16 32

P 32 32 128 32

PD 8/2 16/4 32/8 16/4

For modified D strain (PAO1-20-D), we observed no change in the MIC value of P, while there was a 2-fold increase in the MIC value of PD, which might be the result of acquired resistance toward D. Although showing a minor resistance to PD treatment, PD is a potent treatment for PAO1-20-D by lowering the required D dose by 64 times compared to individual D. Not surprisingly, an

Page | 178 CHAPTER FOUR

increase in the MIC value for PD was observed against PAO1-20-P, where 4 ×

MIC was needed to effectively combat the modified P strain. D, on the other hand, retained the same MIC value against PAO1-20-P. In the case of PAO1-20-

PD, even after 20 passages in the presence of PD, no resistance was observed toward P, whereas 4 × MIC was required to inhibit the bacterial growth using D.

However, this level of resistance to D is considered minimal compared to exclusively D treated PAO1 (PAO1-20-D).

These observations suggest that: (i) the acquisition of resistance toward either

P or D is hindered through coadministration, and (ii) when PD is used against modified strains (PAO1-20-P and PAO1-20-D), P is more efficient in reviving

PAO1 susceptibility to D, rather than vice versa.

4.4 Summary

In summary, we investigated the efficacy of combination therapy involving our synthetic antimicrobial polymer and commercially available antibiotics.

Polymer-drug interactions were studied via the checkerboard assay where two synergistic combinations, containing doxycycline and colistin antibiotics, were detected. Synergistic combinations demonstrated bacteriostatic activity against

Gram-negative bacteria such as P. aeruginosa and E. coli, where the greatest synergism was observed against MDR P. aeruginosa strains. A synergistic effect in planktonic and biofilm killing activity was also achieved using the combination of antimicrobial polymer and doxycycline, which suggests that doxycycline, even though it is a bacteriostatic antibiotic, could potentiate the killing activity of polymer. In addition, serial passaging revealed that this

Page | 179 CHAPTER FOUR

synergistic combination can significantly hinder the generation of resistant mutant strains compared to individual compounds. This study shed valuable information on the potential use of synthetic antimicrobial polymers in combination with specific antibiotics in an effort to combat MDR bacteria.

4.5 References

1. Magiorakos, A. P.; Srinivasan, A.; Carey, R. B.; Carmeli, Y.; Falagas, M. E.; Giske, C. G.; Harbarth, S.; Hindler, J. F.; Kahlmeter, G.; Olsson-Liljequist, B.; Paterson, D. L.; Rice, L. B.; Stelling, J.; Struelens, M. J.; Vatopoulos, A.; Weber, J. T.; Monnet, D. L., Multidrug-resistant, extensively drug-resistant and pandrug-resistant bacteria: an international expert proposal for interim standard definitions for acquired resistance. Clin. Microbiol. Infect. 2012, 18 (3), 268-281. 2. Boucher, H. W.; Talbot, G. H.; Bradley, J. S.; Edwards, J. E.; Gilbert, D.; Rice, L. B.; Scheld, M.; Spellberg, B.; Bartlett, J., Bad Bugs, No Drugs: No ESKAPE! An Update from the Infectious Diseases Society of America. Clin. Infect. Dis. 2009, 48 (1), 1-12. 3. Levy, S. B.; Marshall, B., Antibacterial resistance worldwide: causes, challenges and responses. Nat. Med. 2004, 10, S122-S129. 4. Jones, K. E.; Patel, N. G.; Levy, M. A.; Storeygard, A.; Balk, D.; Gittleman, J. L.; Daszak, P., Global trends in emerging infectious diseases. Nature 2008, 451, 990-993. 5. Sadrearhami, Z.; Nguyen, T.-K.; Namivandi-Zangeneh, R.; Jung, K.; Wong, E. H. H.; Boyer, C., Recent advances in nitric oxide delivery for antimicrobial applications using polymer-based systems. J. Mater. Chem. B 2018, 6 (19), 2945-2959. 6. Serpi, M.; Ferrari, V.; Pertusati, F., Nucleoside Derived Antibiotics to Fight Microbial Drug Resistance: New Utilities for an Established Class of Drugs? J. Med. Chem. 2016, 59 (23), 10343-10382. 7. Davies, J.; Davies, D., Origins and Evolution of Antibiotic Resistance. Microbiol. Mol. Biol. Rev. 2010, 74 (3), 417-433. 8. Costerton, J. W.; Stewart, P. S.; Greenberg, E. P., Bacterial Biofilms: A Common Cause of Persistent Infections. Science 1999, 284 (5418), 1318-1322. 9. D’Costa, V. M.; King, C. E.; Kalan, L.; Morar, M.; Sung, W. W. L.; Schwarz, C.; Froese, D.; Zazula, G.; Calmels, F.; Debruyne, R.; Golding, G. B.; Poinar, H. N.; Wright, G. D., Antibiotic resistance is ancient. Nature 2011, 477, 457-461. 10. Tam, V. H.; Rogers, C. A.; Chang, K.-T.; Weston, J. S.; Caeiro, J.-P.; Garey, K. W., Impact of Multidrug-Resistant Pseudomonas aeruginosa Bacteremia on Patient Outcomes. Antimicrob. Agents Chemother. 2010, 54 (9), 3717-3722. 11. Walsh, C., Molecular mechanisms that confer antibacterial drug resistance. Nature 2000, 406, 775-781. 12. Petchiappan, A.; Chatterji, D., Antibiotic Resistance: Current Perspectives. ACS Omega 2017, 2 (10), 7400-7409. 13. Fischbach, M. A.; Walsh, C. T., Antibiotics for Emerging Pathogens. Science 2009, 325 (5944), 1089-1093.

Page | 180 CHAPTER FOUR

14. González-Bello, C., Antibiotic adjuvants – A strategy to unlock bacterial resistance to antibiotics. Bioorg. Med. Chem. Lett. 2017, 27 (18), 4221-4228. 15. Wright, G. D., Antibiotic Adjuvants: Rescuing Antibiotics from Resistance. Trends Microbiol. 2016, 24 (11), 862-871. 16. Kenawy, E.-R.; Worley, S. D.; Broughton, R., The Chemistry and Applications of Antimicrobial Polymers: A State-of-the-Art Review. Biomacromolecules 2007, 8 (5), 1359-1384. 17. Timofeeva, L.; Kleshcheva, N., Antimicrobial polymers: mechanism of action, factors of activity, and applications. Appl. Microbiol. Biotechnol. 2011, 89 (3), 475-492. 18. Lam, S. J.; Wong, E. H. H.; Boyer, C.; Qiao, G. G., Antimicrobial polymeric nanoparticles. Prog. Polym. Sci. 2018, 76, 40-64. 19. Zasloff, M., Antimicrobial peptides of multicellular organisms. Nature 2002, 415, 389-395. 20. Matsuzaki, K., Control of cell selectivity of antimicrobial peptides. Biochim. Biophys. Acta 2009, 1788 (8), 1687-1692. 21. Jenssen, H.; Hamill, P.; Hancock, R. E. W., Peptide Antimicrobial Agents. Clin. Microbiol. Rev. 2006, 19 (3), 491-511. 22. Fischbach, M. A., Combination therapies for combating antimicrobial resistance. Curr. Opin. Microbiol. 2011, 14 (5), 519-523. 23. Cottarel, G.; Wierzbowski, J., Combination drugs, an emerging option for antibacterial therapy. Trends Biotechnol. 2007, 25 (12), 547-555. 24. Paul, M.; Lador, A.; Grozinsky‑ Glasberg, S.; Leibovici, L., Beta lactam antibiotic monotherapy versus beta lactam‑ aminoglycoside antibiotic combination therapy for sepsis. Cochrane Database Syst Rev 2014, (1), 1-177. 25. Bassetti, M.; Repetto, E.; Righi, E.; Boni, S.; Diverio, M.; Molinari, M. P.; Mussap, M.; Artioli, S.; Ansaldi, F.; Durando, P.; Orengo, G.; Bobbio Pallavicini, F.; Viscoli, C., Colistin and rifampicin in the treatment of multidrug-resistant Acinetobacter baumannii infections. J. Antimicrob. Chemother. 2008, 61 (2), 417-420. 26. Tamma, P. D.; Cosgrove, S. E.; Maragakis, L. L., Combination Therapy for Treatment of Infections with Gram-Negative Bacteria. Clin. Microbiol. Rev. 2012, 25 (3), 450-470. 27. Ejim, L.; Farha, M. A.; Falconer, S. B.; Wildenhain, J.; Coombes, B. K.; Tyers, M.; Brown, E. D.; Wright, G. D., Combinations of antibiotics and nonantibiotic drugs enhance antimicrobial efficacy. Nat. Chem. Biol. 2011, 7, 348-350. 28. Schneider, E. K.; Reyes-Ortega, F.; Velkov, T.; Li, J., Antibiotic–non-antibiotic combinations for combating extremely drug-resistant Gram-negative ‘superbugs’. Essays Biochem. 2017, 61 (1), 115-125. 29. Mouton, J. W., Combination therapy as a tool to prevent emergence of bacterial resistance. Infection 1999, 27 (2), S24-S28. 30. Elphick, H. E.; Scott, A., Single versus combination intravenous anti‑ pseudomonal antibiotic therapy for people with cystic fibrosis. Cochrane Database Syst Rev 2016, (12), 1-41. 31. Brochado, A. R.; Telzerow, A.; Bobonis, J.; Banzhaf, M.; Mateus, A.; Selkrig, J.; Huth, E.; Bassler, S.; Zamarreño Beas, J.; Zietek, M.; Ng, N.; Foerster, S.; Ezraty, B.; Py, B.; Barras, F.; Savitski, M. M.; Bork, P.; Göttig, S.; Typas, A., Species-specific activity of antibacterial drug combinations. Nature 2018, 559 (7713), 259-263. 32. Tian, J.; Zhang, J.; Yang, J.; Du, L.; Geng, H.; Cheng, Y., Conjugated Polymers Act Synergistically with Antibiotics to Combat Bacterial Drug Resistance. ACS Appl. Mater. Interfaces 2017, 9 (22), 18512-18520.

Page | 181 CHAPTER FOUR

33. Ng, V. W. L.; Ke, X.; Lee, A. L. Z.; Hedrick, J. L.; Yang, Y. Y., Synergistic Co- Delivery of Membrane-Disrupting Polymers with Commercial Antibiotics against Highly Opportunistic Bacteria. Adv. Mater. 2013, 25 (46), 6730-6736. 34. Khalil, H.; Chen, T.; Riffon, R.; Wang, R.; Wang, Z., Synergy between polyethylenimine and different families of antibiotics against a resistant clinical isolate of Pseudomonas aeruginosa. Antimicrob. Agents Chemother. 2008, 52 (5), 1635-1641. 35. Nguyen, T.-K.; Lam, S. J.; Ho, K. K. K.; Kumar, N.; Qiao, G. G.; Egan, S.; Boyer, C.; Wong, E. H. H., Rational Design of Single-Chain Polymeric Nanoparticles That Kill Planktonic and Biofilm Bacteria. ACS Infect. Dis. 2017, 3 (3), 237-248. 36. Namivandi-Zangeneh, R.; Kwan, R. J.; Nguyen, T.-K.; Yeow, J.; Byrne, F. L.; Oehlers, S. H.; Wong, E. H. H.; Boyer, C., The effects of polymer topology and chain length on the antimicrobial activity and hemocompatibility of amphiphilic ternary copolymers. Polym. Chem. 2018, 9 (13), 1735-1744. 37. Judzewitsch, P. R.; Nguyen, T.-K.; Shanmugam, S.; Wong, E. H. H.; Boyer, C., Towards Sequence-Controlled Antimicrobial Polymers: Effect of Polymer Block Order on Antimicrobial Activity. Angew. Chem., Int. Ed. 2018, 57 (17), 4559-4564. 38. Nguyen, T.-K.; Selvanayagam, R.; Ho, K. K. K.; Chen, R.; Kutty, S. K.; Rice, S. A.; Kumar, N.; Barraud, N.; Duong, H. T. T.; Boyer, C., Co-delivery of nitric oxide and antibiotic using polymeric nanoparticles. Chem. Sci. 2016, 7 (2), 1016-1027. 39. Namivandi-Zangeneh, R.; Sadrearhami, Z.; Bagheri, A.; Sauvage-Nguyen, M.; Ho, K. K. K.; Kumar, N.; Wong, E. H. H.; Boyer, C., Nitric Oxide-Loaded Antimicrobial Polymer for the Synergistic Eradication of Bacterial Biofilm. ACS Macro Lett. 2018, 7 (5), 592-597. 40. Oo, T. Z.; Cole, N.; Garthwaite, L.; Willcox, M. D. P.; Zhu, H., Evaluation of synergistic activity of bovine lactoferricin with antibiotics in corneal infection. J. Antimicrob. Chemother. 2010, 65 (6), 1243-1251. 41. Willcox, M. D. P.; Hume, E. B.; Schubert, T. L.; Kumar, N., Treatment of Staphylococcus Aureus with Antibiotic and Quorum-Sensing Inhibitor Combinations Reduces Severity of Keratitis. J Ocular Biol 2017, 5 (1), 1-5. 42. Moad, G.; Rizzardo, E.; Thang, S. H., RAFT Polymerization and Some of its Applications. Chem. Asian J. 2013, 8 (8), 1634-1644. 43. Boyer, C.; Bulmus, V.; Davis, T. P.; Ladmiral, V.; Liu, J.; Perrier, S., Bioapplications of RAFT Polymerization. Chem. Rev. 2009, 109 (11), 5402-5436. 44. Rossiter, S. E.; Fletcher, M. H.; Wuest, W. M., Natural Products as Platforms To Overcome Antibiotic Resistance. Chem. Rev. 2017, 117 (19), 12415-12474. 45. Kohanski, M. A.; Dwyer, D. J.; Collins, J. J., How antibiotics kill bacteria: from targets to networks. Nat. Rev. Microbiol. 2010, 8, 423-435. 46. Dutta, D.; Vijay, A. K.; Kumar, N.; Willcox, M. D. P., Melimine-Coated Antimicrobial Contact Lenses Reduce Microbial Keratitis in an Animal Model. Investig. Ophthalmol. Vis. Sci 2016, 57 (13), 5616-5624. 47. Dutta, D.; Cole, N.; Kumar, N.; Willcox, M. D. P., Broad Spectrum Antimicrobial Activity of Melimine Covalently Bound to Contact Lenses. Investig. Ophthalmol. Vis. Sci. 2013, 54 (1), 175-182. 48. Subedi, D.; Vijay, A. K.; Kohli, G. S.; Rice, S. A.; Willcox, M., Association between possession of ExoU and antibiotic resistance in Pseudomonas aeruginosa. PLoS One 2018, 13 (9), 1-14.

Page | 182 CHAPTER FOUR

4.6 Appendix B

Figure B 1. 1H NMR spectra of the Boc-protected P in CDCl3 (a) and the corresponding Boc- deprotected P in D2O (b).

Figure B 2. GPC molecular weight distributions of the Boc-protected P.

Page | 183 CHAPTER FOUR

Table B 1. Checkerboard assay results indicating the interaction between P and selected antibiotics against P. aeruginosa PAO1.

MIC (µg ml-1)

Antibiotic Polymer

In In Antibiotic Alone Alone FICI Combination Combination

1 8 0.38 Doxycycline 8 32 2 8 0.5

Clarithromycin 128 16 32 8 0.38

Azithromycin 64 0.5 32 16 0.51

Gentamicin 0.5 0.125 64 32 0.75

0.25 16 0.75 Tobramycin 0.5 64 0.125 32 0.75

4 4 0.38 Colistin 16 32 2 8 0.38

Ampicillin 256 64 32 16 0.75

Amoxicillin 256 2 64 32 0.51

Ceftriaxone 8 4 32 16 1

0.0312 16 0.75 Ciprofloxacin 0.125 32 0.0625 2 0.56

Page | 184 CHAPTER FOUR

Table B 2. Checkerboard assay results indicating the interaction between P and selected antibiotics against E. coli K12.

MIC (µg ml-1)

Antibiotic Polymer

In In Antibiotic Alone Alone FICI Combination Combination

Doxycycline 2 0. 5 32 8 0.5

Clarithromycin 16 8 32 4 0.63

0.5 16 0.75 Azithromycin 2 32 1 8 0.75

1 8 0.75 Gentamicin 2 32 0.5 16 0.75

0.25 16 0.75 Tobramycin 1 32 0.5 8 0.75

Colistin 32 8 32 8 0.5

Ampicillin 2 1 32 8 0.75

Amoxicillin 2 0.25 32 16 0.63

0.00781 16 0.63 Ceftriaxone 0.0625 32 0.0312 8 0.75

Ciprofloxacin 0.0156 0.0039 64 32 0.75

Page | 185 CHAPTER FOUR

Table B 3. Log10 reduction of viable P. aeruginosa bacterial colonies upon treatment with P, D, C and various concentrations of PD and PC (in µg mL–1), as determined using CFU analysis.

Log10 reductiona

PAO1 PAO1 PA 6294 Entry Entry Entry Planktonic Biofilm Planktonic Biofilm Planktonic Biofilm

P 3.07±0.24 2.21±0.42 P 3.61±0.13 2.81±0.24 P 3.06±0.64 3.02±0.68

- D 0.26±0.28 0.22±0.31 C 0.45±0.28 0.58±0.24 D 0.46±0.19 0.09±0.19

PD PC PD 5.41±0.29 5.47±0.43 5.15±0.59 4.25±0.58 6.18±0.10 3.34±0.08 128/64 128/64 128/64

PD PC PD 5.28±0.61 2.99±0.56 4.38±0.47 3.23±1.56 6.01±0.39 4.44±0.26 128/32 128/32 128/32

PD PC PD 4.79±0.48 2.94±0.84 4.13±0.56 2.70±0.79 5.95±0.87 4.75±0.14 128/16 128/16 128/16 aValues are the mean of three independent experiments ± standard deviation.

Page | 186

CHAPTER FIVE

An Antibiofilm Platform based on the

Combination of Antimicrobial Polymers and Essential Oils

CHAPTER FIVE

5 An Antibiofilm Platform based on the Combination of

Antimicrobial Polymers and Essential Oils

This chapter is based on the work published in Biomacromolecules, 2020, 21,

262–272.

5.1 Introduction

Antibiotic resistance is a critical healthcare issue worldwide.1-3 Unnecessary and prolonged use of antibiotics in the medical and agricultural sectors has caused public health systems to reach the threshold of a postantibiotic era, rendering the treatment of bacterial infections more problematic and costly than it has ever been.4-7

The ability of bacteria to form biofilm on the surface of body tissues and medical devices makes combating bacteria even more challenging, especially against multidrug-resistant (MDR) strains.8, 9 Bacterial biofilms are highly resistant to most of the therapeutic approaches, as they are protected by a matrix

Page | 188 CHAPTER FIVE

of extracellular polymeric substances. This -based matrix, composed of proteins, polysaccharides, and DNA, acts as both physical and chemical barriers to reduce antibiotic influx, while at the same time accelerates the resistance development rate by facilitating the transfer of quorum-sensing molecules and chromosomal plasmids between bacteria cells within the biofilm community.10-13

Development of novel therapeutic platforms that are especially effective against bacterial biofilm might avert the spread of antibiotic resistance.14, 15 An antibiofilm agent can target one or multiple stages in biofilm development by, for instance, inhibiting the formation of bacterial biofilm in the first place,16-19 inducing biofilm dispersal 20-22 or direct killing of biofilm.23-25 Most of the currently available antibiotics, however, lack the efficiency against bacterial biofilm and are highly prone to trigger antibiotic resistance in bacteria even after a few treatments.26, 27 Thus, alternative antimicrobial agents/strategies that are capable of fighting bacterial biofilm and hinder the acquisition of drug resistance will be highly advantageous in new treatment protocols.

Essential oils are natural antimicrobial agents produced as secondary metabolites by many plants.28, 29 These multi-component low molecular weight oils commonly contain terpenes and terpenoids components.30 The antimicrobial activity of essential oils is mostly considered to be associated with the phenolic components, where the nature and relative position of substitutions determine the antimicrobial efficiency.29, 31 Carvacrol and eugenol are major components of oregano and clove oil, respectively.32 These compounds are easy to extract and pose antimicrobial/antibiofilm activity against a wide range of Gram-negative

Page | 189 CHAPTER FIVE

and Gram-positive bacteria.33-35 However, their potential clinical application has been greatly hindered by their poor distribution in aqueous media and low stability. Encapsulation and delivery platforms that are exclusively designed to counter these drawbacks for antimicrobial purposes are of high interest as potential alternatives for current antibiofilm agents.36-38 In this regard, degradable polymeric systems with controllable release of essential oils have demonstrated promising antimicrobial activity against bacterial biofilm.39-42

Synthetic antimicrobial polymers could potentially overcome the challenges associated with the use of essential oils for therapeutic purposes, by acting as both a coagent and a delivery vehicle. Inspired by the fundamental composition of natural antimicrobial peptides,43, 44 synthetic antimicrobial polymers exert their antimicrobial activity through a contact-based membrane disruption mechanism with minimal risk of resistance development in bacteria.45-47 Using advanced polymerization techniques,48-54 bespoke antimicrobial polymers can be synthesized to fulfil the requirements of a delivery system.55-58 In addition to providing supplementary antimicrobial activity, the use of synthetic antimicrobial polymers as delivery vehicles enables the localized delivery of essential oils to bacterial biofilm and, therefore better antimicrobial activity. The electrostatic interactions between the cationic motifs of antimicrobial polymers and the anionic components of extracellular polymeric substances and bacteria cell wall promote the targeted delivery of essential oils to the biofilm.20

In this study, we report an effective antibiofilm platform based on the use of synthetic antimicrobial polymers as delivery vehicles for essential oils. Two antimicrobial polymers in the form of random and block ternary copolymers

Page | 190 CHAPTER FIVE

were synthesized and used in combination with carvacrol and eugenol as oil-in- water emulsions. The potency of these combinations against MDR Pseudomonas aeruginosa biofilm is investigated via biofilm inhibition and killing assays (Figure

5.1). It is worthwhile noting that P. aeruginosa is an opportunistic Gram-negative that contributes to many biofilm-associated diseases such as cystic fibrosis lung infection, contact lens-related keratitis, chronic wound infection, and catheter-associated urinary tract infection. 9, 15, 27 In addition, the World

Health Organization (WHO) has recently urged for the development of new treatments to combat P. aeruginosa infections. 59

Figure 5.1. Combination of synthetic antimicrobial block copolymer and essential oils as oil- in-water emulsion (left) and bacterial biofilm inhibition and eradication induced by this combination (right).

Page | 191 CHAPTER FIVE

5.2 Experimental Section

Materials

Ethylenediamine (Sigma-Aldrich, ≥ 99%), di-tert-butyl dicarbonate (Aldrich,

99%), triethylamine (Scharlau, 99%), acryloyl chloride (Merck, ≥ 96%), oligo(ethylene glycol) methyl ether acrylate (OEGA) Mn 480 g mol–1 (Aldrich), 2- ethylhexyl acrylate (Aldrich, 98%), N-hydroxyethyl acrylamide (Sigma-Aldrich,

97%), benzyl acrylate (99.9%), trifluoroacetic acid (TFA) (Sigma-Aldrich, 99%), dichloromethane (DCM, Merck), chloroform (Merck), tetrahydrofuran (THF,

Merck), hexane (Merck), diethyl ether (Merck), dimethyl sulfoxide (DMSO,

Merck), anhydrous magnesium sulfate (MgSO4, Sigma-Aldrich, ≥99.5%), basic aluminium oxide (Al2O3, Sigma-Aldrich, 99.9%) and 5,10,15,20-tetraphenyl-

21H,23H-porphinezinc (ZnTPP, Sigma-Aldrich) were used as received. 2,2’- azobis (2 methylpropionitrile) (AIBN) (Acros, 98%) was purified by recrystallization from methanol. Milli-Q water with a resistivity of > 18 MΩ⋅cm was obtained from an in-line Millipore RiOs/Origin water purification system.

Essential oils (carvacrol 99% and eugenol 99%) were purchased from Sigma-

Aldrich and were used as received.

Analytical Instruments

Nuclear magnetic resonance (NMR) spectra were obtained using a Bruker

AC300F spectrometer at the sample concentration of about 10-20 mg mL–1 in deuterated solvents D2O, CDCl3 or DMSO (obtained from Cambridge Isotope

Laboratories).

Page | 192 CHAPTER FIVE

Gel permeation chromatography (GPC) analysis was performed using a

Shimadzu liquid chromatography system equipped with a Shimadzu refractive index detector and two MIX C columns (Polymer Lab) operating at 40 °C. DMAc

(containing 0.03% (w/v) LiBr and 0.05% (w/v) 2,6-dibutyl-4-methylphenol (BHT)) was used as the eluent at a flow rate of 1 mL min–1. The system was calibrated with PMMA standards with molecular weights of 200 to 106 g mol–1.

Dynamic light scattering (DLS) and zeta-potential measurements were conducted using a Malvern Zetasizer Nano ZS apparatus equipped with a He-

Ne laser operated at λ = 633 nm and at a scattering angle of 173°. Samples were prepared at a total concentration of ca. 1 mg mL–1 where filtered Milli-Q water

(using 0.45 μm pore size filter) was used as the solvent to solubilize the polymers and prepare the emulsions.

Transmission electron microscopy (TEM) was performed using JEOL-1400 at an accelerating voltage of 100 kV. Samples were prepared by placing a droplet of a 2 mg mL–1 polymer solution in water on a carbon-coated copper grid. Uranyl acetate staining was applied to increase contrast.

UV-Visible spectroscopy was conducted to measure the transmittance of the emulsions at 600 nm using a CARY 3000 spectrometer from Bruker. Samples were prepared at a total concentration of about 4 mg mL–1 where Milli-Q water was used as the solvent to prepare the emulsions. The emulsions were stored in glass vials and kept in the dark at 25 °C during the experiment.

Page | 193 CHAPTER FIVE

Synthesis of antimicrobial and control polymers

Synthesis of cationic monomer

Cationic monomer tert-butyl (4-acrylamidobutyl) carbamate was prepared in the same manner as reported previously.60 Briefly, ethylenediamine (0.3 mol) was dissolved in DCM (400 mL), followed by the dropwise addition of di-tert-butyl dicarbonate (0.03 mol in 100 mL DCM) over 2 h at 0−5 °C. The reaction mixture was stirred overnight at 25 °C. The reaction mixture was washed exhaustively with water using a separation funnel to remove excess diamines. The organic layer was then dehydrated over MgSO4, filtered, and dried using a rotary evaporation unit to yield a pale-yellow oil. Tetrahydrofuran (100 mL) was added to dissolve the intermediate product. Triethylamine (36 mmol) and acryloyl chloride (31.5 mmol) were added dropwise to the solution at 0−5 °C with N2 bubbling. The contents were stirred at 25 °C for 1 h. The byproducts were filtered, and the solvent was removed in vacuo. The crude product was dissolved in chloroform (150 mL) and washed against 0.1 M HCl solution (1 × 75 mL), saturated NaHCO3 (1 × 75 mL), brine (1 × 75 mL), and water (1 × 75 mL) . The organic phase was stirred with MgSO4 and basic Al2O3 for 10 min, filtered, and concentrated in vacuo. The product was further purified by repeated precipitation steps in hexane to yield the tert-butyloxycarbonyl (Boc)-protected monomer as a fine white powder, which was dried in vacuo.

tert-Butyl (2-acrylamidoethyl) carbamate: 1H NMR (300MHz, CDCl3,25 °C), δH

6.56 (br s, 1H, NH), 6.28 (dd, J = 17.1 Hz, 1.5 Hz, 1H, CHH=CH), 6.12 (dd, J = 17.1

Hz, 10.2Hz, 1H, CHH=CH), 5.65 (dd, J = 10.2 Hz, 1.5 Hz, 1H, CHH=CH), 5.05 (br s, 1H, NH), 3.49−3.41 (m, 2H, CH2), 3.34−3.28 (m, 2H, CH2), 1.45 (s, 9H, CH3); 13C

Page | 194 CHAPTER FIVE

NMR (300 MHz, CDCl3,25 °C), δC 166.23, 157.50, 130.88, 126.30, 79.85, 41.05, 40.09,

28.35.

Synthesis of random, block and control copolymers

Random copolymer (RP): The synthesis of amphiphilic ternary random copolymer proceeded in the same manner as reported previously.60 Briefly, The

RAFT agent benzyl dodecyl carbonotrithioate (12 μmol), AIBN (3 μmol), OEGA

(360 μmol), tert-butyl (4-acrylamidobutyl) carbamate (600 μmol), and 2- ethylhexyl acrylate (240 μmol) were dissolved in 1,4-dioxane (such that the total monomer concentration in solvent is 1 M). The solution was purged with N2 for

20 min in an ice bath. The polymerization was conducted for 17 h at 70 °C and then quenched in an ice bath and exposing to air. The polymer was purified by precipitation in a hexane/diethyl ether (7:3) mixture thrice and subsequently dried in vacuo. The monomer composition of the RP was calculated as followed:

ʃa/9 : ʃb/2 : ʃc,d/6 where ʃa, ʃb, and ʃc,d correspond to the integrals of the characteristic protons of cationic monomer (tert-butyl -CH3- groups, δH 1.38-1.52 ppm), OEGA (ester -CH2O- group, 4.0-4.30 ppm) and 2-ethylhexyl acrylate

(methyl -CH3- groups, δH 0.80-0.98 ppm), respectively (Figure C 1).

Block copolymer (BP): To prepare the diblock copolymer, a macroRAFT agent was firstly synthesized in the same manner as mentioned for the random copolymer except that the polymerisation time was reduced to 3.5 h to minimize dead chains. The macroRAFT agent was then chain extended with benzyl acrylate. For the chain extension step, the macroRAFT agent (5 μmol), benzyl acrylate (250 μmol) and AIBN (1.25 μmol) were dissolved in 1,4-dioxane and the solution was degassed by bubbling with N2 for 20 min in an ice bath. Then, the

Page | 195 CHAPTER FIVE

reaction mixture was stirred at 70 °C for 17 h. Polymerization was quenched by placing the flask in an ice bath for 5 min. The polymer was purified by precipitation into a diethyl ether/hexane (3:7) mixture. The precipitate was isolated by centrifugation, dissolved in methanol, and precipitated twice more.

Finally, the polymer was dried in vacuo. The monomer composition of the BP was calculated as followed: ʃa/9 : ʃb/2 : ʃc,d/6 : ʃe/5 where ʃa, ʃb, ʃc,d and ʃe correspond to the integrals of the characteristic protons of cationic monomer

(tert-butyl -CH3- groups, δH 1.38-1.52 ppm), OEGA (ester -CH2O- group, 4.0-4.30 ppm), 2-ethylhexyl acrylate (methyl -CH3- groups, δH 0.80-0.98 ppm) and benzyl acrylate (benzyl group -C6H5- group, δH 7.07-7.4 ppm), respectively (Figure C 4).

Control block copolymer (CP): A control block copolymer was synthesized by replacing the tert-butyl (4-acrylamidobutyl) carbamate monomer with N- hydroxyethyl acrylamide in the macroRAFT agent and polymerization was proceeded using PET-RAFT polymerization50, 51 under green light. The monomer composition of the CP was calculated as followed: ʃa/1 : ʃb/2 : ʃc,d/6 : ʃe/5 where

ʃa, ʃb, ʃc,d and ʃe correspond to the integrals of the characteristic protons of N- hydroxyethyl acrylamide (hydroxyl -OH- group, δH 4.4-5.17 ppm), OEGA (ester

-CH2O- group, 4.0-4.30 ppm), 2-ethylhexyl acrylate (methyl -CH3- groups, δH 0.80-

0.98 ppm) and benzyl acrylate (benzyl -C6H5- group, δH 7.07-7.4 ppm), respectively (Figure C 7).

The Boc protecting groups were removed using TFA in the same manner as reported previously.60 In general, the polymer solution in dichloromethane (ca.

10 wt % polymer) was treated with TFA (20 mol equivalent with respect to the

Boc group) for 3 h at 25 °C. Boc-deprotected polymer was subsequently

Page | 196 CHAPTER FIVE

precipitated into diethyl ether. The precipitate was isolated by centrifugation, dissolved in methanol, and reprecipitated two more times. The polymer was then dried in vacuo and further purified by dialysis against water (Cellu-Sep 3500

MWCO). The aqueous solution was lyophilized to yield the Boc-deprotected polymers.

Preparation of emulsions

Emulsions were prepared by adding the oil phase (10% v/v in DMSO) into aqueous media (containing polymer). The mixtures were emulsified in an ultrasonic bath (Thermoline Scientific, Powersonic410) at full sonic power for 30 s at room temperature.

Critical Micelle Concentration (CMC) determination

The CMC was measured by fluorescence spectroscopy using pyrene as a fluorescent probe. Briefly, 48 μL of a stock solution of pyrene in DCM (2.5 × 10−5

M) was dropped into empty vials, and the solvent was evaporated under reduced pressure. A stock solution of polymer was serially diluted with deionized water to a concentration range of 0.5−512 μgmL−1. A total of 2 mL of each polymer solution was transferred to vials containing pyrene, such that the final pyrene concentration was 6 × 10−7 M in each vial, and the solution was stirred overnight.

Fluorescence measurements were performed using an excitation wavelength of

λ = 333 nm, using a 3 nm slit width for excitation and a 1.5 nm slit width for emission. Emission wavelengths were scanned from 350 to 450 nm. The intensities of the I1 (372 nm) to I3 (383 nm) vibronic bands were evaluated for each sample, and the ratios of these were plotted against the log of the concentration.

Page | 197 CHAPTER FIVE

The CMC was taken as the intersection of two regression lines calculated from the linear portions of the graphs.

Minimum Inhibitory Concentration (MIC) determination

The MIC of the polymers was determined via broth microdilution method according to Clinical and Laboratory Standards Institute (CLSI) guidelines.

Briefly, a single colony was culture in 10 mL of Mueller-Hinton broth (MHB) at

37 °C with shaking at 180 rpm overnight. Subsequently, a subculture was prepared from the overnight culture by diluting 1:100 in 10 mL of MHB and allowed to grow to mid-log phase, then diluted to the appropriate concentration for the MIC test. A 2-fold dilution series of 100 μL of polymers solution or oils emulsion in MHB were added into 96-well microplates followed by the addition of 100 μL of the subculture suspension. The final concentration of bacteria in each well was about 5 × 105 cells mL–1. The plates were incubated at 37 °C for 20 h, and the absorbance at 595 nm was measured with a microtiter plate reader (FLUOstar

Omega, BMG Labtech). MIC values were defined as the lowest concentration of sample that showed no visible growth and inhibited cell growth by more than

90%. Positive controls with no treatment and negative controls without bacteria were included. All assays included two replicates and were repeated in at least two independent experiments.

Biofilm inhibition study

The Gram-negative bacteria strains of P. aeruginosa PAO1 and PA37 were used to grow biofilm for this study. A single colony of PAO1 or PA37 was inoculated in 10 mL of Luria-Bertani medium (LB 10) at 37 °C with shaking at 180 rpm

Page | 198 CHAPTER FIVE

overnight. The overnight culture was diluted 1:200 in freshly prepared M9 supplemented medium which might contain (i) random, block or control copolymers at 4−8 μg mL−1, (ii) carvacrol or eugenol at 300 and 530 μg mL−1, respectively, (iii) different combinations of polymers and essential oils at the abovementioned concentrations, or (iv) no additional compound (i.e., control biofilm). The bacterial suspension was then aliquoted 1 mL per well of tissue- culture treated 24-well plates (Costar, Corning®). The plates were incubated at

37 °C with shaking at 180 rpm in an orbital shaker that does not stop agitation when the door is opened (model OM11, Ratek, Boronia, Australia) and the biofilm cultures were allowed to grow without any disruption. After 6.5 h of growth, the planktonic biomass was quantified by siphoning off the supernatant and measuring the OD595. The remaining biofilm was washed once with PBS (1.0 mL), before adding crystal violet stain (1 mL 0.03% CV stain made from a 1:10 dilution of Gram crystal violet (BD) in PBS). The plates were then incubated on the bench for 20 min before washing the wells twice with PBS (1 mL). The CV- stained biofilms were mixed with 1 mL of 100% ethanol and quantified by measuring the OD595 of the homogenized suspension using a microtiter plate reader (FLUOstar Omega, BMG Labtech). All assays included two replicates and were repeated in at least two independent experiments.

Biofilm imaging

In a similar manner as the biofilm inhibition study, PAO1 biofilms were grown in the presence of different compounds at various concentrations in 35 mm tissue culture dishes (FluoroDish, World Precision Instruments Inc.,

Sarasota, FL, U.S.A.). The supernatant was removed, the biofilm on the culture

Page | 199 CHAPTER FIVE

dish surface was washed with 1 mL of PBS, and then 1 mL of PBS was added into the dish. The dishes were analyzed using a 3D tomographic microscope (3D Cell

Explorer, NanoLive, Lausanne, Switzerland) that was equipped with a digital staining software. All assays were repeated in at least two independent experiments.

Biofilm killing study

To evaluate the bactericidal activity of the selected compounds and combinations, P. aeruginosa PAO1 biofilms were grown in M9 supplemented medium, as mentioned above. After 6.5 h, the planktonic phase was removed and freshly prepared M9 supplemented medium, which contains different compounds at various concentrations, was gently added to each well, and the plates were incubated for 20 min. After treatment, the biofilm viability analysis was determined by a drop plate method. For this, bacteria cells attached on the interior surfaces of the well (surface area 4.5 cm2) were washed once with sterile

PBS to remove loosely attached bacteria, before being resuspended and homogenized in PBS by incubating in an ultrasonication bath (150 W, 40 kHz;

Unisonics, Australia) for 20 min. Resuspended biofilm cells were then serially diluted in PBS and plated onto LB agar. Biofilm colonies were counted and CFU was calculated after a 24 h incubation at 37 °C. All assays included two replicates and were repeated in at least two independent experiments.

Hemolysis study

The hemolytic activity of the individual polymers, essential oils, and combinations was assessed using fresh sheep red blood cells (RBCs) obtained

Page | 200 CHAPTER FIVE

from Serum Australis (Catalog number SD50D). RBCs were diluted 1:20 in PBS

(pH 7.4), pelleted by centrifugation and washed three times with PBS (1000g, 10 min). The RBCs were then resuspended to achieve 5% (v/v) in PBS. Different concentrations of samples (150 μL) were prepared in sterilized tubes, followed by the addition of the RBC suspension (150 μL). PBS buffer was used as a negative control while Triton-X 100 (1% v/v in PBS) was used as a positive hemolysis control. The tubes were incubated for 2 h at 37 °C and 150 rpm shaking speed in an incubator. Following incubation, the tubes were centrifuged (1000g,

8 min) and aliquots of the supernatants (100 μL) were transferred into a 96-well microplate, where the absorbance values were monitored at 485 nm using a microtiter plate reader (FLUOstar Omega, BMG Labtech). The percentage of hemolysis was calculated using the absorbance values and the formula below:

% Hemolysis = (Apolymer − Anegative)⁄(Apositive − Anegative) × 100

5.3 Results and Discussion

5.3.1 Synthesis and characterization of antimicrobial and control polymers

We have previously demonstrated that random ternary copolymers consisting of low-fouling oligo(ethylene glycol), cationic primary amines and hydrophobic groups (in ca. 3:5:2 molar ratio) generally exhibit good antimicrobial activity and biocompatibility,52, 60 either as an individual component or in combination with other antimicrobial agents such as nitric oxide20 or commercially available antibiotics.23 The strategic incorporation of oligo(ethylene glycol), primary amine, and hydrophobic (e.g., ethylhexyl) functionalities into a polymer chain

Page | 201 CHAPTER FIVE

endows the macromolecule with a low-fouling property that minimizes protein complexation in biological media, bacteria binding, and membrane disruption capabilities, respectively. Given the presence of hydrophobic groups (albeit at only ca. 20 mol %), we postulate that an antimicrobial random ternary copolymer may stabilize essential oil droplets to some extent and, as such, was used as an initial reference point in this study. Reversible addition−fragmentation chain transfer (RAFT) polymerization technique48, 49 was employed to synthesize all polymers in this study, including the random ternary copolymer (herein denoted as RP; Figure 5.2). RP was made in the same way as previously described,52, 60 where a Boc-protected amino monomer was copolymerized with oligo(ethylene glycol) acrylate and 2-ethylhexyl acrylate via RAFT followed by Boc removal and purification steps. 1H NMR analysis confirmed the chemical composition of RP which was similar to the molar feed ratio (Figure C 1). The number-averaged molecular weight (Mn) and dispersity (Đ) values of the Boc-protected RP were estimated to be 25400 g mol−1 and 1.19, respectively, based on GPC analysis

(Figure C 2).

To ensure that an antimicrobial polymer can sufficiently stabilize oil droplets and act as a carrier, another antimicrobial polymer in the form of a diblock copolymer (BP) was synthesized by incorporating RP with a second hydrophobic homopolymer block composed of benzyl pendant groups as a representative hydrophobic group (Figure 5.2). This hydrophobic block induces the self- assembly of antimicrobial polymers into a core−shell micelle morphology, which we believe will increase the encapsulation efficiency of essential oils and allow the polymer to perform efficiently as a dual antimicrobial-carrier agent. For this, a macroRAFT agent that is essentially Boc-protected RP, was subsequently chain

Page | 202 CHAPTER FIVE

extended with 50 repeat units of benzyl acrylate. 1H NMR analysis confirmed the incorporation of the second poly(benzyl acrylate) block (Figure C 4). Successful chain extension was also confirmed by GPC analysis as the molecular weight distribution of Boc-protected BP shifted to shorter retention times compared to the macroRAFT agent (Figure C 5). BP have Mn and Đ values of 34500 g mol−1 and 1.22, respectively, based on GPC analysis (Table C 1).

Finally, RP and BP were treated with an excess of TFA to remove the Boc- protecting groups. Successful deprotection of Boc groups was verified by the absence of tert-butyl protons at δH 1.45 ppm in their 1H NMR spectrum (Figure C

1 and Figure C 4).

To decouple the antimicrobial activity of the polymer from its function as a delivery vehicle, we have also synthesized a non-antimicrobial block copolymer, that is control polymer (CP), by simply replacing the Boc-protected cationic monomer with hydrophilic N-hydroxyethyl acrylamide (Figure 5.2). 1H NMR and GPC analysis verified the chemical structure of CP (Figure C 7 and Figure C

8 and Table C 1).

Page | 203 CHAPTER FIVE

Figure 5.2. Chemical structure of the synthesized polymers and essential oils investigated in this study (top). Schematic illustrations of the emulsions containing antimicrobial polymers and essential oils in aqueous media (bottom).

The self-assembly of the synthesized polymers in aqueous solution was followed by fluorescence spectroscopy using pyrene as a fluorescent probe. The critical micelle concentration (CMC), above which the formed micelles are thermodynamically stable, was determined to be 3.31 and 2.47 μg mL−1 for BP and CP, respectively (Figure C 6 and Figure C 9 and Table C 1). Such low CMC values are especially desirable for biological applications, as they require the use of stable micelles at the lowest concentrations possible. However, in the case of

RP we observed relatively minor changes in I1/I3 over the concentration range, compared to BP and CP (Figure C 3 and Table C 1). Driven by intramolecular interactions among the hydrophobic motifs, RP undergoes self-assembly in aqueous media to form single chain polymeric nanoparticles.60 The observed transition in fluorescence spectroscopy might be attributed to the assembly of multi chain nanoparticles, which is most likely induced by the presence of pyrene.

Page | 204 CHAPTER FIVE

5.3.2 Preparation and characterization of polymer-oil combinations

The combinations of synthetic polymers and essential oils were prepared as oil-in-water emulsions by emulsifying essential oils into an aqueous solution containing polymers at various concentrations, using an ultrasonic bath for 30 s

(Figure 5.1). For example, the emulsion containing RP at 32 μg mL−1 and carvacrol (Car) at 300 μg mL−1, referred to as RP/Car at 32/300 μg mL−1, was prepared by emulsifying 3 μL of Car (10% v/v in DMSO) into 1 mL of the aqueous solution of RP at 32 μg mL−1. Emulsions containing individual eugenol (Eug) and

Car in aqueous media were also prepared to serve as control samples. All the emulsions namely, RP/Car, RP/Eug, BP/Car, BP/Eug, CP/Car, CP/ Eug, Car, and

Eug, were freshly prepared 1 h prior to further experiments.

To evaluate the hydrodynamic diameter (Dh) of individual components and their combinations in aqueous media, DLS measurements were conducted in water (Figure 5.3 and Table 5.1). DLS analysis was performed at the overall concentration of 1 mg mL−1, where the weight ratio between polymer and oil for each combination was identical to the weight ratio at the highest concentration used in the killing study (Table C 2). It is worthwhile noting that in all BP containing samples, the polymer concentration was above the corresponding

CMC value. BP and CP showed Dh values of 88 and 34 nm, respectively (Figure

5.3 b and c), which support the formation of micelle morphologies. Since the single chain polymeric nanoparticles are small and very challenging to analyze by DLS, we therefore used transmission electron microscopy (TEM) to analyze the size and morphology of RP (in the dry state). BP was also analyzed by TEM.

Page | 205 CHAPTER FIVE

RP showed an average diameter of 17 ± 3 nm whereas BP formed larger micelles with an average diameter of 62 ± 16 (Figure C 10).

Diameter / nm

Figure 5.3. DLS normalized size distribution of RP/Car, RP/Eug, Car and Eug (a), BP, BP/Car and BP/Eug (b) and CP, CP/Car, CP/ Eug (c) in water.

The Dh for Car and Eug emulsions based on DLS was determined to be 159 and 246 nm, respectively (Figure 5.3 a). We observed very similar Dh values for

Car and RP/Car emulsions and also for Eug and RP/Eug emulsions. Based on these results, we postulate that RP unimer micelles lack the capacity to encapsulate the oil within their hydrophobic cavities and, instead merely interact with the oil at the oil-water interface without altering the size of the original oil droplet (Figure 5.2). Meanwhile for BP/Car, BP/Eug, CP/Car and CP/Eug, the Dh

Page | 206 CHAPTER FIVE

values of these combinations resemble those of BP and CP alone, indicating that the additional hydrophobic block in these block copolymers allows the encapsulation of oil droplets into the hydrophobic core of BP and CP micelles akin to a drug-loading process.

To verify any possible interaction between antimicrobial polymers and essential oils, we followed the phase separation of the emulsions using UV−Vis spectroscopy (Figure C 11). The transmittance of the emulsions at 600 nm was monitored for 8 days and compared to the values of the freshly prepared emulsions (day 1). We observed slower phase separation for all RP and BP containing combinations compared to individual Car and Eug emulsions. This suggests the ability of amphiphilic antimicrobial copolymers to at least partially stabilize oil droplets. In corroboration with DLS data, BP micelles performed better than RP unimer micelles in stabilizing oil droplets.

Table 5.1. Hydrodynamic diameter (Dh) and zeta potential (ζ) values of individual compounds and combinations used in this study.

Entry Dh (nm) ζ (mV) RP - +29 BP 88 +34 CP 34 –17 Car 159 –37 Eug 246 –35 RP/Car 162 +40 RP/Eug 216 +29 BP/Car 77 +41 BP/Eug 75 +25 CP/Car 36 –29 CP/Eug 40 –20

Page | 207 CHAPTER FIVE

In addition to DLS analysis, zeta potential (ζ) measurements were also performed (Table 5.1). Expectedly, RP and BP showed net positive charges of

+29 and +34 mV, respectively, which confirms the cationic character of these polymers due to the presence of primary amino groups, while CP yielded neutral to slightly negative charge of −17 mV. Car and Eug both showed net negative charges of −37and −35 mV, respectively. We observed a net positive charge for all

RP and BP containing combinations while CP/Car and CP/Eug both revealed negative charges. The observed switch in the surface charge from negative to positive upon the emulsification of oils in either RP or BP solutions suggests that the interaction between essential oils and bacteria in RP and BP oil combinations will be higher than oils alone. We believe that RP and BP facilitate the targeted delivery of essential oils to bacteria cells. This is in line to what we observed previously for nitric-oxide loaded RP.20

5.3.3 Determination of the MIC values of synthesised polymers

The antimicrobial activity of the individual polymers was first assessed in vitro against two strains of P. aeruginosa, wild- type PAO1 and MDR PA37, by determining their minium inhibitory concentration (MIC) values (Table 5.2). RP and BP showed bacteriostatic activity against both strains tested, where RP was slightly more potent (one-fold better) than BP. The lower activity of BP might be attributed to the micelle morphology of this polymer in aqueous media which limits its ability to interact with the bacteria cell membrane.61 CP, on the other hand, was inactive, as predicted with MIC values >1024 μg mL−1 because of the absence of cationic moieties.

Page | 208 CHAPTER FIVE

Table 5.2. The antimicrobial activities of synthesized polymers against P. aeruginosa PAO1 and PA37.

MIC (μg mL−1) Entry PAO1 PA37 RP 32 64 BP 64 128 CP >1024 >1024

5.3.4 Biofilm inhibition activity of polymer-oil combinations and individual

components

To investigate the efficiency of the combinations based on antimicrobial polymers and essential oils in combating bacterial biofilm, biofilm inhibition experiments were performed. For this, P. aeruginosa bacterial cultures in M9 media were incubated for 6.5 h with or without the addition of selected compounds at various concentrations, followed by quantification of biofilm formation using the standard crystal violet (CV) staining method.62 Individual polymers and essential oils were included as control samples. A preliminary biofilm inhibition test was used to select the concentrations of individual compounds for this test. Selected concentrations, which are lower than the MIC values for each compound, are intentionally chosen to show low to medium biofilm inhibition activity for ease of detection of any possible synergistic effect in the polymer−oil combinations. Planktonic growth inhibition was also studied by measuring the optical density (OD595) of the bacterial culture after a 6.5 h incubation, in parallel with biofilm inhibition study.

Page | 209 CHAPTER FIVE

Planktonic growth inhibition was found to be in the range of ca. 33-39% for all four combinations. However, the differences between the inhibitions induced by combinations and their individual components were nonsignificant based on

ANOVA analysis at almost all tested concentrations (Figure 5.4 and Table C 3).

Figure 5.4. Inhibition of planktonic bacterial growth. P. aeruginosa PAO1 bacteria were grown in the presence of RP, Car, Eug, RP/Car and RP/Eug (a), and BP, CP, Car, Eug, BP/Car,

BP/Eug, CP/Car and CP/Eug (b). Bacterial growth was quantified by measurement of the

OD595 of the bacterial culture. Data are representative of at least two independent experiments ± SD. Two-way ANOVA test; black asterisks indicate a statistically significant difference of each combination vs individual polymer at the corresponding concentrations.

Blue and red asterisks indicate a statistically significant difference of each combination vs individual Car or Eug, respectively. (**p < 0.01; ***p < 0.001; ****p < 0.0001; ns, nonsignificant

(p > 0.01)).

RP at 4 and 8 μg mL−1 was able to inhibit the biofilm formation by 26 and 52%, respectively, compared to the control biofilm (Figure 5.5 and Table C 3).

Meanwhile, BP showed the same dose dependent trend in biofilm inhibition with

9 and 31% reductions in biofilm biomass at 4 and 8 μg mL−1, respectively, compared to the control biofilm (Figure 5.5 and Table C 3). Lower antibiofilm activity of BP relative to RP is unsurprising considering the core−shell micelle

Page | 210 CHAPTER FIVE

morphology of BP. CP did not show any significant biofilm inhibition. Car and

Eug at respective concentrations of 300 and 530 μg mL−1 led to 43 and 18% reductions in biofilm biomass, respectively, compared to control biofilm (Figure

5.5and Table C 3).

Figure 5.5. P. aeruginosa PAO1 biofilm formation inhibition. Biofilms were grown in the presence of RP, Car, Eug, RP/Car and RP/Eug (a), and BP, CP, Car, Eug, BP/Car, BP/Eug,

CP/Car and CP/Eug (b). Biofilm biomass was quantified by measurement of the OD595 of the crystal violet stained biofilms. Data are representative of at least two independent experiments ± SD. Two-way ANOVA test; black asterisks indicate a statistically significant difference of each combination vs individual polymer at the corresponding concentrations.

Blue and red asterisks indicate a statistically significant difference of each combination vs individual Car or Eug, respectively. (**p < 0.01; ***p < 0.001; ****p < 0.0001; ns, nonsignificant

(p > 0.01)).

Combinations of antimicrobial polymers and essential oils showed the best biofilm inhibition activity compared to all tested compounds. We observed higher biofilm inhibition for each combination in comparison to individual compounds (Figure 5.5 and Table C 3). For example, biofilm formation was reduced by 73% in the presence of RP/Car at the polymer and oil concentrations of 4 and 300 μg mL−1, respectively (denoted as 4/300 μg mL−1). This shows a substantial improvement relative to individual RP and Car, where the biofilm Page | 211 CHAPTER FIVE

inhibition was 26 and 43%, respectively. Despite the lower biofilm inhibition activity of BP compared to RP and also the different potencies of Car and Eug, all four combinations demonstrated similar biofilm inhibition performances resulting in the range of ca. 60-75%, compared to control biofilm.

CP/Car and CP/Eug, on the other hand, were the worst-performing combinations in this test. The biofilm inhibition activity of Car and Eug seems to be masked by the CP polymeric micelles, causing lower biofilm inhibition compared to the oils alone. This observation confirms the advantage of using antimicrobial polymeric micelles to encapsulate essential oils, where the antimicrobial polymer can facilitate the interaction of oil molecules with bacteria biofilm while providing additional antimicrobial effect.

We next evaluated the efficacy of the same compounds against a MDR strain of P. aeruginosa, PA37. Regarding the planktonic bacteria growth inhibition, all tested compounds and combinations demonstrated lower activity relative to

PAO1, and there were nonsignificant differences in activity between the combinations and their individual components (Figure 5.6 and Table C 4).

Page | 212 CHAPTER FIVE

Figure 5.6. Inhibition of planktonic bacterial growth induced. P. aeruginosa PA37 bacteria were grown in the presence of RP, Car, Eug, RP/Car and RP/Eug (a), and BP, CP, Car, Eug,

BP/Car, BP/Eug, CP/Car and CP/Eug (b). Bacterial growth was quantified by measurement of the OD595 of the bacterial culture. Data are representative of at least two independent experiments ± SD. Two-way ANOVA test; black asterisks indicate a statistically significant difference of each combination vs individual polymer at the corresponding concentrations.

Blue and red asterisks indicate a statistically significant difference of each combination vs individual Car or Eug, respectively. (**p < 0.01; ***p < 0.001; ****p < 0.0001; ns, nonsignificant

(p > 0.01)).

In terms of biofilm inhibition activity, we observed slightly lower activity for

RP, BP, and Car against PA37 compared to PAO1 (Figure 5.7 and Table C 4).

However, there was an increase in biofilm inhibition activity for Eug from 18% against PAO1 to 41% against PA37. Interestingly, despite the lower activity for the majority of individual compounds against PA37 compared to PAO1, slightly higher biofilm inhibition capability was attained for the polymer−oil combinations against PA37. All four combinations showed significantly increased biofilm inhibition compared to their individual components, with values ranging from 70 to 85% compared to control biofilm. Generally, the combination of synthetic antimicrobial polymers and essential oils showed good

Page | 213 CHAPTER FIVE

efficacy against P. aeruginosa colonization in the first 6.5 h of incubation, which is the most susceptible period for biofilm development.

Figure 5.7. P. aeruginosa PA37 biofilm formation inhibition. Biofilms were grown in the presence of RP, Car, Eug, RP/Car and RP/Eug (a), and BP, CP, Car, Eug, BP/Car, BP/Eug,

CP/Car and CP/Eug (b). Biofilm biomass was quantified by measurement of the OD595 of the crystal violet stained biofilms. Data are representative of at least two independent experiments ± SD. Two-way ANOVA test; black asterisks indicate a statistically significant difference of each combination vs individual polymer at the corresponding concentrations.

Blue and red asterisks indicate a statistically significant difference of each combination vs individual Car or Eug, respectively. (**p < 0.01; ***p < 0.001; ****p < 0.0001; ns, nonsignificant

(p > 0.01)).

The biofilm inhibition ability of all tested compounds and combinations against PAO1 was also visualized using a 3D tomographic microscope

(Nanolive) equipped with a digital staining software (Figure 5.8). Observation of bacterial biofilms is often challenging because of the low contrast and high transparency of the biofilms, which therefore necessitate complicated preparation and microscopy techniques such as confocal or SEM microscopy. The tomographic microscope used in this study provides a non-invasive approach to depict the biofilm with minimal sample preparation.63 To provide visual aid and

Page | 214 CHAPTER FIVE

quantitative data, the bacteria cells on the surface were digitally coloured via software analysis based on the difference of refractive index between bacteria cells and the background. The biofilm biomass values after 6.5 h incubation were similar to those obtained from CV staining experiments based on 2D and 3D tomographic images.

Figure 5.8. 2D and 3D tomographic microscopy images of P. aeruginosa PAO1 biofilm grown in the presence or absence of individual compounds and combinations for 6.5 h. Images were taken at the center of the culture dish and analyzed using 3D Nanolive Cell Explorer Steve digital staining software (scale bar = 20 μm). Note: Biofilm inhibition values based on this microscopy analysis are presented at the bottom right corner.

Page | 215 CHAPTER FIVE

5.3.5 Biofilm killing activity of polymer-oil combinations and individual

components

The bactericidal activities of the combinations against biofilm were subsequently examined via colony-forming unit (CFU) analysis. Specifically, P. aeruginosa PAO1 biofilms were grown in cell culture media M9 for 6.5 h prior to incubation with selected compounds and combinations at different concentrations for 20 min, and the viability of the treated bacterial biofilm cells was determined by CFU analysis (Figure 5.9and Table C 5).

Figure 5.9. Bactericidal activity of RP, Car, Eug, RP/Car and RP/Eug (a), and BP, CP, Car,

Eug, BP/Car, BP/Eug, CP/Car and CP/Eug (b) against P. aeruginosa PAO1 biofilm. All bactericidal activities were determined by colony-forming unit (CFU) analysis upon 20 min of incubation. Data are representative of at least two independent experiments ± SD. Two- way ANOVA test; black asterisks indicate a statistically significant difference of each combination vs individual polymer at the corresponding concentrations. Blue and red asterisks indicate a statistically significant difference of each combination vs individual Car or Eug, respectively. (**p < 0.01; ***p < 0.001; ****p < 0.0001; ns, nonsignificant (p > 0.01)).

RP and BP demonstrated similar bactericidal efficacy against PAO1 biofilm and with increased activity at higher polymer concentrations. We observed an average of 1.8 ± 0.9 and 1.9 ± 0.9 log10 reductions in CFU at 32 μg mL−1 and 2.9 ±

Page | 216 CHAPTER FIVE

1.0 and 2.3 ± 0.9 log10 reductions in CFU at 64 μg mL−1 for RP and BP, respectively, compared to untreated biofilm (Figure 5.9 and Table C 5). As expected, CP did not result in any significant reduction in bacterial cell viability. Meanwhile, Car and Eug displayed less than 1 log10 reduction in CFU (0.6 ± 0.6 and 0.4 ± 0.9, respectively) relative to untreated biofilm. The differences between the bactericidal activities of RP and combinations RP/Car and RP/Eug were nonsignificant based on ANOVA analysis (Figure 5.9 a). In contrast, BP/Car and

BP/Eug exhibited a remarkable improvement in bactericidal activity in comparison with BP, Car and Eug at all tested concentrations (Figure 5.9 b).

Treatment with BP/Car for 20 min resulted in 3.1 ± 0.6 and 4.3 ± 1.1 log10 reductions in CFU at 32/300 and 64/300 μg mL−1, respectively, compared to untreated biofilm (Figure 5.9 b and Table C 5). Likewise, BP/Eug caused an average of 3.7 ± 1.1 and 4.6 ± 1.0 log10 reductions in CFU at 32/530 and 64/530 μg mL−1, respectively, after 20 min incubation. These results confirm the synergistic interaction between BP and both Car and Eug in terms of their killing activity against PAO1 bacteria biofilm. We postulate that the reason a synergistic interaction was only observed for BP containing combinations is due to the ability of BP micellar nanoparticles to encapsulate essential oils. Positively charged BP micelles with loaded essential oil can effectively deliver and release the payload upon contact with bacterial biofilm, leading to an increased local concentration of essential oils. Simultaneously, BP helps to eradicate the biofilm because of its inherent antimicrobial activity. This delivery capability is most likely missing in RP containing combinations, as the unimer micelles of RP lack the capacity to store the oils. CP/Car and CP/Eug combinations were inactive in

Page | 217 CHAPTER FIVE

terms of bactericidal activity, which was in accordance with results obtained in the biofilm inhibition test.

5.3.6 Hemocompatibility of polymer-oil combinations and individual

components

Finally, hemolysis studies were performed to assess the mammalian cell compatibility of individual compounds and combinations using sheep red blood cells (RBCs). The HC50 value, which is defined as the concentration that causes

50% lysis of RBCs, was evaluated for RP and BP. In the case of Car, Eug and all the combinations, due to the turbidity of the emulsions at high concentrations of the oils, we only determined the hemolysis % at concentrations identical to the concentrations used in the killing study. All the samples were also inspected visually for hemagglutination, which was qualitatively determined by the inability to resuspend RBCs following incubation with the compounds.

We observed a HC50 value of 2048 μg mL−1 for RP with no evidence of hemagglutination, which is consistent with our previous reports (Figure 5.10 and

Table C 6).52, 60 BP, on the other hand, showed higher HC50 value of 4096 μg mL−1 with minor hemagglutination. Car and Eug at respective concentrations of 300 and 530 μg mL−1 led to 99 and 55% lysis of RBCs, respectively, with minor hemagglutination observed for Eug. All RP containing emulsions caused complete lysis of RBCs. BP containing emulsions demonstrated lower hemolytic activity in comparison with individual oils, which is attributed to the encapsulation/shielding of oils inside the hydrophobic core of BP micelles.

Hemolysis % was determined to be in the range of about 28-43% for all BP- containing emulsions with no occurrence of hemagglutination. It is worthwhile

Page | 218 CHAPTER FIVE

nothing that although the coadministration of BP and both oils resulted in a reduction in the hemolytic activities compared to the individual oils, these combinations are still relatively hemolytic and might not be appropriate for intravenous applications. However, these combinations might have potential applications as oral or topical antibiotic.

Figure 5.10. Hemolytic activity of RP, BP, Car, Eug, RP/Car, BP/Car, RP/Eug and BP/Eug.

Data are representative of at least two independent experiments ± SD.

The excellent performance of antimicrobial polymeric micelles in this study suggests the advantages of using antimicrobial block copolymers as potential nanocarriers for antimicrobial/antibiofilm purposes. The efficacy of these antimicrobial polymeric micelles can potentially be further improved by endowing the polymers with cross-linkable functional groups to yield more stable micelles.64-66

Page | 219 CHAPTER FIVE

5.4 Summary

In summary, we have developed an efficient antibiofilm platform involving the combination of synthetic antimicrobial polymers and essential oils, wherein the localized and targeted delivery of essential oils to the bacteria cells was granted by the ability of the antimicrobial polymer to function as a delivery vehicle. We found that the attachment of a dedicated hydrophobic block onto the chain end of an antimicrobial polymer (e.g., ternary random copolymer) crucially allows for the effective encapsulation of essential oils within the micellar cores which ultimately results in high antibiofilm activity (i.e., biofilm formation inhibition and killing of biofilm). Wild-type P. aeruginosa PAO1 biofilm formation was inhibited by >70 % when the bacterial culture was incubated for 6.5 h in the presence of an antimicrobial block copolymer with encapsulated carvacrol or eugenol. Interestingly, these combinations maintained the same biofilm inhibition efficacy against a multidrug-resistant P. aeruginosa strain. In addition, the treatment of P. aeruginosa PAO1 biofilm with polymer−oil combinations that include the block copolymer reduced the viability of bacteria cells by >99.99% after 20 min of incubation time. In essence, the block copolymer acted synergistically with essential oils in killing P. aeruginosa biofilms. This study thus shows the potential application of antimicrobial polymeric micelles that dually function as therapeutic agents themselves as well as delivery vehicles for hydrophobic compounds or drugs, where the targeted delivery of the cargo molecules to the microbial biofilm is desired.

Page | 220 CHAPTER FIVE

5.5 Reference

1. Dengler Haunreiter, V.; Boumasmoud, M.; Häffner, N.; Wipfli, D.; Leimer, N.; Rachmühl, C.; Kühnert, D.; Achermann, Y.; Zbinden, R.; Benussi, S.; Vulin, C.; Zinkernagel, A. S., In-host evolution of Staphylococcus epidermidis in a pacemaker- associated endocarditis resulting in increased antibiotic tolerance. Nat. Commun. 2019, 10 (1), 1-14. 2. Aarestrup, F. M., The livestock reservoir for antimicrobial resistance: a personal view on changing patterns of risks, effects of interventions and the way forward. Philos. Trans. Royal Soc. B. 2015, 370 (1670), 20140085. 3. Levy, S. B.; Marshall, B., Antibacterial resistance worldwide: causes, challenges and responses. Nat. Med. 2004, 10, S122-S129. 4. Boucher, H. W.; Talbot, G. H.; Bradley, J. S.; Edwards, J. E.; Gilbert, D.; Rice, L. B.; Scheld, M.; Spellberg, B.; Bartlett, J., Bad Bugs, No Drugs: No ESKAPE! An Update from the Infectious Diseases Society of America. Clin. Infect. Dis. 2009, 48 (1), 1-12. 5. Hay, S. I.; Rao, P. C.; Dolecek, C.; Day, N. P. J.; Stergachis, A.; Lopez, A. D.; Murray, C. J. L., Measuring and mapping the global burden of antimicrobial resistance. BMC Medicine 2018, 16 (1), 1-3. 6. Walsh, C., Molecular mechanisms that confer antibacterial drug resistance. Nature 2000, 406, 775-781. 7. Rossiter, S. E.; Fletcher, M. H.; Wuest, W. M., Natural Products as Platforms To Overcome Antibiotic Resistance. Chem. Rev. 2017, 117 (19), 12415-12474. 8. Sadrearhami, Z.; Nguyen, T.-K.; Namivandi-Zangeneh, R.; Jung, K.; Wong, E. H. H.; Boyer, C., Recent advances in nitric oxide delivery for antimicrobial applications using polymer-based systems. J. Mater. Chem. B 2018, 6 (19), 2945-2959. 9. Costerton, J. W.; Stewart, P. S.; Greenberg, E. P., Bacterial Biofilms: A Common Cause of Persistent Infections. Science 1999, 284 (5418), 1318-1322. 10. Davies, D., Understanding biofilm resistance to antibacterial agents. Nat. Rev. Drug Discov. 2003, 2 (2), 114-122. 11. Whiteley, M.; Bangera, M. G.; Bumgarner, R. E.; Parsek, M. R.; Teitzel, G. M.; Lory, S.; Greenberg, E. P., Gene expression in Pseudomonas aeruginosa biofilms. Nature 2001, 413 (6858), 860-864. 12. Hall-Stoodley, L.; Stoodley, P., Evolving concepts in biofilm infections. Cell Microbiol. 2009, 11 (7), 1034-1043. 13. Donlan, R. M.; Costerton, J. W., Biofilms: Survival Mechanisms of Clinically Relevant Microorganisms. Clin. Microbiol. Rev. 2002, 15 (2), 167-193. 14. Campoccia, D.; Montanaro, L.; Arciola, C. R., A review of the biomaterials technologies for infection-resistant surfaces. Biomaterials 2013, 34 (34), 8533-8554. 15. Römling, U.; Balsalobre, C., Biofilm infections, their resilience to therapy and innovative treatment strategies. J. Intern. Med. 2012, 272 (6), 541-561. 16. Cegelski, L.; Pinkner, J. S.; Hammer, N. D.; Cusumano, C. K.; Hung, C. S.; Chorell, E.; Åberg, V.; Walker, J. N.; Seed, P. C.; Almqvist, F.; Chapman, M. R.; Hultgren, S. J., Small-molecule inhibitors target Escherichia coli amyloid biogenesis and biofilm formation. Nat. Chem. Biol. 2009, 5, 913-919. 17. Zhang, P.; Li, S.; Chen, H.; Wang, X.; Liu, L.; Lv, F.; Wang, S., Biofilm Inhibition and Elimination Regulated by Cationic Conjugated Polymers. ACS Appl. Mater. Interfaces. 2017, 9 (20), 16933-16938.

Page | 221 CHAPTER FIVE

18. Wang, M.; Shi, J.; Mao, H.; Sun, Z.; Guo, S.; Guo, J.; Yan, F., Fluorescent Imidazolium-Type Poly(ionic liquid)s for Bacterial Imaging and Biofilm Inhibition. Biomacromolecules 2019, 20 (8), 3161-3170. 19. Sadrearhami, Z.; Shafiee, F. N.; Ho, K. K. K.; Kumar, N.; Krasowska, M.; Blencowe, A.; Wong, E. H. H.; Boyer, C., Antibiofilm Nitric Oxide-Releasing Polydopamine Coatings. ACS Appl. Mater. Interfaces. 2019, 11 (7), 7320-7329. 20. Namivandi-Zangeneh, R.; Sadrearhami, Z.; Bagheri, A.; Sauvage-Nguyen, M.; Ho, K. K. K.; Kumar, N.; Wong, E. H. H.; Boyer, C., Nitric Oxide-Loaded Antimicrobial Polymer for the Synergistic Eradication of Bacterial Biofilm. ACS Macro Lett. 2018, 7 (5), 592-597. 21. Duong, H. T. T.; Jung, K.; Kutty, S. K.; Agustina, S.; Adnan, N. N. M.; Basuki, J. S.; Kumar, N.; Davis, T. P.; Barraud, N.; Boyer, C., Nanoparticle (Star Polymer) Delivery of Nitric Oxide Effectively Negates Pseudomonas aeruginosa Biofilm Formation. Biomacromolecules 2014, 15 (7), 2583-2589. 22. Zhou, R.; Zhou, R.; Wang, P.; Luan, B.; Zhang, X.; Fang, Z.; Xian, Y.; Lu, X.; Ostrikov, K. K.; Bazaka, K., Microplasma Bubbles: Reactive Vehicles for Biofilm Dispersal. ACS Appl. Mater. Interfaces. 2019, 11 (23), 20660-20669. 23. Namivandi-Zangeneh, R.; Sadrearhami, Z.; Dutta, D.; Willcox, M.; Wong, E. H. H.; Boyer, C., Synergy between Synthetic Antimicrobial Polymer and Antibiotics: A Promising Platform To Combat Multidrug-Resistant Bacteria. ACS Infect. Dis. 2019, 5 (8), 1357-1365. 24. Sadrearhami, Z.; Yeow, J.; Nguyen, T.-K.; Ho, K. K. K.; Kumar, N.; Boyer, C., Biofilm dispersal using nitric oxide loaded nanoparticles fabricated by photo-PISA: influence of morphology. ChemComm. 2017, 53 (96), 12894-12897. 25. Böttcher, T.; Kolodkin-Gal, I.; Kolter, R.; Losick, R.; Clardy, J., Synthesis and Activity of Biomimetic Biofilm Disruptors. J. Am. Chem. Soc. 2013, 135 (8), 2927-2930. 26. Mah, T.-F. C.; O'Toole, G. A., Mechanisms of biofilm resistance to antimicrobial agents. Trends Microbiol. 2001, 9 (1), 34-39. 27. Bjarnsholt, T.; Tolker-Nielsen, T.; Høiby, N.; Givskov, M., Interference of Pseudomonas aeruginosa signalling and biofilm formation for infection control. Expert Rev Mol Med. 2010, 12, 1-19. 28. Cowan, M. M., Plant products as antimicrobial agents. Clin. Microbiol. Rev. 1999, 12 (4), 564-582. 29. Dorman, H. J. D.; Deans, S. G., Antimicrobial agents from plants: antibacterial activity of plant volatile oils. J. Appl. Microbiol. 2000, 88 (2), 308-316. 30. Bakkali, F.; Averbeck, S.; Averbeck, D.; Idaomar, M., Biological effects of essential oils – A review. Food Chem. Toxicol. 2008, 46 (2), 446-475. 31. Helander, I. M.; Alakomi, H.-L.; Latva-Kala, K.; Mattila-Sandholm, T.; Pol, I.; Smid, E. J.; Gorris, L. G. M.; von Wright, A., Characterization of the Action of Selected Essential Oil Components on Gram-Negative Bacteria. J. Agric. Food Chem. 1998, 46 (9), 3590-3595. 32. Burt, S., Essential oils: their antibacterial properties and potential applications in foods—a review. Int. J. Food Microbiol. 2004, 94 (3), 223-253. 33. Nazzaro, F.; Fratianni, F.; De Martino, L.; Coppola, R.; De Feo, V., Effect of essential oils on pathogenic bacteria. Pharmaceuticals (Basel) 2013, 6 (12), 1451-1474. 34. Silva, L. N.; Zimmer, K. R.; Macedo, A. J.; Trentin, D. S., Plant Natural Products Targeting Bacterial Virulence Factors. Chem. Rev. 2016, 116 (16), 9162-9236. 35. Raut, J. S.; Karuppayil, S. M., A status review on the medicinal properties of essential oils. Ind Crops Prod. 2014, 62, 250-264.

Page | 222 CHAPTER FIVE

36. Chong, Y.-B.; Zhang, H.; Yue, C. Y.; Yang, J., Fabrication and Release Behavior of Microcapsules with Double-Layered Shell Containing Clove Oil for Antibacterial Applications. ACS Appl. Mater. Interfaces. 2018, 10 (18), 15532-15541. 37. Jobdeedamrong, A.; Jenjob, R.; Crespy, D., Encapsulation and Release of Essential Oils in Functional Silica Nanocontainers. Langmuir 2018, 34 (44), 13235- 13243. 38. Scaffaro, R.; Lopresti, F.; D’Arrigo, M.; Marino, A.; Nostro, A., Efficacy of poly(lactic acid)/carvacrol electrospun membranes against Staphylococcus aureus and Candida albicans in single and mixed cultures. Appl. Microbiol. Biotechnol. 2018, 102 (9), 4171-4181. 39. Amato, D. N.; Amato, D. V.; Mavrodi, O. V.; Martin, W. B.; Swilley, S. N.; Parsons, K. H.; Mavrodi, D. V.; Patton, D. L., Pro-Antimicrobial Networks via Degradable Acetals (PANDAs) Using Thiol–Ene Photopolymerization. ACS Macro Lett. 2017, 6 (2), 171-175. 40. Amato, D. N.; Amato, D. V.; Adewunmi, Y.; Mavrodi, O. V.; Parsons, K. H.; Swilley, S. N.; Braasch, D. A.; Walker, W. D.; Mavrodi, D. V.; Patton, D. L., Using Aldehyde Synergism To Direct the Design of Degradable Pro-Antimicrobial Networks. ACS Appl. Bio Mater. 2018, 1 (6), 1983-1991. 41. Li, C.-H.; Chen, X.; Landis, R. F.; Geng, Y.; Makabenta, J. M.; Lemnios, W.; Gupta, A.; Rotello, V. M., Phytochemical-Based Nanocomposites for the Treatment of Bacterial Biofilms. ACS Infect. Dis. 2019, 5 (9), 1590-1596. 42. Landis, R. F.; Li, C.-H.; Gupta, A.; Lee, Y.-W.; Yazdani, M.; Ngernyuang, N.; Altinbasak, I.; Mansoor, S.; Khichi, M. A. S.; Sanyal, A.; Rotello, V. M., Biodegradable Nanocomposite Antimicrobials for the Eradication of Multidrug-Resistant Bacterial Biofilms without Accumulated Resistance. J. Am. Chem. Soc. 2018, 140 (19), 6176-6182. 43. Gabriel, G. J.; Madkour, A. E.; Dabkowski, J. M.; Nelson, C. F.; Nüsslein, K.; Tew, G. N., Synthetic Mimic of Antimicrobial Peptide with Nonmembrane-Disrupting Antibacterial Properties. Biomacromolecules 2008, 9 (11), 2980-2983. 44. Yang, Y.; Cai, Z.; Huang, Z.; Tang, X.; Zhang, X., Antimicrobial cationic polymers: from structural design to functional control. Polym. J. 2018, 50 (1), 33-44. 45. Engler, A. C.; Wiradharma, N.; Ong, Z. Y.; Coady, D. J.; Hedrick, J. L.; Yang, Y.-Y., Emerging trends in macromolecular antimicrobials to fight multi-drug-resistant infections. Nano Today 2012, 7 (3), 201-222. 46. Chin, W.; Zhong, G.; Pu, Q.; Yang, C.; Lou, W.; De Sessions, P. F.; Periaswamy, B.; Lee, A.; Liang, Z. C.; Ding, X.; Gao, S.; Chu, C. W.; Bianco, S.; Bao, C.; Tong, Y. W.; Fan, W.; Wu, M.; Hedrick, J. L.; Yang, Y. Y., A macromolecular approach to eradicate multidrug resistant bacterial infections while mitigating drug resistance onset. Nat. Commun. 2018, 9 (1), 1-14. 47. Konai, M. M.; Bhattacharjee, B.; Ghosh, S.; Haldar, J., Recent Progress in Polymer Research to Tackle Infections and Antimicrobial Resistance. Biomacromolecules 2018, 19 (6), 1888-1917. 48. Moad, G.; Rizzardo, E.; Thang, S. H., RAFT Polymerization and Some of its Applications. Chem. Asian J. 2013, 8 (8), 1634-1644. 49. Boyer, C.; Bulmus, V.; Davis, T. P.; Ladmiral, V.; Liu, J.; Perrier, S., Bioapplications of RAFT Polymerization. Chem. Rev. 2009, 109 (11), 5402-5436. 50. Judzewitsch, P. R.; Nguyen, T.-K.; Shanmugam, S.; Wong, E. H. H.; Boyer, C., Towards Sequence-Controlled Antimicrobial Polymers: Effect of Polymer Block Order on Antimicrobial Activity. Angew. Chem., Int. Ed. 2018, 57 (17), 4559-4564.

Page | 223 CHAPTER FIVE

51. Judzewitsch, P. R.; Zhao, L.; Wong, E. H. H.; Boyer, C., High-Throughput Synthesis of Antimicrobial Copolymers and Rapid Evaluation of Their Bioactivity. Macromolecules 2019, 52 (11), 3975-3986. 52. Namivandi-Zangeneh, R.; Kwan, R. J.; Nguyen, T.-K.; Yeow, J.; Byrne, F. L.; Oehlers, S. H.; Wong, E. H. H.; Boyer, C., The effects of polymer topology and chain length on the antimicrobial activity and hemocompatibility of amphiphilic ternary copolymers. Polym. Chem. 2018, 9 (13), 1735-1744. 53. Perrier, S.; Takolpuckdee, P., Macromolecular design via reversible addition– fragmentation chain transfer (RAFT)/xanthates (MADIX) polymerization. J. Polym. Sci. A 2005, 43 (22), 5347-5393. 54. Mizutani, M.; Palermo, E. F.; Thoma, L. M.; Satoh, K.; Kamigaito, M.; Kuroda, K., Design and Synthesis of Self-Degradable Antibacterial Polymers by Simultaneous Chain- and Step-Growth Radical Copolymerization. Biomacromolecules 2012, 13 (5), 1554-1563. 55. Oda, Y.; Kanaoka, S.; Sato, T.; Aoshima, S.; Kuroda, K., Block versus Random Amphiphilic Copolymers as Antibacterial Agents. Biomacromolecules 2011, 12 (10), 3581-3591. 56. Li, Y.; Liu, G.; Wang, X.; Hu, J.; Liu, S., Enzyme-Responsive Polymeric Vesicles for Bacterial-Strain-Selective Delivery of Antimicrobial Agents. Angew. Chem. Int. Ed. 2016, 55 (5), 1760-1764. 57. Leng, M.; Hu, S.; Lu, A.; Cai, M.; Luo, X., The anti-bacterial poly(caprolactone)-poly(quaternary ammonium salt) as drug delivery carriers. Appl. Microbiol. Biotechnol. 2016, 100 (7), 3049-3059. 58. Eissa, A. M.; Abdulkarim, A.; Sharples, G. J.; Cameron, N. R., Glycosylated Nanoparticles as Efficient Antimicrobial Delivery Agents. Biomacromolecules 2016, 17 (8), 2672-2679. 59. Willyard, C., The drug-resistant bacteria that pose the greatest health threats. Nature 2017, 543 (7643), 15. 60. Nguyen, T.-K.; Lam, S. J.; Ho, K. K. K.; Kumar, N.; Qiao, G. G.; Egan, S.; Boyer, C.; Wong, E. H. H., Rational Design of Single-Chain Polymeric Nanoparticles That Kill Planktonic and Biofilm Bacteria. ACS Infect. Dis. 2017, 3 (3), 237-248. 61. Lam, S. J.; Wong, E. H. H.; Boyer, C.; Qiao, G. G., Antimicrobial polymeric nanoparticles. Prog. Polym. Sci. 2018, 76, 40-64. 62. Barraud, N.; Moscoso, J. A.; Ghigo, J.-M.; Filloux, A., Methods for Studying Biofilm Dispersal in Pseudomonas aeruginosa. In Pseudomonas Methods and Protocols, Filloux, A.; Ramos, J.-L., Eds. Springer New York: New York, NY, 2014; pp 643-651. 63. Nanolive Microbiology. https://nanolive.ch/applications/overview/single-cell- cell-culture-analysis/microbiology/. 64. Kim, Y.; Pourgholami, M. H.; Morris, D. L.; Stenzel, M. H., Effect of Cross- Linking on the Performance of Micelles As Drug Delivery Carriers: A Cell Uptake Study. Biomacromolecules 2012, 13 (3), 814-825. 65. Du, A. W.; Lu, H.; Stenzel, M. H., Core-Cross-Linking Accelerates Antitumor Activities of Paclitaxel–Conjugate Micelles to Prostate Multicellular Tumor Spheroids: A Comparison of 2D and 3D Models. Biomacromolecules 2015, 16 (5), 1470-1479. 66. Landis, R. F.; Gupta, A.; Lee, Y.-W.; Wang, L.-S.; Golba, B.; Couillaud, B.; Ridolfo, R.; Das, R.; Rotello, V. M., Cross-Linked Polymer-Stabilized Nanocomposites for the Treatment of Bacterial Biofilms. ACS Nano 2017, 11 (1), 946-952.

Page | 224 CHAPTER FIVE

5.6 Appendix C

Figure C 1. 1H NMR spectra of the Boc-protected RP in CDCl3 (a) and the corresponding

Boc-deprotected RP in D2O (b).

Page | 225 CHAPTER FIVE

Figure C 2. PC-differential refractive index (DRI) chromatogram of the Boc-protected RP.

Figure C 3. Representative plot of I1/I3 values versus the logarithm of the concentration of the Boc-deprotected RP micellar solution.

Page | 226 CHAPTER FIVE

Figure C 4. 1H NMR spectra of the Boc-protected BP macroRAFT agent in DMSO (a), Boc- protected block copolymer BP in DMSO (b) and the corresponding Boc-deprotected BP in D2O

(c).

Page | 227 CHAPTER FIVE

Figure C 5. GPC-DRI chromatograms of the Boc-protected macroRAFT agent BP and BP block copolymer.

Figure C 6. Representative plot of I1/I3 values versus the logarithm of the concentration of the Boc-deprotected BP micellar solution.

Page | 228 CHAPTER FIVE

Figure C 7. 1H NMR spectra of the CP macroRAFT agent in DMSO (a) and block copolymer

CP in DMSO (b).

Page | 229 CHAPTER FIVE

Figure C 8. GPC-DRI chromatograms of the macroRAFT agent CP and CP block copolymer.

Figure C 9. Representative plot of I1/I3 values versus the logarithm of the concentration of

CP micellar solution.

Page | 230 CHAPTER FIVE

Table C 1. Summary of the number-averaged molecular weight (Mn), dispersity (Ɖ) and

CMC values of the synthesized polymers.

Entry Mn (g mol–1)a Đ a CMC (µg mL–1) RP 25400 1.19 7.48 BP macroRAFT 28400 1.18 - BP 34500 1.22 3.31 CP macroRAFT 26500 1.13 - CP 29900 1.14 2.47 aDetermined via GPC analysis in DMAc solvent.

Table C 2. Concentration of the samples used in the DLS analysis.

Concentration Overall Individual agents Killing study Entry (mg mL–1) (μg mL–1) (μg mL−1) RP 1 1000 64 BP 1 1000 64 CP 1 1000 64 Car 1 1000 300 Eug 1 1000 530 RP/Car 1 175/825 64/300 RP/Eug 1 108/892 64/530 BP/Car 1 175/825 64/300 BP/Eug 1 108/892 64/530 CP/Car 1 175/825 64/300 CP/Eug 1 108/892 64/530

Page | 231 CHAPTER FIVE

Figure C 10. TEM images of RP (a) and BP (b) in water. Samples were prepared using 2 mg mL–1 polymer solution in water.

Figure C 11. Changes in the transmittance (λ = 600 nm) of Car, RP/Car and BP/Car emulsions (a), and Eug, RP/Eug and BP/Eug emulsions (b) as a function of time.

Page | 232 CHAPTER FIVE

Table C 3. Biofilm biomass and planktonic biomass of P. aeruginosa PAO1, and the inhibitions caused by selected compounds at various concentrations. Biofilm biomass and planktonic biomass are quantified by measuring the OD595 of crystal violet stained biofilm and bacterial culture, respectively.

Biofilm Planktonic Concentration Biofilm Planktonic Entry biomass biomass (µg mL–1) Inhibition % Inhibition% (OD 595 )a (OD 595 )a Control - 0.560.02 - 0.210.01 - 4 0.41 0.05 26 0.170.02 18 RP 8 0.27 0.02 52 0.140.01 34 4 0.51 0.06 9 0.180.02 14 BP 8 0.38 0.04 31 0.150.02 26 CP 8 0.66 0.04 -20 0.190.001 8 Car 300 0.32 0.04 43 0.140.002 30 Eug 530 0.45 0.04 18 0.170.004 19 4/300 0.15 0.006 73 0.130.003 37 RP/Car 8/300 0.15 0.002 73 0.130.003 39 4/530 0.23 0.01 59 0.140.003 33 RP/Eug 8/530 0.14 0.002 74 0.130.003 39 4/300 0.22 0.02 60 0.130.005 35 BP/Car 8/300 0.16 0.007 72 0.130.005 37 4/530 0.23 0.018 59 0.140.002 33 BP/Eug 8/530 0.15 0.01 73 0.130.001 39 CP/Car 8/530 0.50 0.06 10 0.150.001 30 CP/Eug 8/530 0.50 0.06 9 0.160.001 24 aValues are the mean of at least two independent experiments ± standard deviation.

Page | 233 CHAPTER FIVE

Table C 4. Biofilm biomass and planktonic biomass of P. aeruginosa PA37 and the inhibitions caused by selected compounds at various concentrations. Biofilm biomass and planktonic biomass are quantified by measuring OD595 of crystal violet stained biofilm and bacteria culture, respectively.

Biofilm Planktonic Concentration Biofilm Planktonic Entry biomass biomass (µg mL–1) Inhibition % Inhibition% (OD 595 ) a (OD 595 ) a Control - 1.0 0.02 - 0.14 0.01 - RP 8 0.63 0.08 37 0.12 0.01 14 BP 8 0.70 0.09 30 0.11 0.004 17 CP 8 0.87 0.04 13 0.13 0.001 6 Car 300 0.65 0.19 36 0.13 0.001 8 Eug 530 0.59 0.02 41 0.13 0.01 5 RP/Car 8/300 0.15 0.001 85 0.12 0.001 12 RP/Eug 8/530 0.19 0.04 81 0.13 0.001 9 BP/Car 8/300 0.17 0.002 83 0.12 0.003 12 BP/Eug 8/530 0.30 0.08 70 0.130.001 8 CP/Car 8/530 0.61 0.02 39 0.12 0.001 9 CP/Eug 8/530 0.71 0.07 29 0.130.001 9 aValues are the mean of at least two independent experiments ± standard deviation.

Page | 234 CHAPTER FIVE

Table C 5. Log10 reduction of viable P. aeruginosa PAO1 bacterial colonies upon treatment with selected compounds and combinations at various concentrations, as determined using CFU analysis.

Entry Concentration (µg mL–1) Log 10 Reductiona 32 1.8 0.9 RP 64 2.9 1.0 32 1.9 0.9 BP 64 2.3 0.9 CP 54 -0.3 0.5 Car 300 0.6 0.6 Eug 530 0.4 0.9 32/300 1.1 0.5 RP/Car 64/300 3.2 0.6 32/530 0.9 0.4 RP/Eug 64/530 3.0 1.2 32/300 3.1 0.6 BP/Car 64/300 4.3 1.1 32/530 3.7 1.1 BP/Eug 64/530 4.6 1.0 CP/Car 64/530 0.2 0.5 CP/Eug 64/530 -0.1 0.8 aValues are the mean of at least two independent experiments ± standard deviation.

Page | 235 CHAPTER FIVE

Table C 6. Hemolysis study results.

Entry Concentration (µg mL–1) Hemolysisa (%) 32 13  3 RP 64 19  2 32 7  1 BP 64 12  6 Car 300 99  9 Eug 530 55  17 32/300 135  2 RP/Car 64/300 135  8 32/530 114  9 RP/Eug 64/530 118  8 32/300 38  12 BP/Car 64/300 28  10 32/530 43  6 BP/Eug 64/530 37  8 aValues are the mean of at least two independent experiments ± standard deviation

Page | 236

CHAPTER SIX

The Effects of Polymer Topology and Chain

Length on the Antimicrobial Activity and

Hemocompatibility of Amphiphilic Ternary

Copolymers

CHAPTER SIX

6 The Effects of Polymer Topology and Chain Length on

the Antimicrobial Activity and Hemocompatibility of

Amphiphilic Ternary Copolymers

This chapter is based on the work published in Polymer Chemistry, 2018, 9, 1735–

1744.

6.1 Introduction

The increase of multidrug resistance in bacteria is now regarded as one of the most pressing healthcare issues worldwide.1-4 Recently, the World Health

Organization (WHO) published a list of “priority pathogens” which indicates the most threatening bacteria to human health that urgently requires new antibiotic treatments.5 At the top of this list are carbapenem-resistant strains of Acinetobacter baumannii and Pseudomonas aeruginosa – both of which are Gram-negative bacteria. While the discovery of a new antibiotic called teixobactin offers hope in overcoming infections caused by Gram-positive bacteria such as methicillin-

Page | 238 CHAPTER SIX

resistant Staphylococcus aureus (MRSA),6 the pipeline for the development of new antimicrobial agents that combat Gram-negative bacteria remains limited.7

Drawing inspiration from (naturally occurring) antimicrobial peptides

(AMPs) and driven by the advances of controlled polymerization techniques,8-13 synthetic polymers have emerged as promising antimicrobial candidates in combating Gram-negative bacteria over the last few years. By mimicking the structural composition of AMPs which primarily consist of cationic and hydrophobic moieties, synthetic antimicrobial polymers can exert their bactericidal properties through physical membrane disruption of the bacteria cell wall (e.g., the outer and/or inner membrane of Gram-negative bacteria).14-33

Unlike conventional antibiotics which target intracellular targets (e.g., inhibition of DNA/RNA synthesis or cell wall synthesis), this particular mechanism hinders resistance development in bacteria towards AMPs and mimics thereof.34 In addition, by combining the synthetic versatility of controlled polymerization techniques, complex antimicrobial polymers with precise macromolecular topologies and functionalities have been generated, some of which display improved biological properties compared to traditional linear analogues.35 For example, Lam et al. recently reported the development of antimicrobial star- shaped polypeptides that exhibited excellent bactericidal properties through a multi-modal mechanism (e.g., extensive membrane pore formation and triggering of immune responses in mice model studies) against various

Gram-negative bacteria, including colistin-resistant strains.36 The architecture was suggested to be a key factor in endowing the star polymers with the multi-modal mechanism and outstanding antimicrobial activity.

Page | 239 CHAPTER SIX

Evidently, investigation into the structure-bioactivity relationship of antimicrobial polymers is crucial, as this will not only improve our understanding on the polymer-bacteria interaction but also aid in the development of more potent polymers. Besides focusing on the antimicrobial activity, the bio- or hemo-compatibility of a polymer is an equally important point to consider especially for potential application in the clinical setting where the polymers are used in topical or intravenous formulations and medical devices coating.37 Given that the cationic groups in antimicrobial polymers may also interact with mammalian cells and induce cytotoxicity, various efforts have been made to reduce this non-specific interaction, for instance, via the use of different types of cationic groups

(e.g., primary vs. secondary amines)38-40 or manipulation of the polymer architecture (e.g., mixed micelles and miktoarm star polymers).41-43 In addition, several groups have also prepared antibacterial polymer coatings and gels with low-fouling properties using zwitterionic or ethylene glycol- based monomers to minimize non-specific interactions with proteins.44-46

Very recently, we described the synthesis of new antimicrobial polymers in the form of single-chain polymeric nanoparticles (SCPNs) that demonstrated good biocompatibility and efficacy against both planktonic and biofilm Gram-negative bacteria.47 These SCPNs were in essence amphiphilic ternary copolymers, and we found that the judicious combination of oligo(ethylene glycol) (OEG), cationic mimic of 2,4- diaminobutyric acid (Dab), and hydrophobic (2-ethylhexyl or 2- phenylethyl) groups, was necessary to achieve optimum antimicrobial activity and biocompatibility.47

Page | 240 CHAPTER SIX

Herein, we report a detailed investigation of various other factors that affect the antimicrobial activity and hemocompatibility of amphiphilic ternary copolymers, including the influence of the polymer chain length and topology

(i.e., random vs block copolymers, and linear vs. hyperbranched polymers). For this study, various polymers are made using reversible addition-fragmentation chain transfer (RAFT) polymerization8, 13 (Figure 6.1), and their antimicrobial activity is assessed based on the minimum inhibitory concentration (MIC) values.

The MIC is defined as the minimum compound concentration that prevents visible bacterial growth. The most common method for determining the biocompatibility of antimicrobial polymers in vitro is by hemagglutination and hemolysis experiments where the HC50 value, defined as the compound concentration at which 50% of red blood cells are lysed, is used as a metric for comparison. Interestingly, polymers with different chain lengths (number- average degree of polymerization (DPn) of 100, 50 and 20) have similar antimicrobial activities but different hemolytic activities. In addition, shorter polymer chains (DPn = 50 and 20) cause hemagglutination. On the other hand, segregation of the hydrophilic OEG and cationic groups from hydrophobic moieties results in the loss of antimicrobial activity. Meanwhile, hyperbranched polymers indicate that branching can improve hemocompatibility (by > 4 times) with only a minor loss of antimicrobial activity. Overall, this study yields valuable information pertaining to the structure-activity correlation of antimicrobial ternary copolymers.

Page | 241 CHAPTER SIX

Figure 6.1. The compositional structures and architectures of the amphiphilic ternary copolymers in this study.

6.2 Experimental Section

Materials

Ethylenediamine (Sigma-Aldrich, ≥ 99%), di-tert-butyl dicarbonate (Aldrich,

99%), triethylamine (Scharlau, 99%), acryloyl chloride (Merck, ≥ 96%), 2- phenylethanol (Aldrich, ≥99%), oligoethylene glycol methyl ether acrylate (OEG acrylate) Mn 480 g mol–1 (Aldrich), 2-ethylhexyl acrylate (Aldrich, 98%), oligoethylene glycol diacrylate Mn 250 g mol–1 (Aldrich), trifluoroacetic acid

(TFA) (Sigma-Aldrich, 99%), chloroform (VWR Chemicals), hexane (Merck), diethyl ether Merck) and basic alumina (Al2O3) (LabChem) were used as received. 2,2’-azobis (2 methylpropionitrile) (AIBN) (Acros, 98%) was purified by recrystallization from methanol. Sodium sulfate (Na2SO4), magnesium sulfate

(MgSO4), sodium hydrogen carbonate (NaHCO3), tetrahydrofuran, and acetone were obtained from Chem-Supply and used as received. Milli-Q water with a

Page | 242 CHAPTER SIX

resistivity of > 18 MΩ⋅cm was obtained from an in-line Millipore RiOs/Origin water purification system. The monomers tert-butyl (2-acrylamidoethyl) carbamate and 2-phenylethyl acrylate were synthesized according to literature procedures.47

Analytical instruments

1H Nuclear magnetic resonance (NMR) spectra were recorded using a Bruker

AC300F spectrometer. Deuterated solvents D2O or CDCl3 (obtained from

Cambridge Isotope Laboratories) were used as reference solvents and samples with a concentration of ca. 10-20 mg mL–1 were prepared. The monomer composition in the polymers that consisted of 2-ethylhexyl acrylate was calculated using the following equation ʃa,b/6 : ʃc/9 : ʃd/2 where ʃa,b, ʃc, and ʃd correspond to the integrals of the characteristic protons of 2-ethylhexyl acrylate

(methyl -CH3- groups, δH 0.80-0.98 ppm), cationic monomer (tert-butyl –CH3- groups, δH 1.38-1.52 ppm) and OEGA (ester -CH2O- groups, 4.10-4.30 ppm), respectively (please refer to Appendix D). For polymers that consisted of 2- phenylethyl acrylate, the monomer composition was determined via the following equation ʃa,b/6 : ʃc/9 : ʃe/5 where ʃe corresponds to the integrals of the characteristic protons of 2-phenylethyl acrylate (aromatic protons, δH 7.10-7.40 ppm).

Gel permeation chromatography (GPC) analysis was performed using a

Shimadzu liquid chromatography system equipped with a Shimadzu refractive index detector and two MIX C columns (Polymer Lab) operating at 40 °C.

Tetrahydrofuran was used as the eluent at a flow rate of 1 mL min–1. The system

Page | 243 CHAPTER SIX

was calibrated with poly (methyl methacrylate) standards with molecular weights of 200 to 106 g mol–1.

Dynamic light scattering (DLS) and zeta-potential measurements were conducted using a Malvern Zetasizer Nano ZS apparatus equipped with a He-

Ne laser operated at λ = 633 nm and at a scattering angle of 173°. All samples were prepared at a concentration of ca. 2 mg mL–1 where filtered Milli-Q water

(using 0.45 μm pore size filter) was used as the solvent to solubilize the polymers.

Synthesis of antimicrobial polymers

Synthesis of cationic monomer

Cationic monomer tert-butyl (4-acrylamidobutyl) carbamate was prepared as previously described.47 Briefly, ethylenediamine (0.3 mol) was dissolved in chloroform (400 mL), followed by the dropwise addition of di-tert-butyl dicarbonate (0.03 mol in 100 mL) over 2 h at 0−5 °C. The reaction mixture was stirred overnight at 25 °C. White precipitates were filtered, and the organic phase was washed exhaustively with water using a separation funnel to remove excess diamines. The organic layer was then dehydrated over MgSO4, filtered, and dried using a rotary evaporation unit to yield a pale-yellow oil. Tetrahydrofuran (100 mL) was added to dissolve the intermediate product. Triethylamine (36 mmol) and acryloyl chloride (31.5 mmol) were added dropwise to the solution at 0−5 °C with N2 bubbling. The contents were stirred at 25 °C for 1 h. The byproducts were filtered, and the solvent was removed in vacuo. The crude product was dissolved in chloroform (150 mL) and washed against brine (1 × 75 mL). The organic phase was stirred with MgSO4 and basic Al2O3 for 10 min, filtered, and concentrated in

Page | 244 CHAPTER SIX

vacuo. The product was further purified by repeated precipitation steps in hexane to yield the tert-butyloxycarbonyl (Boc)-protected monomer as a fine white powder, which was dried in vacuo.

tert-Butyl (2-acrylamidoethyl) carbamate: 1H NMR (300MHz, CDCl3,25 °C), δH

6.56 (br s, 1H, NH), 6.28 (dd, J = 17.1 Hz, 1.5 Hz, 1H, CHH=CH), 6.12 (dd, J = 17.1

Hz, 10.2Hz, 1H, CHH=CH), 5.65 (dd, J = 10.2 Hz, 1.5 Hz, 1H, CHH=CH), 5.05 (br s, 1H, NH), 3.49−3.41 (m, 2H, CH2), 3.34−3.28 (m, 2H, CH2), 1.45 (s, 9H, CH3); 13C

NMR (300 MHz, CDCl3,25 °C), δC 166.23, 157.50, 130.88, 126.30, 79.85, 41.05, 40.09,

28.35.

Synthesis of linear random copolymers

The synthesis of amphiphilic ternary statistical copolymer proceeded as previously reported.47 Briefly, the reversible addition−fragmentation chain transfer (RAFT) agent benzyl dodecyl carbonotrithioate (11.6 μmol), AIBN (4.6

μmol), OEGA (350 μmol), tert-butyl (4-acrylamidobutyl) carbamate (580 μmol), and 2-ethylhexyl acrylate (230 μmol) were dissolved in 1,4-dioxane (such that the total monomer concentration in solvent is 1 M). The solution was purged with N2 for 20 min in an ice bath. The polymerization was conducted for 20 h at 70 °C and then quenched in an ice bath with exposure to air. The polymer was purified by precipitation in a hexane/diethyl ether (7 : 3) mixture thrice and subsequently dried in vacuo.

Synthesis of block copolymers

To prepare the diblock copolymers, a macroRAFT agent was synthesized followed by chain extension with the hydrophobic monomers. Firstly, benzyl

Page | 245 CHAPTER SIX

dodecyl carbonotrithioate (9.0 μmol), AIBN (4.5 μmol), OEGA (112.5 μmol) and tert-butyl (2-acrylamidoethyl) carbamate (337.5 μmol) were dissolved in 1,4- dioxane (such that the total monomer concentration was 1 M). The solution was degassed by bubbling with N2 for 20 min and the reaction mixture was stirred at

70 °C for 3 h before cooling in an ice bath for 5 min. The polymer was purified by dialysis using a dialysis membrane (MWCO of 3.5kDa) against methanol for 2 days and was dried in vacuo. The macroRAFT agent was characterized by 1H

NMR and GPC analysis. For the chain extension step, the macroRAFT agent (4.5

μmol), hydrophobic monomer (135 μmol) and AIBN (2.25 μmol) were dissolved in 1,4-dioxane and the solution was degassed by bubbling with N2 for 30 min in an ice bath. Then, the reaction mixture was stirred at 70 °C for 15 h.

Polymerization was quenched by placing the flask in an ice bath for 5 min. The polymer was purified by precipitation into a diethyl ether/ hexane (3 : 7) mixture.

The precipitate was isolated by centrifugation, dissolved in methanol, and precipitated twice more. Finally, the polymer was dried in vacuo.

Synthesis of hyperbranched copolymers

Hyperbranched polymers were synthesized in the same way as the linear random copolymers but with the addition of a cross-linkable monomer oligoethylene glycol diacrylate (Mn = 250 g mol-1) at 2 and 5 molar equivalents with respect to the RAFT agent.

The Boc protecting groups were removed using TFA in the same manner as reported previously.47 In general, the polymer solution in dichloromethane (ca.

10 wt % polymer) was treated with TFA (20 mol equivalent with respect to the

Boc group) for 3 h at 25 °C. Boc- deprotected polymer was subsequently

Page | 246 CHAPTER SIX

precipitated into diethyl ether/hexane (4:1). The precipitate was isolated by centrifugation, dissolved in methanol, and reprecipitated two more times. The polymer was then dried in vacuo and further purified by dialysis against water

(Cellu-Sep 3500 MWCO). The aqueous solution was lyophilized to yield the Boc- deprotected polymers.

Minimum inhibitory concentration (MIC) determination

The MIC was determined by using the broth microdilution method according to the Clinical and Laboratory Standards Institute (CLSI) guidelines. Briefly, bacterial culture was grown from a single colony in 10 mL of Mueller-Hinton broth (MHB) at 37 °C with shaking at 200 rpm overnight. The subculture was prepared from the overnight culture by diluting 1 : 100 in 5 mL MHB and allowed to grow to the mid-log phase, then diluted to the appropriate concentration for the MIC test. A two-fold dilution series of 50 μL of polymers solution in MHB were added into 96-well microplates followed by the addition of 50 μL of the subculture suspension. The final concentration of bacteria in each well was ca. 5

× 105 cells per mL. The plates were incubated at 37 °C for 20 h, and the absorbance at 600 nm was measured with a microtiter plate reader (FLUOstar Omega, BMG

Labtech). MIC values were defined as the lowest concentration of the sample that showed no visible growth and inhibited cell growth by more than 90%. Positive controls without the polymer and negative controls without bacteria were included. All assays included two replicates and were repeated in at least three independent experiments.

Hemolysis study

Page | 247 CHAPTER SIX

The hemolytic activity of the polymers was assessed using fresh sheep red blood cells (RBCs) obtained from Serum Australis (Catalog number SD50D).

RBCs were diluted 1:20 in PBS (pH 7.4), pelleted by centrifugation and washed three times with PBS (1000g, 10 min). The RBCs were then resuspended to achieve 5% (v/v) in PBS. Different concentrations of polymers (150 μL) were prepared in sterilized tubes, followed by the addition of the RBCs suspension

(150 μL). The highest polymer concentration tested was 2 mg mL–1. PBS buffer was used as a negative control while Triton-X 100 (1% v/v in PBS) was used as a positive hemolysis control. The tubes were incubated for 2 h at 37 °C and 150 rpm shaking speed in an incubator. Following incubation, the tubes were centrifuged

(1000g, 8 min) and aliquots of the supernatants (100 μL) were transferred into a

96-well microplate where the absorbance values were monitored at 485 nm using a microtiter plate reader (FLUOstar Omega, BMG Labtech). The percentage of hemolysis was calculated using the absorbance values and the formula below:

% Hemolysis = (Apolymer − Anegative)⁄(Apositive − Anegative) × 100

Planktonic and biofilm killing study

The laboratory strain P. aeruginosa PAO1 was used to investigate the bactericidal properties. Biofilms were grown as described previously.47 Briefly, in all assays, a single colony of PAO1 was inoculated in 10 mL of Luria Bertani medium (LB 10) at 37 °C with shaking at 200 rpm overnight. The overnight culture was diluted 1 : 100 in freshly prepared M9 minimal medium containing

48 mM Na2HPO4, 22 mM KH2PO4, 9 mM NaCl, 19 mM NH4Cl, pH 7.0, supplemented with 2 mM MgSO4, 100 μM CaCl2 and 20 mM glucose. The bacterial suspension was then aliquoted 1 mL per well of tissue-culture treated

Page | 248 CHAPTER SIX

24-well plates (Costar, Corning®). The plates were incubated at 37 °C with shaking at 180 rpm in an orbital shaker which does not stop agitation when the door is opened (model OM11, Ratek, Boronia, Australia) and the biofilm cultures were allowed to grow for 6.5 h without any disruption. The polymer was then added to the wells and the plates were incubated for 1 h. After treatment, the planktonic and biofilm viability analysis were determined by a drop plate method. For planktonic analyses, free-floating cells in the biofilm supernatant were serially diluted in sterile PBS and plated onto LB agar. For biofilm analysis, cells attached on the interior surfaces of the well (surface area 4.5 cm2) were washed twice with sterile PBS to remove loosely attached bacteria, before being resuspended and homogenized in PBS by incubating in an ultrasonication bath (150 W, 40 kHz;

Unisonics, Australia) for 20 min. Resuspended biofilm cells were then serially diluted and plated onto LB agar. Planktonic and biofilm colonies were counted and the CFU was calculated after 24 h incubation at 37 °C. All assays included two replicates and were repeated in at least three independent experiments.

Biofilm dispersal study

To characterize the effect of the polymer on biofilm dispersal, preformed

PAO1 biofilms were grown for 6.5 h and treated in the same manner as the killing study. Biofilm biomass was quantified by using the crystal violet (CV) staining method as previously described. Briefly, after treatment, the culture supernatant was removed and the biofilm on the well surfaces was washed once with 1 mL of PBS, followed by the addition of 1 mL 0.03% CV stain made from a 1 : 10 dilution of Gram crystal violet (BD) in PBS. The plates were incubated on the bench for 20 min before the wells were washed twice with PBS. The CV stained

Page | 249 CHAPTER SIX

biofilms were mixed with 1 mL 100% ethanol and quantified by measuring the

OD550 of the homogenized suspension using a microtiter plate reader (FLUOstar

Omega, BMG Labtech). All assays included two replicates and were repeated in at least three independent experiments.

Biofilm imaging

To visualize the effect of the polymer on the biofilm, PAO1 biofilms were grown on 35 mm tissue culture dishes (FluoroDish, World Precision Instruments

Inc., Sarasota, FL, USA) in the same way as in the bacteria killing study. The polymer was then added to the well and incubated for 1 h. After treatment, the supernatant was removed and the biofilm on the well surface was washed twice with 2 mL of PBS, followed by the addition of 1 mL PBS. The wells were analysed with a 3D tomographic microscope (3D Cell Explorer, NanoLive, Lausanne,

Switzerland) equipped with a digital staining software. All assays were repeated in at least two independent experiments.

6.3 Results and Discussion

In our previous report, we found that the combination of OEG, primary amine and hydrophobic groups in a single polymer chain, where the functionalities were randomly distributed, was necessary to achieve optimal biological performance as evidenced by evaluating the antimicrobial activity and biocompatibility of the synthesized polymer via several in vitro assays.

Moreover, the synthesized antimicrobial polymer retained the same MIC value after being incubated in DMEM plus 10% FBS medium, which demonstrated the ability of the polymer in in vivo mimicking conditions.47 Specifically, the low- Page | 250 CHAPTER SIX

fouling OEG was essential in preventing the formation of polymer-protein complexes which would hinder the antimicrobial efficacy. In addition, primary amine groups that mimic the amino acid, 2,4-diaminobutyric acid (Dab), yield better biocompatibility than amines that mimic lysine because of the shorter alkyl spacer group from the polymer backbone (2 vs. 4 -CH2-). Furthermore, the incorporation of either 2-ethylhexyl or 2-phenylethyl groups as the hydrophobic component resulted in polymers that effectively cause bacterial membrane wall disruption with minimal resistance development, while maintaining relatively good cytocompatibility with mammalian cells. In theory, besides variation of the chemical composition, other factors such as the polymer chain length and topology could influence the biological properties of an antimicrobial polymer.

Therefore, this study investigates the effect of various macromolecular structural variables on the biological properties of amphiphilic ternary copolymers. Each of these variables is chosen for specific reasons. Firstly, shorter polymer chain lengths (DPn of 50 and 20 compared to 100 in our previous publication) are investigated as they mimic the length of most naturally occurring AMPs (20-40 amino acid residues per peptide chain).34, 48 Secondly, the block copolymer topology is examined as this architecture resembles the amphipathic characteristic of some AMPs (e.g., melittin). Finally, branched architectures are evaluated since other similar examples (e.g., star polymers36, 41) have demonstrated improved bioactivity compared to the linear analogues. All ternary copolymers are synthesized via the RAFT polymerization of OEG acrylate, tert-butyl (4-acrylamidobutyl) carbamate, and 2-ethylhexyl acrylate (or

2-phenylethyl acrylate) monomers, followed by the removal of tert-

Page | 251 CHAPTER SIX

butyloxycarbonyl (Boc) protecting groups with TFA to yield primary amino groups.

6.3.1 Effect of the polymer chain length on biological performance

Three linear random copolymers with DPn of 100, 50 and 20 (denoted as L1, L2 and L3, respectively) that consist of 2-ethylhexyl groups as the hydrophobic component were synthesized to evaluate the effect of the polymer chain length on the antimicrobial activity and hemocompatibility. The molar feed ratio of OEG

: amine : hydrophobic groups was fixed to 3 : 5 : 2 in all three polymers. The Boc- protected polymers produced monomodal molecular weight distributions with dispersity (Ɖ) values of ca. 1.2-1.4 as evidenced by GPC analysis (Figure 6.2 a and

Table 6.1). It is noteworthy that the number-average molecular weight (Mn) values based on GPC analysis were relative to poly(methyl methacrylate) calibration standards and as such serve only as estimates. 1H NMR analysis using the RAFT terminal groups as a reference showed good agreement with experimental and theoretical Mn values. Further analysis of the NMR spectra also confirmed the chemical compositions of the polymers to be identical to the molar feed ratio. The treatment of the polymers with TFA resulted in the quantitative removal of Boc groups, as confirmed by the absence of tert-butyl protons at δH

1.45 ppm (Figure D 1, Figure D 2 and Figure D 3). The hydrodynamic diameter

(Dh) of the Boc-deprotected polymers in water was determined by DLS measurements. Polymers L1, L2 and L3 have an estimated Dh values of 0.8, 1.4 and 1.3 nm, respectively (Figure 6.2 b), which support the formation of single- chain polymeric nanoparticles in accord with our previous publication.47 We postulate that the slightly lower Dh value of L1 in comparison with L2 and L3

Page | 252 CHAPTER SIX

might be associated with the high ability of longer chains to undergo self- assembly to form single-chain polymeric nanoparticles in aqueous media. In addition, zeta potential (ζ) analysis of the polymers revealed that the polymers expectedly have net positive charges (+18.7 to +32.8 mV) due to the presence of primary amine groups.

Figure 6.2. (a) GPC- differential refractive index (DRI) chromatograms of the Boc-protected polymers L1, L2 and L3. (b) DLS normalized volume distributions of Boc-deprotected L1, L2 and L3 polymers in water.

Table 6.1. Polymer Characterization by NMR, GPC, DLS and Zeta Potential Analysis.

Mn Mn Dh Entry Ɖb,c ζ (mV)d (g mol–1)a,b (g mol–1)b,c (nm)d L1 29000 12200 1.4 0.8 18.7 L2 14800 8100 1.2 1.4 24.9 L3 6100 5000 1.2 1.3 32.8 B1 19000 11000 1.7 43.7 36.3 B2 18300 10500 1.5 24.6 28.6 H1 - 12900 1.3 1.7 42.5 H2 - 17200 1.6 1.6 36.1 H3 - 12700 1.3 1.7 49.4 H4 - 13000 1.6 1.8 38.9 aDetermined via 1H NMR analysis. bBased on Boc-protected polymers. cDetermined via GPC analysis. dBased on Boc-deprotected polymers.

Page | 253 CHAPTER SIX

The antimicrobial activity of the polymers was assessed in vitro against Gram- negative bacteria Pseudomonas aeruginosa and Escherichia coli by determining their

MICs (Table 6.2). As mentioned previously, our focus in this study is on Gram- negative pathogens as infections caused by these bacteria are more severe than those caused by Gram-positive bacteria. L1, L2 and L3 have a similar MIC (64 μg mL–1) against E. coli. However, when tested against P. aeruginosa, L1 and L2, which have DPn of 100 and 50, respectively, were slightly more active than L3.

This suggests that polymers with the longer chain length are marginally more bacteriostatic than those with shorter chain lengths against P. aeruginosa.

Table 6.2. Antimicrobial and Hemolytic Activities of Antimicrobial Ternary Copolymers.

MICa (μg mL–1) HC50 (μg mL–1) Selectivityb Entry P. aeruginosa E. coli V. cholerae RBC HC50/MIC L1 32-64 32-64 128 2500 39 L2 64 64 - > 10000 (major)c - L3 128 64 - > 10000 (major)c - B1 >256 >256 - - - B2 >256 >256 - - - H1 64 32-64 128 > 10000 (minor)c - H2 64 64 128 > 10000 >156 H3 >256 - - - - H4 >256 - - - - aThe strains are P. aeruginosa PAO1, E. coli K12, and V. cholerae SIO. bSelectivity is defined as the ratio of HC50 to MIC against P. aeruginosa. cMajor or minor hemagglutination was observed for these samples. The symbol ‘-’ indicates not determined.

Next, we determined the hemocompatibility of L1, L2 and L3 with sheep red blood cells (RBCs) by comparing their HC50 values (Table 6.2). Polymer L1 was more hemolytic than L2 and L3. L1 has a HC50 value of 2 500 µg mL–1 whereas both L2 and L3 resulted in less than 50% lysis of RBCs even when tested at the highest concentration of 10 000 µg mL–1. It is worthwhile noting that the highest

Page | 254 CHAPTER SIX

polymer concentration used for hemolysis experiments in our previous publication was 2 000 µg mL–1.47 At a first glance, L2 and L3 represent polymers with the most optimal biological performance purely based on their MIC and

HC50 values. However, L2 and L3 caused hemagglutination, as evidenced by the inability to resuspend the RBCs following incubation with the polymers (Movie

S1, SI). While a high HC50 value is indicative of good hemocompatibility, it is also imperative for a compound to exhibit non- or low-hemagglutination for clinical applications. Thus, L2 and L3 cannot be considered as better than L1 in terms of the overall biological performance.

Judging from the combined analysis of MIC and HC50 of L1, L2 and L3, we deduced that the ability of linear random copolymers to cause membrane disruption is possibly influenced by the chain length. It is important to note that the ternary random copolymers were shown to cause membrane wall disruption.47 The results here seem to indicate that longer polymer chains are

(slightly) more effective in lysing membrane cell walls (bacteria or erythrocytes) given that L1 has a lower MIC and HC50 than L2 and L3. We postulate that longer polymer chains exhibited more extensive interactions with cell membranes and thus are better at causing membrane disruption events than shorter polymer chains. We are unsure, however, as to why hemagglutination only occurred with shorter polymer chains.

6.3.2 Effect of block copolymer architecture on biological performance

For the synthesis of block copolymers, a macroRAFT agent was first prepared via the random copolymerization of OEG acrylate and tert-butyl (4- acrylamidobutyl) carbamate, where the molar ratio of OEG : cationic monomer

Page | 255 CHAPTER SIX

was set at 1 :3. The polymerization proceeded at 70 °C for 3 h. The monomer conversion was 80% while the DPn was ca. 50 repeat units as determined by 1H

NMR analysis (Figure D 4). GPC analysis revealed monomodal molecular weight distribution with a Ɖ of 1.4 (Figure 6.3 a and Table 6.1).

Figure 6.3. (a) GPC DRI chromatograms of the macroRAFT agent and Boc-protected polymers B1 and B2. (b) DLS normalized volume distributions of Boc-deprotected B1 and

B2 polymers in water. The inset illustrates the formation of micelles via the self-assembly of the block copolymers in water.

Two hydrophobic monomers including 2-ethylhexyl acrylate and 2- phenylethyl acrylate, were subsequently employed in the chain extension steps to yield Boc-protected block copolymers B1 and B2, respectively. 1H NMR analysis revealed that there were ca. 30 repeating units of the hydrophobic moieties per chain in B1 and B2 (Figure D 5 and Figure D 6). Successful chain extension was obtained as the molecular weight distributions of Boc-protected

B1 and B2 shifted to shorter retention times, as observed by GPC analysis (Figure

6.3 a). Following the removal of Boc groups, the self-assembly of the block copolymers in water was followed by DLS. It is worth noting that the molar ratio of the hydrophobic : hydrophilic component was increased from 1 : 4 in the

Page | 256 CHAPTER SIX

random copolymer system to 3 : 5 in this case to ensure micelle formation in aqueous medium. Micelle formation was evident as DLS analysis revealed peaks with Dh values of 43.7 and 24.6 nm for B1 and B2, respectively (Figure 6.3 b).

Zeta-potential (ζ) analysis also confirmed the cationic character of the micelles, with B1 and B2 registering ζ values of +36.3 and +28.6 mV, respectively. The MICs of micelles B1 and B2 were assessed against P. aeruginosa and E. coli. To our surprise, B1 and B2 did not display bacteriostatic activity against both the Gram- negative bacteria even at a polymer concentration of 256 μg mL–1. Our initial hypothesis was that the micelles will first establish interactions with the bacterial cell membrane, followed by micelle disassembly and integration into the membrane lipid bilayer to cause membrane disruption. However, the in vitro antimicrobial tests suggest that micelles B1 and B2 perhaps never underwent micelle disassembly upon contact with the bacteria cells. Instead, we postulate that the micelles preferentially remain in their core-shell morphology rather than integrating with the bacterial membrane lipid bilayer. It is noteworthy that there are contrasting reports in the literature which suggest that block copolymers may or may not possess inherent antimicrobial activity.43 We strongly believe that the antimicrobial properties of block copolymers vary across systems (e.g., different polymer backbones, monomer combinations and type) and specifically in our case, the block copolymers are inactive against Gram-negative bacteria. Given that B1 and B2 were inactive against the bacteria, we did not pursue further hemolysis experiments with these polymers.

Page | 257 CHAPTER SIX

6.3.3 Effect of the hyperbranched architecture on biological performance

The synthesis of hyperbranched polymers was performed in the same fashion as for the linear random copolymers, albeit with the addition of a cross-linkable monomer (OEG diacrylate), similar to literature procedures.49-56 Two hyperbranched polymers which consisted of 2-ethylhexyl groups as the hydrophobic component, labelled as H1 and H2, were prepared using 2 and 5 molar equivalents of OEG diacrylate to the RAFT initiator, respectively. For both polymers, the polymerization was taken to full monomer conversion, as confirmed by 1H NMR analysis (Figure D 7 and Figure D 8). GPC analysis of the

Boc-protected H1 and H2 revealed multimodal molecular weight distributions typically observed for hyperbranched polymers,51 with H2 having a broader distribution and higher molecular weight species than H1 (Figure 6.4 a). After the removal of Boc groups, DLS analysis of the hyperbranched polymers showed that H1 and H2 have Dh values of 1.7 and 1.6 nm, respectively (Figure 6.4 and

Table 6.1). Meanwhile, the zeta potential measurement confirmed the cationic nature of the polymers (ζ > 36 mV). It is worthwhile noting that the zeta potential and hydrodynamic volume of the hyperbranched and linear random copolymers are comparable.

Page | 258 CHAPTER SIX

Figure 6.4. (a) GPC DRI chromatograms of the Boc-protected hyperbranched polymers H1,

H2, H3 and H4. (b) DLS normalized volume distributions of Boc-deprotected H1, H2, H3 and H4 polymers in water.

The MIC of H1 and H2 against P. aeruginosa and E. coli was 64 μg mL–1, which was similar to the linear random copolymers L1 and L2. In terms of hemocompatibility, both H1 and H2 have an HC50 values of > 10 000 μg mL–1, but

H1 caused minor hemagglutination whereas H2 did not (Movie S2, SI). Polymers prepared with a higher amount of OEG diacrylate prevented hemagglutination from occurring. This is especially true considering that major hemagglutination was present in L2. The experiments involving RBCs thus strongly suggest that branching in polymer structures can reduce the occurrence of hemagglutination, though the mechanism is unclear. We also prepared two additional hyperbranched polymers H3 and H4 that were made in the exact same manner as H1 and H2, respectively, but with 2-phenylethyl groups as the hydrophobic component. Surprisingly, H3 and H4 were inactive against the bacteria tested, even at a higher polymer concentration of 256 μg mL–1. This suggests that the ethylhexyl groups in branched polymers have better membrane disruption capabilities compared to phenylethyl groups, thereby leading to higher antimicrobial activity.

Page | 259 CHAPTER SIX

Taken together, hyperbranched polymer H2 has the best overall biological performance in this study. H2 has a selectivity (defined here as the ratio of HC50 to MIC against P. aeruginosa) of > 156, which is > 4 times greater than the selectivity of L1, one of the two lead polymers in our previous study.47 Further tests were performed with H2 (and H1) to ascertain their antimicrobial potency.

H1 and H2 were tested against a highly pathogenic Gram-negative species, Vibrio cholerae. The obtained MIC was 128 μg mL–1 which was the same as L1, thus confirming the ability of H1 and H2 in combating more pathogenic bacteria strains. Next, we investigated the ability of H2 to combat biofilms. Biofilm- related infections are usually hard to treat and often the main cause of chronic inflammations. For this experiment, P. aeruginosa biofilms were grown in the M9 medium for 6.5 h prior to incubation with the polymer for 1 h. Based on colony- forming unit (CFU) analysis, H2 (at a dosage of 64 μg mL–1) demonstrated good bactericidal properties, killing ≥ 99% and 90% of planktonic and biofilm bacteria, respectively. In addition, H2 was also capable of dispersing preformed biofilms.

The treatment of P. aeruginosa biofilms with 64 μg mL–1 of H2 for 1 h resulted in

46% reduction in biofilm biomass compared to untreated controls, as determined by the crystal violet (CV) staining assay. The biofilm biomass that remained following dispersal events was also visualized using a 3D tomographic microscope (NanoLive) with digital staining software (Figure 6.5). Interestingly, it was observed that there was less biofilm in the centre of the well compared to the edges in the H2 treated samples. Approximately, 63% of biofilm biomass was reduced (compared to untreated controls) taking into account the average of these different areas. For the untreated controls, the density of the biofilm was identical throughout the well. It is noteworthy that although the biofilm dispersal

Page | 260 CHAPTER SIX

induced by H2 was not uniform, the majority (90%) of the remaining biofilm biomass are dead bacteria according to the CFU analysis.

Figure 6.5. 2D and 3D tomographic microscopy images of untreated control (a) and H2 treated samples (b-d). Scale bar = 20µm.

6.4 Summary

In summary, a series of amphiphilic ternary copolymers composed of oligoethylene glycol, cationic and hydrophobic functional groups, and with different chain lengths and topologies were synthesized and evaluated for their antimicrobial efficacy and hemocompatibility. Gram-negative bacteria such as P. aeruginosa and E. coli were used to determine the antimicrobial activity of the polymers. The chain length of linear random copolymers has small influence on the antimicrobial activity, with longer chains having slightly higher antimicrobial

Page | 261 CHAPTER SIX

activity. However, we found that shorter polymers chains cause hemagglutination. When the hydrophilic and hydrophobic segments were segregated into two distinct blocks, the block copolymers lost their antimicrobial activity. Importantly, hyperbranched random copolymers that contain 2- ethylhexyl groups were observed to have the best overall biological performance, with HC50 > 10 000 μg mL–1, no hemagglutination, and an MIC of

64 μg mL–1 against P. aeruginosa and E. coli. Although hyperbranched random copolymers have slightly lower antimicrobial activity than linear random copolymers, the branched structures have higher hemocompatibility (by > 4 times). The hyperbranched polymers were also capable of killing planktonic and biofilm bacteria, as well as inducing the dispersal of biofilms. Taken together, this study thus helps in identifying key macromolecular variations that will aid in the design of bio- and hemo-compatible antimicrobial polymers.

6.5 Reference

1. Wright, G. D., Solving the antibiotic crisis. ACS Infect. Dis 2015, 1 (2), 80-84. 2. Taubes, G., The Bacteria Fight Back. Science 2008, 321 (5887), 356-361. 3. Wo, Y.; Brisbois, E. J.; Bartlett, R. H.; Meyerhoff, M. E., Recent advances in thromboresistant and antimicrobial polymers for biomedical applications: just say yes to nitric oxide (NO). Biomater. Sci. 2016, 4 (8), 1161-1183. 4. Song, J.; Jang, J., Antimicrobial polymer nanostructures: Synthetic route, mechanism of action and perspective. Adv. Colloid Interface Sci. 2014, 203, 37-50. 5. Willyard, C., The drug-resistant bacteria that pose the greatest health threats. Nature 2017, 543 (7643). 6. Ling, L. L.; Schneider, T.; Peoples, A. J.; Spoering, A. L.; Engels, I.; Conlon, B. P.; Mueller, A.; Schäberle, T. F.; Hughes, D. E.; Epstein, S., A new antibiotic kills pathogens without detectable resistance. Nature 2015, 517 (7535), 455-459. 7. Payne, D. J.; Gwynn, M. N.; Holmes, D. J.; Pompliano, D. L., Drugs for bad bugs: confronting the challenges of antibacterial discovery. Nat. Rev. Drug Discov. 2007, 6 (1), 29-40. 8. Boyer, C.; Bulmus, V.; Davis, T. P.; Ladmiral, V.; Liu, J.; Perrier, S., Bioapplications of RAFT polymerization. Chem. Rev. 2009, 109 (11), 5402-5436. 9. Hawker, C. J.; Bosman, A. W.; Harth, E., New Polymer Synthesis by Nitroxide Mediated Living Radical Polymerizations. Chem. Rev. 2001, 101 (12), 3661-3688.

Page | 262 CHAPTER SIX

10. Matyjaszewski, K.; Xia, J., Atom Transfer Radical Polymerization. Chem. Rev. 2001, 101 (9), 2921-2990. 11. Matyjaszewski, K.; Tsarevsky, N. V., Nanostructured functional materials prepared by atom transfer radical polymerization. Nat. Chem. 2009, 1 (4), 276-288. 12. Moad, G.; Rizzardo, E.; Thang, S. H., Living Radical Polymerization by the RAFT Process A Second Update. Aust. J. Chem. 2009, 62 (11), 1402-1472. 13. Moad, G.; Rizzardo, E.; Thang, S. H., RAFT polymerization and some of its applications. Chem. Asian J. 2013, 8 (8), 1634-1644. 14. Kuroda, K.; Caputo, G. A.; DeGrado, W. F., The role of hydrophobicity in the antimicrobial and hemolytic activities of polymethacrylate derivatives. Chem. Eur. J. 2009, 15 (5), 1123-1133. 15. Ilker, M. F.; Nüsslein, K.; Tew, G. N.; Coughlin, E. B., Tuning the Hemolytic and Antibacterial Activities of Amphiphilic Polynorbornene Derivatives. J. Am. Chem. Soc. 2004, 126 (48), 15870-15875. 16. Kuroda, K.; DeGrado, W. F., Amphiphilic Polymethacrylate Derivatives as Antimicrobial Agents. J. Am. Chem. Soc. 2005, 127 (12), 4128-4129. 17. Ishitsuka, Y.; Arnt, L.; Majewski, J.; Frey, S. L.; Ratajczak, M.; Kjaer, K.; Tew, G. N.; Lee, K. Y. C., Amphiphilic Poly(phenyleneethynylene)s Can Mimic Antimicrobial Peptide Membrane Disordering Effect by Membrane Insertion [J. Am. Chem. Soc. 2006, 128, 13123−13129]. J. Am. Chem. Soc. 2008, 130 (7), 2372-2372. 18. Palermo, E. F.; Sovadinova, I.; Kuroda, K., Structural Determinants of Antimicrobial Activity and Biocompatibility in Membrane-Disrupting Methacrylamide Random Copolymers. Biomacromolecules 2009, 10 (11), 3098-3107. 19. Palermo, E. F.; Vemparala, S.; Kuroda, K., Cationic Spacer Arm Design Strategy for Control of Antimicrobial Activity and Conformation of Amphiphilic Methacrylate Random Copolymers. Biomacromolecules 2012, 13 (5), 1632-1641. 20. Paslay, L. C.; Abel, B. A.; Brown, T. D.; Koul, V.; Choudhary, V.; McCormick, C. L.; Morgan, S. E., Antimicrobial Poly(methacrylamide) Derivatives Prepared via Aqueous RAFT Polymerization Exhibit Biocidal Efficiency Dependent upon Cation Structure. Biomacromolecules 2012, 13 (8), 2472-2482. 21. Michl, T. D.; Locock, K. E.; Stevens, N. E.; Hayball, J. D.; Vasilev, K.; Postma, A.; Qu, Y.; Traven, A.; Haeussler, M.; Meagher, L., RAFT-derived antimicrobial polymethacrylates: elucidating the impact of end-groups on activity and cytotoxicity. Polym. Chem. 2014, 5 (19), 5813-5822. 22. Mowery, B. P.; Lindner, A. H.; Weisblum, B.; Stahl, S. S.; Gellman, S. H., Structure− activity relationships among random nylon-3 copolymers that mimic antibacterial host-defense peptides. J. Am. Chem. Soc. 2009, 131 (28), 9735-9745. 23. Wang, H.; Shi, X.; Yu, D.; Zhang, J.; Yang, G.; Cui, Y.; Sun, K.; Wang, J.; Yan, H., Antibacterial Activity of Geminized Amphiphilic Cationic Homopolymers. Langmuir 2015, 31 (50), 13469-13477. 24. Punia, A.; Mancuso, A.; Banerjee, P.; Yang, N.-L., Nonhemolytic and Antibacterial Acrylic Copolymers with Hexamethyleneamine and Poly(ethylene glycol) Side Chains. ACS Macro Lett. 2015, 4 (4), 426-430. 25. Zhang, J.; Markiewicz, M. J.; Mowery, B. P.; Weisblum, B.; Stahl, S. S.; Gellman, S. H., C-Terminal Functionalization of Nylon-3 Polymers: Effects of C- Terminal Groups on Antibacterial and Hemolytic Activities. Biomacromolecules 2012, 13 (2), 323-331. 26. Oda, Y.; Kanaoka, S.; Sato, T.; Aoshima, S.; Kuroda, K., Block versus Random Amphiphilic Copolymers as Antibacterial Agents. Biomacromolecules 2011, 12 (10), 3581-3591.

Page | 263 CHAPTER SIX

27. Wang, Y.; Xu, J.; Zhang, Y.; Yan, H.; Liu, K., Antimicrobial and hemolytic activities of copolymers with cationic and hydrophobic groups: A comparison of block and random copolymers. Macromol. Biosci. 2011, 11 (11), 1499-1504. 28. Tew, G. N.; Scott, R. W.; Klein, M. L.; DeGrado, W. F., De novo design of antimicrobial polymers, foldamers, and small molecules: from discovery to practical applications. Acc. Chem. Res. 2009, 43 (1), 30-39. 29. Lienkamp, K.; Tew, G. N., Synthetic Mimics of Antimicrobial Peptides—A Versatile Ring‑ Opening Metathesis Polymerization Based Platform for the Synthesis of Selective Antibacterial and Cell‑ Penetrating Polymers. Chem. Eur. J. 2009, 15 (44), 11784-11800. 30. Lienkamp, K.; Madkour, A. E.; Kumar, K. N.; Nüsslein, K.; Tew, G. N., Antimicrobial Polymers Prepared by Ring‑ Opening Metathesis Polymerization: Manipulating Antimicrobial Properties by Organic Counterion and Charge Density Variation. Chem. Eur. J. 2009, 15 (43), 11715-11722. 31. Lienkamp, K.; Kumar, K. N.; Som, A.; Nüsslein, K.; Tew, G. N., “Doubly selective” antimicrobial polymers: How do they differentiate between bacteria? Chem. Eur. J. 2009, 15 (43), 11710-11714. 32. Takahashi, H.; Caputo, G. A.; Vemparala, S.; Kuroda, K., Synthetic random copolymers as a molecular platform to mimic host-defense antimicrobial peptides. Bioconjugate Chem. 2017, 28 (5), 1340-1350. 33. Porel, M.; Thornlow, D. N.; Phan, N. N.; Alabi, C. A., Sequence-defined bioactive macrocycles via an acid-catalysed cascade reaction. Nat. Chem. 2016, 8 (6), 590-596. 34. Zasloff, M., Antimicrobial peptides of multicellular organisms. Nature 2002, 415 (6870), 389-395. 35. Lam, S. J.; Wong, E. H.; O’Brien-Simpson, N. M.; Pantarat, N.; Blencowe, A.; Reynolds, E. C.; Qiao, G. G., Bionano Interaction Study on Antimicrobial Star-Shaped Peptide Polymer Nanoparticles. ACS Appl. Mater. Interfaces 2016, 8 (49), 33446-33456. 36. Lam, S. J.; O'Brien-Simpson, N. M.; Pantarat, N.; Sulistio, A.; Wong, E. H.; Chen, Y.-Y.; Lenzo, J. C.; Holden, J. A.; Blencowe, A.; Reynolds, E. C., Combating multidrug-resistant Gram-negative bacteria with structurally nanoengineered antimicrobial peptide polymers. Nat. Microbiol. 2016, 1, 16162. 37. Kenawy, E.-R.; Worley, S. D.; Broughton, R., The Chemistry and Applications of Antimicrobial Polymers: A State-of-the-Art Review. Biomacromolecules 2007, 8 (5), 1359-1383. 38. Palermo, E. F.; Kuroda, K., Chemical Structure of Cationic Groups in Amphiphilic Polymethacrylates Modulates the Antimicrobial and Hemolytic Activities. Biomacromolecules 2009, 10 (6), 1416-1428. 39. Mizutani, M.; Palermo, E. F.; Thoma, L. M.; Satoh, K.; Kamigaito, M.; Kuroda, K., Design and Synthesis of Self-Degradable Antibacterial Polymers by Simultaneous Chain- and Step-Growth Radical Copolymerization. Biomacromolecules 2012, 13 (5), 1554-1563. 40. Yuan, W.; Wei, J.; Lu, H.; Fan, L.; Du, J., Water-dispersible and biodegradable polymer micelles with good antibacterial efficacy. Chem. Commun. 2012, 48 (54), 6857- 9. 41. Wong, E. H.; Khin, M. M.; Ravikumar, V.; Si, Z.; Rice, S. A.; Chan-Park, M. B., Modulating antimicrobial activity and mammalian cell biocompatibility with glucosamine-functionalized star polymers. Biomacromolecules 2016, 17 (3), 1170-1178.

Page | 264 CHAPTER SIX

42. Wan, X.; Zhang, Y.; Deng, Y.; Zhang, Q.; Li, J.; Wang, K.; Li, J.; Tan, H.; Fu, Q., Effects of interaction between a polycation and a nonionic polymer on their cross- assembly into mixed micelles. Soft Matter 2015, 11 (21), 4197-207. 43. Lam, S. J.; Wong, E. H. H.; Boyer, C.; Qiao, G. G., Antimicrobial polymeric nanoparticles. Prog. Polym. Sci. 2017, 76, 40-64. 44. Chen, X.; Zhang, G.; Zhang, Q.; Zhan, X.; Chen, F., Preparation and Performance of Amphiphilic Copolymers with Capsaicin-Mimic and PEG Moieties for Protein Resistance and Antibacteria. Ind. Eng. Chem. Res. 2015, 54 (15), 3813-3820. 45. Cheng, G.; Xue, H.; Li, G.; Jiang, S., Integrated antimicrobial and nonfouling hydrogels to inhibit the growth of planktonic bacterial cells and keep the surface clean. Langmuir 2010, 26 (13), 10425-8. 46. Mi, L.; Jiang, S., Synchronizing nonfouling and antimicrobial properties in a zwitterionic hydrogel. Biomaterials 2012, 33 (35), 8928-33. 47. Nguyen, T.-K.; Lam, S. J.; Ho, K. K.; Kumar, N.; Qiao, G. G.; Egan, S.; Boyer, C.; Wong, E. H., Rational Design of Single-Chain Polymeric Nanoparticles That Kill Planktonic and Biofilm Bacteria. ACS Infect. Dis 2017, 3 (3), 237-248. 48. Liu, S. P.; Zhou, L.; Lakshminarayanan, R.; Beuerman, R. W., Multivalent Antimicrobial Peptides as Therapeutics: Design Principles and Structural Diversities. Int. J. Pept. Res. Ther. 2010, 16 (3), 199-213. 49. Luzon, M.; Boyer, C.; Peinado, C.; Corrales, T.; Whittaker, M.; Tao, L.; Davis, T. P., Water‑ soluble, thermoresponsive, hyperbranched copolymers based on PEG‑ methacrylates: Synthesis, characterization, and LCST behavior. J. Polym. Sci., Part A: Polym. Chem. 2010, 48 (13), 2783-2792. 50. O'Brien, N.; McKee, A.; Sherrington, D. C.; Slark, A. T.; Titterton, A., Facile, versatile and cost effective route to branched vinyl polymers. Polymer 2000, 41 (15), 6027-6031. 51. Gao, H.; Matyjaszewski, K., Synthesis of functional polymers with controlled architecture by CRP of monomers in the presence of cross-linkers: From stars to gels. Progress in Polymer Science 2009, 34 (4), 317-350. 52. Slark, A. T.; Sherrington, D. C.; Titterton, A.; Martin, I. K., Branched methacrylate copolymers from multifunctional comonomers: the effect of multifunctional monomer functionality on polymer architecture and properties. J. Mater. Chem. 2003, 13 (11), 2711-2720. 53. Vogt, A. P.; Sumerlin, B. S., Tuning the Temperature Response of Branched Poly(N-isopropylacrylamide) Prepared by RAFT Polymerization. Macromolecules 2008, 41 (20), 7368-7373. 54. Isaure, F.; Cormack, P. A. G.; Sherrington, D. C., Facile synthesis of branched poly(methyl methacrylate)s. J. Mater. Chem. 2003, 13 (11), 2701-2710. 55. Wang, Z.; He, J.; Tao, Y.; Yang, L.; Jiang, H.; Yang, Y., Controlled Chain Branching by RAFT-Based Radical Polymerization. Macromolecules 2003, 36 (20), 7446-7452. 56. Li, C.; He, J.; Li; Cao, J.; Yang, Y., Controlled Radical Polymerization of Styrene in the Presence of a Polymerizable Nitroxide Compound. Macromolecules 1999, 32 (21), 7012-7014.

Page | 265 CHAPTER SIX

6.6 Appendix D

Figure D 1. 1H NMR spectra of Boc-protected linear polymer L1 in CDCl3 (a) and the corresponding Boc-deprotected L1 in D2O (b).

Page | 266 CHAPTER SIX

Figure D 2. 1H NMR spectra of Boc-protected linear polymer L2 in CDCl3 (a) and the corresponding Boc-deprotected L2 in D2O (b).

Page | 267 CHAPTER SIX

Figure D 3. 1H NMR spectra of Boc-protected linear polymer L3 in CDCl3 (a) and the corresponding Boc-deprotected L3 in D2O (b).

Page | 268 CHAPTER SIX

Figure D 4. 1H NMR spectra of Boc-protected macroRAFT agent in CDCl3.

Figure D 5. 1H NMR spectra of Boc-protected block copolymer B1 in CDCl3.

Page | 269 CHAPTER SIX

Figure D 6. 1H NMR spectra of Boc-protected block copolymer B2 in CDCl3.

Figure D 7. 1H NMR spectra of Boc-protected hyperbranched polymer H1 in CDCl3.

Page | 270 CHAPTER SIX

Figure D 8. 1H NMR spectra of Boc-protected hyperbranched polymer H2 in CDCl3.

Page | 271

CHAPTER SEVEN

Concluding Remarks and Future

Perspectives

CHAPTER SEVEN

7 Concluding remarks and future perspectives

7.1 Conclusions

The therapeutic gap caused by the advent of antibiotic resistance and the lack of new antibiotic discovery poses a substantial risk to public health worldwide.

The evidence of resistance acquisition has been observed against all currently used antibiotics, and cases of multidrug-resistant (MDR) infections have been reported in which, bacteria were resistant to almost all available antibiotics. In addition, the formation of multicellular sessile communities of bacterial cells, known as biofilms, further diminishes the efficiency of antibiotics to combat infections. Biofilm bacteria are highly recalcitrant towards antibiotic therapies and host immune responses due to their unique characteristics. Therefore, the development of new and effective antimicrobials to combat MDR and biofilm- related infections is a global priority. The overall goal of this thesis was the development and evaluation of novel therapeutic approaches based on the coadministration of synthetic antimicrobial polymers, composed of oligo(ethylene glycol), cationic and hydrophobic functional groups, along with other antimicrobial agents, namely nitric oxide (NO), antibiotics, and essential oils, against bacterial infections. The synthetic antimicrobial polymer used in this thesis demonstrates strong antimicrobial activity against Gram-negative bacteria in both planktonic and biofilm forms with a significantly low risk of resistance development in bacteria. Moreover, initial biocompatibility assessments against rat H4IIE liver cells and red blood cells (hemolysis assay) suggest high mammalian cell biocompatibility for potential future clinical applications. It is

Page | 273 CHAPTER SEVEN

worthwhile noting that the proposed antimicrobial platform based on the combination of synthetic antimicrobial polymers and other antimicrobial agents is undoubtedly applicable to other antimicrobial polymers with different compositions and architectures. However, any new combination should be carefully assessed to determine the type of interactions between the components.

Similar to the combination of antibiotics, synergistic combinations based on antimicrobial polymers are less prevalent than combinations with additive and antagonistic.

In Chapter three, we introduced a potent antibiofilm agent in the form of a

NO-releasing antimicrobial polymer. Even though the individual antimicrobial polymer has shown antibiofilm activity (i.e., biofilm dispersal and killing effects) to some extent, we aimed to further increase its performance by using it in combination with a robust antibiofilm agent, that is, NO gas. Despite the strong antibiofilm activity, the applicability of NO in biomedical applications is limited by its short half-life and instability in biological environments. Therefore, we inferred that incorporation of NO donor molecules into the structure of the synthetic antimicrobial polymer would assist with the stabilization, controlled release, and delivery of NO to the biofilm site and consequently improve the potency of the antimicrobial polymer against biofilms. In contrast to many of polymeric NO delivery systems, where the polymeric nanoparticles are only used to encapsulate and/or deliver NO, in this study, the polymer was an active antimicrobial agent and also played the role of delivery vehicle for NO. NO loading was conducted in a facile way by the direct reaction of primary amines of the antimicrobial polymer with NO gas. The resulting NO-releasing antimicrobial polymers exhibited a synergistic effect in biofilm dispersal and

Page | 274 CHAPTER SEVEN

could effectively reduce 80% of Pseudomonas aeruginosa (P. aeruginosa) biofilm biomass upon 1 h of treatment at a polymer concentration of 64 μgmL−1. Besides, we observed >99.999% reduction in the viability of planktonic and biofilm P. aeruginosa cells within 1 h of treatment at a polymer concentration of 64 μgmL−1.

Interestingly, this synergistic effect in biofilm dispersal and killing activity was only observed when both NO donors and antimicrobial polymer exist as a single chemical entity since the physical mixture of antimicrobial polymer and a commercially available NO donor (spermine NONOate) did not result in any synergistic antimicrobial activity.

In Chapter four, we explored the potential use of our synthetic antimicrobial polymer in combination with various classes of commercially available antibiotics against Gram-negative planktonic and biofilm bacteria. We used the checkerboard assay to assess the polymer−antibiotic interactions against two

Gram-negative bacteria, namely P. aeruginosa and Escherichia coli. Despite the susceptibility of both tested bacteria to all selected antibiotics and synthetic antimicrobial polymer, we only observed the synergistic interactions for two combinations, involving doxycycline and colistin antibiotics. For all other combinations, the interaction between the antimicrobial polymer and antibiotics was indifference. Synergistic combinations demonstrated superior bacteriostatic activity against all Gram-negative bacteria tested, with the greatest synergism against MDR P. aeruginosa strains, where the minimal inhibitory concentrations

(MICs) of both components were reduced by at least 4-fold. When assessed for bactericidal activity, the synergistic effect was only achieved when the antimicrobial polymer was used in combination with doxycycline. Doxycycline could enhance the bactericidal activity of polymer, despite being bacteriostatic,

Page | 275 CHAPTER SEVEN

resulting in killing >99.999% of planktonic and biofilm P. aeruginosa PAO1 within a 20 min incubation at a polymer concentration of 128 μg mL−1 and doxycycline concentration of 64 μg mL−1. Moreover, resistance study revealed that antimicrobial polymer-doxycycline combination could significantly impede resistance acquisition in bacteria compared with the individual components. This synergistic combination was also capable of reviving the susceptibility to treatment in resistant strains. This study clearly showed the advantages of coadministration of synthetic antimicrobial polymers and antibiotics, in the form of cocktail treatment, as a novel therapeutic approach against superbugs.

Based on our experiences in combination therapies involving synthetic antimicrobial polymers and motivated by the results obtained from those studies, in Chapter five, we assessed the potential synergistic interactions between our synthetic antimicrobial polymer and two antimicrobial essential oils, namely carvacrol and eugenol. Although showing potency as antimicrobial/antibiofilm agents, poor distribution in biological media and low stability limit the practicality of essential oils in the antimicrobial applications. To address these shortcomings and to further potentiate the antimicrobial performance of our synthetic antimicrobial polymer, we co-administrated synthetic antimicrobial polymers, in the form of linear and block copolymers, and essential oils as an oil- in-water emulsion. The self-assembly of the block copolymers into core−shell micelle morphology resulted in the effective encapsulation of essential oils within the hydrophobic cores and the subsequent localized and targeted delivery of essential oils to the bacteria cells. We observed similar biofilm inhibition potencies for combinations that included either of linear or block copolymers, with about 60−75% and 70−85% biofilm inhibition effect against wild-type P.

Page | 276 CHAPTER SEVEN

aeruginosa PAO1 and MDR strain PA37, respectively, upon 6.5 h of treatment.

However, when tested for bactericidal activity, we only observed synergistic interactions between the block copolymer and essential oils. The block copolymer−oil combination could effectively kill >99.99% of biofilm P. aeruginosa

PAO1 upon a 20 min incubation at a polymer concentration of 64 µg mL−1 and essential oil concentration of 300 and 530 µg mL−1 for carvacrol and eugenol, respectively.

The investigation into the structure–activity relationship of synthetic antimicrobial polymers has gained substantial attention over the past two decades. These studies assist with identifying key macromolecular design parameters for the development of potent antimicrobial polymers. In this regard, in Chapter six, we conducted a structure–activity relationship study to evaluate the effect of polymer chain length (DP= 20, 50 and 100) and architecture (linear vs. block vs. hyperbranched) on antimicrobial and hemolytic activities of our ternary antimicrobial polymer. The chain length of linear random copolymers had little impact on the antimicrobial activity, with longer chains exhibiting marginally higher antimicrobial activity against tested Gram-negative bacteria, which was in accordance with the previous reports in the literature. In terms of their hemocompatibility, while the polymers with DP= 20 and 50 had higher HC50 values than the polymer with DP=100, we observed strong hemagglutination effect in the case of short polymer chains (DP= 20 and 50). The segregation of hydrophobic monomers into a distinct block and the subsequent formation of core–shell micelles led to a complete loss of antimicrobial activity for block copolymers. We postulated that the formed micelles failed to efficiently interact with the bacterial membrane through their hydrophobic segments as the micelles

Page | 277 CHAPTER SEVEN

were unable to disassemble upon contact with bacterial cells. We observed the best overall performance for the hyperbranched random copolymers that contained 2-ethylhexyl groups, with a MIC of 64 μg mL−1 against P. aeruginosa and E. coli, HC50 > 10 000 μgmL−1, and no hemagglutination. This polymer also showed bactericidal activity and killed ≥99% of planktonic and 90% of biofilm P. aeruginosa upon 1 h treatment at a concentration of 64 µg mL−1. The hyperbranched architecture seemed to reduce the non-specific interactions with the red blood cells and hence improve the overall biological performance of the antimicrobial polymer.

In terms of comparison, each of the proposed combinations in this thesis has its pros and cons. The combination involving NO (Chapter 3) has shown the highest activity against bacterial biofilm. This combination was able to demonstrate biofilm dispersal and also bactericidal activity at very low concentrations of NO in comparison with the individual NO donor. However, long-term stability and storage of the NO-loaded antimicrobial polymer is an issue regarding the potential clinical application of this combination and requires more investigations for future projects. The combination based on the physical mixture of commercially available antibiotics and antimicrobial polymer

(Chapter 4) was probably the most convenient combination in this thesis in terms of preparation and use, where the stock solutions of each individual components could be prepared and stored for an extended time with no risk of degradation or loss of activity. The synergistic combinations of antimicrobial polymer and antibiotics could effectively fight the MDR strains and reduced the risk of resistance development in bacteria. However, it is noteworthy that the number of the synergistic combinations was relatively low, and the majority of tested

Page | 278 CHAPTER SEVEN

combinations showed indifference interactions with no improvement in antimicrobial performance. Finally, the combination of antimicrobial polymer in the form of polymeric core-shell micelles and essential oils (Chapter 5) represented a potent antibiofilm platform, where the antimicrobial polymer plays a dual role as an antimicrobial agent and also as the delivery vehicle for the localized delivery of hydrophobic components to the bacterial biofilms.

Although the hydrophobic antimicrobial component used in this chapter

(essential oil) was not extremely active, their combination with antimicrobial polymeric micelles showed improved antimicrobial activity. This shows the potential of the antimicrobial polymeric micelles to be used in combination with a wide range of hydrophobic compounds. The stability of the prepared combinations can be further improved through the incorporation of cross- linkable moieties in the structure of the antimicrobial polymer.

Finally, it is worthwhile noting that all the presented results in this thesis are based on in vitro experiments and for any future clinical applications more extensive toxicity assays and also in vivo studies using animal models are required. For instance, as the synthetic antimicrobial polymers used in this thesis are not biodegradable, the investigation of long-term toxicity is necessary.

7.2 Challenges and future perspectives

The field of synthetic antimicrobial polymers is one of the innovative approaches that have been developed in response to the increasing prevalence of

MDR infections. The field has been growing significantly as the result of extensive multidisciplinary research that have been conducted since the first

Page | 279 CHAPTER SEVEN

reports on antimicrobial peptide (AMP)-mimicking polymers in the 2000s.

Synthetic antimicrobial polymers demonstrate comparable biological performance (i.e., antimicrobial activity and biocompatibility) to their natural counterparts (i.e., AMPs), but at a significantly lower synthesis burden, and in comparison with antibiotics are much less probable to induce resistance acquisition in bacteria. The advances in chemistry and controlled polymerization techniques have made it easier than ever to precisely synthesis antimicrobial polymers in significantly short periods, which allows for rapid screening of polymers with various structural features. Combined, all these benefits reinforce the idea that synthetic antimicrobial polymers could be potential alternatives to conventional antibiotics, which are experiencing major failure in the battle against MDR bacteria.

Besides the high potency of synthetic antimicrobial polymers as individual antimicrobial agents, they have also shown great performance as coagents in antimicrobial combination therapies. The mode of action of antimicrobial polymers (i.e., membrane disruption) makes them ideal candidates to be used as adjuvants to potentiate the action of other antimicrobials such as antibiotics.

Moreover, possible synergistic interactions between antimicrobial polymers and various synthetic or natural antimicrobial agents would open up new opportunities to tackle antibiotic resistance. While a great deal of research has been conducted on the synthesis and evaluation of antimicrobial polymers, combination therapies involving antimicrobial polymers have remained relatively unexplored.

Page | 280 CHAPTER SEVEN

Despite their promising potential, the practical clinical application of antimicrobial polymers, either individually or in combination with other antimicrobial agents, requires certain aspects and challenges to be addressed.

Biocompatibility is probably the chief concern regarding the application of antimicrobial polymers in clinical settings. While the lack of target specificity allows antimicrobial polymers to demonstrate a broad-spectrum antimicrobial activity with minimal risk of resistance development, it also leads to non-specific interactions with mammalian cells. The development of strategies to improve the selectivity of antimicrobial polymers toward bacteria over mammalian cells is a crucial aspect that needs further consideration. This could be done through systematic structure-activity relationship studies followed by detailed mechanistic investigations to identify key design parameters affecting the selectivity. Furthermore, in vivo studies are also imperative as antimicrobial polymers might exhibit different biological activity or cause toxicity/adverse effects in the body.

Switching from monotherapy to combination therapy adds more complexity and pitfalls to the therapeutic system. Apart from challenges associated with each individual component, coadministration of components would also raise new questions regarding their formulation and dosing, administration method and pharmacokinetics.

Combination therapy using synthetic antimicrobial polymers is a novel platform to combat MDR bacteria. There have been several potent combinations thus far, and there are many more possible combinations to explore. Advances in polymer science provide the ability to readily manipulate the properties of

Page | 281 CHAPTER SEVEN

antimicrobial polymers according to the characteristics of their coagent(s) and also the requirements of each specific antimicrobial application. The followings are a few suggestions that can potentially advance combination therapy systems involving antimicrobial polymers: (1) using antimicrobial polymers in the form of nanoparticles (e.g., micelles, vesicles, hyperbranched polymers, star polymers, dendrimer, etc.) which allows for encapsulation of the coagent for more efficient delivery to the target site and also would improve the biocompatibility of the polymer, (2) improving treatment efficiency by incorporating targeting moieties into the polymer structure, (3) incorporating stimuli-responsive function to the combination therapy system to achieve controlled-release of components in response to specific environmental cues (e.g., pH, enzymes, temperature, light, etc.).

Page | 282