Novel Antibacterial Biomaterials and Polymers Based on Quorum Sensing Inhibitors

NOVEL ANTIBACTERIAL BIOMATERIALS AND POLYMERS BASED ON QUORUM SENSING INHIBITORS

A thesis submitted in fulfilment of the degree of

Doctor of Philosophy

Aditi Taunk

Supervisors

Prof. Naresh Kumar Prof. Mark D.P. Willcox Prof. David StC. Black

School of Chemistry The University of New South Wales Kensington, Australia

December 2017

This thesis is dedicated to my beloved mother, Dr. Archana Taunk for her unconditional love & support and for always believing in me!

THE UNIVERSITY OF NEW SOUTH WALES Thesis/DissertationSheet

Surname or Family name: TAUNK

Firsi name: ADITI Other name/s:

Abbreviation for degree as given in the Universitycalendar: PhD Faculty: Science

School: School of Chemistry

Title: Novel Antibacterial Biomaterials and Polymers Based on Quorum Sensing Inhibitors

ABSTRACT

Bacterial biofilms on life-saving implanted medical devices are a serious problem in long-term. At present, no effective strategies are available and the emergence of multi-drug resistance has highlighted the need to develop novel antibacterial coatings to combat device- related infections. One approach is to block the bacterial communication pathway or quorum sensing (QS), which is responsible for biofilm formation, by incorporating QS inhibitors (QSls) such as dihydropyrrolones (DHPs) and furanones (FUs) on biomaterial surfaces and polymers. The endogenous biological signalling molecule nitric oxide (NO) is also a potential candidate for prevention of biomedical infections due to its antibiofilm activity.

In this study, DHPs and brominated FUs were immobilized on surfaces via a non-specific nitrene-insertion method. The successful covalent attachment of compounds was confirmed by X-ray photoelectron spectroscopy. The coated surfaces showed excellent in vitro activity against Staphylococcus aureus and Pseudomonas aeruginosa. Interestingly, DHP surfaces at low concentrations (0.17-0.35 % halogen) were found to display similar levels of activity as FUs with higher surface attachment (0.41-0.74 % Br), which was possibly due to change in orientation of DHP during attachment.

The influence of DHP orientation and absence of an exocyclic double bond on the biological activity was then examined by specific covalent attachment using EDC/NHS coupling. The orientation of DHP with free lactam ring exposed to bacterial medium showed higher activity compared to DHPs attached from the nitrogen of the lactam ring. In addition, DHPs lacking the exocyclic double bond were also able to reduce bacterial adhesion without killing both strains of bacteria, indicating DHPs retained their activity even in absence of the exocyclic bond.

This project also focused on developing dual-action surfaces and polymers that were functionalized by DHPs via Michael-addition reaction and diazeniumdiolates (NO donors) derived from the reaction of secondary amines with NO gas. The DHP+NO surfaces demonstrated significantly higher efficacy in reducing colonization of both bacterial strains than the DHP coatings alone, while the hybrid polymer displayed excellent activity by inhibiting 95 % of P. aeruginosa biofilm at all concentrations (42-1 µM) via a non-toxic mechanism.

Therefore, the coatings based on QSls show great potential in reducing device-related infections.

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 Universitylibraries 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.

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FOR OFFICE USE ONLY Date of completion of requirements for Award: CERTIFICATE OF ORIGINALITY

‘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 material which to a substantial extent has 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.’

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‘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.'

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ABSTRACT

Bacterial biofilms on life-saving implanted medical devices are a serious problem in long-term. At present, no effective strategies are available and the emergence of multi- drug resistance has highlighted the need to develop novel antibacterial coatings to combat device-related infections. One approach is to block the bacterial communication pathway or quorum sensing (QS), which is responsible for biofilm formation, by incorporating QS inhibitors (QSIs) such as dihydropyrrolones (DHPs) and furanones

(FUs) on biomaterial surfaces and polymers. The endogenous biological signalling molecule nitric oxide (NO) is also a potential candidate for prevention of biomedical infections due to its antibiofilm activity.

Interestingly, DHP surfaces at low concentrations (0.17–0.35 % halogen) were found to display similar levels of activity as FUs with higher surface attachment (0.41–0.74 %

Br), which was possibly due to change in orientation of DHP during attachment.

In addition, DHPs lacking the exocyclic double bond were also able to reduce bacterial

iii adhesion without killing both strains of bacteria, indicating DHPs retained their activity even in absence of the exocyclic bond.

1 µM) via a non-toxic mechanism.

Therefore, the coatings based on QSIs show great potential in reducing device-related infections.

ACKNOWLEDGEMENTS

I would like to take this opportunity and thank everyone without whom this thesis would not have been written and to whom I am greatly indebted. First and foremost, I would like to express my sincere gratitude to my supervisor Prof. Naresh Kumar for the constant support, patience, motivation and immeasurable intellectual input throughout my PhD. Thank you for encouraging and guiding me with all the invaluable suggestions at all times. I could not have imagined having a better supervisor and mentor for my research. I am also extremely grateful to my co-supervisors, Prof. Mark Willcox and

Prof. David StC Black, for their helpful insights and suggestions.

Besides my supervisors, my sincere thanks also goes to an important member of the

Kumar/Black group, Dr. George Iskander, for his warm encouragement and guidance all these years. Thank you for always helping me out and teaching me different techniques that were very crucial for me to improve my skills in organic synthesis. I would also like to thank Dr. Kitty Ho and Dr. Ren Chen who made me understand my project by always patiently answering my questions and clearing all my doubts. Many thanks for training me with the biology experiments, helping with confocal, XPS and proof reading my thesis.

I would also like to extend my thanks to School of Optometry and Biomedical Imaging

Facility (BMIF) for letting me conduct my microbiology experiments in their lab and providing all the resources necessary for my work. I am grateful for the help I received from Dr. Debarun Dutta and Dr. Ajay Vijay with my biology lab work. I wish to thank

Dr. Michael Carnell for the confocal microscopy training; Dr. Bill Gong and Outi

Mustonen for XPS characterization of my samples; the entire staff of the NMR and

BMSF facility for characterization of my synthesized compounds, especially Dr.

Douglas Lawes for the NMR training; and also Dr. Nancy Scoleri and Dr. Peta Di Bella for the chemistry instrument training.

My PhD would not have been possible without the scholarships I received, the

Australian Postgraduate Award (APA) from the Australian Government and UNSW

Research Excellence Award from UNSW.

I would also like to thank all the past and present members of the Kumar/Black group for creating a positive, healthy workplace and helping me in many ways. I want to mention Samuel, Nripendra, Chris, Eugene, Hao, Jeremy, Vina, Vidia, Daniel, Jacky

Iméze, Murat, Marcin, Oscar, Yilin, Jimmy, Si Khay, An Phan, Rajesh, Shashi,

Basmah, Alice, Chelsea, Sharon, Keith, Jane, Tom, Kenneth and many more. Special thanks to Jeremy, Jacky, Iméze, Daniel, Vina and Vidia for all the fun and chats we had in and outside the lab. A special mention goes to Dr. Samuel Kutty and Dr. Nripendra

Nath Biswas for your wonderful friendship and kind support that helped me tremendously during my rough days in the lab.

A big thank you to my friends in Sydney, Ananya and Daman, for bearing with me during the toughest days of my life, I don’t think I would have survived without your support. A special thank you to Aditya for all the little things that you do to cheer me up and for tolerating my mood swings and to my special friend Manisha for your constant love and encouragement all these years which I truly cherish. A warm thank you to my friends in India and Australia: Presee, Sonal, Raj, Piruz, Dhara, Shubham, Rasha, Lucy,

Zeenat, Rumana, Cathy, Rhiannon, Elaine, Harjot and many more. I would like to thank especially Presee for making this journey enjoyable; Lucy for being a wonderful

vi flatmate by sharing my stress throughout my PhD and Raj for the encouraging and motivational talks for which I am truly grateful

I also owe much gratitude to my dearest grandparents (nani and nana), my brother

Anurag and all my family members for sticking around with me during the darkest phase of my life. A special thank you to my family in Sydney, Vandana mausi, Lalit mausa and my cousins, who helped me settle down in a new country and also supported and guided me at every stage during my stay in Sydney.

Finally, and above all, I am deeply grateful to my mother, the most important person of my life and my inspiration, who always supported my ambitions and encouraged me to be headstrong in life. There are no words for me to express my unfailing gratitude for your endless love, care and sacrifices. I do not think I would have made it this far and reached at this point in my life without your belief and support. I am greatly indebted to you for this life and for everything you have done for me.

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TABLE OF CONTENTS

Certificate of Originality ...... i

Abstract ...... iii

Acknowledgements ...... v

Table of Contents ...... viii

List of Abbreviations ...... xiii

Publications and Presentations ...... xvi

CHAPTER-1: Introduction 1.1 Bacterial Infections ...... - 1 - 1.2 Resistance to Antibiotics ...... - 2 - 1.3 Biomedical Device-Related Infections ...... - 5 - 1.4 Bacterial Biofilm ...... - 6 - 1.5 Quorum Sensing ...... - 10 - 1.6 Quorum Sensing Pathways ...... - 12 - 1.6.1 Quorum Sensing in Gram-Negative Bacteria ...... - 12 - 1.6.2 Quorum Sensing in Gram-Positive Bacteria ...... - 15 - 1.6.3 Quorum Sensing in Gram-Negative and Gram-Positive Bacteria ...... - 17 - 1.7 Quorum Sensing Inhibition...... - 19 - 1.7.1 Natural Furanones ...... - 20 - 1.7.2 Synthetic Furanones ...... - 23 - 1.7.3 1,5-Dihydropyrrol-2-ones ...... - 25 - 1.8 Nitric Oxide ...... - 27 - 1.8.1 Nitric oxide as Antimicrobial and Antibiofilm Agent ...... - 28 - 1.8.2 Nitric Oxide Donors ...... - 30 - 1.8.3 Dual-Action Antibacterial Drugs ...... - 33 - 1.9 Current Antibacterial Surface Coatings...... - 36 - 1.9.1 Antibiotics ...... - 36 - 1.9.2 Silver ...... - 37 - 1.9.3 Quaternary ammonium compounds ...... - 37 - 1.9.4 Antimicrobial Peptides ...... - 38 - viii

1.10 Furanone Coatings...... - 39 - 1.11 Dihydropyrrolone Coatings ...... - 40 - 1.12 Nitric Oxide Releasing Coatings ...... - 43 - 1.13 Thesis Aims ...... - 45 -

CHAPTER-2: Surface Attachment of FU and DHP by Photochemical Activation 2.1 Introduction ...... - 47 - 2.2 Materials and Methods ...... - 51 - 2.2.1 Attachment of 3-Aminopropyltriethoxysilane (APTS) ...... - 51 - 2.2.2 Attachment of 4-Azidobenzoic Acid (ABA) ...... - 51 - 2.2.3 Attachment of FU and DHP via Azide/Nitrene Chemistry ...... - 52 - 2.2.4 X-Ray Photoelectron Spectroscopy (XPS) ...... - 52 - 2.2.5 Contact Angle Measurements ...... - 53 - 2.2.6 Bacterial Adhesion Analysis ...... - 53 - 2.2.7 Statistical Analysis of Data ...... - 55 - 2.3 Results ...... - 55 - 2.3.1 High-Resolution XPS Characterization of Coated Surfaces ...... - 55 - 2.3.2 Contact Angle Measurements ...... - 58 - 2.3.3 Antibacterial Activity ...... - 60 - 2.4 Discussion ...... - 65 - 2.5 Conclusion ...... - 68 -

CHAPTER-3: Role of Orientation of Surface Bound DHP on Biological Activity 3.1 Introduction ...... - 69 - 3.2 Materials and Methods ...... - 73 - 3.2.1 General ...... - 73 - 3.2.2 Synthesis of Acid-Functionalized DHP Derivatives ...... - 74 - 3.2.3 Quorum Sensing Inhibition Assay ...... - 79 - 3.2.4 Attachment of 3-Aminopropyltriethoxysilane (APTS) ...... - 79 - 3.2.5 Plasma Functionalization of FEP by Allylamine ...... - 80 - 3.2.6 Attachment of DHP via EDC/NHS Coupling...... - 81 - 3.2.7 X-Ray Photoelectron Spectroscopy ...... - 82 - 3.2.8 Contact Angle Measurements ...... - 82 - 3.2.9 Bacterial Adhesion Analysis ...... - 82 - 3.2.10 Statistical Analysis of Data ...... - 83 - ix

3.3 Results ...... - 83 - 3.3.1 Synthesis of DHP Derivatives (DHP acids 1-4 and p-acid DHP) ...... - 83 - 3.3.2 Quorum Sensing Inhibitory Activity of Free DHPs ...... - 86 - 3.3.3 XPS Characterization of Glass and FEP Surfaces ...... - 87 - 3.3.4 Contact Angle Measurements ...... - 92 - 3.3.5 Antibacterial Activity ...... - 93 - 3.4 Discussion ...... - 98 - 3.5 Conclusion ...... - 102 -

CHAPTER-4: Antibacterial Activity of DHP Without Exocyclic Double Bond 4.1 Introduction ...... - 103 - 4.2 Materials and Methods ...... - 106 - 4.2.1 General ...... - 106 - 4.2.2 Synthesis of Acid- and Amine-DHP Derivatives ...... - 107 - 4.2.3 Quorum Sensing Inhibition Assay ...... - 112 - 4.2.4 Attachment of 3-Aminopropyltriethoxysilane (APTS) ...... - 112 - 4.2.5 Attachment of Succinic Anhydride (SA)...... - 113 - 4.2.6 Attachment of DHP via EDC/NHS Coupling...... - 113 - 4.2.7 X-ray Photoelectron Spectroscopy (XPS) ...... - 114 - 4.2.8 Contact Angle Measurements ...... - 114 - 4.2.9 Bacterial Adhesion Analysis ...... - 114 - 4.2.10 Statistical Analysis of Data ...... - 115 - 4.3 Results ...... - 115 - 4.3.1 Synthesis of DHP Derivatives (DHP phenyl acid 1-4 and DHP phenyl amine 1-2) ...... - 115 - 4.3.2 Quorum Sensing Inhibition Activity of Free DHPs ...... - 120 - 4.3.3 XPS Characterization ...... - 121 - 4.3.4 Contact Angle Measurements ...... - 124 - 4.3.5 Antibacterial Activity ...... - 125 - 4.4 Discussion ...... - 130 - 4.5 Conclusion ...... - 132 -

CHAPTER-5: Dual-Action Antibacterial Surfaces Based on DHP and Nitric Oxide 5.1 Introduction ...... - 133 - 5.2 Materials and Methods ...... - 138 - 5.2.1 General ...... - 138 - x

5.2.2 Synthesis of Acrylate-Functionalized DHP Derivatives ...... - 139 - 5.2.3 Acrylic Acid Plasma Treatment of FEP ...... - 140 - 5.2.4 Attachment of Spermine Linker by EDC Coupling ...... - 141 - 5.2.5 Attachment of DHP via Michael Addition Reaction ...... - 142 - 5.2.6 Attachment of N-diazeniumdiolate (NONOate) ...... - 142 - 5.2.7 X-ray Photoelectron Spectroscopy (XPS) ...... - 143 - 5.2.8 Contact Angle Measurements ...... - 143 - 5.2.9 Determination of Nitric Oxide Release by Griess Assay ...... - 143 - 5.2.10 Bacterial Adhesion Analysis ...... - 144 - 5.2.11 Statistical Analysis of Data ...... - 145 - 5.3 Results ...... - 145 - 5.3.1 Synthesis of DHP-Acrylate Derivatives (DHP 1-2) ...... - 145 - 5.3.2 High-Resolution XPS characterization ...... - 147 - 5.3.3 Contact Angle Measurements ...... - 152 - 5.3.4 Nitric Oxide Release Study by Griess Assay ...... - 153 - 5.3.5 Antibacterial Activity ...... - 155 - 5.4 Discussion ...... - 162 - 5.5 Conclusion ...... - 165 -

CHAPTER-6: Dual-Action Antibacterial Polymers Based on DHP and Nitric Oxide 6.1 Introduction ...... - 166 - 6.2 Materials and Methods ...... - 171 - 6.2.1 General ...... - 171 - 6.2.2 Synthesis of POEGA Macro-RAFT Agent ...... - 171 - 6.2.3 Synthesis of POEGA-b-PVBC Block Copolymer ...... - 172 - 6.2.4 Conjugation of POEGA-b-PVBC to Spermine (p-sper) ...... - 172 - 6.2.5 Attachment of DHP to Spermine-Conjugated Polymer (p-DHP) ...... - 173 - 6.2.6 Attachment of NONOate to DHP-Conjugated Polymer (p-DHP+NO) .. - 173 - 6.2.7 Determination of Nitric Oxide Release by Griess Assay ...... - 174 - 6.2.8 Characterization by 1H NMR Spectroscopy ...... - 174 - 6.2.9 Size Exclusion Chromatography (SEC) ...... - 176 - 6.2.10 Attenuated Total Reflectance-Fourier Transform Infrared Spectroscopy (ATR-FTIR) ...... - 176 - 6.2.11 X-Ray Photoelectron Spectroscopy (XPS) ...... - 176 - 6.2.12 Biofilm Inhibition Assay ...... - 177 - 6.2.13 Confocal Microscopy Analysis ...... - 178 -

6.2.14 Statistical analysis ...... - 178 - 6.3 Results ...... - 178 - 6.3.1 Synthesis and Characterization of Polymers ...... - 178 - 6.3.2 High-Resolution XPS Analysis of Polymers ...... - 183 - 6.3.3 P. aeruginosa Biofilm Inhibition ...... - 186 - 6.3.4 Confocal Microscopy Analysis...... - 190 - 6.4 Discussion ...... - 193 - 6.5 Conclusion ...... - 196 -

CHAPTER-7: Summary and Future Perspectives 7.1 Summary ...... - 197 - 7.2 Future Work ...... - 199 -

REFERENCES ...... - 201 -

APPENDIX: Journal Publication ...... - 242 -

xii

LIST OF ABBREVIATIONS

ABA 4-Azidobenzoic acid AcOH Acetic acid AHL N-acyl homoserine lactone AI Autoinducer AIBN 2,2′-Azobisisobutyronitrile AIP Autoinducing peptides AMP Antimicrobial peptides ANOVA Analysis of variance APTS 3-Aminopropyltriethoxysilane ATR-FTIR Attenuated total reflectance-Fourier transform infrared spectroscopy Boc tert-butyloxycarbonyl c-di-GMP Cyclic di-guanosine monophosphate

CDCl3 Deuterated chloroform CFU Colony forming unit CLSM Confocal laser scanning microscopy CVD Chemical vapour deposition DCM Dichloromethane DHP Dihydropyrrolone or 1,5-dihydropyrrol-2-one DMAP N,N-dimethylaminopyridine DMSO Dimethyl sulfoxide DPD 4,5-Dihydroxy-2,3 pentanedione

Et2O Diethyl ether EDC 1-Ethyl-3-(3-dimethyl aminopropyl)carbodiimide EPS Extracellular polysaccharide EtOH Ethanol FEP Fluorinated ethylene propylene FU Furanone GFP Green fluorescence protein HAI Hospital-acquired infections

xiii

HRMS High resolution mass spectroscopy J Coupling constant KBr Potassium bromide MeOH Methanol

Mn Average molecular weight M.p. Melting point MRSA Methicillin-resistant Staphylococcus aureus

NaBH(OAc)3 Sodium triacetoxyborohydride NaCl Sodium chloride NaOH Sodium hydroxide

Na2SO4 Sodium sulphate NHS N-Hydroxysuccinimide nmol nanomole NMR Nuclear magnetic resonance NO Nitric oxide - NO2 Nitrite - NO3 Nitrate NONOate N-diazeniumdiolate OD Optical density OEGA oligo(ethylene glycol) methyl ether acrylate p-acid DHP Carboxylic acid at para position of DHP phenyl ring p-DHP DHP-conjugated polymer p-sper Spermine-conjugated polymer PA01 Pseudomonas aeruginosa PA01 PBS Phosphate buffered saline ppm Parts per million QS Quorum sensing RAFT Reversible addition fragmentation chain transfer rpm Revolutions per minute SA Succinic anhydride SA38 Staphylococcus aureus SA38 SEC Size exclusion chromatography TFA Trifluoroacetic acid

xiv

THEO Theoretical THF Tetrahydrofuran TLC Thin liquid chromatography TSB Tryptic soy broth or tryptone soya broth UV Ultraviolet VBC 4-vinylbenzyl chloride XPS X-ray photoelectron spectroscopy µM Micromolar

PUBLICATIONS AND PRESENTATIONS

A part of this research has been submitted for publication as well as presented at the following conferences:

Patent

N. Kumar, M.D.P. Willcox, K.K.K. Ho, A. Taunk, Biofilm-resistant coatings and surfaces. Australian Provisional Patent, June 2017, Application number 2017902462.

Publications

A. Taunk, K.K.K. Ho, G. Iskander, M.D.P. Willcox, N. Kumar, Surface immobilization of antibacterial quorum sensing inhibitors by photochemical activation, Journal of

Biotechnology and Biomaterials 6 (2016) 1000238. doi:10.4172/2155-952X.1000238.

R. Kuppusamy, M. Yasir, T. Berry, C.G. Cranfield, E. Yee, O. Kimyon, A. Taunk,

K.K.K. Ho, B. Cornell, M. Manefield, M.D.P. Willcox, D. Black, N. Kumar, Design and synthesis of short amphiphilic cationic peptidomimetics based on biphenyl backbone as antibacterial agents, European Journal of Medicinal Chemistry 143 (2018)

1702-1722.

A. Taunk, M.D.P. Willcox, N. Kumar, Dual-action biomaterial surface coatings based on dihydropyrrolones and nitric oxide donors to prevent bacterial infections. (In preparation)

B. Almohaywi, A. Taunk, S. Nizalapur, N.N. Biswas, G. Iskander, G. Renate, S.A.

Rice, D. Black, N. Kumar, Design and synthesis of lactams derived from mucochloric

xvi and mucobromic acids as quorum sensing inhibitors of Pseudomonas aeruginosa. (In preparation)

Oral Presentation

A. Taunk, K.K.K. Ho, G. Iskander, M.D.P. Willcox, N. Kumar, Antibacterial biomaterials based on quorum sensing inhibitors. 5th International Symposium of

Surface and Interface of Biomaterials (ISSIB) held in conjunction with the 24th Annual

Conference of the Australasian Society for Biomaterials and Tissue Engineering

(ASBTE), Sydney, Australia, April 2015.

Poster Presentations

A. Taunk, K.K.K. Ho, G. Iskander, M.D.P. Willcox, N. Kumar, Developing antibacterial biomaterials based on quorum sensing inhibitors. Royal Australian

Chemical Institute (RACI) NSW Medicinal Chemistry and Chemical Biology Division

One Day Symposium, School of Chemistry, University of Sydney, Sydney, Australia,

September 2015.

A. Taunk, M.D.P. Willcox, N. Kumar, Immobilizing quorum sensing inhibitors on biomaterial surfaces. 5th Annual Medicinal Chemistry/Drug Discovery Symposium in conjunction with ASCEPT Drug Discovery Special Interest Group, School of

Chemistry, University of New South Wales, Sydney, Australia, November 2015.

A. Taunk, G. Iskander, K.K.K. Ho, M.D.P. Willcox, N. Kumar, Immobilization of quorum sensing inhibitors to develop antibacterial biomaterials. Frontiers in

Bioengineering and Biotechnology, Conference abstract: 10th World Biomaterials

Congress, Montreal, Canada, May 2016. doi: 10.3389/conf.FBIOE.2016.01.02320

xvii

A. Taunk, E. Wong, C. Boyer, M.D.P. Willcox, N. Kumar, Novel dual-action antibacterial biomaterials and polymers based on quorum sensing inhibitors and nitric oxide donors. Solutions for Drug-Resistant Infections (SDRI), Brisbane, Australia, April

2017.

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CHAPTER ONE

Introduction

CHAPTER-1

CHAPTER-1 Introduction

1.1 Bacterial Infections

Many microorganisms such as viruses, bacteria, protozoans, fungi and mycobacteria, cause infections. According to the World Health Organization (WHO), infectious diseases are the second leading cause of death and are responsible for approximately 15 million deaths every year worldwide [1]. Bacteria are responsible for about 90 % of all infections compared to infections of viruses, fungi and protozoa which are less frequent than the bacterial infections [2]. Hospital-acquired infections (HAIs) or nosocomial infections are the frequent causes of bacterial infections which usually begin 48 to 72 hours after hospitalization [3]. About 75 % of these infections are mainly prevalent in developing countries [4]. It has been estimated that 40 % hospitalizations in Asia, Latin

America, sub-Saharan Africa and 5–10 % hospitalizations in North America and Europe result in nosocomial infections [5–7]. This accounts for approximately 1.7 million cases of HAIs and more than 100,000 deaths in the United States alone each year [8,9].

Whereas, in Australia around 200,000 HAIs occur in healthcare facilities annually [10].

The most frequent type of infections in hospitals are related to urinary tract, surgical site, then bloodstream and pneumonia [11–15].

Common nosocomial bacterial pathogens include Streptococcus species, Pseudomonas aeruginosa, Staphylococcus aureus, Bacillus cereus, Escherichia coli and many more

[16–19]. Out of these, S. aureus and P. aeruginosa are considered to be the most virulent pathogens causing HAIs. S. aureus is a Gram-positive cocci which is the second leading cause of nosocomial-onset causing clinical disease in 2 % of all - 1 -

CHAPTER-1

hospitalized patients [20,21]. About 30 % people are intermittent carriers of S. aureus and 20 % carry the bacteria for longer time thereby increasing the risk of developing severe infections [22,23]. Once the bacteria contaminates the tissues through a cut in the skin, surgery or medical implant, it can lead to a range of diseases from minor skin infections to life-threatening endocarditis, pneumonia and sepsis [24,25].

On the other hand, P. aeruginosa is the most common Gram-negative bacteria found in one tenth of all life-threatening infections and is responsible for 11 % of nosocomial infections [23,26]. In hospitals it can be found in moist areas such as sinks, toilets, floor mop, medical equipments, even disinfectants and food [27]. It produces a wide range of virulence factors which can cause various chronic infections such as surgical and wound infections, urinary tract infections, ventilator associated pneumonia and cystic fibrosis particularly in immune compromised patients [28–31].

1.2 Resistance to Antibiotics

The discovery of antibacterial drugs or antibiotics has been one of the most significant health-related events in human history in reducing the threat of infectious diseases and saving countless lives. After the discovery of first antibiotic penicillin, Sir Alexander

Fleming had warned of the threat of bacteria developing resistance due to the misuse and over dependence on antibiotics [32]. The extensive and indiscriminate use of antibiotics for past 70 years has led to rapid development of antibacterial resistance

(Figure 1.1).

- 2 -

CHAPTER-1

Figure 1.1: Timeline of antibiotic discovery and concomitant development of antibiotic

resistance. [33].

Antibiotics can be divided into two classes depending on their mechanism of action, bactericidal, that kill bacterial cells or bacteriostatic, that inhibit bacterial cell growth

[33,34]. Bactericidal antibiotics inhibit various bacterial functions such as DNA replication, RNA transcription, cell wall synthesis and protein translation [33–35]. This process exerts selective pressure on the bacteria to develop resistance in order to survive

[36]. Due to this, most bacterial strains have acquired resistance to at least one antibiotic and 70 % of nosocomial infections are resistant to one or more antibiotics [33].

According to estimates from the Centers for Disease Control and Prevention, in the US, almost 2 million people are infected with bacteria that are resistant to antibiotics, and at least 23,000 people die as a direct result of these infections each year [37]. Also, recent reports suggest that by 2050 drug-resistant infections will be the leading cause of deaths in the world with the death rate going up to 10 million each year (Figure 1.2) [38,39].

- 3 -

CHAPTER-1

Figure 1.2: Estimated number of deaths due to antimicrobial resistance (AMR)

compared to other causes of death [38,39].

The first microbe to develop resistance to penicillin was S. aureus which had developed the ability to produce penicillinase, an enzyme that was capable of hydrolysing the β- lactam ring in penicillin [40–42]. In order to combat penicillin-resistant S. aureus, a new drug methicillin was introduced in 1959, however, within a year S. aureus developed resistance to methicillin as well [43]. Over the years methicillin-resistant S. aureus (MRSA) has emerged as a major nosocomial infection with enhanced virulence factors [44–46]. Currently 90–95 % strains of S. aureus are resistant to penicillin and

40–60 % are methicillin-resistant [17,24]. There are almost 100,000 cases of infections due to MRSA and 19,000 MRSA related deaths in US each year, which is much higher than the deaths from HIV/AIDS [47].

Similarly, P. aeruginosa has also developed resistance toward multiple antibacterial drugs which include β-lactams, aminoglycosides, quinolones and carbapenems [48–51].

- 4 -

CHAPTER-1

Resistance of P. aeruginosa towards the highly potent anti-pseudomonal drug,

Ceftazidime, is increasing at an alarming rate [52,53]. According to a study, resistance was easily developed in around 10 % patients during a course of antibiotic therapy [54].

Therefore, due to high resistance to all tested drugs there are very limited options remaining for mono-therapeutic treatment of P. aeruginosa [55].

1.3 Biomedical Device-Related Infections

Biomedical devices or biomaterials are any matter that interacts with the biological system and is compatible with the biological system so it can be used to replace any non-functional body part and/or organ. In the recent years, the insertion of indwelling or implanted foreign polymer bodies such as cardiovascular devices [56], dental [57,58], orthopaedic [59], cochlear implants [60,61], catheters [62–64], contact lenses [65] and many more have become an indispensable part of modern medical care. They are basically life saving devices that are responsible for significantly improving the quality of life and also increasing the life expectancy of patients [66]. However, the insertion or implantation of medical devices has been associated with a risk of bacterial and fungal infections [67,68].

The contamination of the medical device most likely occurs during implantation via the inoculation of a few microorganisms from the patient’s skin or mucous membranes during implantation [69]. Alternatively, the pathogens may also be acquired from the hands of the surgical or clinical staff [70,71]. Infections of all medical devices are considered life-threatening, with prosthetic valve endocarditis exhibiting the highest risk of mortality (>25 %) [72]. Although the use of aseptic surgical techniques has reduced the level of bacteria in hospitals, microorganisms are still found at the site of

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about 90 % implants [73,74]. Critically, infection of the implanted medical devices comprises of 60–70 % of all HAIs [66,75].

As we proceed further into the 21st century, the emergence of bacterial resistance and the lack of new antibacterial drugs have made device-related infections increasingly more difficult to treat. The ability of bacteria to adhere to materials and form biofilms is an important feature of pathogenicity of the bacteria involved in these infections. The formation of multi-layered and mature bacterial biofilms that are resistant to antibiotics and the host immune system, lead to chronic infections or implant failure, or both [76].

The only remedy, if possible, is to remove the infected medical device which causes a lot of pain and discomfort to the patients along with tremendous increase in medical costs, longer hospital stay, mortality and morbidity [68,77,78]. It was estimated that the economic cost of implant infections in the US in the year 2012 alone was almost $27 billion [79]. Therefore, device-related infections are a major concern facing the medical community today and the key strategy to reduce the incidence of device-related bacterial infections is eradication or inhibition of biofilm formation.

1.4 Bacterial Biofilm

Throughout history, microbiologists believed that microorganisms exist in planktonic form as free flowing cells. The biofilm phenomenon was first discovered in 1674 by

Anton van Leeuwenhoek from dental plaque when he first observed microorganisms attach and grow on tooth surfaces [80,81]. Adherent bacteria can accumulate at surfaces and then grow and divide in a relatively sheltered environment to form a slimy, slippery coat called biofilm. Advanced microscopic techniques led to studies that revealed bacterial biofilms are highly-structured, sessile community of cells held together by a self-produced polymeric matrix and are adherent to inert or living surfaces (Figure 1.3)

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[82–86]. Biofilms can offer considerable advantages to microorganisms, including the facilitation of horizontal transfer of genetic material, the sharing of metabolic by- products, and improving tolerance to environmental challenges, such as antibiotics and the host immune system [81,87,88].

Figure 1.3: Structure of microbial biofilm in dental plaque [89]

A worrying feature of the biofilm-based bacteria is that they can withstand host immune responses and biofilms that are more than seven days old are 500 to 5000 times more resistant to antibiotic treatment compared to their planktonic counterparts [81,90–92].As a result, diseases involving biofilms are generally chronic and extremely difficult to treat. Several mechanisms have been proposed for the increased drug resistance of biofilms [93], which includes the following:

(a) Reduced penetration of the antibiotics through the extracellular matrix. The

matrix acts as a barrier protecting the bacteria inside the biofilm from any

changes in its surrounding by limiting diffusion of antibacterial agents. This

ultimately reduces the exposure of bacterial cells to the antibiotics, and also

provides sufficient time for the cells to prepare antibiotic-degrading enzymes

[81,94].

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(b) Cellular diversity exists throughout the biofilm due to different levels of

nutrients in different areas within a biofilm. Experimental evidences indicate

that the antibiotics do not affect the slow or non-growing cells present in the

nutrient deprived layers deep inside the biofilm since the drugs tend to target the

metabolically active cells [80,95,96].

biofilm, which produces a small fraction of persister cells that are responsible

for reduced susceptibility of biofilms to antibiotics [81,95].

The mode of growth of microbial biofilm and its complexity varies according to different surface protein profiles expressed by different types of bacteria. Despite this, the development cycle of microbial biofilms is the same for both Gram-positive and

Gram-negative bacteria [97]. The biofilm formation cycle involves the following stages: reversible attachment, irreversible attachment, colonization, biofilm maturation, and cell death and dispersion [98–100].

In the initial stage of biofilm development, free-swimming planktonic bacterial cells reversibly adhere to the surface due to weak physical forces such as van der Waals attraction, gravitational force, Brownian motion, electrostatic interactions, hydrogen bonding (Figure 1.4) [83,101–104]. Under favourable conditions, the cells express genes responsible for production of extracellular polysaccharide (EPS) matrix, which plays a role in irreversibly anchoring the cells by providing adhesiveness to the surface and cohesiveness between cells [105,106]. This leads to multiplication of bacteria and formation of microcolonies on the entire surface, which represents the building blocks of the biofilm. The microcolonies also entrap other planktonic bacteria from the surrounding extracellular matrix and form sophisticated bacterial communities.

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Figure 1.4: The biofilm formation cycle [107].

Confocal microscopy has revealed that the matrix of EPS, DNA and proteins together form an elaborate three-dimensional pillar- and mushroom-shaped mature biofilms

[105,108,109]. Depending on the species involved, the multi-layered biofilms mainly consist of microcolonies of sessile bacterial cells (10–15 % by volume) and 85–90 % polymeric matrix material produced by the bacteria [110–112]. As a result, at this stage biofilms show maximum tolerance to antibacterial therapy. The biofilm also aids in providing nutrients and oxygen to the entire community, prevents desiccation and facilitates removal of waste products through the highly developed water-filled channels present within the biofilm [113,114]. The last stage involves cell death and dispersion where after biofilm maturation some single cells are detached due to bacteriophage activity within the biofilm [115]. The dispersed bacterial cells return to their planktonic mode and then spread to another surface location to form new biofilm and the cycle continues.

It has been estimated that 80 % of clinical infections are related to bacterial colonization on surfaces [94]. Apart from clinical equipments and medical implants used in hospitals for treatment of patients, chronic infections are also increasingly common in humans.

Some examples include native valve endocarditis (biofilms on surfaces of the heart and

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heart valves) [110], bacterial prostatitis (biofilms on the surface of prostate gland) [116], cystic fibrosis (biofilms on surfaces of the lungs) [117], otitis media (biofilms on surfaces of the ear) [85,118] and periodontal diseases [119].

Over the past few years bacterial attachment have been problematic and have also caused significant economic damage in many industrial systems such as maritime [120], water storage and distribution systems (including filtration membranes) [121], paper manufacturing systems [122], pipe and rig corrosion [123], oil spoilage [124], heat exchange systems and cooling towers [125]. Biofilms are not only a big threat in the medical and industrial settings but also cause serious problems as food contaminants in the food processing environment [110,126,127].

1.5 Quorum Sensing

For many centuries, scientists believed that bacteria function as individual organisms and multiply under favourable conditions. It was discovered that bacteria could also communicate and convey their presence to one another like other creatures and form communities or biofilms. Nealson et al. described this complex process of bacterial communication in the early 1970s in the bioluminescent marine bacteria, Vibrio fischeri and Vibrio harveyi, which produces light when bacterial cell numbers were high

[128,129]. Further investigation revealed that this process was controlled by small diffusible signaling molecules called autoinducers in a cell density-dependent manner

[128,130]. When the concentration of autoinducers reaches a certain critical number, the accumulated molecules interact with a receptor within the bacterial cell and induce expression of genes required for bioluminescence [131]. This threshold density of cells was termed as ‘quorum’ and the phenomenon was later coined as ‘quorum sensing’

[132]. Therefore, quorum sensing (QS) is a bacterial cell-to-cell communication

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mechanism through which bacteria regulates the expression of genes in response to cell population density [133–136].

QS plays a crucial role in the development of mature biofilms of common bacteria which includes adhesion of bacterial cells to a surface and formation of the EPS matrix

[80,136,137]. For example, P. aeruginosa uses QS to aid the formation of biofilms, which leads to cystic fibrosis in the human lungs and enables bacteria to resist antimicrobial agents and the innate host immune system [117]. In many cases, QS signals contribute directly to pathogenesis through the production of virulence, such as toxins, elastase, hydrogen cyanide, procyanin protease, and other immune-evasive factors [138]. Additionally, myriad of bacterial processes are regulated by QS such as bioluminescence, sporulation, motility, antibiotic production, secondary metabolite production, and many more (Figure 1.5) [139–143].

Estimation of cell numbers/population density Release of bacteria from biofilm

Virulence expression Oxidative stress tolerance

Antibiotic resistance Metabolism

Biofilm formation DNA transfer Quorum Bioluminescence Sensing Sporulation

Nitrogen fixation Bacterial growth

Swarming & motility Production of pigments

Evasion of host defence Survival in hostile conditions

Entry into stationary phase Antibiotic synthesis by antibiotic producers

Figure 1.5: Multiple functions controlled by QS in bacterial communities [144].

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1.6 Quorum Sensing Pathways

QS mechanism depends on the interaction of small, diffusible signalling molecules or autoinducers with a sensor or transcriptional activator to initiate gene expression for coordinated activities. There are three major classes of QS systems that have been studied so far in different bacterial species.

1.6.1 Quorum Sensing in Gram-Negative Bacteria

The QS system of Gram-negative bacteria utilizes the autoinducer-1 (AI-1) type signalling molecules. It is the most widely studied QS pathway that is mediated by signals having a common structural skeleton of N-acyl homoserine lactones (AHLs) and are present in over 70 species of Gram-negative bacteria, most of them are typical human pathogens such as P. aeruginosa, V. fischeri, E. coli, Agrobacterium tumefaciens and Erwinia carotovora [145].

The general mechanism of AHL-based QS pathway involves generation of the AHL signals by the LuxI-type protein in the cell. These signals freely exit across the cell membrane either by diffusion for short chain AHLs or active transport for long chain

AHLs [146]. Once the concentration of the accumulated AHLs reaches a certain threshold in the extracellular environment, the signals bind to its cognate cytoplasmic

LuxR-type receptor protein to form an active dimer. The LuxR-AHL dimer complex interacts with the target promoter sequences which activates transcription of specific set of QS genes (Figure 1.6) [147–149].

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Figure 1.6: Schematic diagram of Gram-negative bacteria QS circuit (LuxI/LuxR

system). Red pentagons denote AHL autoinducers [143].

AHL autoinducer molecules are unique and detected only by the species that produces it. Natural AHLs produced by different bacterial species bear similar structural elements with the same lactone head group but diverse acylated tail chain. The acylated side chain can be of varying length (four to eighteen carbons), saturated or unsaturated, unsubstituted or 3-oxo, 3-hydroxy substituted and may even contain aromatic moieties.

These subtle differences in the signals are important for its specificity (Figure 1.7)

[150–152].

Figure 1.7: Examples of natural autoinducers used by Gram-negative bacteria.

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The QS network of P. aeruginosa consists of two distinct LuxI/LuxR circuits termed

LasI/LasR and RhlI/RhlR, which are named after their influence on the production of elastase and rhamnolipid respectively [153].The homologue proteins LasI and RhlI respectively synthesize two pairs of AHLs, N-(3-oxododecanoyl) homoserine lactone

(3OC12-HSL) and N-butyryl-homoserine lactone (C4-HSL) [154]. LasR-autoinducer complex activates a number of target genes including the expression of RhlI genes, therefore, hierarchally the Rhl system is under the control of las system (Figure 1.8).

The two circuits together stimulate expression of multiple virulence traits such as biofilm formation, motility and toxin production. Additionally, according to a report, P. aeruginosa las and rhl mutant strains that are not able to communicate via QS signals form abnormal biofilms with defective EPS matrix resulting in increased susceptibility to antibiotics [109]. Similarly, in another study, QS mutants of P. aeruginosa showed significantly reduced virulence and chronic infections in animal models [155–157].

These results clearly indicate that the AHL-mediated QS is indeed responsible for P. aeruginosa diseases.

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Figure 1.8: Las and Rhl QS pathways in P. aeruginosa. The two systems function

sequentially and Las-controlled genes are induced before Rhl-activated genes [158].

1.6.2 Quorum Sensing in Gram-Positive Bacteria

In contrast to Gram-negative bacteria, the cell-communication pathway in Gram- positive bacteria such as S. aureus, Enterococcus faecalis, Streptococcus pneumonia, is based on the production and detection of modified oligopeptides known as autoinducing peptides (AIPs) [142]. Majority of AIPs are mainly post-translational oligopeptides, including linear and modified cyclic peptides with diverse structures (Figure 1.9)

[143,159].

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Figure 1.9: AIPs produced by Gram-positive bacteria. The underlined tryptophan in

Bacillus subtilis ComX is isoprenylated [143].

The AIP signal molecules are synthesized as precursor peptides in the cytoplasm, which are then structurally modified, cleaved, and secreted out of the cell. After accumulating at high cell density, the AIP signal binds to its cognate membrane bound two- component histidine-kinase receptor. The complex activates signal transduction via a conserved phosphorylation/dephosphorylation mechanism and passes the sensory information to a cognate cytoplasmic response regulator (Figure 1.10) [147,160]. The phosphorylated response regulators are active and they function as DNA-binding transcription factors to modulate expression of target genes [143,161].

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Figure 1.10: QS pathway in Gram-positive bacteria. Blue octagons denote

processed/modified AIPs. The QS process is regulated by sensor histidine kinase (H),

phosphate (P) and response regulator (D) respectively [143].

1.6.3 Quorum Sensing in Gram-Negative and Gram-Positive Bacteria

The third class of QS system called the autoinducer-2 (AI-2) is shared by both Gram- negative and Gram-positive bacteria and is known to facilitate communication between bacteria of different species to form mixed-species biofilms [127,145]. Bassler et al. first identified this hybrid system in AHL-deficient strain of bioluminescent marine bacterium, V. harveyi, which despite the absence of the natural AHL signalling molecules was able to produce bioluminescence [162]. This implied the presence of a second QS pathway which employs furanosyl borate diester as an autoinducer, a highly unusual structure with no resemblance to other autoinducers (Figure 1.11) [88,163].

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Figure 1.11: Autoinducers for AI-2 mediated QS pathway.

The signals for QS may not be identical but are mainly produced from a common precursor, 4,5-dihydroxy-2,3-pentanedione (DPD), found in more than 70 bacterial species. The DPD molecule, a product of the LuxS gene, is highly unstable and therefore undergoes spontaneous rearrangements and modifications to form a group of cyclic compounds termed as AI-2 signalling molecules [138,164]. AI-2 in conjunction with an AHL or oligopeptide autoinducer regulates biofilm formation and virulence in

V. harveyi, E. coli, Streptococcus mutans, Streptococcus anginosus and other clinically relevant pathogens (Figure 1.12) [148,165,166]. One example of interspecies communication via AI-2 system is the cooperative biofilm growth of two different bacteria in oral plaque, Actinomyces naeslundii and Streptococcus oralis, thus indicating that AI-2 is a universal communication language between different species

[165].

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Figure 1.12: Two QS pathways in V. harveyi employing AI-1 and AI-2 as signalling

molecules. Red pentagons and orange triangles denote AI-1 or AHLs and AI-2

respectively. [161].

1.7 Quorum Sensing Inhibition

QS signalling system is important for bacterial growth, adhesion and biofilm formation resulting in numerous infectious diseases. Coventional antibiotics used for treatement of bacterial infections have lead to development of resistance. An attractive strategy to prevent production of biofilm and virulence factors is blocking or interefering with the bacterial cell-to-cell communication. Therefore, inhibition of QS have led to development of novel class of antimicrobial agents that are less prone to develop bacterial resistance due to its non-growth inhibition mechanism which does not cause survival pressure on bacteria [135].

It has been proposed that an ideal and effective QS inhibitor should meet a few criteria

[90,167]. It should (i) be a small molecule that can reduce QS-regulated gene expression, (ii) be highly specific for the QS-regulator with no toxic effects to the

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bacterial cell or host, (iii) be chemically stable and resistant to host metabolism, (iv) not interfere with bacterial cell metabolic process, and (v) reside in the host for longer time for its effective action.

1.7.1 Natural Furanones

The first example of QS inhibiting agent came from the marine environment. A red marine alga, Delisea pulchra, from the South-East coast of Australia produces a range of halogenated furanone compounds (Figure 1.13A). These compounds are secreted from the vesicles of the alga located near its surface to prevent it from fouling by bacteria and other marine organisms such as epiphytes [168]. In 1993, de Nys et al. isolated around twenty four halogenated furanones from D. pulchra with variable levels of QS inhibitory activity [169,170]. These halogenated furanone derivatives, termed fimbrolides, consist in general a common 4-halo-3-butyl-5-halomethylene-2(5H)- furanone skeleton, with structural variations only in the number and nature of halogen atoms present at C-6 position of the lactone ring and the presence or absence of oxygen containing groups such as acetate or hydroxyl in the C-4 butyl side chain (Figure

1.13B) [167,169,171].

(A) (B)

Figure 1.13: (A) Australian red seaweed, Delisea pulchra; (B) Halogenated furanones

isolated from D. pulchra.

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Natural fimbrolides possess potent antimicrobial effects and are able to influence bacterial behaviour by interfering with the bacterial QS system. Samples of natural furanones have shown to alter AHL-dependent processes such as swarming motility in

Serratia liquefaciens and virulence factors and bioluminescence produced by V. fischeri and V. harveyi [172–174]. Furanones have demonstrated to interfere not only with AHL-based QS pathway but also with the AI-2 bacterial system of V. harveyi, B. subtilis and oral Streptococci [175–177]. In particular, the natural compound, (5Z)-4- bromo-5-(bromomethylene)-3-butyl-2(5H)-furanone, has shown to inhibit AI-2 dependent QS activities, swarming motility and biofilm formation in E. coli without affecting their growth [178].

The exact mechanism of action of QS inhibition by furanones is not completely known, therefore, it was hypothesized that since furanones are structural mimics of AHL

(lactone ring structure) and AI-2 signals (tetrahydrofuran rings), they act as fake signalling molecules to competitively bind to the binding site of the receptor by displacing the cognate autoinducers. The binding of furanones to the LuxR receptor protein of AHL-QS pathway causes conformational changes that enlist the furanone-

LuxR complex into rapid proteolytic degradation (Figure 1.14) [179]. It was postulated by that furanone blocks QS in V. harveyi by impairing the ability of the LuxR to bind

DNA and initiate transcription [180]. Work by Zhang et al. showed that furanone covalently modifies LuxS, the enzyme which produces AI-2 signals, in a highly sequence selective manner and inactivates it, thereby, inhibiting the AI-2 mediated QS mechanism [181].

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Figure 1.14: A schematic representation of AHL-mediated QS system (left) and

disruption of QS system by an antagonist (right) [182].

In addition to the inhibitory effect of furanones towards bacteria, they have also been found to be effective against a range of yeasts as well as fungi [183,184]. In summary, these natural halogenated compounds are capable of inhibiting QS systems of both

Gram-negative and Gram-positive bacterial species and its mode of action as an effective QS inhibitor varies depending on the furanone as well as the bacteria involved.

However, one major disadvantage in the long-term use of naturally occurring furanones

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is that they are mutagenic and toxic to human cells at concentrations required to inhibit the bacteria [185,186]. They were also found to induce toxic effects even at 50 % effective dose in animal model, thus, rendering them unsuitable for treatment of infectious diseases [187].

1.7.2 Synthetic Furanones

The furanones isolated from D. pulchra have displayed excellent broad spectrum QS inhibitory activity. However, most of these compounds are cytotoxic and mutagenic.

Therefore, to overcome these limitations, the fimbrolide scaffold was optimized for the development of improved novel QS inhibitors.

Initially, a series of furanone analogues were synthesized with high degree of structural similarity to natural compounds but with different alkyl chain length and halogen substituiton pattern (Figure 1.15) [188–191]. The inhibitory activity of the synthetic derivatives was investigated and it was revealed that compounds with shorter (two carbon) or no alkyl side chain attached to the lactone ring showed greater antagonist activity than compounds with longer side chain [179,192,193]. The study conducted on these furanones showed that they were able to compete with the AHL signalling molecule and bind to the receptor protein of P. aeruginosa, resulting in inhibition of multi-drug efflux genes and several vireulence factors without affecting the bacterial growth [192,193].

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Figure 1.15: Synthetic furanone analogues derived from natural furanones.

Synthetic compound 4, also known as furanone-30, exhibited enhanced antagonist effect compared to furanone 3 by inhibiting green fluorescent protein (GFP) production at a 10-fold lower dose required by furanone 3. It has also been found that furanone-30 was capable of influencing the expression of approximately 80 % of all QS-controlled genes and significantly increasing the susceptibilty of biofilm to antibacterial drugssuch as tobramycin [193]. Recently, He at al. reported the interference of furanone-30 with

QS-controlled adhesion and biofilm accumulation of S. mutans, which led to formation of thinner and diffused biofilms on tooth surfaces [194]. Moreover, furanone 3 and 4 have also shown to inhibit the AHL-regulated pathogenicity in the lungs of mice infected with P. aeruginosa [193,195].

More recently, bicyclic and thio-incorpotated derivatives of furanones have been synthesized (Figure 1.16). The bicyclic derivatives exhibited enhanced QS inhibitory activity against biofilms of P. aeruginosa and E. coli with no growth inhibition and reduced ctyotoxicity, while the thio-furanones were capable of disrupting already formed B. subtilis biofilm and thereby increasing the efficiency of antibiotics [196,197].

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Figure 1.16: Chemical structures of bicyclic furanones and thio-furanones.

1.7.3 1,5-Dihydropyrrol-2-ones

Investigation of furanone analogues was further extended by varying the lactone structural motif, the core nucleus of furanones, to generate structurally similar compounds with 1,5-dihydropyrrol-2-one ring system. The main difference between the two structures is the replacement of the lactone oxygen atom of the heterocyclic ring with the nitrogen atom to obtain a lactam motif (Figure 1.17). The lactam ring system is hydrolytically more stable than the lactone of the parent furanone making them less susceptible to ring opening reactions (lactonolysis) under physiological conditions

[198,199]. This difference in stability could be possibly due to higher resonance stabilization of the five-membered pyrrol-2-one ring compared to lactone due to greater availability of the nitrogen lone pair of electrons.

Figure 1.17: Substitution of bivalent -O- of lactone with -N- to form lactam.

Due to the excellent biological performance of the brominated furanones, the initial synthesis of the lactam analogues mainly focused on retaining similar essential features and halogen pattern. Our research group synthesized a range of brominated pyrrol-2-one

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analogues with similar bromination pattern present in furanones via a novel and efficient lactone to lactam conversion reaction (Figure 1.18) [200]. The preliminary biological screening showed that the N-substituted dihydropyrrol-2-one derivatives obtained from lactamization of furanones retained their potent QS inhibitory activity against E. coli [201].

Figure 1.18: Dihydropyrrol-2-ones synthesized via lactone to lactam ring conversion

reaction [200,201].

Furthermore, novel derivatives with an aryl substituent at C-4 position were also synthesized and have shown high inhibitory activity against QS-controlled processes, virulence and biofilm formation of P. aeruginosa and S. epidermidis (Figure 1.19)

[202]. In another study, direct conversion of potent furanones to pyrrol-2-one ring

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system was employed for the synthesis of novel lactam analogues with terminal azide and alkyne functionality in the core nucleus. The alkyne derivative 13a was found to inhibit approximately 55 % of GFP produced by a modified reporter strain of P. aeruginosa [203]. Overall the results indicate that 1,5-dihydropyrrol-2-ones have the ability to serve as potential leads for further development of novel QS inhibitors as antimicrobial therapeutics.

Figure 1.19: Synthetic dihydropyrrol-2-one derivatives.

1.8 Nitric Oxide

Nitric oxide [nitrogen monoxide, •NO, IUPAC: oxidonitrogen(•)] is a colourless diatomic, uncharged gas formed during lightning or electrical storms in the troposphere.

NO is a vital signalling molecule in eukaryotes for which it was declared as the

‘molecule of the year’ in 1992 by the Science magazine [204]. NO is synthesized, in mammals, by the enzyme nitric oxide synthase (NOS) from L-arginine. Over the years it has gained recognition for its vital role in regulation of diverse physiological functions in the human body such as inhibition of platelet aggregation, neurotransmission, smooth muscle relaxation and angiogenesis [205–207]. In addition,

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NO also plays multiple key roles in the bacterial life cycle such as regulation of dentrification, iron acquisition and detoxification [208,209].

Research has shown growing evidence that NO and the QS systems are inter-connected and that NO plays a very important role in controlling the QS-dependent activities such as biofilm formation, motility and virulence of different bacterial species. Studies have shown that enzymes involved in the dentrification process within the bacterial cells, are highly expressed in biofilms of P. aeruginosa compared to planktonic cells, and their overexpression is controlled by QS [210–212]. Furthermore, NO derived from the anaerobic growth of P. aeruginosa, controls the QS system and virulence expression.

Recent studies have also revealed that NO participates in the QS pathways of other bacterial systems such as V. harveyi and V. fischeri and is responsible for concentration- dependent increase in bioluminescence, biofilm formation, iron uptake and motility

[213–215]. Thus, QS and NO inter-connection is evident from the above data and NO has emerged as an important factor in many processes that are essential for formation of biofilm.

1.8.1 Nitric oxide as Antimicrobial and Antibiofilm Agent

NO is a ubiquitous biologically active signalling molecule that is endogenously produced in mammals and plays a critical role in mammalian innate immune response against numerous pathogenic microorganisms such as viruses, fungi, bacteria and parasites. Experiments were conducted on mice lacking the NOS gene, that is responsible for biosynthesis of NO, and the mice were found to be highly susceptible to various infectious diseases including malaria and tuberculosis, thereby highlighting the importance of endogenous NO [216,217]. The mechanism of the NO antimicrobial action is not known, but it largely depends on the concentration and dosage of NO. - 28 -

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At high concentrations, NO reacts to form reactive toxic by-products such as peroxynitrite, dinitrogen trioxide and the hydroxyl radical, which are able to induce significant oxidative, deamination and nitrostative stress on various bacterial systems

[218–220]. The reactive intermediates chemically alter the cell membrane and intracellular proteins, DNA, lipid membranes and other macromolecules, resulting in bacterial death [221,222]. Webb et al. found that high levels of peroxynitrite and other reactive species were present in 7-day old mature biofilms of P. aeruginosa, which induce oxidative and/or nitrostative stress resulting in biofilm eradication and cell death

[115]. In another study, a genetically modified strain of P. aeruginosa capable of generating large amount of NO exhibited enhanced biofilm dispersion, whereas another

P. aeruginosa mutant that lacked the ability to produce NO did not disperse biofilm

[223].

Studies have also shown that at low non-toxic concentrations, NO can induce biofilm dispersal of various bacterial species including a common human pathogen, P. aeruginosa, as well as multi-species biofilms and yeast biofilms from water pipes and filtration membranes [224,225]. It was revealed that the mechanism for biofilm dispersal at low NO concentrations involves the secondary messenger cyclic di- guanosine monophosphate (c-di-GMP) [226]. Further, exogenous, sublethal doses of

NO, was able to increase the sensitivity of various biofilms to antimicrobial treatments, by transforming bacteria in resistant biofilm into sensitive planktonic mode [225]. For instance, addition of NO has induced biofilm inhibition and dispersal of various Gram- negative bacteria, such as P. aeruginosa, E. coli and Vibrio cholerae as well as Gram- positive bacterium, S. aureus [227–230].

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Recently, sustained exposure of NO was shown to prevent MRSA biofilm formation in vitro and in vivo on an animal model [231]. Therefore, the results suggest that NO can be used as an effective antimicrobial therapeutic agent to control bacterial biofilm formation and pathogenicity.

1.8.2 Nitric Oxide Donors

One major challenge in employing NO as an effective antibiofilm agent is delivering

NO in a sustained and controlled manner to the biological system. The exogenous delivery of NO is difficult due to limited solubility of NO gas in water, high reactivity and a very short half-life of 0.1–5 seconds [232]. To overcome these problems, investigators have developed a wide range of novel molecules capable of releasing NO under specific conditions. Some of the common NO donating molecules which have been explored include nitrates, nitrites, S-nitrosothiols, N-diazeniumdiolates, N- nitrosoamines and metal-NO complexes, to name a few (Figure 1.20) [233,234].

Figure 1.20: Selected examples of NO donors [234].

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Diazeniumdiolates or diazen-l-ium-l,2-diolates are anionic compounds containing the

[N(O)NO]- functional group that have been useful for the reliable generation of NO.

The two nitrogens and oxygens are arranged as shown in the resonating structures with one nitrogen bonded to another atom or molecule (X) (Figure 1.21) [235]. A range of diazeniumdiolates that have been prepared are bound to carbon, nitrogen, oxygen or sulphur and depending on the atom it is attached to they are classified as C- diazeniumdiolates, N-diazeniumdiolates, O-diazeniumdiolates and S-diazeniumdiolates respectively [235]. They all showed similar structural and spectral properties however, their reactivity was much more diverse [233].

Figure 1.21: Possible resonance structures of diazeniumdiolate.

Of all the NO donors mentioned, N-diazeniumdiolate is of special interest that is capable of releasing significant amount of NO. The first N-bound diazeniumdiolate was the diethylamine-NO adduct prepared in 1960 by Drago and co-workers [236–238]. A wide range of N-diazeniumdiolates are commercially available which were synthesized in general by exposing an appropriate amine, preferably secondary amine, to NO gas under high atmospheric pressure (5 atm) (Figure 1.22) [237,239–241].

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Figure 1.22: Structures of selected N-diazeniumdiolates along with their half-lives in

0.1 M phosphate buffer (pH 7.4) at 37 °C [242].

The NO release from N-diazeniumdiolates is considered to be a pH-dependent pseudo first order reaction. The mechanism generally involves protonation of the amine nitrogen attached to the diazeniumdiolate, followed by release of the parent amine and two molecules of NO (Figure 1.23) [243,244].

Figure 1.23: NO release mechanism by N-diazeniumdiolates.

In contrast to other NO donors, there are two main advantageous features of these compounds that set them apart. First, they release NO at first order rates on dissolution in buffered fluids or cell culture media without any redox activation. Second is that each compound generates two molecules of NO under physiological conditions with half- lives ranging from 1 min to 1 day depending on the structure of the substrate. This - 32 -

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allows researchers to determine the effects of NO at different release rates for a variety of applications [242,245]. For instance, to bring a momentary drop in blood pressure only for a few seconds, a rapid but brief infusion of NO from DEA/NO 17 diazeniumdiolate having half-life of 2 min would be suitable [246]. Similarly, for constant release of NO for many hours or days for treatment of prolonged cytostasis or cerebral vasospasm, a longer NO releasing DETA/NO 20 (20 hour half-life) would be preferable [247–249].

1.8.3 Dual-Action Antibacterial Drugs

The failure of conventional antibiotics to treat biofilms and infections necessitates the development of new antibacterial agents. As mentioned earlier, NO is a promising therapeutic agent for treatment of bacterial biofilm-related infections. Therefore, a combination of existing drugs with NO donors represents an attractive strategy for NO delivery that combines the existing mode of action of drug with the biological activity of NO. This is particularly important in improving the efficacy and reducing ill effects of a drug [182]. For example, hybrids of antibiotic compounds such as cephalosporin, ketaconazole, aspirin and many more, combined with different NO donors have been reported to improve the efficiency of the drugs (Figure 1.24).

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Figure 1.24: NO hybrid antibacterial drugs.

Cephalosporin in conjugation with diazeniumdiolates was able to disperse biofilms of

P. aeruginosa by releasing NO only after cleavage of the β-lactam ring by bacterial enzyme β-lactamase [250]. Antibiotics such as tobramycin and ciprofloxacin that are no longer effective in eradicating biofilms were also combined with NO donors, and the hybrid drugs showed enhanced antibiofilm activity compared to the antibiotic alone

[251]. Similarly, ketaconazole-NO hybrid showed antifungal activity against a broad range of fungal species [252].

Another antibiofilm strategy involves the use of NO donors in combination with a QS inhibiting agent to eradicate biofilm-related infections and virulence expression of bacteria. A range of novel hybrid compounds was developed by combining analogues of furanones with NO donors such as nitrates, diazeniumdiolates and S-nitrosothiols

(Figure 1.25). The hybrid molecules were found to retain their potent QS inhibitory activity with significant improvement in activity than the base furanone alone [253].

Similarly, hybrid compounds based on dihydropyrrolones (DHP) and NO donors also showed potent QS inhibitory activity. Furthermore, the QS signalling molecules - 34 -

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commonly found in Gram-negative bacteria, AHLs, were also conjugated with NO donors such as nitrate, which transformed them into antagonists capable of controlling

QS activities and bacterial virulence expression [182,254].

Figure 1.25: Hybrid QS inhibitors based on NO donors.

In summary, these newly developed QS inhibitors based on NO donating compounds have the potential to treat biofilm-related infections without exerting selective pressure on bacteria and therefore, they are less likely to develop resistance.

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1.9 Current Antibacterial Surface Coatings

Biomaterial-associated bacterial infections are a major complication in the modern healthcare system. Developing antibacterial surfaces by applying an antibacterial material on the surface of the biomaterials is one of the widely used methods. These generally involve either the release of an active compound from the device or the covalent immobilization of an antibacterial compound to the surface. Various strategies to control formation of biofilm on medical devices with different antibacterial agents including antibiotics, silver and antimicrobial peptides have been examined.

1.9.1 Antibiotics

A number of surface coatings based on incorporation of conventional antibiotics have been described in literature. The main advantage is that local delivery of high doses of antibiotics at a specific site can be administered without exceeding the systemic toxicity of the drug [255]. Another important issue is the release kinetics of the drug. The drug at an appropriate dose must act before the EPS matrix is formed around the bacterial colony, for it to act in an effective manner [255].

Coatings based on numerous antibiotics such as minocycline-rifampin, vancomycin, norfloxacin and daptomycin have been reported [256–259]. These antibiotics-based coatings have been successful in preventing biofilm formation on various biomaterials

[260–262]. For example, vancomycin was covalently bound to titanium alloy implants.

The modified titanium surfaces significantly inhibited biofilm formation of

Staphylococcus epidermidis [263]. However, use of certain aminoglycoside antibiotics such as tobramycin have led to enhanced biofilm formation [264] and even the threat of multiple drug resistant strains of bacteria and the varying efficacy of antibiotics against different species of bacteria has limited the use of these coatings.

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1.9.2 Silver

Silver is one of the strongest antibacterial agents. Due to its strong bactericidal activity it has been incorporated onto various biomedical devices such as catheters, contact lenses, vascular graft, surgical sutures and fracture-fixation devices [265,266]. Silver releases silver ions, which disrupts the normal function of bacterial DNA, enzymes and metabolic proteins resulting in cell death. Thus, silver has been found to be effective against a broad spectrum of pathogens, including P. aeruginosa, E. coli, S. aureus, S. epidermidis and Enterococcus species [267–269].

However, medical devices coated with silver or silver ions have not shown promising results in clinical studies. In a clinical trial, central venous catheters impregnated with silver had no significant effect in controlling bacterial colonization, bloodstream infections and ICU mortality [270]. Another study revealed that silver coated catheters lost their antibacterial activity within 28 days [271]. This was probably due to inactivation of silver mediated activity after coming in contact with physiological fluids and also the coating wearing off from the surface [272]. Moreover, silver-resistant bacteria have been reported and also a high dose silver has displayed genotoxic and cytotoxic effects on human cells by accelerating the formation of thrombin and platelet activation leading to high risk of thrombosis [273–275]. All these factors have made silver impregnated coatings undesirable.

1.9.3 Quaternary ammonium compounds

Quaternary ammonium compounds are widely utilised as antibacterial agents. The positively charged quaternary ammonium compounds can be incorporated into biomaterials by copolymerization techniques or covalently binding them to the surface

[276,277]. The surfaces having high positive charge exhibit contact based antibacterial

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activity on negatively charged bacterial cells [278]. Tiller et al. showed that surfaces containing ammonium compounds exert their activity against Gram-negative and Gram- positive bacteria by disrupting the bacterial cell membrane [279]. In a study, Gottenbos et al. developed silicone rubber coatings by covalently grafting quaternary ammonium groups, which showed both in vitro and in vivo antibacterial properties against adhering bacteria, S. aureus, S. epidermidis, E. coli and P. aeruginosa [280]. Polymer derivatives with quaternary ammonium salts were found to be biologically active against various pathogenic bacteria as well as fungus, Trichophyton rubrum [277]. Despite high broad spectrum activity, bacterial resistance towards these compounds and their toxicity to human cells constitute the major disadvantages of this approach [281,282].

1.9.4 Antimicrobial Peptides

Another strategy that has been investigated is the incorporation of antimicrobial peptides (AMPs) on biomaterial surfaces. Several studies in the literature have reported the immobilization of various AMPs such as melimine, ultrashort lipopeptides, melittin, buforin 2 and tritrpticin on different surfaces using diverse approaches [283–286]. The broad spectrum activity of AMPs (against bacteria, fungi and viruses) and their efficacy at extremely low peptide concentrations make them excellent candidates for coating medical devices and implants. However, their reduced activity upon tethering to solid supports can significantly compromise their effectiveness as biomedical coating materials [287]. Furthermore, pH sensitivity, local toxicity, allergy after repeated applications and the cost of synthesis are some the drawbacks of AMP-coated devices

[288]. Another aspect that needs to be considered is the accumulation of dead bacteria on the AMP-coated surfaces. These dead bacteria may promote bacterial accumulation by serving as breeding grounds for other bacteria, thus reducing the efficiency of the coated surfaces. - 38 -

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1.10 Furanone Coatings

The strategies described earlier for developing antibacterial surfaces are associated with several drawbacks such as bacterial resistance towards antibiotics and silver, cytotoxicity of silver and quaternary ammonium compounds and low activity of AMPs after surface attachment. Therefore, to overcome these limitations, an effective strategy involves attachment of furanone compounds on biomaterial surfaces.

Several strategies have been employed to coat furanones on surfaces to inhibit biofilm formation. A furanone derivative was physisorbed on different polymer surfaces commonly used for medical devices (Figure 1.26) [289]. The resulting surfaces showed reduction in bacterial adhesion and slime production of S. epidermidis. However, a gradual loss of the furanone was observed due to its slow release from the biomaterial surface into the surrounding solution.

Figure 1.26: Synthetic furanone physically adsorbed on biomaterial surfaces [289].

To overcome this limitation, two new strategies were developed to covalently bind furanones onto polymer substrates. First, copolymers based on styrene and furanone monomers were synthesized by radical polymerization [290]. The furanone-incorporated disks generated by this approach showed a dose-dependent reduction in S. epidermidis bacterial adhesion. The disks with 5 % and 8.5 % furanone showed reduction in biofilm by 67 % and 89 % respectively (Figure 1.27).

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Figure 1.27: In vitro bacterial load on control and furanone disks [290].

Alternatively, the same furanone derivative was covalently grafted onto the surface of a catheter via an azide/nitrene based approach [290]. The coated surfaces were found to inhibit biofilm formation of S. epidermidis in vitro by 78 % and were also able to control infection up to 65 days in an in vivo sheep model. The excellent activity indicates that even after covalent attachment, the furanones retain their QS inhibitory activity. Moreover, the results also suggested that the QS receptors are located at the bacterial cell membrane. Therefore, diffusion of active compounds into the cell is not required in order to interfere with the bacterial QS. However, the exact mode of action is not completely understood. In another approach, Al-Bataineh et al. successfully immobilized furanones on biomaterial surfaces in the presence of photoreactive azide groups. The covalent grafting of the furanones was demonstrated using various surface analysis techniques, but the antibacterial activity of the coated samples was not reported.

1.11 Dihydropyrrolone Coatings

Another promising candidate for reduction of biofilm formation on medical implants is dihydropyrrolone (DHP) compound. Analogues of DHPs have been successfully grafted

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onto glass and polystyrene substrates by Michael addition reaction. The surfaces were found to reduce the production of biofilm of P. aeruginosa and S. aureus without increased killing for both strains of bacteria [291,292]. The same attachment strategy was employed for incorporation of DHP onto polyacrylamide beads to investigate the in vivo activity. The in vivo results suggest that DHP-coated polymeric substrates were able to reduce the bacterial load and pathogenicity in a subcutaneous S. aureus infection model (Figure 1.28) [293]. The surfaces were found to reduce the production of biofilm of P. aeruginosa and S. aureus in vitro and in vivo.

Blank DHP surface

Figure 1.28: Attachment of DHPs on polymer beads by Michael addition reaction and

in vivo results showing DHP-incorporated beads are effective against Staphylococcal

infections [293].

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Another study focused on investigating the mechanism of action of surface bound

DHPs, which was achieved by covalently grafting DHPs on the surface by click chemistry [294]. The modified surfaces were not only able to reduce the adhesion of P. aeruginosa and S. aureus but also significantly reduced the adhesion of a QS reporter strain, P. aeruginosa MH602. The modified reporter strain produces GFP when the QS system is active. The surface bound DHPs were able to repress the expression of GFP by 72 % without affecting the cell viability, thereby suggesting that DHPs inhibit QS systems by interacting with receptors on the cell membranes of bacteria (Figure 1.29).

Such excellent activity indicates that DHPs retained their activity when covalently bound, and were able to block QS of the reporter even after attachment. This finding was further confirmed when DHP-incorporated polystyrene substrates significantly reduced the number of P. aeruginosa QS reporter bacteria [292]. Overall, the results indicate that any of the attachment strategies employed did not hinder the biological function of DHPs. However, only a few DHP compounds have been explored, while there are many potent DHPs yet to be attached and tested.

Figure 1.29: Reduction in GFP-expressing cells displayed by DHP-coated surface

[294].

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1.12 Nitric Oxide Releasing Coatings

As mentioned earlier, NO has potential to be used as biofilm dispersal agent for treatment of existing biofilms. Several NO releasing biomaterial coatings have been developed and used for antibiofilm applications. For example, gold nanoparticles that are biocompatible and employed for a range of biomedical applications, were functionalized with N-diazeniumdiolate conjugated macromolecules [295]. The gold nanoparticles exhibited low biofouling along with controlled NO release for 6 days.

These NO releasing nanoparticles were found to be effective in eradicating P. aeruginosa biofilm in a dose dependent manner where 2.5 ppm and 10 ppm of nanoparticles significantly reduced biofilm formation by 67 % and 83 % respectively.

Similar studies were conducted on a biomedical thermoplastic, silicone-polycarbonate- urethane doped with S-nitroso-N-acetylpenicillamine (SNAP), which was found to be highly effective in reducing bacterial adhesion of S. epidermidis and P. aeruginosa

[296]. Moreover, NO releasing catheters have also been reported to exhibit potent biofilm inhibitory activity against S. aureus both in vitro and in vivo [231,297].

Lately, there has been considerable interest in developing polymeric systems loaded with common NO donor moieties such as N-diazeniumdiolates (NONOates), S- nitrosothiols (RSNOs) including S-nitrosoglutathione (GSNO) and S-nitroso-N- acetylpenicillamine (SNAP), for storage and sustained delivery of NO [298–300].

Therefore, conjugation of NO donors to different polymeric systems such as star polymers, dendrimers and micelles has become an area of increasing interest [301–303].

In 2014, Duong et al. reported a star polymer incorporated with N-diazeniumdiolate NO donor that is able to release NO in a sustainable manner and showed great efficacy in

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inhibiting bacterial attachment and biofilm formation of P. aeruginosa in a non-growth inhibitory manner [304].

Delivery of NO through nanoparticles can also be combined with other therapeutic agents to eradicate bacterial biofilm-related infections. Recently, dual-action polymers were reported which were designed by incorporating an aminoglycoside antibiotic, gentamicin, which is capable of killing the bacteria and NO donor compounds that would aid in dispersion of antibiotic sensitive bacteria [305]. The N-diazeniumdiolate

NO donor was obtained by reacting gentamicin with NO gas to yield the hybrid complex. Both the released agents from the hybrid nanoparticles acted synergistically and were found to be highly effective in dispersing biofilms of P. aeruginosa (Figure

1.30).

Figure 1.30: Reduced P. aeruginosa biofilm exhibited by 10 µM of gentamicin-based

NO hybrid polymer. Viable and non-viable bacteria appear green and red, as well as

those stained both green/red, respectively. Scale bar = 50 µm [305].

Furthermore, the bactericidal activity of gentamicin strongly decreased the viability of bacterial cells in both biofilm and planktonic mode by 90 % and 95 % respectively. The results suggest the simultaneous release of gentamicin and NO from a polymeric matrix appears to be a highly promising strategy that can be applied as surface coatings on - 44 -

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clinical implants. However, unlike coatings based on furanone and DHP compounds, the hybrid nanoparticles displayed enhanced bacterial killing effect that may induce selective survival pressure on bacteria thereby increasing the chances of developing resistance.

1.13 Thesis Aims

Bacterial infections associated with implants and medical devices have emerged as a significant clinical problem. Due to the increasing frequency of multi-drug resistant bacterial strains and limited therapeutic options for treatment, device-related infections have become difficult to combat. Therefore, surface coatings that combine desirable features such as strong antibacterial efficacy, low toxicity towards bacteria, biocompatibility and ease of fabrication are in strong demand.

The use of QS inhibitors is an attractive strategy, as they are less prone to resistance and have potential to suppress virulence factor production and biofilm formation. The literature review shows that the field of antibacterial coatings based on QS inhibitors has achieved considerable success with encouraging results. However, further research is still required for the establishment of a more general approach to prevent device- related infections. Therefore, this project will explore different attachment strategies of a range of QS inhibitors, in order to develop the best broad-spectrum coating.

Chapter 2 demonstrates the non-specific covalent surface attachment of furanones and

DHPs based on azide/nitrene chemistry. The in vitro efficacies of the coated surfaces were examined against two common pathogens, P. aeruginosa and S. aureus.

Chapter 3 examines the biological activity of DHPs when attached on the surface from different points within the same molecule. Carboxylic acid-functionalized DHP

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analogues were synthesized for specific covalent attachment by carbodiimide chemistry and surfaces were tested for their antibacterial efficacy.

Chapter 4 investigates the efficacy of DHPs in absence of the exocyclic vinyl group at the C-5 position. This chapter outlines synthesis of functionalized DHPs and covalent immobilization via carbodiimide chemistry. The activity of resulting samples was examined.

Chapter 5 evaluates the activity of hybrid dual-action biomaterial developed by immobilizing DHP derivatives by Michael addition reaction and reacting the surfaces with NO gas to form N-diazeniumdiolate groups. The NO release profile of the surfaces was determined followed by its antibacterial activity.

Chapter 6 focuses on design and synthesis of dual-action polymeric structure by incorporating DHP and encapsulating NO donating moiety within the same polymer.

The NO release properties of the polymer were examined followed by its biofilm dispersal activity.

Chapter 7 provides an overall conclusion of the thesis and suggestions for future development of DHP coatings.

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CHAPTER TWO

Surface Attachment of FU and DHP by Photochemical Activation

CHAPTER-2

CHAPTER-2 Surface Attachment of FU and DHP by Photochemical Activation

2.1 Introduction

In the recent years, biomedical devices such as stents, heart valves and contact lenses have become an integral part of the healthcare system [306]. Bacterial attachment and colonization on surfaces of various medical implants results in the formation of biofilm

[82,84,102,158]. The bacterial communication mechanism or quorum sensing (QS) is known to control surface attachment, extracellular polymeric matrix production and release virulence factors [127,307]. Therefore, an antibacterial coating developed by incorporating QS inhibiting compound to prevent biofilm formation of pathogenic bacteria has emerged as a desirable method for prevention of device-related infections.

The main advantage of these coatings is that it would less likely induce bacterial toxicity and hence prevent development of drug resistance.

A wide range of synthetic furanones (FU), which have been derived from natural FUs, have exhibited excellent antibacterial properties [177,193]. They have also shown to possess good biofilm inhibitory activities in vitro and in vivo after attachment on biomaterial surfaces [290,308]. Similarly, structural analogues of FUs, dihydropyrrolones (DHPs) have also been successful in inhibiting bacterial QS systems in solution as well as after specific covalent attachment on surfaces [200–202,291,294].

However, a direct comparison of the activity of FUs and DHPs after attachment on the surface is lacking. Therefore, to better understand the effectiveness of FU and DHP after attachment, immobilization strategies using the same techniques are required to evaluate their activity. - 47 -

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The immobilization of FU and DHP compounds onto a substrate depends on the chemical structure of the compound. For example, a compound containing carboxylic acid functionality can be directly grafted onto a surface containing free amine groups by carbodiimide chemistry. However, the general structure of FUs and DHPs do not contain any active functional group for covalent surface attachment. Reports investigating the structure activity relationship of FUs have shown that the exocyclic double bond at the C-5 position is an important structural element responsible for its antibacterial activity [309,310]. Similarly, some studies have also indicated that the substituents at C-3, C-4 and the heterocyclic lactone ring are essential for imparting antagonist activity [173,195,202]. It was therefore hypothesized that structural modification of the FU scaffold to introduce a reactive chemical substituent may interfere with its activity.

Hume et al. had reported an azide/nitrene based method for covalent immobilization of

FU that lack a suitable reaction site on the surface of a catheter. The biological results showed that the FU compound retained its activity against S. epidermidis after covalent attachment. A detailed surface analysis, using X-ray photoelectron spectroscopy (XPS) and time-of-flight secondary ion mass spectrometry (ToF-SIMS), was performed by Al-

Bataineh et al. after incorporating FUs on to different azide functionalized surfaces

[311–314]. The spectroscopy data revealed a high density of FUs were immobilized on the surfaces via azide/nitrene route. However, the antibacterial activity of the surfaces was not reported.

Due to the absence of a suitable reactive functionality and difficulty in introducing an active site because of low stability of FU ring in acidic and alkaline conditions, the azide/nitrene chemistry approach appears to be an attractive strategy for surface

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immobilization of FUs. For a direct comparison of the attachment efficiency and efficacy of FUs with DHPs, DHP molecules were also surface immobilized via azide reaction. In this chapter, three potent FU and DHP compounds were selected for covalent surface immobilization (Figure 2.1). Glass was chosen as a model surface due to its relative ease of functionalization for attachment and characterization by various techniques such as XPS, contact angle measurements and fluorescence microscopy.

Furthermore, coupling strategies performed on glass can be easily reproduced on different types of biomaterial surfaces such as Teflon, titanium, silicone etc.

Figure 2.1: Chemical structures of selected FUs and DHPs used in this study.

For attachment, a suitable azide containing molecule was required. Aryl azides were first introduced by Fleet and co-workers as photoaffinity labelling agents which are used for binding biomolecules after photo-activation [315–319]. Phenyl azides such as

4-azidobenzoic acid (ABA) are commonly used for surface functionalization, crosslinking and protein attachment due to its fast kinetics, high reaction efficiency and high storage stability [320–324]. After irradiation under UV light, azides forms a highly reactive intermediate, nitrene, which is capable of inserting into C-H and N-H bonds to form stable covalent bonds [325–328].

In this work, an amino silane linker, 3-aminopropyltriethoxysilane (APTS), was first reacted with the glass surface by chemical vapour deposition to introduce free amine - 49 -

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groups. Following this, the amine surface was coupled with ABA to develop azide terminated glass surfaces. The FUs and DHPs were subsequently immobilized on the azide glass by photo-activation of the azide groups under UV light. Since nitrenes undergo C-H/N-H cleavage-insertion reactions, it was also expected that it would react with different parts of the molecule resulting in non-specific covalent attachment of FUs and DHPs through various positions (Figure 2.2). The coated surfaces were characterized by X-ray photoelectron spectroscopy (XPS) and contact angle measurements, and the antibacterial efficacy of surface-bound FUs and DHPs was assessed against two common bacterial pathogens, Pseudomonas aeruginosa and

Staphylococcus aureus.

Figure 2.2: Non-specific photo-immobilization of FU and DHP molecules via

azide/nitrene chemical route.

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2.2 Materials and Methods

2.2.1 Attachment of 3-Aminopropyltriethoxysilane (APTS)

Glass coverslips (No. 1, diameter 13 mm D 263 M glass, ProSciTech, Australia) were first cleaned in freshly prepared piranha solution (3:1 v/v concentrated sulphuric acid to

30 % hydrogen peroxide) at 100 °C for 1 h. After thorough rinsing with distilled water, the clean coverslips were rinsed once with absolute ethanol and air-dried. The substrates were then silanized according to the previously developed method [291]. Briefly, the clean substrates were placed on steel mesh within a glass vessel that contained a 3- aminopropyltriethoxysilane (APTS) solution (10 % v/v in dry toluene; 1 ml). The glass vessel was sealed and heated at 140 °C for 18 h (Figure 2.3). The coverslips were rinsed with dry toluene (×2), absolute ethanol and air-dried.

Figure 2.3: Chemical vapour deposition of 3-aminopropyltriethoxysilane (APTS) on

blank glass coverslip.

2.2.2 Attachment of 4-Azidobenzoic Acid (ABA)

The APTS-coated coverslips were then immersed in a solution of 4-azidobenzoic acid

(ABA; 49.0 µM), 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride

(EDC, 245.2 µM) and N-hydroxysuccinimide (NHS, 98.0 µM) in absolute ethanol (1.5 ml), and agitated overnight at room temperature under dark room conditions. The ABA-

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functionalized surfaces were rinsed twice with absolute ethanol and once with MilliQ water, air-dried and stored under dark conditions before use.

2.2.3 Attachment of FU and DHP via Azide/Nitrene Chemistry

The synthetic halogenated FU compounds (FU-1, -2 and -3) were synthesized as described [329]. Similarly, DHP compounds (DHP-1, -2 and -3) were synthesized following the method developed previously by Kumar and Iskander [202].

Stock solutions of FU (25 mg/ml in dichloromethane) and DHP (25 mg/ml in acetone) were prepared and 200 µl of the FU or DHP solution was placed onto the ABA glass surface. After complete evaporation of the solvent, the surfaces were irradiated under

UV at 320 nm for 10 min in a CL-1000 Crosslinker (Ultra-Violet Products Ltd, Upland,

CA, USA) (Figure 2.4). The unreacted FU and DHP were removed by extensively washing the samples with dichloromethane and acetone respectively, MilliQ water and absolute ethanol, then air dried and stored in clean sterile container.

Figure 2.4: Immobilization of FUs and DHPs via photo-activation of azide groups on

glass surface.

2.2.4 X-Ray Photoelectron Spectroscopy (XPS)

X-ray photoelectron spectroscopy measurements were performed on an ESCALAB

220iXL. Monochromatic Al Kα X-rays (1486.6 eV) incident at 58° to the analyzer lens were used to excite electrons from the sample. Emitted photoelectrons were collected on - 52 -

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a hemispherical analyzer with a multichannel detector at a take-off angle of 90° from the plane of the sample surface. The analyzing chamber operated below 10–8 Torr, and the spot size was approximately 1 mm2. The resolution of the spectrometer was ~0.6 eV. All energies are reported as binding energies in eV taken from three samples of each type and referenced to the C 1s signal (corrected to 285.0 eV). Survey scans were carried out at 100 ms dwell time and an analyzer pass energy of 100 eV. High- resolution scans were run with 0.1 eV step size, dwell time of 100 ms, and analyzer pass energy set to 20 eV. After background subtraction using the Shirley routine, spectra were fitted with a convolution of Lorentzian and Gaussian profiles as described by

Ciampi et al. [330].

2.2.5 Contact Angle Measurements

Contact angles were determined using a contact angle goniometer (Rame-Hart, Inc NRL

USA, Model no. 100-00). Multiple drops of deionized water were placed on each surface using a micro-syringe. The angle between the droplet and the surface was measured using a 50 mm Cosmicar Television Lens (Japan). Rame-Hart Imaging software was used to calculate the contact angle. A minimum of fifteen measurements were made on five samples of each FU and DHP.

2.2.6 Bacterial Adhesion Analysis

Bacteria (Staphylococcus aureus SA38 and Pseudomonas aeruginosa PA01) from frozen stock (–80 °C) were streaked on chocolate agar (Oxoid, UK) and incubated at 37

°C overnight. A colony of the bacteria was taken from the plate and cultured overnight at 37 °C in 15 ml tryptone soya broth (TSB; Oxoid, UK). The bacteria were washed twice with fresh TSB by centrifugation. The optical density (OD) of the culture was

8 adjusted to OD660 = 0.1 which corresponds to 10 CFU/ml.

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In a 12-well plate, the surfaces to be tested were first sterilized with 70 % w/v ethanol for 30 min, then thoroughly washed with sterile phosphate buffered saline (PBS) three times and finally placed in 4 ml of the adjusted bacterial culture. The surfaces were incubated at 37 °C for 24 h. The media was then replaced by fresh TSB (4 ml) and further incubated for 24 h at 37 °C. Subsequently, the samples were washed twice with

PBS before examination by fluorescence microscopy.

The glass samples with adherent bacterial cells were stained with Live/Dead BacLight

Bacterial Viability Kit (Molecular Probes, Inc, Eugene, OR, USA) according to the manufacturers’ procedure and as described in the literature for analysis of biofilms on surfaces [223,291,294,331]. Briefly, 2 µl of the two components were mixed thoroughly in 1 ml of PBS; 100 µl of the solution was then placed on each sample and allowed to incubate at room temperature in the dark for 15 min. The excess stain was gently washed by PBS (200 µl). Bacteria were fixed by adding 100 µl of 4 % formaldehyde on each sample and then placed on the glass microscopy slide.

Microscopic observation and image acquisition were performed with Olympus FV1200

Confocal Inverted Microscope. The bacterial cells that were stained green were considered to be viable, while those that stained red or both green and red were considered to be dead [332]. Images from 10 representative areas on each triplicate samples for each surface from three experimental repeats were taken and analysed using

ImageJ software [333]. The results were reported as the average percentage coverage of live and dead cells of the fields of view.

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2.2.7 Statistical Analysis of Data

Data were analyzed by the one-way analysis of variance (ANOVA) using GraphPad

Prism 7.03 software. Differences between the groups were determined using post hoc multiple comparison with Tukey correction, and significance of results was set at 5 %.

2.3 Results

2.3.1 High-Resolution XPS Characterization of Coated Surfaces

The surfaces were characterized by XPS at each step of the immobilization sequence to ensure surfaces were successfully modified. The XPS data collected for the blank

(untreated), APTS, ABA, FU and DHP coated surfaces are summarized in Table 2.1.

Changes in the elemental composition of carbon, nitrogen and halogen indicate successful immobilization of FU and DHP on the surfaces (Table 2.1). The blank glass contained 7.6 % carbon and 0.5 % nitrogen. After functionalization with APTS, the carbon and nitrogen percentages increased to 43.9 % and 7.1 % respectively when compared to the blank. Both the carbon and nitrogen content increased even further to

46.2 % and 8.1 % respectively when ABA was coupled with the amine surface. The subsequent attachment of FU and DHP was confirmed by a further increase in carbon percentage by 2.6–3.5 % and 1.7–2.5 % respectively. Finally, the detection of halogens from the FU and DHP compounds in the XPS data, which are used as a marker, further confirmed the attachment of FUs and DHPs. Based on the halogen content, FUs (0.74–

0.41 % Br) exhibited higher surface attachment compared to DHP compounds (0.17 %

Br for DHP-1, 0.32 % F for DHP-2 and 0.35 % Br for DHP-3).

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Table 2.1: XPS analysis of blank APTS, ABA, FU and DHP coated surfaces.

Surface % C % N % Halogen

Blank 7.6 0.5 - APTS 43.9 7.1 - ABA 46.2 8.1 - FU-1 48.8 7.3 0.74% Br FU-2 49.0 7.7 0.41% Br FU-3 49.7 7.1 0.65% Br DHP-1 47.9 7.7 0.17% Br DHP-2 48.5 8.7 0.32% F DHP-3 48.7 8.5 0.35% Br

Analysis of the high-resolution C 1s spectra of the APTS surface demonstrated the presence of three distinct components C-H/C-C, C-N and C=O at binding energies

284.9 eV, 286.1 eV and 288.2 eV respectively. The N 1s spectrum of the APTS surface

+ showed two peaks at 399.6 eV and 401.4 eV corresponding to –NH2 and –NH3 respectively (Figure 2.5A). After the subsequent attachment of ABA on the glass surface, two new additional peaks in a 2:1 ratio emerged at 400.2 eV and 404.6 eV in the N 1s scan, which is a set of characteristic peaks attributable to the azide functionality (Figure 2.5B) [334–336]. The peak at 400.2 eV was assigned to the two terminal nitrogen atoms of the azide and the peak at 404.6 eV was assigned to the central nitrogen atom because of its low electron density compared to the terminal nitrogen atoms. In addition, the intensity of the N-H peak at 399.6 eV reduced and a

+ slight shift in the peak from 401.4 eV to 402.1 for –NH3 . Furthermore, an additional peak at 289.0 eV (N-C=O) in the carbon narrow scan was observed indicating that the

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coupling reaction between the amine-terminated surface and carboxylic acid of ABA successfully formed an amide bond.

The characteristic azide peaks were not observed after the subsequent treatment of the

ABA surface with FU or DHP, suggesting the azide functional groups were consumed for the covalent linkage of FUs and DHPs. Instead, the N 1s spectra showed a peak corresponding to N-H at 399.5 eV which is consistent with the formation of N-H group on photo-activating the azide and also due to various side reactions of phenyl azides under UV light [320,321,326,327,337]. Furthermore, a slight shift in the peak for N-

C=O (from 289.0 eV to 288.7 eV) in the C 1s spectra was also observed for all FU and

DHP coated surfaces along with broadening of the band, possibly due to addition of C-

Br or C-F, indicating successful attachment of FU and DHP.

(A) APTS glass

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(B) ABA glass

Figure 2.5: XPS high resolution N 1s spectra of (A) APTS glass and (B) ABA glass

surfaces.

2.3.2 Contact Angle Measurements

The surfaces were also characterized by determining the water contact angle values after every modification step. The contact angle measures the changes in hydrophobicity of a surface following attachment of APTS, ABA, FUs and DHPs. The contact angle values of all the glass substrates are shown in Table 2.2.

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Table 2.2: Water contact angle measurements of blank APTS, ABA, FU and DHP

coated surfaces.

Contact Surface Angle (°) (±1) Blank 20 APTS 73 ABA 67 FU-1 70 FU-2 60 FU-3 65 DHP-1 71 DHP-2 69 DHP-3 74

The blank glass has a very low contact angle of 20° which indicates the surface is highly hydrophilic due to the free hydroxyl groups present on the surface. A significant change in contact angle was observed after attachment of APTS (from 20° to 73°), indicating an increase in surface hydrophobicity due to the aliphatic carbon chain of APTS which is hydrophobic in nature. The contact angle or hydrophobicity of the surface remained approximately the same (67°) after surface attachment of ABA, which is expected due to the presence of the hydrophobic aromatic ring in ABA. The subsequent attachment of

FU and DHP did not result in any significant changes in the contact angle values from the ABA surface, due to the presence of different hydrophobic moieties (alkyl chain, phenyl ring and halogen atoms) on the FU and DHP compounds.

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2.3.3 Antibacterial Activity

The ability of FU and DHP to prevent bacterial colonization of S. aureus and P. aeruginosa on the modified surfaces was evaluated using fluorescence microscopy, and representative images are shown in Figure 2.6 and Figure 2.7. The total surface area covered by bacteria and the amount of live and dead bacteria (stained green and red respectively using the Live/Dead staining kit) was determined by performing image analysis. The results showing relative proportions of live and dead bacterial cells for S. aureus and P. aeruginosa are shown in Figure 2.8 respectively.

Figure 2.6 shows microscopy images of live (green) and dead (red) S. aureus cells adhered to the coated and control surfaces, where extensive colonization indicated by the high bacterial coverage can be seen on the APTS and ABA process controls (Figure

2.6A and B). Figure 2.7 shows images for adhesion of P. aeruginosa on the surfaces.

The bacterial coverage of both strains on all the FU and DHP coated surfaces was significantly lower than the process control.

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S. aureus

A – APTS B – ABA

C – FU-2 D – DHP-3

Figure 2.6: Fluorescence microscopic images of glass surfaces after adhesion of S. aureus to APTS and ABA process control (A and B); FU-2 and DHP-3 coated surfaces

(C and D). Live bacterial cells stained green and dead bacteria stained red.

Magnification 200×. Scale bar = 100 µm.

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P. aeruginosa

A – APTS B – ABA

C – FU-2 D – DHP-3

Figure 2.7: Fluorescence microscopic images of glass surfaces after adhesion of P. aeruginosa to APTS and ABA process control (A and B); FU-2 and DHP-3 coated surfaces (C and D). Live bacterial cells stained green and dead bacteria stained red.

Magnification 200×. Scale bar = 100 µm.

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The quantitative image analysis showed that both strains of bacteria displayed high percentage coverage on the control surfaces. The percentage of adherent bacteria on

ABA for S. aureus were found to be 21.4 ± 2.3 % and for P. aeruginosa 11.8 ± 0.7 % coverage was observed (Figure 2.8). The adhesion of both strains of bacteria on blank and APTS glass did not vary significantly and was similar to that reported in literature

[291].

Results of image analysis for S. aureus showed significant reductions in overall bacterial coverage of 75.4 ± 5.0 %, 80.9 ± 4.1 %, and 74.8 ± 4.2 % for FU 1-3 compared to the ABA control surface (p < 0.001) (Figure 2.8A). Similarly, image analysis results showed DHP 1-3 coated surfaces were able to reduce adherent cells by

75.6 ± 5.8 %, 89.9 ± 1.6 %, 93.4 ± 1.1 % (p < 0.001), where DHP-2 and DHP-3 displayed comparatively lower bacterial coverage than DHP-1 coated surface (p < 0.05).

There was no significant difference in the percentage of bacterial cells stained red (dead bacteria) between the controls and modified surfaces.

The attachment of P. aeruginosa on the FU and DHP coated surfaces was found to be significantly lower than the control, with reductions of 54.8 ± 2.2 %, 68.7 ± 1.7 %, 52.9

± 3.0 % for FU 1-3 and 55.9 ± 2.8 %, 54.3 ± 2.1 %, 71.2 ± 1.4 % for DHP 1-3 (p <

0.001) respectively (Figure 2.8B). In this case, FU-2 and DHP-3 gave maximum reduction in bacterial attachment compared to other FU and DHP surfaces (p < 0.05).

Similar to S. aureus, no significant difference was observed in the percentage of dead cells between the control and modified surfaces.

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A Bacterial coverage - S. aureus 27 24 21

18 Dead 15 bacteria 12 Live 9 ^ bacteria 6 * * * ^ Bacterial covergae (%) Bacterial * * 3 * 0 APTS ABA FU-1 FU-2 FU-3 DHP-1 DHP-2 DHP-3

B Bacterial coverage - P. aeruginosa 14

10 Dead 8 bacteria

6 * * * * Live × ^ bacteria

4 * * Bacterial Coverage(%) Bacterial 2

0 APTS ABA FU-1 FU-2 FU-3 DHP-1 DHP-2 DHP-3

Figure 2.8: Percentage bacterial coverage of live and dead bacteria for (A) S. aureus and (B) P. aeruginosa (mean ± standard error of the mean); *indicates p < 0.001 compared to APTS and ABA control; ×indicates p < 0.05 compared to FU surfaces;

^indicates p < 0.05 compared to DHP surfaces.

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2.4 Discussion

Tens of millions of medical devices are used each year, and in spite of advances in biomaterial technologies, a significant proportion of the devices are colonized by bacterial biofilms, resulting in device failure and infections. The formation of biofilms on biomedical devices is therefore a serious problem. In the present study, various potent QS inhibiting compounds, FUs and DHPs, were covalently immobilized on glass surface by a non-specific attachment strategy and the antibacterial efficacy of the resultant surfaces was assessed.

XPS analysis indicated the successful attachment of FUs and DHPs via the described photo-activation strategy with FUs having slightly higher attachment efficiency compared to the DHPs. All the covalently bound FUs and DHPs were able to significantly reduce colonization of both Gram-positive (S. aureus) and Gram-negative

(P. aeruginosa) bacteria. Some differences in the activity of the coated surfaces were observed. Surfaces immobilized with DHP-2 and DHP-3 were found to be the most potent against bacterial adhesion of S. aureus, whereas for P. aeruginosa, the most active surfaces were coated with FU-2 and DHP-3. While all compounds were effective in reducing bacterial adhesion, FU-2 and DHP-3 displayed the best broad spectrum antibacterial activity.

The high level of reduction in adherent bacteria displayed by DHP-3 (93.4 % and 71.2

% of reduction against S. aureus and P. aeruginosa respectively) is consistent with reports of its higher activity compared to other DHPs as a free compound and even after surface attachment [202,291]. In a previous study, an acrylate-functionalized derivative of DHP-3 was covalently attached onto glass surface via Michael addition reaction

[291]. A higher surface concentration of DHP-3 was achieved via the non-specific azide

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reaction (0.35 % Br) described in this study than by the Michael addition reaction (0.21

% Br). Further, the DHP-3 coatings via Michael addition reaction also showed comparatively lower activity in reducing bacterial coverage (76.5 % for S. aureus and

58.9 % for P. aeruginosa), suggesting the azide coatings increased the efficacy of DHP-

3 by 17 % and 12 % against S. aureus and P. aeruginosa respectively.

Numerous studies have demonstrated that an increase in surface concentration of an active compound leads to better antibacterial activity [290,294,338]. Surprisingly in this study, DHP surfaces have displayed potent activity even at low concentration. Among all the compounds used in this study, DHP-1 gave the least attachment (0.17 % Br) to the surface but displayed a similar level of activity as FU-1 which gave a maximum attachment (0.74 % Br). This discrepancy could be due to the orientation of DHP on the surface, making it more available for antibacterial activity compared to a similar concentration of FU. Similarly, FU-3 was expected to display maximum efficacy amongst all the FUs due to its high activity in solution and high surface attachment efficiency (0.65 % Br) [179,193,195]. Instead, FU-2 displayed the best activity out of all FUs at lower surface concentration (0.41 % Br) with reductions of 80.9 % and 68.7

% of adherent S. aureus and P. aeruginosa respectively, while FU-3 displayed reductions of 74.8 % and 52.9 % for S. aureus and P. aeruginosa respectively.

Several strategies have been explored in the past to immobilize QS inhibiting compounds on the surfaces to inhibit biofilm formation [289,294,339]. For example, a furanone derivative has been physically adsorbed on various biomaterial surfaces [289].

However, such a non-uniform coating is highly prone to leaching and gradual loss of the active compound. In another approach, FUs and DHPs were coated on surfaces via specific attachment strategies [291,294,339]. Although this attachment strategy

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overcomes the limitations of uneven coating and leaching, it requires extensive modification of the compound for surface attachment. As mentioned earlier, any structural change or modification of the active compound may also result in decrease in activity. The non-specific attachment strategy employed in this study does not require structural modification or functionalization of the compound. Also, unlike the previous attachment strategies, the azide reaction described in this study is much faster, making it easier and more convenient to implement. This study is the first to investigate the antimicrobial activity of photo-immobilized DHPs on surfaces.

DHPs act by interfering with the bacterial QS system. In particular, DHPs are able to disrupt the N-acylhomoserine lactones (AHL) regulated QS system in Gram-negative bacteria [202]. The mechanism through which DHPs inhibit QS is postulated to be similar to that of FUs, that is, via displacing the AHL signal from the receptor site without affecting bacterial growth [172,173,179,340]. Surfaces immobilized with DHPs were capable of interfering with the AHL regulated las QS system in P. aeruginosa, thereby inhibiting biofilm formation [294]. In the current study, about 97 % of adherent bacteria on the coated surfaces were alive, supporting the previous data that FUs and

DHPs act without killing the bacteria and exerting no selective pressure on the bacteria to develop resistance [291,308]. Therefore, it is likely that the FU and DHP surfaces generated in this study act through the same mechanism of action for Gram-negative bacteria.

On the other hand, for Gram-positive bacteria, the mode of action of FU and DHP is still not fully understood. Research investigating the effect of the AHL, N-(3- oxododecanoyl)-L-homoserine lactone (3-oxo-C12-HSL), on S. aureus showed that the mode of action of 3-oxo-C12-HSL involves inhibition of the agr-dependent QS system

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by binding to the cytoplasmic membrane of S. aureus [341]. Similarly, two new classes of compounds recently identified, one derived from 3-oxo AHLs and other from 3-acyl tetronic acids, have displayed agr QS inhibitory activity in S. aureus [342]. In another study, the mechanism of action of a derivative of AHL was found to be through the dissipation of the membrane potential and pH gradient of S. aureus and Bacillus cereus

[343]. Therefore, it is possible that FUs and DHPs, which are structurally related to

AHLs, inhibit QS of Gram-positive bacteria via an indirect approach through the interaction with the bacterial cell membrane.

2.5 Conclusion

In the current study, an effective and versatile technique for the immobilization of QS inhibitors as an antibacterial coating was demonstrated. All the FU and DHP coated surfaces were able to reduce the adhesion of S. aureus and P. aeruginosa, the most common pathogens associated with biomaterial infections. This suggests that the non- specific attachment of FUs and DHPs to the highly reactive azide groups does not impair the antibacterial activity of the compounds. Additionally, some of the DHP surfaces displayed enhanced efficacy despite low attachment probably due to the change in DHP orientation after attachment, which needs to be further investigated. Since functionalization of compounds was not needed, this strategy was found to be a fast and easy technique for developing novel coatings for prevention of infections of biomedical devices.

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Role of Orientation of Surface Bound DHP on Biological Activity

CHAPTER-3

CHAPTER-3 Role of Orientation of Surface Bound DHP on Biological Activity

3.1 Introduction

A promising strategy to prevent infections of biomedical devices is to coat their surface with a suitable antibacterial agent. Compared to conventional antibiotics, QS inhibitors such as dihydropyrrol-2-ones (DHPs) are excellent coating agents for medical implants since they are less susceptible to induce the development of antibacterial resistance and have exhibited broad spectrum antibacterial activity with low cytotoxicity [202,344].

A recent study has reported that the surface concentration of covalently-attached DHPs had a great impact on the activity of coated surfaces, with higher amounts of specifically-bound DHPs leading to increased biofilm inhibition [294]. However, it was observed in the previous chapter that the activity of modified surfaces was not greatly affected by the concentration of DHPs that were grafted by non-specific attachment.

These surfaces were highly effective in preventing bacterial colonization at low concentrations against two common pathogens, Staphylococcus aureus and

Pseudomonas aeruginosa. This difference in the efficacy of DHPs could be due to changes in the orientation of DHP when immobilized on the surface via non-specific attachment strategy, by exposing the more active part of molecule to the bacteria.

For the development of efficient and long-lasting antibacterial devices and implants, it is important to understand the effect of the orientation of the attached active molecule on its activity. To date, no systematic study has been carried out to investigate the effect of different orientations of DHP on its antibacterial activity. In this chapter, potent DHP

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compounds that have been reported in previous studies were covalently linked to the surface via a specific attachment strategy from various positions of the DHP scaffold.

By exposing different parts of the same active molecule to bacteria, the effect of compound orientation on biological activity could be explored.

In the first approach, a series of potent DHP analogues having a similar structural skeleton with different substituents at the phenyl C-4 position were functionalized with a free carboxylic acid group at the N-1 position according to a modified method developed by Kumar and Iskander [202] (Figure 3.1A). This generates a site of attachment at the N-atom of the lactam ring with minimal change to the molecular structure of the DHP. In the second approach, a carboxylic acid group was introduced at the para position of the C-4 pendant phenyl ring (Figure 3.1B) following a literature procedure with a few modifications [202].

Figure 3.1: Functionalization of active DHP molecule with carboxylic acid group at

(A) N-1 position and (B) C-4 phenyl ring.

In both the compounds, the exocyclic vinyl group at the C-5 position is untouched since it has been found to be crucial for antibiofilm activity from the structural activity relationship of the analogous furanone compounds [309,310]. Attachment of the DHPs

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at the C-5 position could therefore impair the biological activity of the compound. The activity of resulting free DHP compounds was investigated on the QS system of Gram- negative bacteria, a QS reporter strain P. aeruginosa MH602, which produces GFP when the QS system is active. The level of GFP expression reflects the level of QS in the bacterial culture, and the compounds that inhibit QS should result in lowered expression of GFP.

The acid-DHP derivatives were then subsequently grafted onto amino-functionalized glass substrates by the EDC/NHS coupling reaction (Figure 3.2).

Figure 3.2: Specific covalent attachment of DHPs on amine-functionalized surfaces

using the EDC/NHS coupling reaction.

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The efficacy of DHPs was also investigated on a clinically relevant biomaterial surface, since glass is not a biomaterial and is used only as test surface. For this, a substrate extensively used in the biomedical fields was needed and therefore a long-lasting synthetic biomaterial, fluorinated ethylene propylene (FEP, also known as Teflon FEP) was selected for attachment (Figure 3.3). The N-substituted DHP (phenyl substituent, R

= H) and the para-acid substituted phenyl DHP were attached onto the new biomaterial surface using the same attachment strategy developed on glass. Due to the non-reactive nature and chemical inertness of FEP, plasma-mediated surface functionalization is often used to introduce reactive chemical groups such as carboxyl, amine, hydroxyl groups on the surface [345–347]. The FEP surfaces in this chapter were functionalized with free primary amine groups for attachment using the allylamine plasma treatment process.

Figure 3.3: Chemical structure of FEP.

Overall, four DHPs with carboxylic acid groups at the N-1 position of the lactam ring

(DHP acids 1-4) and one DHP analogue with a carboxylic group at the C-4 pendant phenyl ring (p-acid DHP) were synthesized (Figure 3.4). They were attached onto

APTS-functionalized glass and allylamine-treated FEP surfaces by the EDC/NHS coupling reaction. The functionalized glass and FEP surfaces were characterised by X- ray photoelectron spectroscopy (XPS) and contact angle measurements and tested for their biological activity against S. aureus and P. aeruginosa.

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Figure 3.4: Chemical structures of DHP derivatives synthesized.

3.2 Materials and Methods

3.2.1 General

All chemical reagents were purchased from commercial sources (Alfa-Aesar and Sigma

Aldrich) and used without further purification. Solvents were sourced from commercial sources and used as obtained. Reactions were performed using oven-dried glassware under an atmosphere of nitrogen and in anhydrous conditions (if required). Room temperature refers to the ambient temperature (22–24 °C). Yields refer to chromatographically and spectroscopically pure compounds unless otherwise stated.

Reactions were monitored by thin layer chromatography (TLC) precoated with Merck silica gel 60 F254. Visualization was performed by the quenching of short or long wavelength UV fluorescence or by staining with potassium permanganate or ninhydrin solution. Flash chromatography was carried out using Grace Davison LC60A 6–35 micron silica gel. Preparative thin layer chromatography was carried out on 3 × 200 ×

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reported as wavelength (λ) in nm and absorption coefficient (ε) in cm-1M-1. High resolution mass spectrometry (HRMS) was performed by the Bioanalytical Mass

Spectrometry facility, UNSW. Melting points were obtained using Mel-Temp melting point apparatus and are uncorrected. Proton and carbon NMR was recorded in designated solvents using Bruker DPX 300 or a Bruker Avance 400 spectrometer as designated. Chemical shifts (δ) are quoted in parts per million (ppm), to the nearest 0.01 ppm and internally referenced relative to the solvent nuclei. 1H NMR spectral data are reported as follows: chemical shift in ppm; multiplicity in broad (br), singlet (s), doublet

(d), triplet (t), quartet (q), multiplet (m) or a combination of these (e.g. dd, dt etc.)]; coupling constant (J) in hertz, integration, proton count and assignment.

3.2.2 Synthesis of Acid-Functionalized DHP Derivatives

Tert-butyl-2-(5-(2-(tert-butoxy)-2-oxoethoxy)-5-methyl-2-oxo-4-phenyl-1,5-dihydro-

2H-pyrrol-1-yl)acetate

5-Hydroxy-5-methyl-4-phenyl-1,5-dihydro-2H-pyrrol-2-one

(1.0 g, 2.58 mmol) was added to a solution of potassium hydroxide (0.6 g, 10.57 mmol) in dry DMSO (5 ml) and the mixture was stirred at room temperature for 20 min. To the solution, tert-butyl chloroacetate (1.6 g, 10.57 mmol) was added and stirred at room temperature for 24 h. The crude mixture was washed with water and extracted into ethyl acetate. The extracted organic layer was dried over sodium sulphate, evaporated under vacuum and flash chromatographed to yield a white

1 solid (1.36 g, 61 %). M.p. 128 °C; H NMR (300 MHz, CDCl3) δ 1.46 (d, J = 8.6 Hz,

21H, 7 x CH3), 4.01 (s, 2H, CH2), 4.34 (s, 2H, CH2), 6.26 (d, J = 1.2 Hz, 1H, CH), 7.44

13 (brs, 5H, ArH); C NMR (100 MHz, CDCl3) δ 23.2 (CH3), 28.0 (6 x CH3), 41.6 (CH2),

60.9 (CH2), 82.4 (2 x C-O), 96.9 (CH), 121.0 (C), 128.6 (2 x ArCH), 129.4 (ArCH), - 74 -

CHAPTER-3

131.9 (2 x ArCH), 145.1 (ArC), 150.7 (C), 167.2 (C=O), 168.9 (C=O), 172.6 (C=O); IR

-1 (ATR): υmax 3071, 2972, 1725, 1703, 1389, 1227, 1150, 1084, 916, 837, 769, 695 cm ;

-1 -1 UV (ACN): λmax 207 nm (ε 8970 cm M ), 218 (9762), 261 (11,848); HRMS (ESI) m/z

+ calcd for C23H31NO6Na 440.2044 [M+Na] , found 440.2042.

2-(5-Methylene-2-oxo-4-phenyl-1,5-dihydro-2H-pyrrol-1-yl)acetic acid

(DHP acid-1)

Trifluoroacetic acid (2.5 ml) was added to a solution of tert-butyl-2-(2-

(2-(tert-butoxy)-2-oxoethoxy)-2-methyl-5-oxo-3-phenyl-2,5-dihydro-

1H-pyrrol-1-yl) acetate (1.1 g, 2.76 mmol) in dichloromethane (15 ml) and the mixture was stirred at room temperature for 24 h. The reaction mixture was washed with saturated sodium bicarbonate and water. The ethyl acetate layer was separated, dried over sodium sulphate and chromatographed to

1 yield a white solid (0.38 g, 69 %). M.p. 165-166 °C; H NMR (300 MHz, CDCl3) δ 4.52

(s, 2H, CH2), 4.98 (dd, J = 2.7 and 1.7 Hz, 1H, =CH2), 5.08 (d, J = 2.7 Hz, 1H, =CH2),

13 6.30 (s, 1H, CH), 7.44-7.49 (m, 5H, ArH); C NMR (100 MHz, CDCl3) δ 40.5 (CH2),

97.4 (CH2), 120.8 (CH), 128.6 (4 x ArCH), 129.6 (ArCH), 131.6 (ArC), 144.7 (C),

151.6 (C), 171.4 (C=O), 175.0 (C=O); IR (ATR): υmax 2905, 2828, 2725, 2601, 2533,

- 1732, 1650, 1625, 1431, 1341, 1211, 1146, 864, 765, 704, 700 cm 1; UV (ACN): λmax

-1 -1 276 nm (ε 13,011 cm M ); HRMS (ESI) m/z calcd for C13H11NO3Na 252.0631

[M+Na]+, found 252.0632.

Other derivatives (DHP acid 2-4) were synthesized by following the same method from the corresponding tert-butyl acetate DHPs.

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2-(5-Methylene-2-oxo-4-(2-fluorophenyl)-1,5-dihydro-2H-pyrrol-1-yl)acetic acid

(DHP acid-2)

1 M.p. 150 °C; H NMR (400 MHz, CDCl3) δ 4.51 (s, 2H, CH2), 4.94-4.97

(m, 2H, =CH2), 6.39 (s, 1H, CH), 7.16-7.24 (m, 2H, ArH), 7.35-7.45 (m,

13 2H, ArH); C NMR (100 MHz, CDCl3) δ 40.5 (CH2), 97.1 (CH2), 116.2

(CH), 116.4 (ArCH), 123.5 (ArC), 124.2 (ArCH), 130.9 (ArCH), 131.3

(ArCH), 144.5 (C), 145.0 (C), 161.0 (ArCF), 169.0 (C=O), 171.5 (C=O);

IR (ATR): υmax 1549, 2916, 2603, 2537, 2104, 1931, 1727, 1624, 1486, 1432, 1352,

-1 -1 -1 1208, 1085, 882, 832, 759 cm ; UV (ACN): λmax 272 nm (ε 10,352 cm M ); HRMS

+ (ESI) m/z calcd for C13H10FNO3Na 270.0537 [M+Na] , found 270.0535.

2-(5-Methylene-2-oxo-4-(4-fluorophenyl)-1,5-dihydro-2H-pyrrol-1-yl)acetic acid

(DHP acid-3)

1 M.p. 201-202 °C; H NMR (400 MHz, CDCl3) δ 4.50 (s, 2H, CH2), 4.96

(t, J = 2.7 Hz, 1H, =CH2), 5.01 (d, J = 2.7 Hz, 1H, =CH2), 6.26 (s, 1H,

CH), 7.13-7.17 (m, 2H, ArH), 7.41-7.44 (m, 2H, ArH); 13C NMR (100

MHz, CDCl3) δ 40.4 (CH2), 97.2 (CH2), 115.8 (CH), 116.1 (2 x ArCH),

120.9 (ArC), 130.4 (2 x ArCH), 144.7 (C), 150.0 (C), 162.0 (ArCF),

168.9 (C=O), 172.0 (C=O); IR (ATR): υmax 2886, 2729, 2606, 2538, 2116, 1733, 1655,

-1 1628, 1501, 1430, 1351, 1212, 1144, 916, 882, 845, 770 cm ; UV (ACN): λmax 275 nm

-1 -1 + (ε 10,846 cm M ); HRMS (ESI) m/z calcd for C13H10FNO3Na 270.0537 [M+Na] , found 270.0534.

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2-(5-Methylene-2-oxo-4-(4-bromophenyl)-1,5-dihydro-2H-pyrrol-1-yl)acetic acid

(DHP acid-4)

1 M.p. 191 °C; H NMR (300 MHz, Acetone-d6) δ 4.49 (s, 2H, CH2), 5.04

(d, J = 2.4 Hz, 1H, =CH2), 5.19 (s, 1H, =CH2), 6.4 (s, 1H, CH), 7.51 (d,

J = 8.4, 2H, ArH), 7.7 (d, J = 8.4, 2H, ArH); 13C NMR (75.5 MHz,

Acetone-d6) δ 40.0 (CH2), 96.1 (CH2), 121.3 (CH), 123.2 (ArCBr),

130.5 (2 x ArCH), 131.5 (C), 132.0 (2 x ArCH), 144.7 (ArC), 149.3 (C),

167.9 (C=O), 168.7 (C=O); IR (ATR): υmax 3350, 2918, 2733, 2528, 2110, 1908, 1719,

-1 1664, 1483, 1433, 1396, 1349, 1217, 1199, 1069, 873, 824 cm ; UV (ACN): λmax 222

-1 -1 nm (ε 10,897 cm M ), 278 (11,112); HRMS (ESI) m/z calcd for C13H10BrNO3Na

329.9736 [M+Na]+, found 329.9735.

(Z)-4-(1-Carboxy-3-oxobut-1-en-2-yl)benzoic acid

Phosphoric acid (15 ml) was added to a solution of 4-(2- oxopropyl)benzoic acid (0.35 g, 1.5 mmol) and glyoxylic acid (0.37 g, 4 mmol) and the mixture was heated at 75 – 80

°C for 5 h. The mixture was cooled to room temperature, extracted into 1:1 DCM/ether layer and dried over MgSO4. The solvent was evaporated under vacuum and purified by flash chromatography with 5:1 ethyl acetate/methanol to obtain a pale yellow solid

1 (0.18 g, 50 %). M.p. 114-116 °C; H NMR (300 MHz, DMSO-d6) δ 1.68 (s, 3H, CH3), δ

6.8 (s, 1H, =CH), δ 8.0-8.05 (m, 4H, ArH), δ 13.1 (brs, 1H, COOH).

5-Hydroxy-5-methyl-4-(4-carboxyphenyl)-1,5-dihydro-2H-pyrrol-2- one

(Z)-4-(1-Carboxy-3-oxobut-1-en-2-yl)benzoic acid was treated with trifluoroacetic acid (5 ml) at room temperature for 1 h. The solvent was

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removed under vacuum to yield a brown solid, presumably 5-hydroxy-5-methyl-4-(4- carboxyphenyl)-2(5H)furanone (0.1 g, 0.42 mmol), which was dissolved in thionyl chloride (5 ml) and stirred at room temperature for 24 h. The excess thionyl chloride was removed from the reaction mixture under high vacuum. The residue was stirred with ice-cold water for few minutes and then with aqueous ammonia (7 ml, 30 %) for 3 h. The excess ammonia was removed in vacuo and the residue was acidified with 2M

HCl (3 ml) to obtain a brown precipitate which was filtered under vacuum and subjected to flash chromatography with EtOAc/MeOH (9:1) to yield the title product as

1 a white solid (0.035 g, 35 %). M.p. 198-200 °C; H NMR (300 MHz, DMSO-d6) δ 1.56

(s, 3H, CH3), 3.7 (brs, 1H, OH), 6.7 (s, 1H, CH), 7.5 (s, 1H, ArH), 7.9 (m, 3H, ArH),

13 8.5 (s, 1H, NH), 13.0 (brs, 1H, COOH); C NMR (75.5 MHz,DMSO-d6) δ 26.7 (CH3),

107.0 (C), 117.0 (CH), 128.9 (2 x ArCH), 130.2 (ArC), 130.5 (2 x ArCH), 133.6 (ArC),

164.5 (C), 168.0 (C=O), 170.0 (C=O); HRMS (ESI) m/z calcd for C12H11NO4Na

256.0580 [M+Na]+, found 256.0580.

4-(4-Carboxy phenyl)-5-methyelene-1,5-dihydro-2H-pyrrol-2-one

(p-acid DHP)

A mixture of 5-hydroxy-5-methyl-4-(4-carboxyphenyl)-1,5-dihydro-

2H-pyrrol-2-one (0.2 g, 0.85 mmol) in borontrifluoride dietherate (2 ml) was stirred at room temperature for 24 h. The resulting reaction mixture was filtered under vacuum, washed with cold water and dried.

The residue was chromatographed on silica gel using EtOAc/MeOH (5:1) to yield the final product as a pale yellow solid (0.10 g, 50 %). M.p. 145-146 °C; 1H NMR (300

MHz, DMSO-d6) δ 4.91 (s, 1H, CH), 5.19 (s, 1H, =CH2), 6.42 (s, 1H, =CH2), 7.46-7.59

(m, 2H, ArH), 7.95-8.08 (m, 2H, ArH), 10.2 (1H, COOH); 13C NMR (75.5 MHz,

DMSO-d6) δ 98.0 (CH2), 123.7 (CH), 128.3 (2 x ArCH), 128.8 (2 x ArCH), 134.6 - 78 -

CHAPTER-3

(ArC), 135.3 (C), 144.1 (ArC), 149.3 (C), 167.7 (C=O), 170.1 (C=O); IR (ATR): υmax

3457, 3361, 3200, 2358, 2257, 2109, 1634, 1600, 1409, 1189, 1033, 924, 848, 770, 741

-1 -1 -1 cm ; UV (THF): λmax 236 nm (ε 8817 cm M ), 281 (3935); HRMS (ESI) m/z calcd for

+ C12H9NO3H 216.0655 [M+H] , found 216.0655.

3.2.3 Quorum Sensing Inhibition Assay

The QS inhibitory activity of the free DHP compounds was determined using the P. aeruginosa MH602 lasB reporter strain (PlasB::gfp(ASV)) by following the protocol developed by Hentzer et al. [192]. The QS monitor strain carries a plasmid containing a green fluorescent protein (GFP) reporter gene (gfp(ASV)) [348], which responds to the

AHL 3-oxo-dodecanoyl homoserine lactone (3oxo-C12-HSL) and expresses GFP when

QS is active [192]. To each well of the top row in a 96-well plate, 160 μl of Luria–

Bertani broth medium (LB10) and 40 μl of 5 mM test compound in DMSO were added.

The test compound was diluted by two-fold in LB10 in all subsequent wells. Then, 100

μl of a 100-times diluted overnight culture of P. aeruginosa MH602 in LB10 was added to all wells, and the final volume in each well was 200 μl. The plates were incubated at

37 °C for 15 h in a microplate reader (Wallac Victor, Perkin-Elmer), and every 30 min the plates were briefly shaken and measured for GFP expression (fluorescence: excitation 485 nm, emission 535 nm) and cell growth (OD600). Furanone 30 was used as a positive control due to its high QS inhibitory activity. The inhibitory effect of a

DMSO control (1 % of total volume) was also examined in similar fashion but no inhibitory effect on either GFP expression or cell growth was observed.

3.2.4 Attachment of 3-Aminopropyltriethoxysilane (APTS)

Glass coverslips (No. 1, diameter 13 mm D 263 M glass, ProSciTech, Australia) were first cleaned in freshly prepared piranha solution (3:1 v/v concentrated sulphuric acid to

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30 % hydrogen peroxide) at 100 °C for 1 h. After thorough rinsing with distilled water, the clean coverslips were rinsed once with absolute ethanol and air-dried.

The coverslips were then silanized using a previously developed method [291]. Briefly, the clean substrates were placed on a steel mesh within a glass vessel that contains a 3- aminopropyltriethoxysilane (APTS) solution (10 % v/v in dry toluene; 1 ml). The glass vessel was sealed and heated at 140 °C for 18 h. The coverslips were rinsed with dry toluene (×2), absolute ethanol and air-dried.

3.2.5 Plasma Functionalization of FEP by Allylamine

Prior to use, FEP sheets were cleaned with absolute ethanol and then rapidly dried with a jet of nitrogen. The FEP samples were plasma-activated using allylamine in a custom- built plasma reactor according to a previously established procedure (Figure 3.5) [349].

Figure 3.5: Plasma coating of allylamine monomer on blank FEP sheets.

Briefly, the reactor was comprised of a cylindrical glass chamber (height = 350 mm, diameter = 170 mm). The reactor contained a horizontal disc electrode of diameter 150 mm on the bottom and a 6 mm rod electrode on the top, separated by 150 mm. The substrates were placed on the lower rectangular electrode of the plasma reactor. The allylamine monomer (Sigma-Aldrich, 98 %) was degassed 5 times prior to deposition.

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The plasma deposition was carried out twice for 25 s with an initial pressure of 0.2 mbar

(200 kHz, 20 W). The activated FEP samples with primary amine groups were stored in a sterile container and used within a week.

3.2.6 Attachment of DHP via EDC/NHS Coupling

A solution containing DHP (20.2 µM), 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide hydrochloride (EDC, 101.2 µM), N-hydroxysuccinimide (NHS, 40.5 µM) and a small crystal of 4-dimethylaminopyridine (DMAP) in 1:1 ethanol/water was prepared. The amine-functionalized APTS glass (Figure 3.6A) or FEP (Figure 3.6B) surface was immersed in 1.5 ml of this solution and agitated overnight. The unreacted DHP was removed by extensively washing the samples with MilliQ water and absolute ethanol, and the surfaces were then air dried and stored in a clean sterile container.

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Figure 3.6: Specific attachment of acid-functionalized DHPs via EDC/NHS coupling

reaction on amine-functionalized (A) glass and (B) FEP surfaces.

3.2.7 X-Ray Photoelectron Spectroscopy

The attachment of DHP was analyzed by XPS using the same instrument setup as described in section 2.2.4.

3.2.8 Contact Angle Measurements

Contact angles were measured using the same method described in section 2.2.5. A minimum of fifteen measurements were made of five samples of each type.

3.2.9 Bacterial Adhesion Analysis

The bacterial strains used for this study were Staphylococcus aureus SA38 and

8 Pseudomonas aeruginosa PA01. Bacterial cultures were adjusted to 10 CFU/ml (OD660

= 0.1) before incubation of samples and stained with Live/Dead BacLight Viability Kits

L-7007 according to the same experimental procedures described in section 2.2.6.

Bacteria were fixed by adding 50 µl of 4 % formaldehyde to each sample and placed on the glass microscopy slide.

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Microscopic observation and image acquisition were performed with an Olympus

FV1200 Confocal Inverted Microscope. The bacterial cells that were stained green were considered to be viable, while those that stained red or both green and red were considered to be dead. Images from 10 representative areas on each of triplicate samples for each surface from a minimum of three independent experiments were taken and analysed using ImageJ software [333]. The results were reported as the average percentage coverage of live and dead cells of the fields of view.

3.2.10 Statistical Analysis of Data

Further analysis of the data was done by one-way analysis of variance (ANOVA) using

GraphPad Prism 7.03 software. Post hoc multiple comparisons were done using Tukey correction, and significance of results was set at 5 %.

3.3 Results

3.3.1 Synthesis of DHP Derivatives (DHP acids 1-4 and p-acid DHP)

The acid-functionalized DHP analogues (DHP acid 1-4) were synthesized via a tert- butyl acetate DHP intermediate which was in turn obtained by reacting various 5- hydroxy-5-methyl-4-aryl-1,5-dihydro-2H-pyrrol-2-ones 1 with tert-butyl chloroacetate

2 in presence of potassium hydroxide (Scheme 3.1). The proton NMR spectra of the intermediates 3, confirmed the presence of two tert-butyl acetate groups in each compound by the appearance of a singlet at 1.46 ppm for the CH3 groups. In the next step, the intermediate compounds were treated with trifluoroacetic acid, which facilitated cleavage of ether at C-5 resulting in removal of the ester group followed by dehydration to form the characteristic C-5 vinylic double bond. At the same time, the N-

1 ester group underwent hydrolysis in presence of trifluoroacetic acid to convert to carboxylic acid. The disappearance of the peaks for the C-5 acetate group and N-1 tert- - 83 -

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butyl group in the NMR spectra indicated completion of the reaction. The crude products were purified to obtain 2-(5-methylene-2-oxo-4-aryl-1,5-dihydro-2H-pyrrol-1- yl)acetic acid 4 (DHP acids 1-4) in 26–33 % yields.

Scheme 3.1: Synthesis of acid derivatives of DHPs (DHP acids 1-4).

The p-acid DHP compound was synthesized through a series of steps as shown in

Scheme 3.2. 4-(2-Oxopropyl)benzoic acid 5 was reacted with glyoxylic acid 6 at 75–80

°C for 5 h to generate the di-acid compound in 50 % yield after purification. The identity of the di-acid product, (Z)-4-(1-carboxy-3-oxobut-1-en-2-yl)benzoic acid 7, was confirmed by proton NMR spectroscopy. Cyclization of the di-acid compound to form the corresponding furanone compound 8 with a hydroxyl group at C-5 was

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accomplished in the next step by using trifluoroacetic acid. The reaction was monitored by TLC which confirmed the formation of the hydroxyl furanone after 1 h.

Scheme 3.2: Synthetic route for p-acid DHP.

The next step involved a lactone to lactam ring conversion which was initially attempted by reaction of the hydroxyl furanone with aqueous NH3. However, this was unsuccessful due to the formation of multiple unexpected products. Instead, the C-5 hydroxyl group was replaced with chlorine by using thionyl chloride as the chlorinating agent. To avoid concomitant chlorination of the carboxylic acid group, the reaction was carried out at room temperature. The reaction was continuously monitored by TLC which indicated the completion of reaction after 24 h. The 5-chlorofuranone compound

9 successfully underwent the lactone to lactam conversion with aqueous ammonia giving the DHP product, 5-hydroxy-5-methyl-4-(4-carboxyphenyl)-1,5-dihydro-2H- pyrrol-2-one 10, after 3 h. The formation of the DHP was confirmed by proton NMR, - 85 -

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which exhibited a characteristic broad peak at 8.5 ppm representing the –NH group of the lactam ring.

The final step involved dehydration of the hydroxyl DHP 10 to regenerate the methylene group at the C-5 position. Dehydration was carried out using borontrifluoride etherate as the dehydrating agent. The proton NMR analysis of the product showed the presence of two singlet peaks at 5.19 ppm and 6.42 ppm corresponding to the two protons of the C-5 double bond, indicating successful dehydration to form 4-(4-carboxy phenyl)-5-methyelene-1,5-dihydro-2H-pyrrol-2-one 11 (p-acid DHP).

3.3.2 Quorum Sensing Inhibitory Activity of Free DHPs

To determine whether free DHPs are capable of inhibiting QS system of P. aeruginosa, planktonic cultures of P. aeruginosa MH602 harbouring the lasB-gfp(ASV) fusion were incubated with DHPs at different concentrations. The percentage of QS inhibition of the

DHP compounds are presented in Table 3.1.

Table 3.1: QS inhibitory activity (%) against P. aeruginosa MH602 lasB reporter strain at different concentrations of DHP derivatives; aGrowth inhibition ≤ 10%; bGrowth inhibition 11–30 %.

QS Inhibition (%) Compound 250 µM 125 µM 62.5 µM

DHP acid-1 56.3 ± 0.4b 40.4 ± 1.2a 27.6 ± 1.7a

DHP acid-2 55.3 ± 3.8a 34.2 ± 2.3a 33.5 ± 1.2a

DHP acid-3 57.5 ± 0.7b 35.2 ± 2.0a 33.0 ± 1.8a

DHP acid-4 53.6 ± 1.5a 35.1 ± 1.1a 31.0 ± 1.7a

p-acid DHP 68.6 ± 1.1b 38.5 ± 1.1b 35.0 ± 2.4a

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The expression of GFP was suppressed by all the DHPs in a dose-dependent manner indicating that the free compounds interfered with the QS system of P. aeruginosa

MH602. The p-acid DHP showed the highest percentage inhibition of 68.6 % at 250 µM amongst the synthesized DHPs, followed by DHP acid-3 and DHP acid-1 which showed

57.5 % and 56.3 % inhibition. At 125 µM, DHP acid-1 (40.4 %) and p-acid DHP (38.5

%) had maximum QS inhibitory effect while the remaining DHPs displayed similar level of inhibition. The inhibition values did not vary significantly for halogenated

DHPs (DHP acid 2-4) at all concentrations (62.5–250 µM). Overall, p-acid DHP displayed maximum inhibition.

3.3.3 XPS Characterization of Glass and FEP Surfaces

DHP surfaces were subsequently developed by incorporating the active DHP compounds by EDC/NHS coupling reaction. To determine the surface composition,

XPS analysis was carried out after every modification step. The elemental composition of the glass surfaces are shown in Table 3.2 respectively.

The changes in the carbon, nitrogen and halogen composition on the glass surface before and after attachment of APTS and DHPs indicated successful surface modification (Table 3.2). The carbon and nitrogen concentration increased drastically from 6.6 % and 0.6 % to 45.4 % and 8.1 %, respectively, after functionalization of surface by APTS [291,350]. The subsequent attachment of DHP acids 1-4 and p-acid

DHP further increased carbon content by 1.8–6.3 % and 4.8 % respectively compared to the APTS control. Similarly, the nitrogen concentration also increased by 0.7–2.7 % after reaction with DHP acids and by 1.1 % for p-acid DHP. Furthermore, the halogens detected in the XPS analysis confirmed the attachment of halogenated DHP acids 2-3 on the APTS glass surface. As indicated by the halogen content, the highest attachment

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efficiency was displayed by DHP acid-4 (0.91 % Br) which was approximately 2-5 times higher than the fluorine substituted DHPs. This was followed by DHP acid-2 (0.4

% ortho-F) which had roughly twice the amount of fluorine as DHP acid-3 (0.17 % para-F). In absence of halogen the surface coverage of DHP acid-1 and p-acid DHP was determined by the carbon and nitrogen values which indicate both the non-halogenated

DHPs have similar coverage on the APTS glass surface.

Table 3.2: XPS elemental composition of glass surfaces.

Surface % C % N % Halogen Blank 6.6 0.6 - APTS 45.4 8.1 - DHP acid-1 48.6 8.9 - DHP acid-2 49.3 8.8 0.40 % F DHP acid-3 47.2 10.8 0.17 % F DHP acid-4 51.7 10.3 0.91 % Br p-acid DHP 50.2 9.2 -

The same attachment strategy was then employed to incorporate DHP acid-1 and p-acid

DHP on FEP substrates. The blank FEP surfaces were found to contain 31.8 % carbon and 67.9 % fluorine with traces of oxygen (0.2 %) (Table 3.3). After treatment with allylamine plasma, the polymer surface showed the presence of nitrogen (12.3 %) while the carbon and oxygen content increased to 74.0 % and 13.5 % respectively and the fluorine percentage dropped down to 0.2 %. The presence of oxygen after non-oxygen plasma treatment is a common phenomenon, possibly due to the formation of free radicals on the surface which react with oxygen present in the chamber during plasma treatment or with atmospheric oxygen when exposed to air [346,347,351,352]. The

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decrease in the fluorine content after plasma deposition could either be due to defluorination of the polymer surface induced by secondary electrons created during

XPS analysis [353,354] or the presence of a thick coating of allylamine on the FEP surface [345,355]. The attachment of DHPs onto allylamine-coated FEP surfaces was confirmed by increase in carbon content by 3.0–3.9 %, decrease in nitrogen and oxygen by 0.8–2.1 % and 1.8–2.1 % respectively. The values indicated both the DHPs have similar attachment efficiency.

Table 3.3: XPS elemental composition of FEP surfaces.

Surface % C % N % O % F

FEP-Blank 31.8 - 0.2 67.9 FEP-Amine 74.0 12.3 13.5 0.2 FEP-DHP acid-1 77.0 11.5 11.4 0.1 FEP-p-acid DHP 77.9 10.2 11.7 0.2

The curve fitting results for C 1s and N 1s regions and proposed assignments based on chemical shifts are shown in Table 3.4. The C 1s spectrum for APTS glass showed three carbon species assigned to an aliphatic carbon at 284.9 eV, C-N at 285.9 eV and

C=O at 287.9 eV. For the DHP-treated surfaces, a new peak emerged at ~288.6 eV corresponding to the amide bond (N-C=O), indicating successful reaction of DHPs with the APTS surface. In the case of FEP surfaces, the blank showed the presence of four distinct carbon species, C-C, CF2, CF3 and O-CF2 at 284.8 eV, 290.5 eV, 292.4 eV and

294.4 eV respectively. The amine plasma treatment generated additional carbon components assigned to C-C (284.8 eV), C-N (285.8 eV), C-O (286.3 eV), C=O (287.8 eV) and N-C=O (288.8 eV). The oxidation reactions on the FEP surface during plasma treatment formed oxygen functional groups (C-O and C=O). Finally, the successful

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attachment of DHPs on the amine FEP surface was determined by the changes in the C

1s spectra. The increase in peak area for the N-C=O bond confirmed the attachment of

DHPs.

Table 3.4: XPS binding energies for C 1s and N 1s and proposed assignments with percentage peak intensities.

C 1s N 1s Binding Peak Binding Peak Surface energy Assignment area energy Assignment area (eV) (%) (eV) (%)

284.9 C-C 63.0 399.5 NH2 80.2

APTS + 285.9 C-N 25.5 401.4 NH3 , Tertiary N 19.7 287.9 C=O 11.4

284.8 C-C 57.0 399.5 NH2 34.0 DHP acid-1 285.8 C-N 21.0 400.2 N-C=O 56.6 + 287.8 C=O 11.6 401.6 NH3 , Tertiary N 9.4

288.6 N-C=O 10.4

284.8 C-C 56.7 399.5 NH 30.9 DHP 2 acid-2 285.8 C-N 21.2 400.2 N-C=O 58.3

+ 287.8 C=O 12.1 401.6 NH3 , Tertiary N 10.8

288.6 N-C=O 10.0

284.8 C-C 55.8 399.4 NH2 21.7 DHP 285.8 C-N 24.2 400.0 N-C=O 67.6 acid-3 + 287.8 C=O 10.0 401.6 NH3 , Tertiary N 10.7 288.7 N-C=O 10.0

284.8 C-C 60.0 399.4 NH2 36.8 285.8 C-N 22.8 400.0 N-C=O 59.7 DHP + acid-4 287.6 C=O 8.8 401.4 NH3 , Tertiary N 3.5 288.6 N-C=O 8.4

p-DHP 284.8 C-C 67.2 399.6 NH2 61.6 acid 285.8 C-N 12.0 400.1 N-C=O 31.3 - 90 -

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+ 287.9 C=O 14.1 401.5 NH3 , Tertiary N 7.1 288.4 N-C=O 6.6

284.8 C-C 0.8

FEP- 290.5 CF2 3.5 Blank 292.4 CF3 91.3

294.4 O-CF2 4.4

284.8 C-C 67.7 399.4 NH2 84.4

FEP- 285.8 C-N 5.1 400.3 N-C=O 8.5 Amine + 286.3 C-O 19.0 401.4 NH3 , Tertiary N 7.1 287.8 C=O 7.4

288.8 N-C=O 0.8

284.7 C-C 67.5 399.1 NH2 81.9 FEP- DHP 286.1 C-O 23.7 400.0 N-C=O 18.1 acid-1 287.7 C=O 7.3 288.9 N-C=O 1.5

284.7 C-C 62.7 399.1 NH2 76.0 FEP- p-acid 286.1 C-O 23.7 400.1 N-C=O 24.0 DHP 287.7 C=O 10.0 288.8 N-C=O 3.6

The N 1s high resolution scan for all the glass samples showed peaks at around 399.5

+ eV and 401.5 eV which were assigned to an amine bond and NH3 /tertiary nitrogen respectively (Table 3.4). For the DHP glass surfaces, the peak at ~400 eV was attributed to the nitrogen from the amide bond (N-C=O), which is an indication of surface modification by the DHP compounds. The ratio of the nitrogen species for all the glass surfaces changed after DHP modification, notably with an increase in peak area for the amide bond. For the FEP-amine surface, the N 1s signal at 399.4 eV is an indication for presence of amine groups on the surface. The high density of the amine - 91 -

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groups on the surface formed after allylamine deposition is consistent with literature

[345]. The amide group at 400.3 eV (8.5 %) is also present on the FEP-amine surface due to post-plasma oxidation that converts amines to amides [356]. The reaction of the

DHP acid-1 and p-acid DHP on the FEP-amine surface was confirmed by the changes in the proportion of the nitrogen species where the peak area for amide bond increased from 8.5 % to 18.1 % and 24.0 % respectively.

3.3.4 Contact Angle Measurements

Static water contact angle measurement was employed to determine the changes in hydrophobicity of the surface after each modification step. The hydrophobicity of materials is a useful parameter that is correlated with cell-biomaterial interfacial interactions [357,358]. The contact angle of the uncoated glass surface was 19° (Table

3.5). After modification with APTS, a significant increase in surface hydrophobicity was observed, with a contact angle value of 72°. The contact angle values remained approximately the same (68–75°) after subsequent attachment of DHPs due to the relatively hydrophobic nature of the DHPs.

Table 3.5: Contact angle measurements of glass and FEP substrates.

Contact angle Surface (°) (±1) Blank 19 APTS 72 DHP acid-1 71 DHP acid-2 73 DHP acid-3 74 DHP acid-4 75 p-acid DHP 68

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FEP-Blank 108 FEP-Amine 78 FEP-DHP acid-1 81 FEP-p-acid DHP 77

In contrast to blank glass, the FEP-blank surface was found to be highly hydrophobic with high contact angle value of 108° which is in accordance with reported values [359–

361] (Table 3.5). The contact angle of the surface decreased drastically to 78° after allylamine plasma treatment, however, the surface is still very hydrophobic. This finding is consistent with reports where functionalization of untreated FEP (blank) resulted in reduction in hydrophobicity i.e. decrease in water contact angle value [361–

363]. The surface coated with DHP acid-1 (81°) and p-acid DHP (77°) showed similar contact angle to the FEP-amine surface.

3.3.5 Antibacterial Activity

The adhesion of S. aureus and P. aeruginosa to the modified surfaces was investigated using fluorescence microscopy by staining the surfaces using the BacLight Live/Dead

Bacterial Viability kit. Representative micrographs of S. aureus and P. aeruginosa adhesion on control and DHP-coated surfaces are shown in Figure 3.7 and Figure 3.8 respectively. Extensive bacterial colonization was observed on untreated controls,

APTS and FEP-amine, by both S. aureus and P. aeruginosa. Reduction in cell adhesion for both bacterial strains was observed on all DHP substrates compared to their respective control samples. The images were analyzed to determine the relative proportion of live and dead bacteria (stained green and red) on each surface and the results for S. aureus and P. aeruginosa are shown in Figure 3.9 and Figure 3.10 respectively.

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S. aureus

A – APTS D – FEP-Amine

B – DHP acid-1 E – FEP-DHP acid-1

C – p-acid DHP F – FEP-p-acid DHP

Figure 3.7: Confocal microscopic images of S. aureus adhered to APTS and FEP-amine process control (A and D), DHP acid-1 surfaces (B and E) and p-acid DHP surfaces (C and F). Live bacterial cells stained green and dead bacteria stained red. Magnification

200×. Scale bar = 100 µm.

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P. aeruginosa

A – APTS D – FEP-Amine

B – DHP acid-1 E – FEP-DHP acid-1

C – p-acid DHP F – FEP-p-acid DHP

Figure 3.8: Confocal microscopic images of P. aeruginosa adhered to APTS and FEP- amine process control (A and D), DHP acid-1 surfaces (B and E) and p-acid DHP surfaces (C and F). Live bacterial cells stained green and dead bacteria stained red.

Magnification 200×. Scale bar = 100 µm.

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Results of image analysis showed high coverage of S. aureus on the untreated surfaces, blank glass and FEP (Figure 3.9). The APTS and FEP-amine control surfaces displayed similar levels of adhesion of more than 13 %, suggesting that APTS and allylamine modification do not have a significant effect on bacterial adhesion. In contrast, the

DHP-coated glass surfaces displayed significantly lower bacterial coverage compared to the APTS-coated glass surface. Specifically, the DHP acids 1-4 and p-acid DHP displayed reductions in surface coverage by 64.2–75.8 % and 68.2 % respectively compared to APTS (p < 0.001). Among the various DHP acids, DHP acid-1 (75.8 ± 0.8

%), DHP acid-2 (75.8 ± 0.5 %) and DHP acid-4 (74.2 ± 0.8 %) showed slightly higher bacterial reduction than DHP acid-3 (64.2 ± 0.4 %), however significant difference in activity was not observed. The p-acid DHP coated glass also showed similar efficacy compared to other DHPs, which reduced 68.2 ± 0.4 % of S. aureus adhesion.

For the FEP surfaces, DHP acid-1 and p-acid DHP displayed reduction of 52.7 ± 0.4 % and 70.0 ± 0.8 % compared to the FEP-amine control (p < 0.001), in which the p-acid

DHP performed significantly better than DHP acid-1 (p < 0.05) (Figure 3.9). There was no significant increase in the proportion of dead (red-staining) cells for all modified samples indicating the surfaces inhibit bacterial growth rather than killing, and therefore it is unlikely for bacteria to develop resistance to the DHPs [33,364].

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Bacterial coverage - S. aureus

18 16 14 12 Dead 10 bacteria × 8 * * * * Live 6 * * * bacteria

4 Bacterial coverage (%) coverage Bacterial 2 0

Figure 3.9: Percentage surface coverage of live and dead bacteria for S. aureus (mean

± standard error of mean); *indicates p < 0.001 compared to control; ×indicates p <

0.05 compared to FEP-DHP acid-1 modified surface.

Similar to S. aureus, P. aeruginosa showed high bacterial coverage on APTS and FEP- amine control surfaces (Figure 3.10), as well as blank glass and FEP surfaces

Significant reductions in adherent bacterial cells was observed for all the DHP-modified glass surfaces when compared to the APTS control (50.4–71.3 % reduction for DHP acids 1-4 and 60.1 % for p-acid DHP; p < 0.001). Out of these, the most potent compound was DHP acid-3 which showed the highest reduction of 71.3 ± 0.2 % amongst all the glass surfaces (p < 0.05). There were no statistical differences between the DHP acid-1 and p-acid DHP (60.1 ± 0.3 %) modified glass samples, however when coupled on the FEP-amine surface, p-acid DHP exhibited significantly greater reduction

(70.0 ± 0.3 %; p < 0.05) in bacterial adhesion compared to FEP-DHP acid-1 (50.3 ± 0.1

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%). The percentage of dead bacteria (red-staining cells) did not increase for any of the modified samples.

Bacterial Coverage - P. aeruginosa

8 Dead * * * bacteria 6 * ^ * × * * Live 4 bacteria

Bacterial coverage (%) coverage Bacterial 2

Figure 3.10: Percentage surface coverage of live and dead bacteria for P. aeruginosa

(mean ± standard error of mean); *indicates p < 0.001 compared to control; ^indicates p < 0.05 compared to DHP acids and p-acid DHP-modified glass surfaces; ×indicates p

< 0.05 compared to FEP-DHP acid-1 modified surface.

3.4 Discussion

In the current study, the antibacterial efficacy of DHPs attached from different points on the molecule to glass and FEP surfaces was investigated. The DHP surface coatings were tested for their ability to reduce the adherent bacterial cells of S. aureus and P. aeruginosa, while the free DHPs were also tested for their QS inhibitory activity.

Additionally, the DHP-coated surfaces were characterized using XPS and contact angle measurements. - 98 -

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The DHP derivatives synthesized for surface attachment were found to reduce QS signalling of the reporter strain, P. aeruginosa MH602, in a concentration-dependent manner without affecting the bacterial growth. The results indicate that the modification of DHPs via nitrogen of lactam ring or C-4 phenyl ring did not affect the activity of the compounds. It was observed that p-acid DHP resulted in higher QS inhibition compared to N-substituted DHPs. This can be attributed to the presence of free heterocyclic lactam ring of p-acid DHP that resulted in higher antibacterial activity.

The glass surfaces were then modified with DHP acids 1-4 and p-acid DHP using a specific attachment strategy via the EDC/NHS coupling reaction. XPS analysis of the coatings demonstrated that DHPs were successfully attached onto the surfaces. Analysis of the halogen content indicated that DHP acid-4 had the highest attachment efficiency, followed by DHP acid-2 and DHP acid-3. Similar to the QS inhibition results, the DHP acids after attachment showed similar activity, except DHP acid-3, which exhibited higher activity against P. aeruginosa. The activity of the surfaces modified with DHP acid-1 and DHP acid-4 was consistent with a previous study where analogues of DHPs studied in this chapter were functionalized with an acrylate group at the N-1 position and grafted onto APTS glass via Michael addition reaction [291]. Similarly, the efficacy of fluorinated DHP acids was also found to be comparable with the same fluorinated

DHPs immobilized by Michael addition reaction. The para-fluoro DHP even at low surface concentration exhibited higher activity against P. aeruginosa compared to ortho-fluoro compound when attached on the surface by EDC coupling and Michael addition reaction.

Furthermore, covalently attached DHPs via non-specific azide chemistry exhibited higher reduction in bacterial adhesion when compared to DHPs specifically attached via

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the EDC coupling reaction. For instance, the para-bromo DHP compound attached via azide chemistry exhibited low surface attachment (0.35 % para-Br) but displayed high bacterial reductions (93.4 % and 71.2 % against S. aureus and P. aeruginosa respectively) [350], while the same para-bromo DHP (DHP acid-4) when attached specifically via EDC/NHS coupling had nearly three-fold higher surface attachment

(0.91 % para-Br) but exhibited only 74.2 % and 55.0 % reductions of S. aureus and P. aeruginosa adhesion. This indicates that the choice of attachment methodology used, which affects the orientation of the DHP on the surface, could have a significant influence on antibacterial efficacy.

In order to investigate the effect of molecular orientation on the surface, the acid- functionalized DHP molecule (p-acid DHP) was attached on the surface through the pendant phenyl ring. On the glass substrates, the p-acid DHP exhibited significant reductions of 68.2 % for S. aureus and 60.1 % for P. aeruginosa cells (p < 0.001).

However, the activity of p-acid DHP did not vary significantly from DHP acids 1-4, except against P. aeruginosa where DHP acid-3 sample was found to be significantly better than p-acid DHP (p < 0.05). The results suggests that despite high activity of p- acid DHP in solution, the covalent attachment reduced the activity of the compound, and the antibacterial efficacy of a coated surface may not always be consistent with the activity of a free compound.

To further investigate the activity of surface bound p-acid DHP on different substrates and reproducibility of above results, p-acid DHP and DHP acid-1 were grafted onto a new biomaterial, FEP surfaces. DHP acid-1 was chosen from the group of N-substituted

DHP acid compounds due to its broad-spectrum antibacterial activity on glass and structural similarity to p-acid DHP (i.e., both compounds do not contain halogens). Both

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the DHP compounds showed similar attachment efficiency on the FEP surface as indicated by their elemental composition in the XPS analysis data. However, the p-acid

DHP sample exhibited higher activity than DHP acid-1 at reducing adhesion of both strains of bacteria on the FEP surface (p < 0.05). The difference in efficacy for DHP acid-1 and p-acid DHP on glass and FEP could be due to the difference in attachment efficiency that varies from one surface to another. Also, changes in the surface physical properties such as surface roughness is known to have a strong influence on the rate and extent of bacterial adhesion [80,365,366].

Research has also shown that bacteria adhere more easily to non-polar surfaces than to hydrophilic substrates [367,368], that is, higher surface hydrophobicity results in higher biofilm development [369,370]. However, bacterial adhesion was significantly reduced after DHP attachment, even though the glass and FEP DHP-coated substrates were hydrophobic. This result implies that the reduction in bacterial colonization was caused by the antibacterial activity of the surface attached DHP compounds and not due to the surface hydrophobicity.

Taken together, the modification of the DHP at the lactam N-1 position was less favourable than modification at the phenyl ring, both in solution as well as after covalent attachment on FEP. However, in the context of the DHP modified glass surfaces, the change in molecular orientation of DHP resulted in similar biological activity. This suggests that the free lactam and C-4 phenyl ring of the DHP exposed towards the bacteria are equally effective in reducing bacterial adhesion on glass surfaces. This would explain the high activity of DHP-coated glass surfaces generated in Chapter 2 despite low surface concentration.

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3.5 Conclusion

In this chapter, the effect of molecular orientation of surface-immobilized DHPs on its biological activity was evaluated by attaching different parts of the molecule to glass and FEP surfaces using a specific attachment strategy. The N-substituted DHP acids 1-4 and the p-acid DHP were equally effective in reducing bacterial colonization of S. aureus and P. aeruginosa on glass substrates, while higher activity was obtained when

DHP was attached to the FEP surface via the C-4 pendant phenyl ring, thereby exposing the potent lactam ring of DHP towards the bacteria. This DHP-coated surface can be used as a base for examining other parameters such as effect of flexibility and length of the surface linker that could affect the activity of DHPs after surface attachment.

Additionally, the structure-activity relationship of surface-bound DHP is still not very clear and requires further investigation for development of DHPs as antibacterial surface coatings.

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Antibacterial Activity of DHP Without Exocyclic Double Bond

CHAPTER-4

CHAPTER-4 Antibacterial Activity of DHP Without Exocyclic Double Bond

4.1 Introduction

The structural modification of existing QS inhibitors to generate more diverse and potent antibacterial compounds is a popular strategy. All active DHPs reported to date have been mainly generated from the parent scaffold by modifying the C-3 alkyl chain, modifying the substituents at C-4 and N-1, or altering the bromination pattern of the C-5 methylene group (Figure 4.1) [182,200–202,344]. Considerable effort has been made to study the activity of these compounds, but to date no study has focused on determining the effect of the absence of the exocyclic double bond at C-5 on the biological activity of the lactams.

Figure 4.1: Structural modifications of synthetic DHPs.

Recently, studies were conducted on brominated furanones in order to identify the structural elements that are important for inhibiting bacterial biofilms. Structure-activity

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relationship analysis showed that furanones lacking an exocyclic vinyl group displayed very low or no activity [309,310]. Molecular docking of furanones by Sabbah et al. against the LuxR model suggested that biological activity was dependent upon interactions between the exomethylene group and the residues Trp94 and Leu118

[310,371]. In the absence of this group, furanones cannot form these interactions with

LuxR and hence their inhibitory activity would be lost. Therefore, in order to determine whether DHPs behave in a similar manner to the furanone compounds, analogues of

DHPs without an exocyclic double bond were synthesized (Figure 4.2) and tested for their antibacterial activity after covalently immobilizing them on the surface.

Figure 4.2: Structural modification of DHPs at the C-5 position of the lactam ring.

One way to generate DHP analogues without the C-5 methylene group is to convert mucohalic acids into pyrrol-2-ones (lactone to lactam conversion) by reductive amination using a suitable amine and reducing agent (Figure 4.3A). Mucohalic acids are inexpensive starting materials with multiple functional groups. They are known to exist in both the pseudo-unsaturated γ-lactone form 1 and the open chain aldehyde-acid form 2, and can react as either tautomeric form depending on the conditions (Figure

4.3B) [372,373]. These highly functionalized compounds are versatile building blocks in organic synthesis and have recently attracted attention for synthesizing various antibacterial compounds [374–376], anticancer agent [377], anti-seizure agent [378] and anti-inflammatory drugs [379]. In this work, six analogues of N-substituted DHPs were generated from mucohalic acids and acid/amine-substituted aryl amines in a single step or through an intermediate according to a literature method with some modifications

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[380]. The synthesized DHPs without the conjugated double bond system at C-5 were functionalized with acid or amine groups for covalent attachment on surfaces.

(A)

(B)

Figure 4.3: (A) Lactone to lactam conversion of mucohalic acids by reductive

amination reaction; (B) Equilibria of mucohalic acids.

To allow covalent attachment of DHPs, surfaces were first functionalized with 3- aminopropyltriethoxysilane (APTS) to form amine groups. To develop surfaces with acid groups, succinic anhydride was coupled with the amine groups of the APTS surface [381–383]. The acid and amine-functionalized glass surfaces were reacted with their respective amine/acid DHPs by EDC coupling (Figure 4.4). The resulting DHP- coated surfaces were characterised by X-ray photoelectron spectroscopy (XPS) and contact angle measurements. The QS activity of free compounds and bacterial adhesion of the surfaces was subsequently analysed in order to assess the importance of the exomethylene group on biological activity.

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Figure 4.4: Immobilization of acid- and amine-functionalized DHPs on amine and

acid-terminated glass coverslips respectively via EDC/NHS coupling reaction.

4.2 Materials and Methods

4.2.1 General

All chemical reagents were purchased from commercial sources (Alfa-Aesar and Sigma

Aldrich) and used without further purification. Solvents were sourced from commercial sources and used as obtained. Reactions were performed using oven-dried glassware under an atmosphere of nitrogen and in anhydrous conditions (as required). Room temperature refers to the ambient temperature (22–24 °C). Yields refer to chromatographically and spectroscopically pure compounds unless otherwise stated.

200 mm glass plates coated with Merck 60GF254 silica gel. Infrared spectra were - 106 -

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recorded using a Cary 630 FTIR spectrophotometer. Ultraviolet spectra were measured using a Cary 100 Bio UV-visible spectrophotometer in the designated solvents and data reported as wavelength (λ) in nm and absorption coefficient (ε) in cm-1M-1. High resolution mass spectrometry was performed by the Bioanalytical Mass Spectrometry

Facility, UNSW. Melting points were obtained using Mel-Temp melting point apparatus and are uncorrected. Proton and carbon NMR was recorded in designated solvents using

Bruker DPX 300 or a Bruker Avance 400 spectrometer as designated. Chemical shifts

(δ) are quoted in parts per million (ppm), to the nearest 0.01 ppm and internally referenced relative to the solvent nuclei. 1H NMR spectral data are reported as follows: chemical shift in ppm; multiplicity in broad (br), singlet (s), doublet (d), triplet (t), quartet (q), multiplet (m) or a combination of these (e.g. dd, dt etc.)]; coupling constant

(J) in hertz, integration, proton count and assignment.

4.2.2 Synthesis of Acid- and Amine-DHP Derivatives

N-(4-carboxyphenyl)-3,4-dichloro-1,5-dihydro-2H-pyrrol-2-one

(DHP phenyl acid-1)

The title compound was synthesized by first dissolving mucochloric acid (1 g, 5.91 mmol) in 5:3 v/v dichloromethane/glacial acetic acid

(12 ml). To this solution, p-aminobenzoic acid (0.81 g, 5.91 mmol) in dichloromethane (8 ml) was added followed by sodium triacetoxyborohydride (3.76 g, 17.75 mmol). The reaction mixture was stirred at room temperature for 18 h, during which a yellow precipitate was evident. The mixture was filtered under vacuum and washed with dichloromethane and distilled water to yield a

1 yellow solid (0.6 g; 37 %). M.p. 229 °C; H NMR (300 MHz, DMSO-d6) δ 4.91 (s, 2H,

13 CH2), 7.84-7.88 (m, 2H, ArH), 7.96-8.01 (m, 2H, ArH), 12.8 (brs, 1H, COOH); C

NMR (75.5 MHz, DMSO-d6) δ 53.9 (CH2), 118.3 (2 x ArCH), 124.2 (ArCN), 126.7 - 107 -

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(ArC), 130.9 (2 x ArCH), 142.4 (CCl), 142.8 (CCl), 162.8 (C=O), 167.1 (C=O); IR

(ATR): υmax 2939, 2814, 2659, 2537, 1679, 1604, 1516, 1425, 1375, 1279, 1042, 934,

-1 -1 -1 860, 769, 741, 718 cm ; UV (THF): λmax 236 nm (ε 5338 cm M ), 289 (2736); HRMS

+ (ESI) m/z calcd for C11H7Cl2NO3Na 293.9695 [M+Na] , found 293.9696.

N-(3-carboxyphenyl)-3,4-dichloro-1,5-dihydro-2H-pyrrol-2-one

(DHP phenyl acid-2)

The title compound was synthesized by reacting mucochloric acid (1 g, 5.91 mmol), m-aminobenzoic acid (0.81 g, 5.91 mmol) and sodium triacetoxyborohydride (3.76 g, 17.75 mmol) in 5:3 v/v dichloromethane/glacial acetic acid (12 ml) at room temperature for 18 h, during which a yellow precipitate was evident. The mixture was filtered under vacuum and the filtered solid was purified by flash chromatography. The solid was then recrystallized in methanol after chromatography to yield the pure title product as a white solid (0.54 g; 34 %). M.p. 214-216 °C; 1H NMR (300 MHz, DMSO- d6) δ 4.93 (s, 2H, CH2), 7.55 (t, J = 7.98 Hz, 1H, ArH), 7.74 (tt, J = 7.98 and 1.48 Hz,

1H, ArH), 7.89-7.93 (m, 1H, ArH), 8.34 (t, J = 1.8 Hz, 1H, ArH), 13.13 (brs, 1H,

13 COOH); C NMR (75.5 MHz, DMSO-d6) δ 54.0 (CH2), 120.0 (ArCH), 123.4 (ArCN),

125.8 (ArC), 129.9 (ArCH), 132.1 (ArCH), 138.9 (ArCH), 142.4 (2 x CCl), 162.7

(C=O), 167.3 (C=O); IR (ATR): υmax 2969, 2824, 2654, 2539, 1699, 1586, 1491, 1433,

-1 1382, 1313, 1272, 1158, 1051, 938, 899, 818, 757, 739, 677 cm ; UV (THF): λmax 236

-1 -1 nm (ε 20671 cm M ), 288 (32,346); HRMS (ESI) m/z calcd for C11H7Cl2NO3Na

293.9695 [M+Na]+, found 293.9698.

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N-(4-carboxyphenyl)-3,4-dibromo-1,5-dihydro-2H-pyrrol-2-one

(DHP phenyl acid-3)

Mucobromic acid (3 g, 11.63 mmol), p-aminobenzoic acid (1.59 g,

11.63 mmol) and sodium triacetoxyborohydride (7.39 g, 34.9 mmol) in 5:3 v/v dichloromethane/glacial acetic acid (30 ml), were stirred and gently heated at 30 °C for 3 h during which time a precipitate was evident. The mixture was filtered under vacuum and the solid was recrystallized in

1:9 acetone/methanol to get the desired product as a white solid (0.296 g, 7 %). M.p.

1 221 °C; H NMR (300 MHz, DMSO-d6) δ 4.9 (s, 2H, CH2), 7.83-7.98 (m, 4H, ArH),

13 12.8 (brs, 1H, COOH); C NMR (75.5 MHz, DMSO-d6) δ 57.2 (CH2), 118.2 (2 x

ArCH), 120.0 (ArCN), 126.6 (ArC), 130.9 (2 x ArCH), 137.4 (CBr), 142.5 (CBr), 163.7

(C=O), 167.1 (C=O); IR (ATR): υmax 2811, 2659, 2535, 2112, 1679, 1601, 1516, 1423,

-1 1371, 1275, 1188, 1145, 1017, 929, 889, 757, 704 cm ; UV (THF): λmax 245 nm (ε

-1 -1 14,965 cm M ), 288 (15,575); HRMS (ESI) m/z calcd for C11H7Br2NO3Na 381.8685

[M+Na]+, found 381.8684.

N-(3-carboxyphenyl)-3,4-dibromo-1,5-dihydro-2H-pyrrol-2-one

(DHP phenyl acid-4)

The title compound was synthesized by following the same method for DHP phenyl acid-2 from the corresponding mucobromic acid to afford the product as pale yellow solid (0.36

1 g, 26 %). M.p. 186 °C; H NMR (300 MHz, DMSO-d6) δ 4.93

(s, 2H, CH2), 7.55 (t, J = 8.0 Hz, 1H, ArH), 7.73 (tt, J = 8.0 and 1.47 Hz, 1H, ArH),

7.89-7.93 (m, 1H, ArH), 8.33 (t, J = 1.8 Hz, 1H, ArH), 13.09 (brs, 1H, COOH); 13C

NMR (75.5 MHz, DMSO-d6) δ 57.3(CH2), 119.9 (ArCH), 123.3 (ArCN), 125.7 (ArC),

129.8 (ArCH), 136.9 (ArCH), 139.0 (ArCH), 146.0 (2 x CBr), 163.5 (C=O), 167.4 - 109 -

CHAPTER-4

(C=O); IR (ATR): υmax 2821, 2551, 2321, 1698, 1584, 1490, 1425, 1380, 1312, 1289,

-1 - 1227, 1151, 1032, 939, 901, 840, 757, 673 cm ; UV (THF): λmax 239 nm (ε 15,683 cm

1 -1 + M ), 291 (6639); HRMS (ESI) m/z calcd for C11H7Br2NO3Na 381.8685 [M+Na] , found 381.8685.

N-(4’-tert-butylphenylcarbamate)-3,4-dichloro-1,5-dihydro-2H-pyrrol-2-one

(boc-DHP phenyl amine-1)

Mucochloric acid (0.9 g, 5.36 mmol), N-boc-p-phenylenediamine

(1.1 g, 5.36 mmol) and sodium triacetoxyborohydride (3.38 g, 15.98 mmol) in 5:3 v/v dichloromethane/glacial acetic acid (12 ml) were stirred at room temperature for 3 h. The reaction mixture was washed with water and brine and then extracted into ethyl acetate. The organic layer was dried over sodium sulphate and evaporated under vacuum to yield the title compound as a dark red solid (1.16 g, 64 %). M.p. 130-131 °C; 1H NMR (400 MHz,

DMSO-d6) δ 1.48 (s, 9H, 9 x CH3), 4.8 (s, 2H, CH2), 7.46-7.59 (m, 4H, ArH), 9.38 (s,

13 1H, NH); C NMR (100 MHz, DMSO-d6) δ 28.5 (3 x CH3), 54.1 (CH2), 82.0 (C-O),

118.9 (2 x ArCH), 120.3 (ArC), 124.2 (ArC), 133.1 (2 x ArCH), 136.9 (CCl), 141.5

(CCl), 153.2 (C=O), 162.2 (C=O); IR (ATR): υmax 3345, 2923, 2321, 1698, 1589, 1519,

-1 1385, 1312, 1228, 1151, 1025, 932, 831, 758 cm ; UV (ACN): λmax 245 nm (ε 13,990

-1 -1 + cm M ); HRMS (ESI) m/z calcd for C15H16Cl2N2O3Na 365.0430 [M+Na] , found

381.0426.

N-(4-aminophenyl)-3,4-dichloro-1,5-dihydro-2H-pyrrol-2-one

(DHP phenyl amine-1)

The boc group was cleaved by treating the above DHP intermediate

(0.53 g) with trifluoroacetic acid (5 ml) at room temperature for 1 h

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and then evaporating it under high vacuum. The residue was washed with saturated solution of sodium bicarbonate and the solid obtained was filtered under vacuum to

1 yield a red solid (0.25 g, 64 %). M.p. 106 °C; H NMR (300 MHz, DMSO-d6) δ 4.7 (s,

13 2H, CH2), 5.13 (brs, 1H, NH2), 6.55-6.6 (m, 2H, ArH), 7.24-7.29 (m, 2H, ArH); C

NMR (75.5 MHz, DMSO-d6) δ 54.6 (CH2), 114.2 (2 x ArCH), 121.5 (ArC), 122.3 (2 x

ArCH), 127.5 (ArC), 140.6 (CCl), 146.9 (CCl), 163.0 (C=O); IR (ATR): υmax 3297,

3205, 2921, 1697, 1636, 1515, 1400, 1389, 1300, 1175, 1043, 928, 812 cm-1; UV

-1 -1 (ACN): λmax 247 nm (ε 10,817 cm M ), 307 (5783); HRMS (ESI) m/z calcd for

+ C10H8Cl2N2ONa 264.9906 [M+Na] , found 264.9909.

N-(4’-tert-butylphenylcarbamate)-3,4-dibromo-1,5-dihydro-2H-pyrrol-2-one

(boc-DHP phenyl amine-2)

The title compound was synthesized by reacting mucobromic acid

(1 g, 3.87 mmol), N-boc-p-phenylenediamine (0.8 g, 3.87 mmol) and sodium triacetoxyborohydride (2.46 g, 11.63 mmol) in 5:3 v/v dichloromethane/glacial acetic acid (10 ml) at room temperature for

18 h. The mixture was washed with water and brine and then extracted into ethyl acetate. The organic layer was dried over sodium sulphate and chromatographed on silica gel to yield the desired product as yellow solid (0.6 g, 36 %). 1H NMR (300 MHz,

CDCl3) δ 1.54 (s, 9H, 9 x CH3), 4.5 (s, 2H, CH2), 6.51 (s, 1H, NH), 7.39-7.42 (m, 2H,

13 ArH), 7.55-7.58 (m, 2H, ArH); C NMR (75.5 MHz, CDCl3) δ 28.3 (3 x CH3), 57.2

(CH2), 80.8 (C-O), 119.2 (2 x ArCH), 119.9 (ArC), 121.5 (ArC), 132.9 (2 x ArCH),

133.3 (CBr), 135.5 (CBr), 152.6 (C=O), 163.1 (C=O); IR (ATR): υmax 3348, 3099,

2973, 1768, 1688, 1605, 1518, 1430, 1364, 1283, 1232, 1146, 1020, 846, 739, 680 cm-1;

+ HRMS (ESI) m/z calcd for C15H16Br2N2O3Na 452.9420 [M+Na] , found 452.9422.

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N-(4-aminophenyl)-3,4-dibromo-1,5-dihydro-2H-pyrrol-2-one

(DHP phenyl amine-2)

The title compound was synthesized by following the same method used to synthesize DHP phenyl amine-1 to afford a yellow solid

1 (0.13 g, 28 %). M.p. 163 °C; H NMR (300 MHz, DMSO-d6) δ 4.7 (s,

2H, CH2), 5.16 (brs, 1H, NH2), 6.56-6.59 (m, 2H, ArH), 7.25-7.28 (m,

13 2H, ArH); C NMR (75.5 MHz, DMSO-d6) δ 57.0 (CH2),115.4 (2 x ArCH), 121.6 (2 x

ArCH), 127.0 (ArC), 129.4 (ArC), 132.4 (CBr), 144.2 (CBr), 164.2 (C=O); IR (ATR):

υmax 3305, 3208, 2920, 2287, 1693, 1611, 1512, 1442, 1383, 1280, 1150, 1023, 900, 830

-1 -1 -1 cm ; UV (ACN): λmax 243 (ε 8973 cm M ), 306 (3430); HRMS (ESI) m/z calcd for

+ C10H9Br2N2O 330.9076 [M+H] , found 330.9075.

4.2.3 Quorum Sensing Inhibition Assay

To evaluate the effectiveness of the synthesized DHP derivatives on QS signals, the P. aeruginosa MH602 PlasB::gfp(ASV) reporter strain, which harbors a chromosomal fusion of the lasB promoter to an unstable GFP gene and responds to the AHL 3-oxododecanoyl homoserine lactone (3oxo-C12-HSL), was used [192]. The QS inhibitory activity of free DHPs was determined by following the protocol described in section

3.2.4

4.2.4 Attachment of 3-Aminopropyltriethoxysilane (APTS)

Glass coverslips (No. 1, diameter 13 mm D 263 M glass, ProSciTech, Australia) used in this experiment were first cleaned by soaking in freshly prepared piranha solution (3:1 v/v concentrated sulphuric acid to 30 % hydrogen peroxide) at 100 °C for 1 h. After thorough rinsing with distilled water, the clean coverslips were rinsed once with absolute ethanol and air-dried.

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The substrates were then silanized according to the previously developed method [291].

Briefly, the clean glass coverslips were placed on steel mesh within a glass vessel that contained a 3-aminopropyltriethoxysilane (APTS) solution (10 % v/v in dry toluene; 1 ml). The glass vessel was sealed and heated at 140 °C for 18 h. The coverslips were rinsed with dry toluene (×2), absolute ethanol and air-dried.

4.2.5 Attachment of Succinic Anhydride (SA)

In order to generate the carboxylic acid surface, the APTS-coated coverslips were agitated overnight in 1 ml solution of succinic anhydride in DMSO (160 mg/ml) at room temperature. The APTS-SA functionalized surfaces were rinsed twice with

DMSO and once with ethanol, air-dried and stored before use.

4.2.6 Attachment of DHP via EDC/NHS Coupling

The APTS and APTS-SA coated coverslips were respectively placed in a 1:1 ethanol/water solution of acid- or amine-functionalized DHP (18.3 µM), containing 1- ethyl-3-(3-dimethylaminopropyl)carbodiimide hydrochloride (EDC, 92.0 µM), N- hydroxysuccinimide (NHS, 36.8 µM) and a small crystal of 4-dimethylaminopyridine

(DMAP). The coverslips were agitated overnight in their respective DHP solutions.

Following this, the treated surfaces were rinsed with MilliQ water and absolute ethanol and stored in sterile containers (Figure 4.5).

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Figure 4.5: Covalent attachment of DHPs on their respective surfaces.

4.2.7 X-ray Photoelectron Spectroscopy (XPS)

The attachment of DHP was analyzed by XPS using the same instrument setup as described in section 2.2.4.

4.2.8 Contact Angle Measurements

Water contact angles were measured using the same method described in section 2.2.5.

A minimum of fifteen measurements were made of five samples of each type.

4.2.9 Bacterial Adhesion Analysis

Staphylococcus aureus SA38 and Pseudomonas aeruginosa PA01 were used for this

8 study. Bacterial cultures were adjusted to 10 CFU/ml (OD660 = 0.1) before incubation of samples and stained with Live/Dead BacLight Viability Kits L-7007 according to the same experimental procedures described in section 2.2.6.

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Bacteria were fixed by adding 50 µl of 4 % formaldehyde to each sample and placed on the glass microscopy slide. Microscopic observation and image acquisition were performed with an Olympus FV1200 Confocal Inverted Microscope. The bacterial cells that were stained green were considered to be viable, while those that stained red or both green and red were considered to be dead. Images from 10 representative areas on each of triplicate samples for each surface from a minimum of three independent experiments were taken and analysed using ImageJ software [333]. The image analysis results were reported as the average percentage coverage of live and dead cells in the fields of view.

4.2.10 Statistical Analysis of Data

Data were analyzed the one-way analysis of variance (ANOVA) using GraphPad Prism

7.03 software. Post hoc multiple comparisons were done using Tukey correction, and results with p < 0.05 were considered significant.

4.3 Results

4.3.1 Synthesis of DHP Derivatives (DHP phenyl acid 1-4 and DHP phenyl amine 1-2)

The desired acid-functionalized DHP analogues in this chapter were synthesized by a one-step reductive amination reaction of commercially available mucohalic acids. The mucohalic acids underwent the lactone to lactam conversion by reacting with substituted aryl amines. The acid- and amine-functionalized DHP analogues synthesized in this chapter are summarized in Figure 4.6.

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Figure 4.6: Chemical structures of acid and amine-functionalized DHP derivatives.

Synthesis of DHP phenyl acid-1

The DHP phenyl acid-1 was synthesized successfully by reacting mucochloric acid with p-aminobenzoic acid in 5:3 v/v dichloromethane/glacial acetic acid, in presence of sodium triacetoxyborohydride at room temperature (Scheme 4.1). The proton NMR spectrum of the product obtained confirmed the insertion of the N-phenyl moiety, as indicated by the appearance of signals in the aromatic region from 7.84–8.01 ppm attributed to the four phenyl protons. A characteristic singlet corresponding to the newly formed CH2 group at 4.91 ppm and a broad peak at 12.8 ppm corresponding to the carboxylic acid group further confirmed the formation of the desired product.

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Scheme 4.1: Synthesis of DHP phenyl acid-1

As described previously, mucohalic acids can exist in an open chain or in a cyclic form.

Previous spectral analyses indicate that the cyclic lactone form predominates in solid state and solution [384,385]. In order to increase the rate of the reaction, acetic acid was employed as a weak acid additive to increase the amount of the open α,β-unsaturated aldehyde form and to maintain its stability and solubility [386,387]. Sodium triacetoxyborohydride, a mild and effective reducing agent, was used in the reactions due to its remarkable selectivity in reducing the aldehyde moiety of the open structure over other carbonyl groups [386,388]. A possible mechanism of the reaction between mucochloric acid and p-aminobenzoic acid proposed by Zhang et al. is illustrated in

Scheme 4.2 [389]. DHP phenyl acid-2 and DHP phenyl acid-4 were also synthesized by following the same reaction route.

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Scheme 4.2: Possible mechanism of reductive amination of mucochloric acid [389].

Synthesis of DHP phenyl acid-3

For the synthesis of DHP phenyl acid-3, the same reaction route as described above was initially attempted using mucobromic acid and p-aminobenzoic acid. However, unexpected multiple side products were obtained along with unreacted mucobromic acid and unreduced lactam. The reaction was successful when repeated at a higher temperature of 30 °C (Scheme 4.3). Proton NMR analysis indicated completion of reaction after 3 h, yielding the desired product after recrystallization.

Scheme 4.3: Synthesis of DHP phenyl acid-3.

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Synthesis of DHP phenyl amine-1

The reductive amination reaction between mucochloric acid and p-phenylenediamine was initially carried out under similar reaction conditions for the synthesis of DHP phenyl acids. However, due to low overall yields and formation of multiple products, the reaction was performed with boc-protected p-phenylenediamine instead of p- phenylenediamine, followed by cleavage of the boc protecting group in the next step.

Mucochloric acid and N-boc-p-phenylenediamine were reacted by the same procedure as described earlier (Scheme 4.4). The reaction was completed within 3 h as determined by TLC and the structure was confirmed by proton NMR which displayed a singlet for the boc protecting group at 1.48 ppm and a double of doublets for the phenyl protons at

7.46 ppm. The CH2 group and the boc-protected amine appeared as singlets at 4.8 ppm and 9.38 ppm respectively. The next step involved deprotection of the boc group to form the free primary amine, which was achieved by treating boc-DHP phenyl amine-1 with trifluoroacetic acid for 1 h. The formation of DHP phenyl amine-1 was confirmed by proton NMR spectrum. A new singlet at 5.13 ppm appeared in the NMR spectrum indicating formation of the free primary amine group.

Scheme 4.4: Synthesis of DHP phenyl amine-1.

Similarly, DHP phenyl amine-2 was synthesized by following the same reaction procedure.

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4.3.2 Quorum Sensing Inhibition Activity of Free DHPs

The free DHP compounds synthesized for surface attachment were also tested for their

QS inhibitory activity at different concentrations using P. aeruginosa MH602 reporter strain as a model organism [192]. This reporter strain produces an unstable green florescence protein (GFP-ASV) during normal growth of the bacteria due to an increase in AHL signalling molecules. The level of GFP expression can thus be used as a marker to investigate the effect of compounds on bacterial QS. Compounds that inhibit QS pathways are expected to reduce GFP expression in the reporter strain. The results of the assay are presented in Table 4.1.

Table 4.1: Percentage quorum sensing inhibition activity against P. aeruginosa MH602 lasB reporter strain at different concentrations of DHP derivatives; aNo growth inhibition; bGrowth inhibition ≤ 10 %; NA = no activity.

QS Inhibition (%) Compound 250 µM 125 µM 62.5 µM

DHP phenyl acid-1 19.8 ± 3.3a 13.2 ± 3.9a 6.9 ± 1.8a

DHP phenyl acid-2 35.8 ± 1.8a 25.9 ± 2.5a 15.4 ± 0.8a

DHP phenyl acid-3 13.3 ± 3.0a 6.2 ± 3.5a NA

DHP phenyl acid-4 28.1 ± 3.1a 16.3 ± 1.9a 9.2 ± 1.1a

DHP phenyl amine-1 58.8 ± 5.7b 37.0 ± 3.1a 22.9 ± 3.2a

DHP phenyl amine-2 42.6 ± 3.3b 26.1 ± 3.0b 13.6 ± 2.6a

The results showed that the DHP derivatives were able to reduce the expression of GFP-

ASV in a concentration-dependent manner with little or no effect on bacterial growth.

The most active QS inhibitors were DHP phenyl amine-1 and DHP phenyl amine-2,

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which showed 58.8 % and 42.6 % QS inhibition at 250 µM respectively, with ≤10 % growth inhibition for both compounds. Amongst the acid-functionalized compounds,

DHP phenyl acid-2 and DHP phenyl acid-4 displayed higher QS inhibitory activity at all concentrations than the other acid DHPs. No growth inhibition was observed at any concentration for all the acid-functionalized DHPs.

4.3.3 XPS Characterization

XPS analysis was performed to determine the elemental composition of the surface after each step. The data for all the surfaces is shown in Table 4.2.

Table 4.2: XPS elemental composition of blank, APTS, APTS-SA and DHP modified

glass surfaces.

Surface % C % N % Halogen

APTS 47.5 8.1 -

DHP phenyl acid-1 50.3 9.5 0.72% Cl

DHP phenyl acid-2 55.5 9.4 1.28% Cl

DHP phenyl acid-3 52.7 8.6 0.98% Br

DHP phenyl acid-4 52.7 8.9 0.46% Br

APTS-SA 49.9 8.1 -

DHP phenyl amine-1 53.0 8.4 0.49% Cl

DHP phenyl amine-2 52.0 8.6 0.14% Br

Successful functionalization of the glass by APTS, APTS-SA and DHPs was revealed by the changes in the carbon, nitrogen and halogen composition. The attachment of

APTS on the blank glass was confirmed by the increase in carbon and nitrogen percentage (47.5 % C and 8.1 % N) which was found to be consistent with previous

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results [291,350]. The carbon and nitrogen composition further increased by 2.8–8.0 % and 0.5–1.4 % respectively after immobilization of DHP phenyl acids on glass.

Additionally, the presence of halogens in the XPS data (0.72–1.28 % Cl and 0.98–0.46

% Br) could be used as a marker to confirm the successful attachment of DHP phenyl acids.

Similarly, incorporation of succinic anhydride linker on APTS glass was determined by increase in carbon content from 47.5 % to 49.9 %, and subsequent coupling of APTS-

SA with DHP phenyl amines further increased the carbon composition by 2.1–2.2 % and the nitrogen content by 0.3–0.5 %. Also, the detection of 0.49 % Cl for DHP phenyl amine-1 and 0.14 % Br for DHP phenyl amine-2 in the XPS scan further confirmed the covalent attachment of DHPs. The DHP phenyl amines have displayed lower halogen content compared to their acid analogues implying lower coating efficiency, which could be possibly due to less number of acid groups on the surface for attachment.

The high resolution XPS scan of the C 1s and N 1s region of all the surfaces is shown in

Table 4.3.

Table 4.3: XPS binding energies for C 1s and N 1s and proposed assignments with percentage peak intensities.

C 1s N 1s Binding Peak Binding Peak Surface energy Assignment area energy Assignment area (eV) (%) (eV) (%)

284.8 C-C 63.1 399.5 NH2 82.4 APTS 285.8 C-N 25.5 401.4 NH + 17.6 3 287.9 C=O 11.4

284.7 C-C 54.5 399.6 NH 80.4 DHP 2 + phenyl 285.7 C-N 22.9 401.4 NH3 , Tertiary N 19.6 acid-1 287.6 C=O 10.8

288.5 N-C=O 11.8

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284.7 C-C 54.0 399.7 NH2 80.9 DHP + phenyl 285.7 C-N 21.6 401.5 NH3 , Tertiary N 19.1 acid-2 287.6 C=O 10.8

288.6 N-C=O 13.6

284.8 C-C 57.0 399.1 NH 78.3 DHP 2 + phenyl 285.8 C-N 20.8 401.6 NH3 , Tertiary N 21.7 acid-3 287.9 C=O 10.2

288.5 N-C=O 12.0

284.7 C-C 58.8 399.5 NH 77.3 DHP 2 + phenyl 285.7 C-N 20.1 401.3 NH3 , Tertiary N 22.7 acid-4 287.6 C=O 10.0

288.6 N-C=O 11.1

284.7 C-C 62.6 399.3 NH2 84.5 + APTS- 285.7 C-N 18.0 401.5 NH3 , Tertiary N 15.4 SA 287.8 C=O 12.0

288.9 N-C=O 7.4

284.7 C-C 62.4 399.2 NH 79.4 DHP 2 + phenyl 285.6 C-N 15.2 401.5 NH3 , Tertiary N 20.6 amine-1 287.8 C=O 12.6

288.6 N-C=O 9.8

284.7 C-C 62.5 399.2 NH 77.8 DHP 2 + phenyl 285.7 C-N 17.2 401.5 NH3 , Tertiary N 22.1 amine-2 287.8 C=O 11.4

288.6 N-C=O 8.8

The C 1s high resolution spectrum of APTS surface showed three common carbon species, C-C, C-N and C=O at 284.8 eV, 285.8 eV and 287.9 eV respectively, while the

+ N 1s spectrum showed peaks for NH2 and NH3 at 399.5 eV and 401.4 eV respectively.

On addition of DHP phenyl acids on the surface, a new peak at ~288.5 eV corresponding to the amide bond (N-C=O) emerged in C 1s scan, confirming the reaction between APTS and the acid functional group of DHP. The nitrogen species also displayed some changes due to tertiary nitrogen from the DHP. - 123 -

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Similarly, attachment of succinic anhydride on APTS surface was confirmed by the presence of peak at 288.9 eV assigned to amide bond. The C 1s and N 1s XPS scan showed some changes after reacting APTS-SA substrate with DHP phenyl amines. A slight shift in the peak from 288.9 eV to 288.6 eV was also observed due to C-Br and C-

Cl bond from the DHPs.

4.3.4 Contact Angle Measurements

The change in hydrophobicity of the surfaces after each modification step was determined by measuring the contact angle of all the surfaces. The contact angle values of all samples are shown in Table 4.4.

Table 4.4: Contact angle values of blank, APTS, APTS-SA and DHP modified surfaces.

Contact angle Surface (°) (±1) Blank 20 APTS 78 DHP phenyl acid-1 72 DHP phenyl acid-2 76 DHP phenyl acid-3 79 DHP phenyl acid-4 73 APTS-SA 66 DHP phenyl amine-1 71

DHP phenyl amine-2 71

In contrast to low contact angle of 20° for blank glass, the contact angle value increased significantly to 78° for the amine-terminated APTS surface. However, APTS-SA displayed a lower contact angle value (67°) due to the less hydrophobic nature of

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carboxylic acids compared to amines [381,390,391]. The coupling of DHPs on APTS or

APTS-SA coated surfaces resulted in similar contact angles (71–79°) due to the addition of hydrophobic aromatic moiety and halogen atoms.

4.3.5 Antibacterial Activity

The ability of DHP coated surfaces to prevent colonisation of S. aureus and P. aeruginosa was evaluated by fluorescence microscopy. Live/dead staining was used to investigate bacterial adhesion and biofilm formation on all the surfaces. Representative images of the DHP surfaces and its corresponding control surface for S. aureus and P. aeruginosa are shown in Figure 4.7 and Figure 4.8 respectively. To evaluate the total of the surface covered by bacteria and he relative proportion of live bacteria stained green and dead bacteria stained red, quantitative image analysis was performed and the results are shown in Figure 4.9.

The APTS and APTS-SA control surfaces showed extensive colonization and biofilm formation by S. aureus and P. aeruginosa. The adhesion of both strains of bacteria on blank glass (data not shown) was similar to that reported in literature [291,350].

Significant reduction in bacterial cell adhesion was observed for both bacterial strains on the DHP-coated surfaces compared to the control samples. (Figure 4.7 and Figure

4.8).

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S. aureus

A – APTS B – DHP phenyl acid-4

C– APTS-SA D – DHP phenyl amine-1

Figure 4.7: Confocal microscopic images of S. aureus adhered to APTS and APTS-SA process control (A and C), DHP phenyl acid-4 and DHP phenyl amine-1 surfaces (B and D). Live bacterial cells stained green and bacteria with damaged membranes stained red. Magnification 200×. Scale bar = 100 µm.

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P. aeruginosa

A – APTS B – DHP phenyl acid-4

C – APTS-SA D – DHP phenyl amine-1

Figure 4.8: Confocal microscopic images of P. aeruginosa adhered to APTS and APTS-

SA process control (A and C), DHP phenyl acid-4 and DHP phenyl amine-1 surfaces (B and D). Live bacterial cells stained green and bacteria with damaged membranes stained red. Magnification 200×. Scale bar = 100 µm.

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For S. aureus, image analysis of blank (10.5 ± 0.6 %), APTS (10.8 ± 0.4 %) and APTS-

SA (12.0 ± 1.0 %) glass showed similar level of bacterial coverage, while surfaces coated with DHP exhibited significantly lower bacterial coverage compared to the blank and process control samples (APTS and APTS-SA) (Figure 4.9A). The DHP phenyl acids 1-4 reduced coverage by 38.0–46.6 % compared to APTS control (p < 0.01), whereas DHP phenyl amines 1-2 showed reductions of 42.0–50.0 % compared to the

APTS-SA control surface (p < 0.01). DHP phenyl acid-2 (46.6 ± 0.3 % reduction) and

DHP phenyl acid-4 (44.8 ± 0.2 % reduction) were slightly more effective than the other two DHP phenyl acid surfaces, although the difference between them was not statistically significant. On the other hand, the DHP phenyl amine-1 surface showing

50.0 ± 0.2 % reduction performed better than the DHP phenyl amine-2 surface, but the surfaces were not found to be significantly different from each other. There was no significant increase in the proportion of dead (red stained) cells for all modified samples.

For P. aeruginosa, the blank, APTS and APTS-SA controls again showed high bacterial coverage of 9.4 ± 0.4 %, 10.6 ± 1.0 % and 12.1 ± 0.8 % respectively. In comparison, the

DHP-coated substrates showed significantly fewer adherent bacterial cells (Figure

4.9B). The reductions observed for DHP phenyl acids 1-4 varied from 35.0–48.5 % (p <

0.01), while surfaces coated with DHP phenyl amines 1-2 displayed 48.3–55.4 % reduction in bacterial adhesion (p < 0.01). Of these, DHP phenyl acid-4 showed slightly higher bacterial reduction (48.5 ± 0.3 %) than the other DHP phenyl acids and DHP phenyl amine-1 (55.4 ± 0.3 %) was found to be more effective compared to DHP phenyl amine-2, however, similar to S. aureus, significant difference was not observed between the DHP treated surfaces. Moreover, the proportion of dead cells did not increase significantly for any of the modified samples. - 128 -

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A Bacterial coverage - S. aureus 14 12 10 * 8 * * Dead * * * bacteria 6 Live 4 bacteria

Bacterial coverage (%) coverage Bacterial 2 0

B Bacterial coverage - P. aeruginosa 14

10 * 8 * * * * Dead Bacteria * 6 Live Bacteria

4 Bacterial coverage (%) coverage Bacterial 2

Figure 4.9: Percentage surface coverage of live and dead bacteria for (A) S. aureus and

(B) P. aeruginosa (mean ± standard error of mean); *indicates p < 0.01 compared to control.

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4.4 Discussion

Medical implant-associated infections, most frequently caused by S. aureus and P. aeruginosa, are of increasing concern in modern medicine. The synthetic analogues of furanones, DHPs, are promising candidates for treating microbial infections as they are able to inhibit biofilm formation in several bacterial species. However, limited research has been performed to determine the changes in activity of DHPs in the absence of the exocyclic double bond. In this chapter, analogues of halogenated DHPs lacking an exocyclic double bond were synthesized and analysed for their bacterial inhibition activity after covalent attachment on glass substrates.

The target DHPs were successfully synthesized by a simple reductive amination reaction of mucohalic acids with different aryl amines. The halogenated DHP acids were covalently coupled onto the amine-functionalized glass by EDC/NHS reaction. The successful attachment of DHPs to the APTS surface was confirmed by XPS analysis.

The highest surface concentration was obtained with DHP phenyl acid-2 (1.28 % Cl) compared to other acid-functionalized DHPs. The ring-opening reaction of succinic anhydride with APTS generated the acid-functionalized APTS-SA surfaces, which were subsequently used for coupling of amine-functionalized DHPs. The DHP phenyl amines displayed lower attachment efficiency compared to DHP phenyl acids (0.49 % Cl for

DHP phenyl amine-1 and 0.14 % Br for DHP phenyl amine-2). This may be a result of lower surface concentration of carboxylic acid groups on APTS-SA surface indicating some amine groups remained unreacted after the initial reaction with succinic anhydride.

All covalently bound halogenated DHPs were able to inhibit bacterial colonization of S. aureus and P. aeruginosa with no significant difference between them. Overall, higher

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efficacy was displayed by the surface coated with DHP phenyl amine-1. It also displayed the highest QS inhibitory activity and ≤10 % growth inhibition against P. aeruginosa MH602 in solution at all concentrations (250–62.5 µM). Moreover, the proportion of dead (red-stained) cells on the surface did not increase significantly with the DHP substrates, which indicates that similar to furanones the DHP compounds in solution and surface act without killing the bacteria [170,392].

Surprisingly, DHP phenyl acid-1 and -3 displayed the least reduction in adhesion (≤40

%) for both bacterial strains despite their high surface attachment (0.72 % Cl for DHP phenyl acid-1 and 0.98 % Br for DHP phenyl acid-3). Their structural analogues DHP phenyl acid-2 and -4, both meta-substituted acid compounds gave ≥42 % reduction of bacterial adhesion. It is interesting to note that the two meta-acid compounds had vastly different attachment efficiency (1.28 % Cl for DHP phenyl acid-2 and 0.46 % Br for

DHP phenyl acid-4) but displayed similar levels of bacterial reduction. The enhanced activity displayed by DHP phenyl acid-4 despite its low concentration could be attributed to the presence of the bromine atom in its lactam ring, which is known to play an important role in maintaining the biological activity of furanones as well as lactams

[309,393,394]. Further improvement in activity was observed even at low surface concentration, when the carboxylic acid group of DHP phenyl acid-1 and -3 was replaced by an amine functional group at the para position (p < 0.01). These results were consistent with the QS inhibition assay where the amine-functionalized DHPs displayed higher QS inhibitory activities (42.6–58.8 % QS inhibition) in solution than the acid-functionalized DHPs (13.3–35.8 % QS inhibition).

In general, the coatings generated from DHPs bearing an exocyclic double bond have displayed higher activity compared to surfaces produced by DHPs in this study. The

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DHP substrates produced from photoactivation of azide surface (Chapter 2) and Michael addition reaction exhibited reductions of up to 90 % and 70 % against S. aureus and P. aeruginosa respectively [291,350]. While surfaces developed by DHPs lacking exocyclic C-5 bond displayed reductions of no more than 50 % against S. aureus and 55

% against P. aeruginosa. This reduced activity of DHP phenyl acids and amines could be attributed to their lack of exomethylene functionality. Previous studies on furanones have demonstrated complete loss of activity in absence of C-5 double bond thus indicating that C-5 bond is required to confer inhibitory activity to furanones [309,310].

However, in contrast to the furanones, the absence of C-5 double bond does not have a drastic effect on the activity of the DHPs. This could be possibly due to higher stability of the lactam ring compared to the lactone ring of furanones.

4.5 Conclusion

In this study, novel DHP compounds lacking an exocyclic double bond were successfully synthesized and tested for their antibacterial and QS inhibitory activities.

The results indicated that all the DHPs could inhibit QS and reduce bacterial adhesion on surfaces. It should be noted that, although the DHPs showed low activity against S. aureus and P. aeruginosa, complete loss of activity was not observed for the modified surfaces. From these biological results, it can be concluded that in the absence of the exocyclic methylene group there may be additional factors responsible for maintaining the activity of DHPs that needs to be investigated. Moreover, further structural modifications of the DHPs studied in this chapter might improve the biological activity of these compounds for generation of novel surface coatings to prevent device-related infections.

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Dual-Action Antibacterial Surfaces Based on DHP and Nitric Oxide

CHAPTER-5

CHAPTER-5 Dual-Action Antibacterial Surfaces Based on DHP and Nitric Oxide

5.1 Introduction

The drastic increase in antibiotic resistance has created a demand for alternate antibacterial therapies for combating bacterial infections as well as for protecting biomedical devices. Another promising strategy for the design of antibacterial biomedical devices is to use nitric oxide (NO) to control biofilm formation. NO possesses excellent biofilm inhibitory activity against both Gram-positive and Gram- negative bacteria [228,230]. Additionally, NO plays a very important role in controlling bacterial quorum sensing (QS) and virulence expression in different bacterial species

[215].

Given the potent biofilm inhibitory activity of NO, novel dual-action hybrid molecules have been previously synthesized by our group by combining various NO donors such as N-diazeniumdiolate (NONOate), S-nitrosothiol (S-NO) and nitrates with potent QS inhibiting compounds, such as furanones (forming “furanone+NO”) and DHPs (forming

“DHP+NO”) (Figure 5.1) [182]. With continuous NO release, the hybrid compounds were more successful at eradicating biofilm of P. aeruginosa than the QS inhibitor alone [253]. Additionally, a number of materials such as gold nanoparticles, catheters and polyurethane have been immobilized with NO donors for preventing bacterial infections. The NO releasing surfaces were found to be effective in eradicating in vitro biofilms of P. aeruginosa, S. epidermidis and S. aureus as well as reducing S. aureus infections in vivo [231,295–297]. Developing dual-action NO-releasing biomaterials

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therefore represents an exciting opportunity for the control and prevention of bacterial infections.

Figure 5.1: Dual-action furanone+NO and DHP+NO molecules synthesized by Kutty

[182].

In this chapter, novel dual-action surfaces were developed by incorporating both DHP and a suitable NO donor onto a biomaterial surface. This could potentially be achieved either by directly attaching the previously tested DHP+NO compounds on the surface, or by reacting furanone+NO compounds with amine-functionalized surfaces to form the corresponding DHP (via lactone to lactam conversion) with concomitant attachment

(Figure 5.2). However, both of these approaches are limited by the sensitivity of the hybrid molecules. It is likely that the direct immobilization of DHP+NO molecules onto a surface or the reaction of furanone+NO compounds with an amine-functionalized surface would be accompanied by thermal decomposition of NO donors, resulting in excessive NO release before biological testing.

Figure 5.2: Dual-action surface based on lactone (furanone+NO) to lactam

(DHP+NO) conversion on surface

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Therefore, an alternate strategy to develop these surfaces was adopted, in which the

DHP compound was first covalently linked to the surface via specific attachment, followed by subsequent derivatization of the attached DHP to introduce the NO donor moiety. Recently, Ho et al. successfully grafted DHPs onto glass surfaces and polymer beads by introducing the acrylate functionality at the N-1 position of the heterocyclic lactam ring of the DHP. The acrylate-functionalized DHP reacted via Michael addition with the primary amine groups present on the surface and beads (Figure 5.3). The resulting DHP-modified substrates reduced the formation of biofilm of P. aeruginosa and S. aureus both in vitro and in vivo [291,293].

Figure 5.3: Covalent immobilization of DHP-acrylates via Michael addition reaction

[291].

The same attachment strategy was employed in this study. Two potent DHP molecules were selected and modified with an acrylate group according to a method developed by

Kumar and Iskander (Figure 5.4) [202]. The DHP-acrylate compounds were then covalently linked onto a primary amine-functionalized surface via Michael addition reaction, and the resulting DHP coatings were subsequently treated with NO gas to form the NONOate moiety on the surface. For incorporation of NONOate groups, the direct reaction of NO with an amine is the only useful method of preparation. However,

NONOates derived from primary amines are not stable, resulting in low NO storage and short NO release duration profiles [237,395,396]. Therefore, a surface containing both

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primary and secondary amine groups is needed for reaction with DHPs and NO respectively.

Figure 5.4: Functionalization of 4-aryl substituted DHPs with an acrylate group.

The attachment strategies were performed on fluorinated ethylene propylene (FEP) surfaces. The surface was first functionalized with free carboxylic acid groups using an acrylic acid plasma treatment process. A physiologically important polyamine, spermine, was coupled to the carboxylated surface via the EDC coupling reaction.

Spermine was selected as the linker because it offers both free primary amine groups for reaction with DHP-acrylates as well as secondary amines to react with NO to generate

NO-donating moieties (Figure 5.5). To evaluate the efficacy of DHP+NO surfaces, spermine-coated FEP in absence of DHP was also reacted with NO to from spermine+NO surface and then compared with DHP+NO.

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Figure 5.5: Schematic representation of (A) dual-action FEP surfaces developed by

incorporating DHP-acrylate derivatives and NONOate NO donors and (B) attachment

of NONOate to spermine surface.

This is the first time that dual-action NO-releasing biomaterials based on QS inhibitors have been developed, which are expected to have more potential in inhibiting bacterial adhesion compared to single-action surfaces. The resulting DHP, DHP+NO and spermine+NO-donating FEP coatings were characterized by X-ray photoelectron spectroscopy (XPS) and the NO releasing profile was determined by the Griess assay, followed by bacterial adhesion analysis using fluorescence microscopy against

Staphylococcus aureus and Pseudomonas aeruginosa.

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5.2 Materials and Methods

5.2.1 General

All chemical reagents were purchased from commercial sources (Alfa-Aesar and Sigma

Aldrich) and used without further purification. Solvents were sourced from commercial sources and used as obtained. Reactions were performed using oven-dried glassware under an atmosphere of nitrogen and in anhydrous conditions (as required). Room temperature refers to the ambient temperature (22–24 °C). Yields refer to chromatographically and spectroscopically pure compounds unless otherwise stated.

200 mm glass plates coated with Merck 60GF254 silica gel. Infrared spectra were recorded using a Cary 630 FTIR spectrophotometer. Ultraviolet spectra were measured using a Cary 100 Bio UV-visible spectrophotometer in the designated solvents and data reported as wavelength (λ) in nm and absorption coefficient (ε) in cm-1M-1. High resolution mass spectrometry was performed by the Bioanalytical Mass Spectrometry

Facility, UNSW. Melting points were obtained using Mel-Temp melting point apparatus and are uncorrected. Proton and carbon NMR was recorded in designated solvents using

Bruker DPX 300 or a Bruker Avance 400 spectrometer as designated. Chemical shifts

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quartet (q), multiplet (m) or a combination of these (e.g. dd, dt etc.)]; coupling constant

(J) in hertz, integration, proton count and assignment.

5.2.2 Synthesis of Acrylate-Functionalized DHP Derivatives

5-Methyl-1-(prop-2-enoyl)-5-(prop-2-enoyloxy)-4-phenyl-dihydropyrrol-2-one

To a solution of 5-hydroxy-5-methyl-4-phenyl-dihydropyrrol-2- one (0.23 g, 1.22 mmol) in 1:10 dry THF/dichloromethane (11 ml), was added triethylamine (1.5 ml, 10.76 mmol) along with a few crystals of hydroquinone while stirring the mixture in an ice bath. A solution of acryloyl chloride in dry dichloromethane (3 ml) was added dropwise over a period of 10 min and stirred further for 3 h. The solvent from the crude mixture was removed by evaporation under vacuum and the residue was purified by flash chromatography using dichloromethane as an eluent to yield the title compound as a

1 pale yellow solid (0.21 g, 75 %). H NMR (300 MHz, CDCl3): δ 2.11 (s, 3H, CH3),

5.84-5.92 (m, 2H, =CH2), 6.10–6.16 (m, 1H,–CH=), 6.43-6.51 (m, 3H, =CH2 and -

CH=), 7.19 (s, 1H, CH), 7.43–7.46 (m, 3H, ArH) and 7.46–7.66 (m, 2H, ArH).

5-Methylene-1-(prop-2-enoyl)-4-phenyl-dihydropyrrol-2-one (DHP-1)

The title compound was synthesized by treating a solution of 5- methyl-1-(prop-2-enoyl)-5-(prop-2-enoyloxy)-4-aryl-dihydropyrrol-2- one (0.44 g, 1.48 mmol) in dichloromethane (9 ml) with trifluoroacetic acid (1 ml). The reaction mixture was stirred at room temperature for 2 h. The resultant mixture was neutralized with saturated sodium bicarbonate and water and then extracted into dichloromethane. The organic layer was dried over sodium sulphate and chromatographed on a silica column using dichloromethane to obtain pure title product as white solid (0.18 g, 51 %). M.p. 134-135 °C; 1H NMR (300 MHz,

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CDCl3): δ 5.42 (d, J = 0.9 Hz, 1H, =CH2), 5.93 (dd, J = 1.8, 10.5 Hz, 1H, =CH2), 6.16

(d, J = 0.9 Hz, 1H, =CH2), 6.58 (dd, J = 17.1 and 1.8 Hz, 1H, =CH2), 6.70 (t, J = 1.2 Hz,

13 1H, -CH=), 7.45-7.53 (m, 6H, ArH and CH); C NMR (75 MHz, CDCl3): δ 108.5

(CH2), 119.5 (CH), 128.7 (4 x ArCH), 130.1 (CH2), 130.4 (CH), 130.9 (CH), 141.8 (C),

155.5 (C), 163.4 (C), 165.8 (C=O), 168.6 (C=O); HRMS (ESI) m/z calcd for

+ C14H11NO2Na 248.0682 [M+Na] , found 248.0684.

5-Methylene-1-(prop-2-enoyl)-4-(2-fluorophenyl)-dihydropyrrol-2-one (DHP-2)

The title compound was synthesized by following the same method used for synthesis of DHP-1 to yield a pale yellow solid (0.23 g, 72

1 %). M.p. 121-122 °C; H NMR (CDCl3): δ 5.25 (d, J = 0.6 Hz, 1H,

=CH2), 5.93 (dd, J = 10.2 and 1.8 Hz, 1H, =CH2), 6.24 (d, J = 0.9 Hz,

1H, =CH2), 6.58 (dd, J = 16.5 and 1.8 Hz, 1H, =CH2), 6.66 (t, J = 1.2 Hz, 1H, -CH=),

7.21-7.28 (m, 2H, ArH), 7.33-7.39 (m, 1H, ArH), 7.47-7.55 (m, 2H, ArH and CH); 13C

NMR (75 MHz, CDCl3): δ 108.2 (CH2), 116.3 (CH), 122.2 (CH), 124.3 (CH), 130.4

(CH), 130.8 (CH), 131.1 (CH2), 131.9 (CH), 141.7 (C), 149.2 (C), 158.1 (C), 161.4

(CF), 165.8 (C=O), 168.6 (C=O); HRMS (ESI) m/z calcd for C14H10FNO2Na 266.0593

[M+Na]+, found 266.0590.

The complete spectral details of all compounds have been previously reported in the literature [202,291].

5.2.3 Acrylic Acid Plasma Treatment of FEP

Before use, FEP sheets were cleaned with absolute ethanol then rapidly dried with a jet of nitrogen. The FEP samples were plasma-activated using acrylic acid in a custom- built plasma reactor according to a previously established procedure (Figure 5.6) [349].

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Figure 5.6: Plasma coating of acrylic acid monomer on blank FEP sheets.

Briefly, the reactor used was comprised of a cylindrical glass chamber (height = 350 mm, diameter = 170 mm). The reactor contained a horizontal disc electrode of diameter

150 mm on the bottom and a 6 mm rod electrode on the top, separated by 150 mm. The substrates were placed on the lower rectangular electrode of the plasma reactor. The acrylic acid monomer (Sigma-Aldrich, 98 %) was degassed 5 times prior to deposition.

The plasma deposition was carried out twice for 25 s with an initial pressure of 0.2 mbar

(200 kHz, 20 W). The activated FEP samples with carboxylic acid groups were stored in a sterile container and used within a week.

5.2.4 Attachment of Spermine Linker by EDC Coupling

The carboxylic acid FEP surfaces were immersed in a solution of EDC (30 mg/ml) in

0.1 M sodium acetate buffer solution at pH 5.0 for 30 min with gentle shaking. The surfaces were then treated with a solution of spermine (3 mg/ml) in MilliQ water and left to react overnight at room temperature. The resultant amine-functionalized surfaces were washed with MilliQ water three times.

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5.2.5 Attachment of DHP via Michael Addition Reaction

The DHPs were immobilized on the amine-terminated FEP according to a previously developed method (Figure 5.7) [291]. Briefly, a solution of DHP in absolute ethanol was prepared (6 mg/ml) and each amine-functionalized surface was immersed in 500 µl of the DHP solution. The surfaces were left to react overnight with agitation at room temperature. The resultant DHP-immobilized surfaces were rinsed with absolute ethanol three times, air-dried and stored in sterile containers.

5.2.6 Attachment of N-diazeniumdiolate (NONOate)

The spermine and DHP-immobilized FEP surfaces were placed in a Parr apparatus and clamped. The apparatus was then purged and evacuated with nitrogen three times and pressurized with NO gas to 5 atm at 25 °C for 48 h to form NONOate NO donors

(Figure 5.7).The surfaces were purged with nitrogen gas and then left under vacuum for

24 h to remove excess NO. The NONOate surfaces were stored at 4 °C until required for further analysis.

Similarly, the blank FEP surfaces were also reacted with NO and used as a control to confirm the attachment of NONOate on spermine and DHP substrates.

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Figure 5.7: Covalent attachment of DHPs and reaction with NO gas to form NONOate

NO donors.

5.2.7 X-ray Photoelectron Spectroscopy (XPS)

The surfaces were characterized with XPS using the same instrument setup as described in section 2.2.4.

5.2.8 Contact Angle Measurements

Contact angles were determined using the same method described in section 2.2.5. A minimum of fifteen measurements were made of five samples of each type.

5.2.9 Determination of Nitric Oxide Release by Griess Assay

NO released from the surface was determined using a standard Griess reagent kit

(Molecular Probes® Life Technologies™). The surfaces to be tested were placed in

PBS (1 ml) for 24 h. Then, a 150 µl aliquot from the PBS test solution was added to 20

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µl of Griess reagent, freshly prepared by mixing equal volumes of reagents A and B, in each well of a 96-well plate. The mixture was topped up with 130 µl of PBS to make up a total volume of 300 µl, and then left to incubate at room temperature for 30 min with agitation at 500 rpm. After 30 min, the absorbance of the wells was measured at 580 nm using a FLUOstar® Omega Microplate Reader.

Solutions of 0 to 100 µM sodium nitrite were used to prepare a standard curve of nitrite absorbance versus concentration under the same experimental conditions. Extrapolation from the standard curve gave the concentration of nitrite (µM) generated from different surfaces.

5.2.10 Bacterial Adhesion Analysis

The bacterial strains, Staphylococcus aureus SA38 and Pseudomonas aeruginosa PA01 were used for this study. Samples with adherent bacteria were prepared and stained with

Live/Dead BacLight Viability Kits L-7007 according to the same experimental procedures described in section 2.2.6. Bacteria were fixed by adding 50 µl of 4 % formaldehyde to each sample and placed on the glass microscopy slide.

Microscopic observation and image acquisition were performed with an Olympus

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5.2.11 Statistical Analysis of Data

Further analysis of data was done by the one-way analysis of variance (ANOVA) using

GraphPad Prism 7.03 software. Statistical differences were analyzed using post hoc

Tukey correction. Results were considered significant for p < 0.05.

5.3 Results

5.3.1 Synthesis of DHP-Acrylate Derivatives (DHP 1-2)

The DHP-acrylate derivatives (DHP 1-2) used in this project were successfully synthesised according to a literature procedure [202]. The hydroxyl DHP compound 1 was reacted with acryloyl chloride 2 in the presence of triethylamine and hydroquinone to form a di-acrylate DHP intermediate 3 (Scheme 5.1). As the C-5 hydroxyl group of

DHP is more reactive than the N-1 amine group, an excess of acryloyl chloride was used in order to ensure that an acrylate would form at N-1. The proton NMR spectrum of the purified product showed peaks for two acrylate groups, indicating successful synthesis of the di-acrylate 3, (5-methyl-1-(prop-2-enoyl)-5-(prop-2-enoyloxy)-4-aryl- dihydropyrrol-2-one), as the major product.

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Scheme 5.1: Synthesis of DHP-acrylate derivatives.

The next step involved the selective hydrolysis of the acrylate group at C-5, which was achieved by treating the di-acrylate compounds 3 with trifluoroacetic acid at room temperature. Under these conditions, the di-acrylate compounds undergo acid-catalyzed ester hydrolysis to form hydroxyl DHPs with acrylate only at N-1, followed by dehydration to form the C-5 exocyclic double bond. The reaction was monitored by thin layer chromatography which indicated completion of reaction after 2 h. The proton

NMR of the product obtained after flash column chromatography exhibited peaks for the characteristic vinyl bond at C-5 and acrylate functionality intact at N-1, indicating successful formation of 4 (5-methylene-1-(prop-2-enoyl)-4-aryl-dihydropyrrol-2-one, where aryl = phenyl for DHP-1 and aryl = ortho-fluorophenyl for DHP-2). The spectral

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data of all compounds was consistent with that previously reported in the literature

[202,291].

5.3.2 High-Resolution XPS characterization

XPS analysis of the surfaces was performed to assess the elemental composition before and after modification. The data for untreated FEP (blank), acid, spermine, DHP and

NO coated surfaces are summarized in Table 5.1. The high resolution curve-fitting results and proposed assignments for C 1s and N 1s are shown in Table 5.2 and Table

5.3 respectively.

The blank FEP surfaces were found to contain mainly 31.3 % carbon and 68.5 % fluorine, and a small amount of oxygen (0.2 %) as contamination (Table 5.1). The acrylic acid plasma deposition on blank surfaces resulted in a polymer layer that was rich in carbon (73.1 %), oxygen (23.8 %) along with traces of nitrogen (0.7 %). The fluorine percentage decreased to 2.4 % suggesting either presence of a thick acrylic acid coating or defluorination of the surface during XPS analysis [345,353–355]. The attachment of the aliphatic diamine linker spermine on the acid surface was confirmed by increase in carbon percentage to 75.0 % and nitrogen percentage to 2.1 %.

Coupling of DHP-acrylates in the next step further increased the carbon and nitrogen composition (77.7–78.8 % C and 3.1–3.2 % N). Both DHPs displayed similar attachment efficiency on the spermine surface, as indicated by the similar values of carbon, nitrogen and oxygen contents for DHP-1 and DHP-2. However, the fluorine content decreased from 1.6 to 1.2 % for DHP-1 whereas for DHP-2 it increased from

1.6 to 2.3 %, which could be attributed to the presence of the ortho-fluorine group in

DHP-2. The incorporation of NO-donating moiety on spermine and DHP 1-2 surfaces resulted in further increases in nitrogen and oxygen values. - 147 -

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Table 5.1: XPS elemental composition of blank, acid, spermine, DHP and NO modified

FEP surfaces.

Surface % C % N % O % F

Blank 31.3 - 0.2 68.5 Acid 73.1 0.7 23.8 2.4 Spermine 75.0 2.1 21.3 1.6 Spermine+NO 66.4 3.0 23.6 7.0 DHP-1 78.8 3.1 16.9 1.2 DHP-1+NO 72.7 5.5 19.4 2.4 DHP-2 77.7 3.2 16.8 2.3 DHP-2+NO 70.9 4.6 20.5 4.0

The deconvoluted peaks for the high resolution C 1s spectrum are presented in Table

5.2. The blank FEP surface revealed the presence of C-C at 284.8 eV, CF2 at 290.5 eV,

CF3 at 292.4 eV and O-CF2 at 294.5 eV as shown in Chapter 3. After acrylic acid plasma deposition, peaks for new species emerged at 286.3 eV, 287.7 eV and 289.1 eV.

These peaks were assigned to C-O/C-N, C=O and O-C=O respectively, confirming the presence of carboxylic acid groups on the surface. The next step involved spermine conjugation, which resulted in the appearance of a peak at 288.7 eV corresponding to the newly formed amide bond (N-C=O). The percentage of the surface acid groups (O-

C=O at 289.1 eV) also decreased from 2.2 to 1.2 % as expected after spermine attachment. These results indicated successful reaction of spermine with the carboxylic acid groups on the FEP substrate to form an amine-functionalized surface.

The coupling reaction of DHP-1 and DHP-2 with the spermine linker showed further increase in peak intensity for the C-O/C-N, C=O and N-C=O signals. Additionally, complete attenuation for the acid peak (O-C=O) was observed, suggesting a thick - 148 -

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coating of DHPs had formed on the surface. The addition of NONOate groups slightly changed the ratio of the carbon components for DHP+NO and spermine+NO.

Table 5.2: XPS binding energies for C 1s and proposed assignments with percentage

peak intensities.

C 1s

Surface 284.8 eV 286.3 eV 287.7 eV 288.7 eV 289.1 eV 292.2 eV

C-C C-O/C-N C=O N-C=O O-C=O CF3

Acid 76.0 15.1 6.7 - 2.2 -

Spermine 74.8 16.6 6.5 0.9 1.2 -

Spermine+NO 67.7 16.3 6.6 - 2.7 6.8

DHP-1 71.7 17.0 7.4 3.9 - -

DHP-1+NO 66.2 19.1 9.1 5.6 - -

DHP-2 70.1 17.8 8.1 4.0 - -

DHP-2+NO 67.1 19.1 8.3 5.5 - -

The native N 1s scan of spermine and DHP samples showed the presence of two

+ common nitrogen species assigned to N-H (399.7 eV) and tertiary N/NH3 (401.8 eV)

(Table 5.3). The N 1s binding energy scan of DHP-1 before and after attachment of NO is shown in Figure 5.8. Upon coupling of the N-H groups present on the surface with

NO to form NONOate, the intensity of the N-H peak was reduced and a new peak arose at 405.8 eV with a small shoulder at 407.6 eV. By reference to published values, these values were assigned to NO3 and gaseous NO respectively from the NONOate donors

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attached on the surface [397–400]. Furthermore, there was also an increase in the intensity for the peak at 401.8 eV which was assigned to nitrolysis species, such as R-

(NO)2 or NO dimers [401,402]. The N 1s survey scan therefore indicated successful incorporation of NO on the DHP-1 coated surface. Similar changes in intensity of N 1s peaks were observed upon NO attachment for spermine+NO and DHP-2+NO modified surfaces.

Table 5.3: XPS binding energies for N 1s and proposed assignments with percentage

peak intensities.

N 1s

Surface 399.7 eV 401.8 eV 405.8 eV 407.6 eV

N-H Tertiary N/ NO3 Gaseous + NH3 /R-(NO)2 NO

Spermine 91.1 8.9 - -

Spermine+NO 47.6 17.5 28.5 6.3

DHP-1 88.7 11.3 - -

DHP-1+NO 58.5 14.5 23.0 3.9

DHP-2 75.8 24.2 - -

DHP-2+NO 54.7 14.9 25.9 4.5

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(A) Before NO reaction - DHP-1 surface

(B) After NO reaction - DHP-1+NO surface

Figure 5.8: XPS N 1s narrow scan spectra of DHP-1 surfaces (A) before treatment with

NO and (B) after treatment with NO.

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5.3.3 Contact Angle Measurements

The change in hydrophobicity of the surfaces is used to determine successful modification of the surface. The water contact angle values for all the surfaces were measured after each step. The larger the water contact angle value, the higher the hydrophobicity of the surface.

The hydrophobicity of the surface decreased markedly after acid functionalization of the blank FEP surface. The contact angle values were 108° and 59° for the blank and acid

FEP surfaces respectively (Table 5.4). A further decrease in hydrophobicity was observed after reaction with spermine (41°). The DHP-modified surfaces showed slight increases in hydrophobicity (49° for DHP-1 and 52° for DHP-2) due to addition of hydrophobic components from the DHPs. The subsequent reaction of spermine and both the DHP surfaces with NO resulted in further increase in contact angle values of 3 to 6°.

Table 5.4: Contact angle measurements of FEP substrates.

Contact angle Surface (°) (±1) Blank 108 Acid 59 Spermine 41 Spermine+NO 47 DHP-1 49 DHP-1+NO 54 DHP-2 52 DHP-2+NO 55

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5.3.4 Nitric Oxide Release Study by Griess Assay

The NO release from the surfaces was assessed by the Griess assay, a method that was first reported by Johann Peter Griess in 1879 and which is now the most frequently used colorimetric technique to quantify metabolites of NO (nitrite and nitrate) [250,251]. It should be noted that the primary decomposition product of NO in oxygen-containing

- - aqueous media is nitrite (NO2 ) and the oxidation of NO to NO2 can be represented as follows [403,404]:

- - - However, nitrates (NO3 ) can only be measured after reduction of NO3 to NO2 by chemical reductants such as cadmium, zinc and hydrazine or a bacterial nitrate reductase

(NR) enzyme. For example, enzymatic reduction of nitrate catalyzed in presence of

NADPH (nicotinamide adenine dinucleotide phosphate oxidase) as a co-factor is shown below [405]:

In the Griess assay, an aromatic sulfanilamide (reagent B) is first diazotised under acidic

- conditions by nitrite (NO2 ), followed by reaction with a coupling reagent, N-naphthyl- ethylenediamine (reagent A), to form an intense purple-coloured water soluble azo dye

(Figure 5.9) [406]. The absorbance maximum of the azo compound occurs at around

548 nm, which can be easily measured by a UV-visible spectrophotometer.

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(λmax = 548 nm)

Figure 5.9: Diazotization reaction between Griess reagent B and nitrite, followed by

coupling to reagent A to form an azo dye.

A standard curve obtained for 0 to 100 µM sodium nitrite solutions was prepared, which showed a good linear fit between 0 and 100 µM (Figure 5.10). The concentration of NO released in PBS from the surfaces (quantitated as nitrite ions) could then be determined using the standard curve.

Standard Curve – Sodium Nitrite 1.2

0.8 (548 nm)(548 0.6 y = 0.0094x + 0.0658 0.4 R² = 0.9961

Absorbance 0.2

0 0 20 40 60 80 100 Nitrite concentration (µM)

Figure 5.10: Sodium nitrite standard curve obtained using Griess assay.

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The Griess assay results indicated the successful formation of NONOate donors on spermine and DHP-modified surfaces. After 24 h of incubation in PBS, spermine+NO,

DHP-1+NO and DHP-2+NO exhibited much higher NO release of 547.2 ± 1.8, 577.7 ±

1.2 and 636.1 ± 0.7 nmol/mm2 of the surface, respectively compared to the blank surface (52.7 ± 1.0 nmol/mm2) (Table 5.5). This could be attributed to the fact that the blank does not possess any amine groups that could react with NO to form the

NONOate functionality. Additionally, the similar nitrite concentrations (NO release) exhibited by the spermine, DHP-1 and DHP-2 surfaces suggested that the attachment efficiency of NO to form NONOate is comparable with the XPS data for the three surfaces.

Table 5.5: Nitrite concentration (µM) and NO release (nmol/mm2) after 24 h in PBS, as

determined using the Griess assay.

Nitrite NO Release Surface Concentration (µM) (nmol/mm2)

Blank+NO 1.9 ± 1.5 52.7 ± 1.0

Spermine+NO 19.7 ± 2.5 547.2 ± 1.8

DHP-1+NO 20.8 ± 1.9 577.7 ± 1.2

DHP-2+NO 22.9 ± 1.1 636.1 ± 0.7

5.3.5 Antibacterial Activity

The DHP coatings generated were tested for their antibacterial efficacy against both

Gram-positive S. aureus and Gram-negative P. aeruginosa. In order to determine the adhesion and viability of bacteria on the surfaces, samples were incubated at 37 °C for

48 h in an adjusted bacterial culture (108 CFU/ml), followed by Live/Dead staining,

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which stains live bacteria green and dead bacteria red. Representative images for controls (acid and spermine-treated), and selected DHP and DHP+NO samples are shown in Figure 5.11 and Figure 5.12 for S. aureus and P. aeruginosa respectively.

The relative proportion of live (green-stained) and dead bacteria with damaged membranes (red-stained) on the modified surfaces was evaluated by image analysis and the results for S. aureus and P. aeruginosa are shown in Figure 5.13 and Figure 5.14.

Extensive bacterial colonization was observed on acid and spermine control surfaces by

S. aureus (Figure 5.11A and B) and P. aeruginosa (Figure 5.12A and B), as indicated by the high coverage of the green-stained (live) bacteria. In contrast to the control samples, the surfaces modified by DHP-1 showed considerably less bacterial coverage

(Figure 5.11C and Figure 5.12C). Further reduction in adherent bacteria was observed on surface coated with DHP-1+NO compared to control and DHP-1 (Figure 5.11D and

Figure 5.12D).

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S. aureus

A – Acid B – Spermine

C – DHP-1 D – DHP-1+NO

Figure 5.11: Confocal images of S. aureus adhered to acid and spermine control (A and

B), DHP-1 and DHP-1+NO-modified surfaces (C and D). Live bacterial cells stained green and bacteria with damaged membranes stained red. Magnification 200×. Scale bar = 100 µm.

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P. aeruginosa

A – Acid B – Spermine

C – DHP-1 D – DHP-1+NO

Figure 5.12: Confocal images of P. aeruginosa adhered to acid and spermine control

(A and B), DHP-1 and DHP-1+NO-modified surfaces (C and D). Live bacterial cells stained green and bacteria with damaged membranes stained red. Magnification 200×.

Scale bar = 100 µm.

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To compare the amount of live and dead bacteria on the samples, quantitative image analysis was performed on the images, which confirmed the high coverage of S. aureus on the blank (10.5 ± 0.5 %), acid control (14.1 ± 1.1 %), and spermine control (15.2 ±

1.4 %) surfaces (Figure 5.13). In contrast, the DHP and DHP+NO coatings displayed significantly lower bacterial coverage compared to the control surfaces. The DHP-1 and

DHP-2 surfaces displayed coverage values of 4.4 ± 0.9 % and 6.0 ± 1.0 % respectively, which was over 3-fold lower than the spermine control (p < 0.001).

Subsequent coupling with NO resulted in further decrease in coverage (2.5 ± 0.4 % for

DHP-1+NO and 4.3 ± 0.5 % for DHP-2+NO), with more pronounced reductions of

71.4–83.1 % compared to spermine (p < 0.001). Importantly, both DHP-1+NO (p <

0.05) and DHP-2+NO (p < 0.01) were significantly more active than their corresponding

DHP parent coatings indicating addition of NO enhanced the inhibitory activity of the surfaces. However, spermine+NO showed only a small and not statistically significant decrease in surface coverage compared to the spermine control. Amongst all the surfaces, DHP-1+NO was found to be the most effective in reducing bacterial colonization of S. aureus probably due to high surface and solution activity of DHP-1

[202,203,291]. A significant increase in the proportion of dead (red-stained) cells was not observed for the modified samples relative to the control samples indicating the surfaces acted without bacterial killing and that the coatings are not engendering bacterial resistance.

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Bacterial coverage - S. aureus

18 16 14 12 Dead bacteria 10 # * × 8 * Live ^ * 6 bacteria

4 * Bacterial coverage (%) Bacterial 2 0

Figure 5.13: Percentage bacterial coverage for S. aureus SA38 (mean ± standard error of mean); *indicates p < 0.001 compared to spermine control; ^indicates p < 0.05 compared to DHP-1; ×indicates p < 0.01 compared to other DHP-2; #indicates p < 0.01 compared to other DHP modified surfaces.

In case of P. aeruginosa, the image analysis showed that the blank (13.7 ± 1.5 %), acid

(13.1 ± 1.5 %) and spermine (14.8 ± 1.1 %) surfaces exhibited high density of bacterial colonization (Figure 5.14), whereas the DHP-1 (5.6 ± 0.4 %) and DHP-2 (6.3 ± 0.3 %) coated surfaces exhibited significantly fewer adherent bacteria than the spermine control

(p < 0.001). Further reductions in percentage bacterial coverage was observed for both the NO-treated DHP samples when compared to spermine (64.6–77.1 %).

Moreover, the DHP-1+NO with fewer adherent bacteria (3.3 ± 0.3 %) was significantly better than the DHP-1 sample (p < 0.05), however, DHP-2+NO sample did not exhibit significant reduction in bacterial colonization than DHP-2 coating. Interestingly, against

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P. aeruginosa the spermine+NO modified surface reduced colonization that appeared to be marginally significant (p < 0.1) compared to the spermine control surface. Similar to

S. aureus, the DHP-1+NO sample displayed maximum reduction against P. aeruginosa

(p < 0.01). The modified samples did not show significant increase in percentage of dead (red-stained) cells compared to the control samples.

Bacterial coverage - P. aeruginosa

18 16 14 × 12 Dead 10 # * Bacteria 8 * ^ * 6 * Live

4 bacteria Bacterial coverage (%) Bacterial 2 0

Figure 5.14: Percentage bacterial coverage for P. aeruginosa PA01 (mean ± standard error of mean); *indicates p < 0.001 and ×indicates p < 0.1 compared to spermine control; ^indicates p < 0.05 compared to DHP-1; #indicates p < 0.01 compared to other

DHP modified surfaces.

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5.4 Discussion

In the current investigation, the combined effect of NO and DHPs on bacterial colonization of surfaces was systematically examined. Acrylate analogues of DHPs

(DHP-1 and DHP-2) were synthesized and then covalently grafted onto amine- functionalized FEP surfaces via the Michael addition reaction. The secondary amine groups of the spermine polyamine linker were reacted with NO to form NONOate moieties. This is the first study to develop dual-action biomaterials by grafting DHPs and NO donors and investigating its effect in reducing bacterial adhesion.

The successful attachment of DHPs and the incorporation of NO donors on the FEP surface was monitored by XPS analysis. Both the DHP-acrylate derivatives gave similar attachment efficiency as indicated by their percentage of elemental composition. Further treatment with NO resulted in the incorporation of NONOate donors which was confirmed by the presence of characteristic peaks for NO in the XPS N 1s narrow scan.

The contact angle values also increased by 3 to 6° after addition of NONOate groups on the surfaces.

In order to evaluate the effect of the new DHP and NONOate-conjugated surfaces, the surfaces were tested for their ability to release NO and reduce bacterial adhesion. The efficacy of NO-releasing surfaces is greatly dependent on the concentration of NO produced under physiological conditions. Studies have shown that polyamine- conjugated NONOates undergo a first order decomposition reaction in solution, producing approximately 2 equiv of NO per mole of NONOate [245]. In this project, it was found using the Griess assay that spermine+NO, DHP-1+NO and DHP-2+NO surfaces had total NO release of 547.2 ± 1.8, 577.7 ± 1.2 and 636.1 ± 0.7 nmol/mm2 of the surface respectively after 24 hours. The NONOate surfaces showed slow rate of

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dissociation instead of spontaneous decomposition in presence of buffered solution. An extended NO release profile is important for biomedical applications, as to protect materials from bacterial fouling over long periods of time.

The DHP and DHP-NONOate FEP surfaces exhibited high antibacterial activity in reducing colonization of S. aureus and P. aeruginosa. With covalently-bound DHPs alone, bacterial colonization was significantly reduced compared to the spermine control (p < 0.001). Amongst the two DHP compounds, the unsubstituted DHP compound (DHP-1) was more active against bacterial adhesion (reduction of 70.9 % against S. aureus and 61.9 % against P. aeruginosa) compared to the fluorine- substituted DHP (60.3 % against S. aureus and 56.7 % against P. aeruginosa). This correlates with previous findings where DHP-1 showed the best activity on glass substrates, with reductions of up to 79.3 % and 65.8 % against S. aureus and P. aeruginosa respectively, as well as on polymer beads with 5-log unit reduction of S. aureus bacterial count [291,293]. As expected, the addition of NO donors on the DHP- modified samples resulted in further improvements in activity compared to the DHP coatings alone. Overall, the DHP-1+NO coating displayed the best broad spectrum antibacterial activity (83.1 % against S. aureus and 77.1 % against P. aeruginosa) amongst all the modified samples.

To verify the effect of NO-releasing groups on their own, the efficacy of spermine+NO surface was also evaluated against both bacterial strains. The spermine+NO surface was marginally effective at reducing surface adhesion of only P. aeruginosa compared to the spermine control, which was consistent with previous data showing that spermine- conjugated NONOate donors could prevent cell attachment and biofilm formation of P. aeruginosa [304]. However, against S. aureus, there was no significant difference in

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bacterial adhesion between spermine+NO and the spermine control. This is contrary to certain reports where NO alone has shown to significantly repress S. aureus biofilm formation [230,231,296]. Taken together, these data indicate that NO alone is not sufficient for imparting activity, and the broad-spectrum efficacy displayed by DHP-

NONOate surfaces was due to the combined effect of both DHPs and NO.

In addition to gradual release of NO, another important criterion for biomedical application is the nature of the surface obtained after complete release of NO. In the case of DHP+NO substrates, the complete release of NO would leave the DHP compounds still grafted on the surface, which are potent even without NO as shown in the data above. In contrast, the decomposition of spermine+NO would only leave the spermine linker attached to the surface, which does not have any antibacterial activity on its own. Furthermore, the image analysis of the confocal microscopy images did not show any significant increase in the proportion of the dead bacteria on the DHP 1-2 as well as DHP 1-2+NO samples. This indicates that neither DHP alone nor DHP in concert with NO had an additional killing effect on bacteria. The non-growth inhibitory activity displayed by both DHP and DHP+NO samples is consistent with previous DHP and NO-releasing coatings which exhibited bacterial reduction without bacterial killing

[291,304,305,350]. This non-toxic mechanism of action is important, as it may not stimulate resistance.

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5.5 Conclusion

In this project, novel NONOate-donating DHP biomaterial surfaces were successfully developed. The combined effect of DHP and NO resulted in significant improvement in reducing surface colonization of common pathogenic bacteria, S. aureus and P. aeruginosa, compared to the base DHP-coated surfaces, without exhibiting toxicity towards bacteria. The DHP-1+NO surface in particular gave the best broad-spectrum activity, with about 80 % reduction in bacterial adhesion compared to the control. The results strongly indicate that the novel dual-action hybrid surface coatings have high potential to be used for prevention of bacterial cell attachment, and therefore these hybrid coatings can be readily transferred to other biomaterial surfaces as well as polymers.

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Dual-Action Antibacterial Polymers Based on DHP and Nitric Oxide

CHAPTER-6

CHAPTER-6 Dual-Action Antibacterial Polymers Based on DHP and Nitric Oxide

6.1 Introduction

In the past few years, the use of polymers as drug delivery vehicles has become an integral part of polymer therapeutics and nanomedicine. Duong et al. reported the synthesis of NO-releasing cross-linked star polymer which exhibited enhanced stability and efficacy in preventing biofilm development process of P. aeruginosa [304]. In addition to the increased stability of NO donors in the polymer matrix, the main advantage of NO-incorporated polymers compared to small molecule NO donors is its ability to protect the donor moieties from multiple triggers such as heat, light and enzymes. Therefore, incorporation of NO into polymers is a promising avenue to treat biofilm-related infections.

In the previous chapter, a potent QS inhibiting compound, dihydropyrrolone (DHP), was combined with NO-releasing N-diazeniumdiolate (NONOate) donors to develop novel and highly efficient dual-action biomaterials, with the DHP-1+NO surface exhibiting excellent broad-spectrum antibacterial activity. It was envisaged that a similar positive synergy could be generated by combining QS inhibitors with the delivery of NO through polymeric structures, thereby allowing for the concomitant delivery of multiple payloads. Therefore, by employing the same attachment method as for surfaces, novel dual-action polymeric carriers were designed in this work by encapsulating acrylate-functionalized DHP compound (DHP-1) and NONOate-donating molecules within the same polymer.

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It was hypothesized that by attaching to a single macromolecule, this strategy could offer a high DHP and NO loading capacity, resulting in significant increase in the local concentration of DHP and NO hence leading to a great improvement in antibacterial efficacy. Moreover, these hybrid polymers may retain their activity even after complete

NO release, making them highly versatile antibacterial agents. Further advantages of polymer based delivery systems over administration of antibacterial drugs are improved pharmacokinetic properties, reduction in side effects, prolonged release of incorporated drug and prevention of developing multi-drug resistance [407–410].

Polymers are mechanically and chemically robust, and possess a high degree of flexibility that allows the introduction of a variety of functional groups. The approach that was used in this project for the synthesis of polymers was reversible addition fragmentation chain transfer (RAFT) polymerization (a type of living polymerization) coupled with the post-modification of polymers [411,412]. RAFT polymerization was first developed in 1990 by a CSIRO team [413–415]. This method of polymerization was chosen for its ability to control molecular weight and polydispersity index, as well as its ability to produce a wide range of stars, combs, brush and block copolymers with ease compared to other polymerization techniques [416,417]. In general, the RAFT process involves a RAFT agent which facilitates the addition of monomer units into its

C-S bond (Figure 6.1), to generate a polymeric chain known as a macro-RAFT agent

[415]. As the macro-RAFT agent contains an active RAFT end-group, it can undergo further polymerization in the presence of a new monomer to form a block polymer

[418].

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Figure 6.1: The overall RAFT polymerization technique [415].

The general procedure for the synthesis of a dual-action DHP and NO-releasing polymer is shown in Scheme 6.1. The RAFT agent employed in this chapter was a trithiocarbonate (RAFT), which was first polymerized with the monomer oligo(ethylene glycol) methyl ether acrylate (OEGA) using 2,2′-azobisisobutyronitrile

(AIBN) as the initiator, to form the macro-RAFT agent, poly[oligo(ethylene glycol) methyl ether acrylate] (POEGA). The advantage of a trithiocarbonate-based RAFT agent is its easy preparation even in the presence of other functional groups [418]. The

POEGA macro-RAFT was then subjected to polymerization reaction with 4- vinylbenzyl chloride (VBC) to introduce chlorine pendant functional groups into the polymer. The block copolymer (POEGA-b-PVBC) was subsequently modified by conjugating with a di-amino functionalized linker, spermine, via nucleophilic substitution of the chlorine groups to form spermine-conjugated polymer (p-sper). The acrylate-functionalized DHP was then coupled with the primary amino groups of the spermine linker by Michael addition reaction to form the DHP polymer (p-DHP).

Finally, the secondary amines present in p-DHP polymer chain were reacted with NO gas under pressure to form DHP-NONOate polymer (p-DHP+NO). The p-sper was also reacted with NO to form p-sper+NO polymer, which was used as a control to compare the NO release properties and biological efficacy of p-DHP+NO. This non-toxic, biocompatible multi-delivery p-DHP+NO polymer was tested for its biofilm inhibition activity against P. aeruginosa by crystal violet staining and confocal microscopy. - 168 -

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Scheme 6.1: Synthetic approach for the preparation of DHP-NONOate polymer (p-

DHP+NO) via RAFT polymerization.

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6.2 Materials and Methods

6.2.1 General

Oligo(ethylene glycol) methyl ether acrylate (480 g/mol, OEGA), 2-

(dodecylthiocarbonothioylthio)propionic acid (350.60 g/mol, 97 %, RAFT) and 4- vinylbenzyl chloride (152.62 g/mol, 90 %, VBC) were purchased from Sigma-Aldrich and used as received. Spermine (202.35 g/mol, 97 %) was purchased from Alfa Aesar and used as received. 2,2´-Azobisisobutyronitrile (164.21 g/mol, AIBN) was purchased from Sigma-Aldrich, crystallized from methanol and stored at 0 °C before use. Toluene, diethyl ether, methanol, petroleum spirit and acetone were used without further purification. Deuterated solvents, CDCl3 and DMSO-d6 were obtained from Cambridge

Isotope Laboratories, Inc. High purity N2 (Linde gases) was used for degassing.

Ultrapure deionized water was obtained using a MilliQ purification system.

6.2.2 Synthesis of POEGA Macro-RAFT Agent

A mixture of OEGA (9 g, 18.7 × 10-3 mol), RAFT (0.26 g, 7.5 × 10-4 mol) and AIBN

(0.024 g, 1.5 × 10-4 mol) with the ratio of [OEGA]:[RAFT]:[AIBN] = 25:1.0:0.1 was prepared in toluene (7.5 ml) in a 25 ml round bottom flask, equipped with a magnetic stirrer. The flask was sealed with a rubber septum and purged with nitrogen gas for 30 min at 0 °C. The degassed solution was immersed in a pre-heated oil bath at 70 °C for 5 h. The monomer conversion was determined by 1H NMR analysis. The reaction was then cooled to 4 °C overnight to terminate polymerization. The crude polymer was precipitated in excess of petroleum spirit and centrifuged (6000 rpm for 1 min). The centrifugation step was repeated two times by adding fresh petroleum spirit to remove any traces of unreacted monomer and the resultant polymer was dried under vacuum at room temperature. The samples were stored at 4 °C until required for further chain

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extension. POEGA was analyzed by 1H NMR and size exclusion chromatography

(SEC).

6.2.3 Synthesis of POEGA-b-PVBC Block Copolymer

POEGA was used as a macro-RAFT agent for chain extension with VBC.

-4 [POEGA]:[VBC]:[AIBN] = 1:25:0.25. POEGA (Mn(NMR) = 9950 g/mol; 8.0 g, 8.0 × 10 mol), VBC (3.06 g, 20.1 × 10-3 mol), AIBN (0.03 g, 20.1 × 10-5 mol) and toluene (7.5 ml) were prepared in a 20 ml round bottom flask, equipped with a magnetic stirrer bar.

The flask was sealed with a rubber septum and purged vigorously with nitrogen gas for

30 min at 0 °C. The reaction mixture was immersed in a pre-heated oil bath at 70 °C for

42 h and monomer conversion monitored by 1H NMR. The reaction was then cooled to

4 °C to terminate polymerization. The polymer was purified by precipitation in excess of diethyl ether and centrifugation (6000 rpm for 1 min). The centrifugation step was repeated two times by adding fresh ether and then the resultant polymer was dried under vacuum at room temperature. The block copolymer was characterized by 1H NMR and

SEC.

6.2.4 Conjugation of POEGA-b-PVBC to Spermine (p-sper)

-5 POEGA-b-PVBC copolymer (Mn(NMR)= 12392 g/mol; 1.0 g, 8.0 × 10 mol) was dissolved in methanol (5.5 ml) containing spermine (1.8 g, 9.0 × 10-3 mol) in 7:1 spermine/polymer mole ratio in a 10 ml round bottom flask, equipped with a magnetic stirrer bar. The reaction mixture was stirred at room temperature for 24 h and monitored by 1H NMR. After completion, the reaction medium was purified by dialysis using a cellulose membrane in 0.1 M NaCl and MilliQ water. The dialyzed polymer (p-sper) was freeze-dried for 3–4 days and analyzed by 1H NMR.

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6.2.5 Attachment of DHP to Spermine-Conjugated Polymer (p-DHP)

1-Acryloyl-4-(4-phenyl)-5-methylene-1,5-dihydro-2H-pyrrol-2-one (DHP-1; 225.08 g/mol) was incorporated into the spermine-conjugated POEGA-b-PVBC (Mn(THEO) =

15629 g/mol) copolymer. A solution of DHP-1 (0.09 g, 4.2 × 10-4 mol) and p-sper (0.4 g, 2.5 × 10-5 mol) in a 16:1 DHP-1/polymer mole ratio was prepared in absolute ethanol

(7 ml) and stirred at room temperature for 24 h. The reaction was monitored by TLC which indicated completion of reaction by lack of free DHP compound in the reaction mixture. The solvent was evaporated under vacuum and the resulting polymer was dried under reduced pressure at room temperature. The polymer, p-DHP, was analyzed by 1H

NMR and used for further reaction with NO gas to introduce NO releasing NONOate moiety.

6.2.6 Attachment of NONOate to DHP-Conjugated Polymer (p-DHP+NO)

The p-DHP (0.2 g; Mn(THEO) = 19231 g/mol) was dissolved in methanol/THF (10 ml) and placed in a Parr apparatus and clamped. The apparatus was then purged and evacuated with nitrogen three times and pressurized with NO gas to 5 atm at 25 °C for

48 h to form NONOate NO donors. Excess NO was then vented through purging with nitrogen gas and solvent was evaporated under vacuum. The DHP-NONOate polymer

(p-DHP+NO; Mn(THEO) = 21151 g/mol) was then dried under reduced pressure and then stored at 4 °C until required for further analysis.

Similarly, the spermine-conjugated polymer (0.2 g) was dissolved in methanol/THF (5 ml) and reacted with NO gas as described above (p-sper+NO; Mn(THEO) = 17550 g/mol).

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6.2.7 Determination of Nitric Oxide Release by Griess Assay

NO released from the polymer at specified time intervals was determined using a standard Griess reagent kit (G-7921, Molecular Probes), which is normally used for nitrite determination [419]. NONOate readily releases NO upon contact with water at physiological pH. Typically, 2 mg p-DHP+NO polymer sample was dissolved in 1 ml of phosphate buffered saline (PBS). The solution was enclosed in a sealed dialysis membrane (Cellu-Sep 3500 MWCO) that allows free diffusion of NO. The membrane was then immersed in 6 ml PBS and incubated at 37 °C. At various time points, a 150 µl aliquot from the PBS solution was taken for determining the concentration of NO. In a

96-well plate, the test aliquot was added to 20 µl of Griess reagent freshly prepared by mixing equal volumes of reagent A and B. The mixture was topped up with 130 µl of

PBS to make up a total volume of 300 µl and then incubated for 30 min at 500 rpm. The

UV-Vis absorbance of the solution was measured at 548 nm using a FLUOstar® Omega

Microplate Reader (BMG Labtech). The procedure was repeated at different time points and the total nitrite concentration in the sample solutions was calculated from a standard curve prepared using sodium nitrite.

6.2.8 Characterization by 1H NMR Spectroscopy

1H NMR spectra were recorded using Bruker Avance 400 or a Bruker AVANCE DMX

600 spectrometers. CDCl3 and DMSO-d6 were used as solvents. All chemical shifts are quoted in parts per million (ppm) and referenced to residual the residual solvent

1 frequencies ( H NMR: CDCl3 = 7.26 ppm, DMSO-d6 = 2.50 ppm).

The OEGA monomer conversion into a polymer chain was determined from 1H NMR spectroscopy by the following equation: αPOEGA = 1 – [(ʃ5.8-6.2 ppm/ʃ3.4 ppm) × 100] where ʃ5.8-6.2 ppm is the peak integral of the monomer protons (vinyl proton CH=CH2,

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3H) and ʃ3.4 ppm is the peak integral from the monomer/polymer (methoxy proton

OCH3, 3H).

The average number of monomer units integrated into the polymer chain (nPOEGA) was

POEGA POEGA calculated as follows: n = α × [OEGA]0, where [OEGA]0 is initial monomer concentration.

The experimental molecular weight, Mn(NMR), of the POEGA polymer was calculated from 1H NMR by adding the molecular weight of the OEGA units integrated into the polymer chain (n × MWOEGA) to the molecular weight of the RAFT agent (MWRAFT), as follows: Mn(NMR) = (n × MWOEGA) + MWRAFT

The theoretical molecular weight, Mn(THEO), of POEGA was calculated from the following equation: Mn(THEO) = [OEGA]0/[RAFT]0 × MWOEGA + MWRAFT, with [OEGA]0 and [RAFT]0 corresponding to the initial concentrations of OEGA and RAFT respectively.

VBC monomer conversion was determined via 1H NMR analysis by the following equation: αVBC = 1 – [ʃ5.7 ppm/(ʃ4.5 ppm/2)], where ʃ5.7 and ʃ4.5 represents the peak integrals of monomer (vinyl proton, 1H) and the monomer/polymer (benzylic proton,

2H).

The average number of VBC units integrated into the polymer chain (nVBC) was calculated as follows: nVBC = (ʃ4.5 ppm/ʃ4.1 ppm) × nPOEGA, where ʃ4.5 ppm and ʃ4.1 ppm correspond to the integration of signals for the benzylic proton (–CH2Cl, 2H) and the ester (–OCH2, 2H).

The experimental molecular weight, Mn(NMR), of the POEGA-b-PVBC polymer was

1 VBC calculated from H NMR analysis as follows: Mn(NMR) = (n × MWVBC) + MWPOEGA, - 175 -

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where MWVBC and MWPOEGA correspond to the molecular weights of VBC and

POEGA, respectively.

6.2.9 Size Exclusion Chromatography (SEC)

SEC analysis of polymer samples was performed in tetrahydrofuran (THF, HPLC grade) at 50 °C at a flow rate of 1 ml min-1 with a Shimadzu modular system comprising an SIL-10AD automatic injector, a Polymer Laboratories 5.0 µl bead-size guard column

(50 × 7.8 mm) followed by four linear PL (Styragel) columns (105,104,103 and 500 Å) and an RID-10A differential refractive-index detector. The SEC calibration was performed with narrow-polydispersity polystyrene standards ranging between 104 and 2

× 106 g/mol. Polymer solutions at 2–3 mg ml-1 were prepared in the eluent and filtered through 0.45 µm filters prior to injection.

6.2.10 Attenuated Total Reflectance-Fourier Transform Infrared Spectroscopy (ATR-

FTIR)

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. The polymer samples were pre-dried as a thin film for ATR-FTIR analysis.

6.2.11 X-Ray Photoelectron Spectroscopy (XPS)

The polymers were characterized using X-ray photoelectron spectroscopy

(ESCALAB220-iXL, VG Scientific, West Sussex, England). The instrument employs a monochromatised Al Kα X-ray source and the photo-energy was 1486.6 eV with a source power of 120 W. Vacuum pressure was ≤10-8 mbar.

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6.2.12 Biofilm Inhibition Assay

The laboratory strain P. aeruginosa PA01 was used to characterize the effects of NO and/or DHP-conjugated polymers on biofilm formation. Biofilms were grown as previously described with some modifications [250]. Briefly, in all assays, overnight cultures in TSB medium were diluted to an OD600 of 0.005 in 1 ml M9 minimal medium

(containing 48 mM Na2HPO4, 22 mM KH2PO4, 9 mM NaCl, 19 mM NH4Cl, 2 mM

MgSO4, 20 mM glucose, 100 mM CaCl2, pH 7.0) in 24-well plates (Costar, Corning®).

Various treatments including DHP-NONOate polymer (p-DHP+NO), spermine-

NONOate polymer (p-sper+NO), DHP-conjugated polymer (p-DHP) and free DHP were added to the wells to obtain final concentrations of 42 µM, 21 µM, 10.5 µM, 5.2

µM, 2.5 µM and 1 µM in dry DMSO. The plates were incubated at 37 °C with shaking at 180 rpm in an orbital shaker Ratek model OM11 (Boronia, Australia) for 6 h to allow biofilms to grow for 6 h on the wall and/or bottom of the culture wells.

After 6 h, the planktonic biomass was quantified by siphoning off the supernatant and measuring its OD600. The remaining biofilm biomass or cells adhered to the wells were determined by crystal violet staining. The biofilm on the well surfaces was first washed with 1 ml PBS, before adding 0.03 % crystal violet stain (1 ml) made from a 1:10 dilution of Gram crystal violet (0.3 %) in PBS. The plates were incubated for 20 min, and the wells were washed twice with PBS. Photographs of the stained biofilms were recorded using a digital camera. The amount of crystal violet-stained biofilm was quantified by adding 1 ml of 100 % ethanol and measuring the OD550 of the homogenized suspension using FLUOstar® Omega Microplate Reader (BMG Labtech).

OD measurements of control wells where no bacteria were added at the beginning of the experiment were subtracted from all values.

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6.2.13 Confocal Microscopy Analysis

For biofilm inhibition analysis, P. aeruginosa biofilms were grown in 24-well plates containing sterile glass in presence of polymer samples. After 6 h incubation, biofilms were rinsed twice with PBS before being stained with LIVE/DEAD® BacLight™ bacterial viability kit reagents (L-7007, Molecular Probes) according to the manufacturers’ procedure. Bacteria were then fixed by adding 50 µl of 4 % formaldehyde to each sample. Microscopic observation and image acquisition were performed with an Olympus FV1200 Confocal Inverted Microscope. Images from 10 representative areas on each of triplicate samples for each sample were taken. Cells that were stained green were considered to be viable, those that stained red and stained both green and red were considered to be non-viable.

6.2.14 Statistical analysis

All assays included at least two replicates and were repeated in two independent experiments. Statistical analysis were performed with GraphPad Prism 7.03 (GraphPad

Software) using one-way ANOVA followed by Tukey multiple comparison test comparing treated samples to the untreated control.

6.3 Results

6.3.1 Synthesis and Characterization of Polymers

The synthesis of block polymer was achieved by RAFT polymerization as shown in

Scheme 6.1. The macro-RAFT agent, poly[oligo(ethylene glycol) methyl ether acrylate]

(POEGA), was first synthesized using olig(oethylene glycol) methyl ether acrylate

(OEGA) as the monomer and 2-(dodecylthiocarbonothioylthio)propionic acid (RAFT) as the chain transfer RAFT in presence of a radical initiator 2,2′-azobisisobutylonitrile

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(AIBN), with a ratio of [OEGA]:[RAFT]:[AIBN] = 25:1:0.1 in toluene at 70 °C. The monomer conversion percentage was determined from the NMR spectrum of the crude polymer, by comparing the sum of signals for the vinyl group protons of the unreacted monomer (5.8–6.2 ppm) with the signals for the terminal methoxy group at 3.4 ppm of the monomer/polymer (Figure 6.2A). The conversion was about 90 % after 5 h of

1 reaction. The average molecular weight based on H NMR (Mn(NMR)) with approximately 20 repeating units was found to be 9950 g/mol. Size exclusion chromatography (SEC) was also performed which indicated average molecular weight,

Mn(SEC) of 9400 g/mol and a low polydispersity index (PDI) of 1.12. The average molecular weight obtained from NMR and SEC was found to be lower than the theoretical molecular weight value corresponding to 25 monomer units (Mn(THEO) =

12350 g/mol).

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Figure 6.2 : 1H NMR spectra of (A) POEGA, (B) POEGA-b-VBC, (C)p-sper and (D) p-

DHP polymers.

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The POEGA homopolymer chain was extended with 4-vinylbenzyl chloride (VBC) as the monomer using a molar ratio of [POEGA]:[VBC]:[AIBN] = 1:25:0.25. VBC was polymerized for 42 h at 70 °C in toluene to form the block copolymer, POEGA-b-

PVBC. 1H NMR analysis indicated ~74 % VBC conversion with approximately 16 repeating units of VBC per polymer chain, with the presence of a singlet for the benzyl group at 4.5 ppm and signals for aromatic protons from 6.4–7.1 ppm confirming the incorporation of VBC (Figure 6.2B). The pure block copolymer was characterized by

SEC, which revealed a significant shift in molecular weight, from 9400 g/mol to 11 420 g/mol, as well as a PDI of 1.16, which are consistent with successful extension of the polymer chain. ATR-FTIR analysis was also performed on the polymer sample, which showed a characteristic signal at 690 cm-1 that was attributed to the C-Cl bond present in

POEGA-b-PVBC [295,420].

The benzyl chloride (–CH2Cl) group of POEGA-b-PVBC was conjugated with the polyamine spermine linker via nucleophilic substitution by reacting in methanol at room temperature for 24 h, followed by successive dialysis against 0.1 M NaCl and water.

The successful formation of the spermine polymer (p-sper) was confirmed from 1H

NMR spectroscopy, which showed the disappearance of the benzyl chloride proton at

4.5 ppm and the appearance of new peaks at 1.2–1.7 ppm and 3.5 ppm attributed to the spermine groups (Figure 6.2C). FTIR spectroscopy also revealed the disappearance of the characteristic peak for chlorine at 690 cm-1, and showed the presence of a broad absorption band at around 3486 cm-1 corresponding to secondary amine (N-H). This was consistent with the presence of secondary amine groups in the polymer after successful spermine conjugation [295,304].

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In the next step, the DHP molecule was incorporated into the spermine polymer chain to form the DHP-conjugated polymer (p-DHP) via Michael addition reaction between the acrylate-functionalized DHP and the primary amine group of spermine. After stirring together in absolute ethanol for 24 h at room temperature, the successful addition of the

DHP compound was confirmed by the appearance of signals at 7.4–7.5 ppm in the 1H spectrum corresponding to the aromatic protons of the DHP, respectively (Figure

6.2D). The FTIR spectrum showed the appearance of a small peak at around 1670 cm-1 for the carbonyl group of the DHP moiety.

The secondary amine groups present in the spermine and DHP-immobilized polymers were subsequently reacted with NO gas for 48 h at 75 psi (5 atm) to form NONOate groups, to give p-sper+NO and p-DHP+NO polymers, respectively [421]. The FTIR spectra of the NONOate-conjugated polymers showed a decrease in intensity of the secondary amine N-H signal at 3486 cm-1, and the appearance of a new signal at 1374 cm-1 for p-DHP+NO and around 1350 cm-1 for p-sper+NO corresponding to N-O, confirming successful attachment of NONOate on the polymer chain [304].

The NO release profile from the NONOate donors was performed using the Griess assay in phosphate buffered saline (PBS), pH 7.4 at 37 °C to mimic physiological conditions [422–424]. The NO (nitrite) concentration was calculated from the standard curve obtained from 0–100 µM sodium nitrite solutions. As shown in Figure 6.3, NO release from the p-DHP+NO polymer was initially slow, with a very low nitrite concentration in the first few hours, however, the NO release accelerated after 16 h and the nitrite concentration reached to 47 µM at 48 h. In contrast, the NO release from p- sper+NO was initially higher compared to the p-DHP+NO, which could be due to NO donors being attached to primary as well as secondary amines. However, after 24 h, the

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nitrite concentration of p-sper+NO was less than p-DHP+NO, and at 48 h the maximum nitrite released was around 33 µM.

NO release 60

20 p-DHP+NO

10 p-sper+NO Nitrite concentration (µM) concentration Nitrite

0 0 1 3 8 16 24 45 48

Figure 6.3: Nitrite concentration (µM) from p-DHP+NO and p-sper+NO polymers at

pH 7.4 at 37 °C as determined using the Griess assay. The concentration of both the

polymers is 2 mg/ml (experiments were performed in duplicates).

6.3.2 High-Resolution XPS Analysis of Polymers

The polymers were characterized by XPS after every step to determine the changes in the elemental composition after successful attachment of DHP and NONOate (Table

6.1).

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Table 6.1: XPS analysis of polymers after every modification step

Polymer % C % N % O % Cl

POEGA 69.1 - 30.9 - POEGA-b-PVBC 72.1 - 26.7 1.2 p-sper 69.8 2.5 27.3 0.4 p-sper+NO 68.8 2.7 28.2 0.3 p-DHP 71.7 2.3 25.7 0.3 p-DHP+NO 72.5 3.2 24.0 0.3

The XPS data for the POEGA macro-RAFT agent showed the presence of carbon (69.1

%) and oxygen (30.9 %), which is consistent with its molecular structure. The subsequent incorporation of VBC units was confirmed by the appearance of a chlorine signal (1.2 %). After the benzyl chloride of the block co-polymer was reacted with spermine, the chlorine intensity decreased to 0.4 % while there was an emergence of 2.5

% of nitrogen, consistent with the presence of the amine groups of the incorporated spermine linker. However, complete attenuation of the chlorine signal was not obtained, which indicates that not all the VBC groups reacted with spermine.

Attachment of the DHP molecule subsequently led to an increase in the carbon content from 68.8 % to 71.7 % along with slight decrease in values for nitrogen and oxygen.

Finally, the successful incorporation of NONOate groups into the p-sper and p-DHP polymers was confirmed by increase in nitrogen composition along with appearance of a peak at ~406.2 eV in the N 1s narrow scan spectrum corresponding to the NONOate groups. The XPS N 1s scan for p-DHP is shown in Figure 6.4A which shows the

+ presence of three species, namely N-H, N-C=O and NH3 /tertiary nitrogen. After

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reaction with NO gas, a new peak arose at 406.2 eV corresponding to NO incorporated in the polymer chain (Figure 6.4B).

(A) p-DHP polymer

(B) p-DHP+NO polymer

Figure 6.4: XPS N 1s narrow scan of (A) p-DHP polymer and (B) p-DHP+NO

polymer. Appearance of NO peak at 406 eV was observed for p-DHP+NO.

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6.3.3 P. aeruginosa Biofilm Inhibition

The biofilm inhibition activity of the dual-action DHP-NONOate polymer was investigated against a biofilm-forming Gram-negative bacterium, P. aeruginosa PA01.

For biological testing, biofilms of P. aeruginosa were grown in M9 minimal media for

6 h in the presence of 1 to 42 µM solutions of various treatments including DHP-

NONOate polymer (p-DHP+NO), spermine-NONOate polymer (p-sper+NO), DHP- conjugated polymer (p-DHP) and free DHP-acrylate compound (DHP). The effect of the samples on biofilm inhibition was assessed by crystal violet staining, while the amount of planktonic biomass was evaluated by determination of OD600 values. The biofilm biomass determined from crystal violet staining assay for all the samples is shown in Figure 6.5A and the planktonic biomass data is shown in Figure 6.5C.

Representative photograph images of stained biofilms treated with different concentrations of p-DHP+NO polymer are shown in Figure 6.5B.

The DHP-NONOate dual-action polymer (p-DHP+NO) was found to exhibit strong biofilm inhibition after 6 h at all concentrations, resulting in 95.4–96.1 % reduction in biofilm biomass compared to the untreated biofilm control (p < 0.001) (Figure 6.5A).

Figure 6.5B shows the effect of the polymer by very low staining of the wells treated with 1–42 µM p-DHP+NO compared to the control well. The NO-releasing polymer without the active DHP i.e. spermine-NONOate (p-sper+NO) also prevented biofilm formation in a dose-dependent manner, though it was less active than p-DHP+NO, with

84.0 % reduction of biofilm formation at concentrations equal or greater than 10.5 µM

(p < 0.001), but only 67.2 % inhibition at 2.5 µM (p < 0.05) and 25.6 % inhibition at 1

µM. The DHP-conjugated polymer without NONOate donor (p-DHP) induced 81.0 % reduction in biofilm biomass (p < 0.001) at 42 µM, however, inhibition declined to

10.6–48.1 % at concentrations between 2.5 and 21 µM. The free DHP compound - 186 -

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showed activity similar to the p-DHP polymer, with 89.2 % dispersion of the bacterial cells at 42 µM (p < 0.001) and less than 40 % biofilm inhibition at concentrations of

10.5 µM or lower. However, at a sample concentration of 1 µM, both p-DHP polymer and free DHP compound were found to be ineffective in preventing biofilm formation.

The planktonic biomass of all the samples was also investigated. A high number of bacterial cells in planktonic mode is an indication of more number of cells being dispersed from the biofilm. As shown in Figure 6.5C, the number of planktonic cells in the supernatant solution of the wells treated with p-DHP+NO was 5.3–32.0 % higher compared to untreated culture across all concentrations (1–42 µM), indicating high transition from biofilm to planktonic mode which is due to the non-toxic effect of DHP and NO. Similarly, the NO-donating spermine polymer (p-sper+NO) induced between

9.7–34.2 % increase in planktonic cells across all concentrations (1–42 µM). The lack of toxicity for p-sper+NO is consistent with a previous report on a spermine-conjugated

NONOate star polymer, which also did not have a killing effect against the P. aeruginosa biofilm [304]. The p-DHP polymer caused a small and insignificant decrease in the planktonic biomass of around 3.9–10.3 % at all concentrations, except at

42 µM which showed a 35.1 % increase in planktonic bacteria which is consistent with high antibiofilm activity of 42 µM p-DHP. The biofilm treatment with DHP led to reduction in planktonic growth by 24.8–34.1 % across all concentrations compared to the control, indicating that under these growth conditions free DHP might have exhibited some level of toxicity.

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(A)

Biofilm Biomass - P. aeruginosa

) 550 3.5

2.5 p-DHP+NO

2 p-DHP Free DHP 1.5 p-sper+NO 1

0.5

Biofilm Biomass (Crystal Violet, Violet, OD (Crystal Biofilm Biomass 0 Control 1 2.5 5.2 10.5 21 42 Compound concentration (µM)

(B)

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(C)

Planktonic Biomass - P. aeruginosa 0.4

) 0.35 600 0.3

0.25 p-DHP+NO

0.2 p-DHP Free DHP 0.15 p-sper+NO

0.1 Planktonic Biomass(OD Planktonic 0.05

0 Control 1 2.5 5.2 10.5 21 42 Compound concentration (µM)

Figure 6.5: (A) Biofilm biomass analyzed by crystal violet staining in presence or absence of 1–42 µM DHP-NONOate polymer (p-DHP+NO), spermine polymer (p- sper+NO), DHP-conjugated polymer (p-DHP) and free DHP compound; (B)

Photograph images of stained biofilms treated with the indicated concentrations of p-

DHP+NO polymer; (C) Planktonic biomass in presence of polymers and DHP determined by measuring the supernatant at OD600.

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6.3.4 Confocal Microscopy Analysis

The ability of the p-DHP+NO polymer to prevent bacterial colonization and inhibit biofilm development of P. aeruginosa was also evaluated by confocal fluorescence microscopy. The biofilm in the cell culture wells were stained with Live/Dead staining kit, where the live bacteria are stained green and dead bacteria are stained red. The confocal microscopic images of control, free DHP, p-DHP, p-sper+NO and p-DHP+NO at 1 µM concentration are shown in Figure 6.6. The wells treated with control, free

DHP, p-DHP and p-DHP+NO exhibited high density of biofilm formation that was visually more compact and dense (Figure 6.6A, B, C and D) compared to biofilm grown in presence of p-DHP+NO (Figure 6.6E).

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P. aeruginosa PA01

A – Control B – Free DHP (1 µM)

C – p-DHP (1 µM) D – p-sper+NO (1 µM)

E – p-DHP+NO (1 µM)

Figure 6.6: Representative confocal micrographs showing P. aeruginosa biofilm stained with Live/Dead dye kit. Viable and non-viable bacteria appear green and red, respectively. Scale bar = 100 µm.

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The image analysis illustrated that bacterial cells grown in presence of 1 µM free DHP compound, DHP polymer, and spermine-NONOate polymer exhibited similar biofilm formation compared to the untreated control, with no statistical difference between them

(Figure 6.7). In contrast, cultures treated with 1 µM p-DHP+NO displayed excellent activity by markedly reducing biofilm formation compared to the other samples and control (p < 0.001). The percentage of dead cells did not vary significantly between the samples and control indicating there was no significant bactericidal activity. The results obtained from confocal microscopy are in agreement with biofilm inhibition assay determined by crystal violet staining.

Bacterial coverage - P. aeruginosa 24

16 Dead bacteria 12

Live 8 bacteria

Bacterial coverage (%) coverage Bacterial 4 *

0 Control p-DHP+NO p-DHP Free DHP p-sper+NO

Compound concentration 1µM

Figure 6.7: Percentage bacterial coverage for P. aeruginosa PA01 (mean ± standard

error of mean); *indicates p < 0.001 control.

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6.4 Discussion

The use of QS inhibitors and NO has potential in preventing biofilm formation, particularly in the context of medical implant or device-related infections. In this work, novel dual-action polymers were designed for the simultaneous delivery of DHP and

NO, which showed promising activity in inhibiting the formation of biofilms of P. aeruginosa, which plays a critical role in causing lung infections and infectious diseases related to biomedical devices.

The DHP polymer, spermine-NONOate polymer and DHP-NONOate were synthesized and characterized by various techniques such as NMR, FTIR and XPS spectroscopy. In particular, the formation NONOate groups in the polymer was confirmed by characteristic peaks at ~406.2 eV in XPS and around 1374 cm-1 in FTIR. NO release from the polymers under physiological-like conditions was quantitatively evaluated by testing 2 mg/ml of samples using the Griess assay, which showed controlled release of

NO for at least 2 days. While the p-sper+NO polymer initially showed higher NO release than p-DHP+NO, the final concentration of NO at 48 h was higher for p-

DHP+NO (47 µM) than for p-sper+NO (32 µM). According to literature reports, spermine-NONOates (SPER/NO) are usually fast NO donors having very short half- lives of less than an hour at pH 7.4 at 37 °C [245,425]. However, rapid release of NO in the first hour was not observed for both p-sper+NO and p-DHP+NO polymers in this study, in contrast to various NO donors reported in literature such as pyrrolidine-

NONOate (PYRRO/NO) or diethylamine-NONOate (DEA/NO) which have half-life time of only few minutes [242]. The more gradual and prolonged release of NO for the

NO-donating polymers prepared in this study could possibly be attributed to the

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encapsulation of NONOate groups in the core of the polymer, leading to enhanced stability.

The biofilm inhibition assay performed using crystal violet staining revealed that all the samples were able to significantly inhibit biofilm formation at concentrations of 21 µM or greater. The free DHP compound alone dispersed 74.4 % and 89.2 % of biofilm at 21 and 42 µM respectively, although surprisingly it also decreased the number of bacteria in the planktonic phase by about 30 % across all concentrations, suggesting some level of toxicity. Conjugating DHP into a polymer (p-DHP) also led to significantly reduced activity, with less than 50 % reduction in biofilm biomass at concentrations of 21 µM or below. However, it is important to note that unlike the free DHP compound, p-DHP induced only a small and insignificant toxic effect towards planktonic bacterial growth at concentrations of 1–21 µM.

The addition of NO-donating molecules to the polymer chain greatly improved the antibiofilm activity of the polymers at concentrations below 21 µM. Treatment with p- sper+NO polymers induced 84 % reduction in biofilm biomass at 10.5 µM, while increasing planktonic cells by 29.9 % indicating its lack of toxic effects. This non-toxic effect appears to be consistent with data published previously where spermine-

NONOate star polymer exhibited biofilm dispersion without affecting the planktonic cells [304]. Moreover, the 10.5 µM p-sper+NO was also found to be more effective in dispersing biofilm compared to the commercially available SPER/NO which led to only

30 % reduction in biofilm at 50 µM [305]. However, the activity of p-sper was reduced at lower sample concentrations of 1–5.2 µM. In contrast, the p-DHP+NO polymer exhibited excellent antibiofilm efficacy at all concentrations, with 95 % of biofilm eradicated even at a very low concentration of 1 µM, concomitant with 32.9 % increase

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in planktonic biomass. The p-DHP+NO at 1 µM was more effective than all concentrations of p-sper+NO and p-DHP. These results therefore suggest that the potency of the p-DHP+NO polymer was enhanced due to the combined dual-action effect of both DHP and NO, while still not being toxic to planktonic cells. The high efficacy of p-DHP+NO at preventing biofilm formation via a non-toxic mechanism at 1

µM compared to control and other samples was confirmed via confocal microscopy.

Moreover, the dual-action polymers developed in this chapter displayed significant improvement in activity compared to the dual-action FEP surfaces (Chapter 5) which were able to reduce surface colonization of P. aeruginosa by 77.1 %. The enhanced activity of the polymer could be attributed to its high degree of flexibility in delivering the desired amount of antibacterial agents (DHP and NO) in a sustained manner to a specific target site. The synthesis and biological activity of dual-action polymeric nanoparticles developed via the combination of an antibiotic, gentamicin, along with

NONOate donors was previously reported in literature [305]. Although these polymeric nanoparticles reduced 83 % of P. aeruginosa biofilm biomass, they also massively decreased planktonic viability by 94 % compared to control, which could trigger the development of resistance. In contrast, the non-bactericidal property of DHP-NONOate polymer is a desirable feature for antibacterial agents to avoid drug resistance in the long term.

Overall the biological data presented in this study demonstrated that the novel DHP-

NONOate hybrid polymers retained their efficacy to prevent biofilm attachment whilst remaining non-toxic to bacteria, potentially making them superior to existing NO donors for long-term use in preventing biomedical device-related bacterial infections.

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6.5 Conclusion

In this project, a novel polymeric system was synthesized for the co-delivery of potent

DHP compound and NONOate to the bacterial medium. These DHP+NO hybrid polymers showed positive synergistic effects in inhibiting biofilm formation of P. aeruginosa, and were much more effective than the conventional antibiotics and/or NO- donating compounds. The results suggest that the hybrid polymer was able to simultaneously exploit QS inhibition and NO release for controlling biofilm formation without exerting any bacterial toxicity. Hence, harnessing the combined effect of both

DHP and NO, encapsulated within the polymeric matrix, is an attractive strategy in combating bacterial infections in clinical settings.

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CHAPTER SEVEN

Summary and Future Perspectives

CHAPTER-7

CHAPTER-7 Summary and Future Perspectives

7.1 Summary

In this thesis, novel strategies were employed for immobilization of synthetic derivatives of QS inhibitors to develop antibacterial coatings. The first strategy involved non-specific covalent attachment of FU and DHP compounds on azide-terminated glass surfaces via azide/nitrene chemistry. Successful attachment was obtained by photoactivating the azide groups in presence of FU or DHP. All the coated surfaces were found to be effective in reducing colonization of S. aureus and P. aeruginosa, with

FU-2 and DHP-3 displaying the best activity. It was observed that the activity of DHP surfaces was not greatly affected by the surface concentration. The surfaces displayed potent activity even at low concentration. This difference in efficacy was possibly due to the change in orientation of the molecule during surface immobilization, by exposing the more active region to the bacteria.

To understand the effect of orientation of DHP after attachment, active DHP compounds were covalently linked to the surface from different points within the same molecule via a specific attachment strategy. In chapter 2, two types of DHP compounds were synthesized by introducing a carboxylic acid group at the N-atom of the lactam ring and the para position of the pendant phenyl ring. The compounds were attached by carbodiimide chemistry on amine-functionalized glass and FEP, thereby exposing different parts of the molecule to the bacterial medium. While all coated glass surfaces significantly reduced bacterial adhesion, the activity of the DHP attached from the phenyl ring (p-acid) did not vary significantly from N-substituted DHPs (DHP acid 1-4).

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CHAPTER-7

This shows that different molecular orientations of DHP (lactam and phenyl ring) are equally active in reducing bacterial colonization on glass. However, on FEP surfaces, p- acid DHP exhibited higher bacterial reduction of around 70 % against both S. aureus and P. aeruginosa than DHP acid-1 (52.7 % against S. aureus and 50.3 % against P. aeruginosa). This difference in activity indicated that exposing the free lactam to bacteria plays an important role in improving the antibacterial efficacy.

The next chapter focused on determining the antibacterial activity of DHP in absence of the characteristic exocyclic double at the C-5 position. Halogenated DHP analogues lacking an exocyclic double bond were synthesized having acid- and amine- functionalization and attached on amine- and acid-functionalized surfaces respectively using EDC/NHS. The results indicated that all the DHPs could reduce bacterial adhesion on surfaces but displayed comparatively lower activity than DHPs having an exocyclic double bond. However, it should be noted that complete loss of activity was not observed for any of the coated surfaces, with some compounds displaying ≥50 % reductions, suggesting that the C-5 methylene group may not be an essential pre- requisite for maintaining the activity of DHPs and further structural modifications might improve the activity of the compounds.

The efficacy of the surfaces was further examined by incorporating DHP and NO donating moieties on the surface in Chapter 5. It was hypothesized that these dual- action surfaces would improve the antibacterial performance of the surface coatings.

The coatings were prepared by immobilizing DHPs onto FEP substrates via Michael addition reaction and subsequently reacted with NO to incorporate N-diazeniumdiolate

(NONOate) groups. The dual-action surface coatings displayed significant improvement in activity compared to DHP coatings alone, with DHP-1+NO exhibiting best

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CHAPTER-7

antibacterial activity (83.1 % against S. aureus and 77.1 % against P. aeruginosa), while DHP-1 reduced 70.9 % of S. aureus cells and 61.9 % P. aeruginosa cells.

Furthermore, the low activity of NO coatings in absence of DHP confirmed that high efficacy of DHP+NO coatings was due to the combined effect of both DHP and NO.

The results also showed that the reduction displayed by DHP+NO coatings did not have any additional killing effect on both bacterial cells which is particularly important as the coatings are less prone to stimulating resistance.

In chapter 6, the same attachment strategies employed in chapter 5 were used to develop dual-action polymers. The biological data demonstrated that the novel p-DHP+NO polymer was highly effective in eradicating 95 % of P. aeruginosa biofilms at all concentrations including low concentration of 1 µM. Importantly, polymer with DHP alone and free DHP compound were found to be active only at high concentrations (42–

21 µM). While polymer incorporated with only NO showed biofilm dispersion activity in a dose-dependent manner from 42–1 µM. The results clearly showed that the potency of p-DHP+NO hybrid polymer was enhanced due to the dual-action effect of DHP and

NO against P. aeruginosa via a non-toxic mechanism, making them desirable for long- term use in preventing deice-related infections.

From the findings in this thesis, it can be concluded that surface bound DHPs and DHP-

NONOates show promising antibacterial activity and they can be applied as antibacterial coatings on surfaces of biomedical devices and implants.

7.2 Future Work

The DHP as well as DHP+NO coatings and polymers have shown excellent antibacterial activity. Though a previous study has demonstrated the ability of surface bound DHP in interfering with the AHL-regulated las QS system of P. aeruginosa - 199 -

CHAPTER-7

[294], the precise mechanism is still not very clear. A detailed transcriptome analysis of the changes in gene expression of relevant bacterial strains induced by surface attached

DHPs would be helpful in identifying the affected pathways and indicate mechanisms by which biofilm formation is controlled. The study also indicated the presence of membrane-based receptors on bacteria which possibly interact with surface bound

DHPs [294]. In order to elucidate the membrane-based pathway, a surface bound natural signalling molecule, AHL, will be required which theoretically will enhance the formation of biofilm if a membrane-based receptor is present. Similarly, for Gram-positive bacterium, S. aureus, the mode of action is still not fully understood. Therefore, additional genetic tools such as QS mutants of S. aureus will be required to investigate the mechanism of action of DHP on the S. aureus QS circuit.

Apart from this, there are a few points that have not been completely addressed in this work. The stability and efficacy of the coatings and polymers for long-term usage need to be investigated. Further studies determining the in vivo efficacy along with cytotoxicity and immune response studies are also important that should be conducted to fully understand the effect of DHPs and NO after attachment.

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APPENDIX

Journal Publication

nolog ch y & Taunk et al., J Biotechnol Biomater 2016, 6:3 te io B io B f m DOI: 10.4172/2155-952X.1000238 o

a e

i r

u l

o J Journal of Biotechnology & Biomaterials ISSN: 2155-952X

Research Article Open Access Surface Immobilization of Antibacterial Quorum Sensing Inhibitors by Photochemical Activation Aditi Taunk1, Kitty Ka Kit Ho1, George Iskander1, Mark DP Willcox2 and Naresh Kumar1* 1School of Chemistry, University of New South Wales, Sydney, NSW 2052, Australia 2School of Optometry and Vision Science, University of New South Wales, Sydney, NSW 2052, Australia

Abstract Infection of implanted medical devices is one of the major causes of nosocomial infections. A significant proportion of the devices become colonized by bacterial biofilms, thus resulting in high morbidity and risk of mortality. This study focuses on the non-specific covalent attachment of potent quorum sensing (QS) and biofilm inhibiting compounds, furanones (FUs) and dihydropyrrol-2-ones (DHPs), onto glass surfaces by azide/nitrene chemistry. The attachment of FUs and DHPs was confirmed by X-ray photoelectron spectroscopy (XPS) and contact angle measurements. The modified surfaces were then assessed for their antibacterial efficacy against Staphylococcus aureus and Pseudomonas aeruginosa using confocal laser scanning microscopy (CLSM). Both FU and DHP coated surfaces were able to significantly reduce bacterial adhesion p( <0.001) with p-bromophenyl substituted DHP giving maximum reductions of up to 93% and 71% against S. aureus and P. aeruginosa, respectively. Therefore, photo- immobilization of QS inhibitors is an effective technique to produce novel antibacterial biomaterial surfaces.

Keywords: Biomaterial; Antibacterial; Quorum sensing; Surface mortality [15]. Thus, there is an urgent need to develop new strategies modification; Furanone; Dihydropyrrolone;Pseudomonas aeruginosa; to prevent bacterial infection on biomedical devices. Staphylococcus aureus A marine alga, Delisea pulchra, from Australia produces Abbreviations halogenated furanone (FU) compounds [16,17] that have the ability to inhibit fouling by other marine organisms by blocking the HAI: Hospital-Acquired Infection; DHP: Dihydropyrrol-2-one; bacterial communication pathway known as quorum sensing (QS). FU: Furanone; QSI: Quorum Sensing Inhibitor; AI: Autoinducer; APTS: QS is a process where bacteria use various autoinducers (AI) or small 3-Aminopropyltriethoxysilane; ABA: 4-Azidobenzoic Acid; EDC: signalling molecules to communicate with each other. This process 1-Ethyl-3-(3-Dimethyl Aminopropyl)Carbodiimide Hydrochloride; plays an important role in controlling behavioural activities of bacteria NHS: N-Hydroxysuccinimide; XPS: X-Ray Photoelectron Spectroscopy; such as the formation of biofilm and virulence factors. Halogenated TSB: Tryptone Soya Broth; PBS: Phosphate Buffered Saline; OD: FUs competitively bind to receptor proteins and displace the signalling

Optical Density; AHL: N-Acyl Homoserine Lactone; 3-Oxo-C12-HSL: molecules [18-20]. This can result in inhibition of biofilm formation N-(3-Oxododecanoyl)-L-Homoserine Lactone of Gram-negative bacteria. However, most of the natural FUs are toxic to human cells, thus limiting their use [21]. Therefore, a range Introduction of FU analogues having low cytotoxicity have been synthesized which Infection of commonly used medical devices such as catheters, maintain excellent activity against Gram-negative and Gram-positive cardiac pacemakers, intraocular lenses, dental implants, accounts for bacteria [22,23]. A few synthetic FUs have also been immobilized via 60-70% of all hospital acquired infections (HAIs) [1]. The cost for covalent attachment on biomaterial surfaces and which showed good treatment ranges between $28-45 billion per annum in United States biofilm inhibitory activitiesin vitro and in vivo [24,25]. Halogenated alone [2]. Duration of hospital stay, mortality and morbidity are also FUs have also been attached to surfaces by a non-specific covalent increased when infections are caused by multi-drug resistant bacteria attachment strategy; however, the activity of the compounds after [3-5]. With no effective therapies currently available, device-related attachment was not reported [26,27]. infections are extremely difficult to treat. It has been estimated that Structural analogues of FUs, dihydropyrrol-2-ones (DHPs) [28], about 80% of these infections are associated with biofilm formation also displayed excellent QS inhibiting activity with low cytotoxicity in on medical devices [2]. Biofilms on implants are upto 1000-fold more solution as well as after specific covalent attachment on surfaces [29- resistant to antibiotics when compared to their planktonic counterpart 31]. However, a direct comparison of the activity of FUs and DHPs [6,7]. Therefore, the prevention of biofilm formation on biomaterials is is lacking. Therefore, to better compare the effectiveness of FU and a preferable strategy than treatment. Various strategies to control the formation of biofilm on medical devices have been examined, including coating antibiotics such as *Corresponding author: Naresh Kumar, School of Chemistry, The University of New South Wales, Sydney, NSW 2052 Australia, Tel: 61 2 9385 4698; Fax: 61 2 9385 6141; norfloxacin [8], minocycline-rifampin [9], impregnating chlorhexidine E-mail: [email protected] [10], silver [11,12], and gendine [13] on the surface of the implants. However, these coatings may reduce infections only over a relatively Received July 21, 2016; Accepted July 30, 2016; Published August 06, 2016 short time frame as they are commonly dependent on the release of Citation: Taunk A, Ho KKK, Iskander G, Willcox MDP, Kumar N (2016) Surface the antimicrobial for activity. A comparative study revealed that Immobilization of Antibacterial Quorum Sensing Inhibitors by Photochemical Activation. J Biotechnol Biomater 6: 238. doi:10.4172/2155-952X.1000238 chlorhexidine, silver and minocycline-rifampin coated catheters lost their antibacterial activity within 28 days [14]. In a clinic trial, central Copyright: © 2016 Taunk A, et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted venous catheters impregnated with silver had no significant effect in use, distribution, and reproduction in any medium, provided the original author and controlling bacterial colonization, bloodstream infections and ICU source are credited.

J Biotechnol Biomater Volume 6 • Issue 3 • 1000238 ISSN: 2155-952X, an open access journal Citation: Taunk A, Ho KKK, Iskander G, Willcox MDP, Kumar N (2016) Surface Immobilization of Antibacterial Quorum Sensing Inhibitors by Photochemical Activation. J Biotechnol Biomater 6: 238. doi:10.4172/2155-952X.1000238

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DHP immobilization strategies, the same techniques should be used to The angle between the droplet and the surface was measured using evaluate their activities. a 50 mm Cosmicar Television Lens (Japan). Rame-Hart Imaging software was used to calculate the contact angle. A minimum of fifteen In this study, DHPs and FUs were immobilized on azide- measurements were made on five samples of each FU and DHP. functionalized surfaces by photoactivation under UV light. The antibacterial efficacy of the coated surfaces was assessed against Antibacterial activity two common pathogenic bacteria, Pseudomonas aeruginosa and Staphylococcus aureus. The attachment efficiency and antibacterial Bacteria (Staphylococcus aureus SA38 and Pseudomonas aeruginosa activity of the resulting surfaces were also compared with previously PA01) from frozen stock (-80°C) were streaked on chocolate agar developed coatings where DHPs were attached via Michael addition (Oxoid, UK) and incubated at 37°C overnight. A colony of the bacteria reaction. was taken from the plate and cultured overnight at 37°C in 15 ml tryptone soya broth (TSB; Oxoid, UK). The bacteria were washed Materials and Methods twice with fresh TSB by centrifugation. The optical density (OD) of the culture was adjusted to OD =0.1 which corresponds to 1 × 108 cfu/ml. Attachment of 4-azidobenzoic acid (ABA) 660 In a 12-well plate, the surfaces to be tested were first sterilized Glass coverslips (No. 1, diameter 13 mm D 263 M glass, ProSciTech, with 70% w/v ethanol for 30 min, then thoroughly washed with sterile Australia) were first cleaned in freshly prepared piranha solution (3:1 phosphate buffered saline (PBS) three times and finally placed in 4 ml v/v concentrated sulphuric acid to 30% hydrogen peroxide) at 100°C of the adjusted bacterial culture. The surfaces were incubated at 37°C for 1 h. After thorough rinsing with distilled water, the clean coverslips for 24 h. The media was then replaced by fresh TSB (4 ml) and further were rinsed once with absolute ethanol and air-dried. The substrates incubated for 24 h at 37°C. Subsequently, the samples were washed were then silanized according to the previously developed method [30]. twice with PBS before examination by fluorescence microscopy. Briefly, the clean substrates were placed on steel mesh within a glass vessel that contained a 3-aminopropyltriethoxysilane (APTS) solution Bacterial adhesion analysis (10% v/v in dry toluene; 1 ml). The glass vessel was sealed and heated at 140°C for 18 h. The coverslips were rinsed with dry toluene (x2), The glass samples with adherent bacterial cells were stained with absolute ethanol and air-dried. The APTS-coated coverslips were then Live/Dead BacLight Bacterial Viability Kit (Molecular Probes, Inc., immersed in a solution of 4-azidobenzoic acid (ABA; 49.0 µM), 1-ethyl- OR, USA) according to the manufacturers’ procedure and as described 3-(3-dimethylaminopropyl) carbodiimide hydrochloride (EDC, 245.2 in the literature for analysis of biofilms on surfaces [30,31]. Bacteria µM) and N-hydroxysuccinimide (NHS, 98.0 µM) in absolute ethanol were then fixed by adding 100 µl of 4% formaldehyde on each sample. (1.5 ml), and agitated overnight at room temperature under dark room Microscopic observation and image acquisition were performed with conditions (Figure 1B). The ABA-functionalized surfaces were rinsed Olympus FV1200 Confocal Microscope. Images from 10 representative twice with absolute ethanol and once with MilliQ water, air-dried and areas on each of triplicate samples for each surface were taken and stored under dark conditions before use. analysed using ImageJ software [33]. The image analysis results were reported as the average percentage coverage of live and dead cells in Attachment of FU and DHP the fields of view. The synthetic halogenated FU compounds (FU-1, 2 and 3; Figure 1A) were synthesized as described [32]. Similarly, DHP compounds A Br (DHP-1, 2 and 3; Figure 1A) were synthesized following the method developed previously by Kumar and Iskander [29]. Br F Br H Br Stock solutions of FU (25 mg/ml in dichloromethane) and DHP (25 n O Br O Br Br NH O N O N mg/ml in acetone) were prepared and 200 µl of the FU or DHP solution O O O H H was placed onto the ABA glass surface. After complete evaporation of n = 3 (FU-1) FU-3 DHP-1 DHP-2 DHP-3 n = 1 (FU-2) the solvent, the surfaces were irradiated under UV at 320 nm for 10 min in a CL-1000 Cross-linker (Ultra-Violet Products Ltd, Upland, B CA, USA) (Figure 1B). The unreacted FU and DHP were removed by OH extensively washing the samples with dichloromethane and acetone O OH APTS O Si NH2 respectively, MilliQ water and absolute ethanol, then air dried and O 140°C,18 hr stored in clean sterile container. OH X-ray photoelectron spectroscopy (XPS)

The surfaces were characterized using X-ray photoelectron O O NH NH C N NH C NH spectroscopy (XPS; ESCALAB220-iXL, VG Scientific, West Sussex, 2 3 O O HOOC N NH 3 England). The X-ray source was monochromated Al Kα and the photo- 2 NH C N3 NH C NH energy was 1486.6 eV with a source power of 120 W. The vacuum O UV O NH EDC/NHS 2 NH C N NH C pressure was ≤ 10-8 mbar. dark conditions 3 NH = FU or DHP Contact angle measurements Figure 1: (A) Chemical structures of FUs and DHPs used in this study (B) Contact angles were determined using a contact angle goniometer Chemical vapour deposition of 3-aminopropyltriethoxysilane (APTS) on blank (Rame-Hart, Inc. NRL USA, Model no. 100-00). Multiple drops of glass coverslip followed by immobilization of FUs and DHPs via photoactivation of azide groups on glass surface. deionized water were placed on each surface using a micro-syringe.

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Statistical analysis of data attachment of ABA, which is expected due to the presence of the hydrophobic aromatic ring in ABA. Subsequent attachment of FU and Further analysis of the data was done by the one-way analysis of DHP resulted in similar contact angles to the ABA surfaces, due to the variance (ANOVA) using GraphPad Prism 6.05 software. Post hoc presence of different hydrophobic moieties (alkyl chain, phenyl ring multiple comparisons were done using Tukey correction. Statistical and halogen atoms) on the FU and DHP compounds. significance was set at 5%. Antibacterial activity Results The adhesion of S. aureus and P. aeruginosa on the modified surfaces XPS characterization of the coated surfaces were evaluated using fluorescence microscopy, and representative The surfaces were characterized by XPS at each step of the images are shown in Figure 2. The total surface area covered by bacteria immobilization sequence to ensure surfaces were successfully modified. and the relative proportion of live and dead bacteria (stained green and The XPS data collected for the blank (untreated), APTS, ABA, FU and red respectively) were evaluated by image analysis and the results are DHP coated surfaces are summarized in Table 1. shown in Figure 3. Changes in the elemental composition of carbon, nitrogen and Figures 2A and 2B show microscopy images of live (green) and halogen indicate successful immobilization of FU and DHP on the dead (red) S. aureus cells adhered to coated and control surfaces, where surfaces (Table 1). After functionalization with APTS, the carbon and extensive colonization and biofilm formation can be seen on the ABA nitrogen percentages increased to 43.9% and 7.1% respectively when control (Figure 2A). Figures 2C and 2D show images for adhesion of compared to the blank glass (4.9% C, 0.5% N). Both the carbon and P. aeruginosa on the surfaces. Both strains of bacteria displayed similar nitrogen content increased even further to 46.2% and 8.1% respectively level of bacterial colonization on the ABA control surface (Figures 2A when ABA was coupled with the amine surface. The subsequent and 2C). The adhesion of both strains of bacteria on blank and APTS attachment of FU or DHP was confirmed by a further increase in glass (data not shown) was similar to that reported in literature [30]. carbon percentage. Finally, the detection of halogens from the FU and The bacterial coverage of both strains on all the FU and DHP coated DHP compounds further confirmed the attachment of FUs and DHPs surfaces was significantly lower than the process control. (0.74-0.41% Br for FUs, 0.17% Br for DHP-1, 0.32% F for DHP-2 and 0.35% Br for DHP-3). % C % N % Halogen Contact Angle (°) Blank 4.9 0.5 - 20 Analysis of the high-resolution C1s spectra of the APTS surface APTS 43.9 7.1 - 73 demonstrated the presence of three distinct components C-H/C-C, ABA 46.2 8.1 - 67 C-N and C=O at binding energies 284.9 eV, 286.1 eV and 288.2 eV FU-1 48.8 7.3 0.74% Br 70 respectively. The N1s spectrum of the APTS surface showed two peaks at FU-2 49.0 7.7 0.41% Br 60 + 399.6 eV and 401.4 eV corresponding to –NH2 and –NH3 respectively. FU-3 49.7 7.1 0.65% Br 65 After the subsequent attachment of ABA, two new additional peaks DHP-1 47.9 7.7 0.17% Br 71 in a 2:1 ratio emerged at 400.2 eV and 404.6 eV, which is a set of DHP-2 48.5 8.7 0.32% F 69 characteristic peaks attribute to the azide functionality [34–36]. The DHP-3 48.7 8.5 0.35% Br 74 peak at 400.2 eV was assigned to the two terminal nitrogen atoms of Table 1: XPS analysis and contact angle measurements of blank, ABA, FU and the azide and the peak at 404.6 eV was assigned to the central nitrogen DHP coated surfaces. atom because of its low electron density compared to the terminal nitrogen atoms. Furthermore, an additional peak at 289 eV (N-C=O) in the carbon narrow scan was observed indicating that the coupling S. aureus P. aeruginosa reaction between the amine-terminated surface and carboxylic acid A – ABA C – ABA of ABA successfully formed an amide bond. The characteristic azide peaks were not observed after the subsequent treatment of the ABA surface with FU or DHP, suggesting the azide functional groups were consumed for the covalent linkage of FUs and DHPs. Instead, the N1s spectra showed a peak corresponding to N-H at 399.5 eV which is consistent with the formation of an –NH2 group on photo-activating the azide, and also due to various side reactions of arylazides under UV light [37]. Furthermore, a shift in the peak for N-C=O (from 289.0 eV to B – DHP-3 D – DHP-3 288.7 eV) in the C1s spectra was also observed for all FU and DHP coated surfaces along with broadening of the band, possibly due to addition of C-Br or C-F, indicating successful attachment of FU and DHP. Contact angle measurements The modified surfaces were also characterized by determining the contact angle after every modification step (Table 1). A significant change in contact angle was observed for APTS surface (from 20° Figure 2: Fluorescence microscopic images of glass surfaces after adhesion to 73°), indicating an increase in surface hydrophobicity due to the of S. aureus and P. aeruginosa to ABA process control (A and C); DHP-3 aliphatic carbon chain of APTS which is hydrophobic in nature. The coated surfaces (B and D). Live bacterial cells stained green and dead bacteria stained red. Magnification 200x. Scale bar=100 µm. contact angle remained approximately the same (67°) after surface

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The image analysis results forS. aureus showed significant specific attachment strategy and the antibacterial efficacy of the resultant reductions in overall bacterial coverage of 75.4 ± 5.0%, 80.9 ± 4.1%, surfaces was assessed. XPS analysis indicated the successful attachment 74.8 ± 4.2% for FUs 1-3 and 75.6 ± 5.8%, 89.9 ± 1.6%, 93.4 ± 1.1% for of FUs and DHPs via the described photoactivation strategy with FUs DHPs 1-3 respectively compared to the ABA control surface (p<0.001) having slightly higher attachment efficiency compared to the DHPs. All (Figure 3A). Amongst these, the most effective compounds were FU- the covalently bound FUs and DHPs were able to significantly reduce 2, DHP-2 and DHP-3, which displayed comparatively lower bacterial colonization of both Gram-positive (S. aureus) and Gram-negative (P. coverage than other coated surfaces (p<0.05). There was no significant aeruginosa) bacteria on the surfaces. Surfaces immobilized with FU-2, difference in the percentage of bacterial cells stained red (dead bacteria) DHP-2 and DHP-3 were found to be the most potent against bacterial between the controls and modified surfaces. adhesion of S. aureus, whereas for P. aeruginosa, the most active surfaces were coated with FU-2 and DHP-3. While all compounds were The attachment ofP. aeruginosa on the FU and DHP coated surfaces effective in reducing bacterial adhesion, FU-2 and DHP-3 displayed the was found to be significantly lower than the control, with reductions of best broad spectrum antibacterial activity. The high level of reduction 54.8 ± 2.2%, 68.7 ± 1.7%, 52.9 ± 3.0% for FU 1-3 and 55.9 ± 2.8%, 54.3 in adherent bacteria displayed by DHP-3 (93% and 71% of reduction ± 2.1%, 71.23 ± 1.4% for DHP 1-3 respectively (p<0.001) (Figure 3B). against S. aureus and P. aeruginosa respectively) is consistent with In this case, FU-2 and DHP-3 gave maximum reduction in bacterial reports in the literature [29,30]. A previous study has also examined the attachment compared to other FU and DHP surfaces (p<0.05). Similar efficacy of DHP-3 by covalently grafting it on the surface via a Michael to S. aureus, no significant difference was observed in the percentage of addition reaction [30]. A higher surface concentration of DHP-3 was dead cells between the control and modified surfaces. achieved via the non-specific azide reaction (0.35% Br) described in Discussion this study than by the Michael addition reaction (0.21% Br). Tens of millions of medical devices are used each year, and in spite Numerous studies have demonstrated that an increase in surface of advances in biomaterial technologies, a significant proportion of the concentration of an active compound leads to better antibacterial devices are colonized by bacterial biofilms, resulting in device failure activity [25,38]. Surprisingly, in this study DHP surfaces have displayed and infections. The formation of biofilms on biomedical devices is potent activity even at low concentration. Among all the compounds therefore a serious problem that is very difficult to treat. used in this study, DHP-1 gave the least attachment (0.17% Br) to the surface but displayed a similar level of activity as FU-1 (p<0.05) which In the present study, various potent QS inhibiting compounds, FUs gave a maximum attachment (0.74% Br). This discrepancy could be and DHPs, were covalently immobilized on glass surfaces by a non- due to the orientation of DHP on the surface, making it more available for antimicrobial activity compared to a similar concentration of FU. Similarly, FU-3 was expected to display maximum efficacy amongst all A the FUs due to its high activity in solution and also high attachment efficiency (0.65% Br) [22,39]. Instead, FU-2 displayed the best activity out of all FUs at lower surface concentration (0.41% Br) with reductions Bacterial coverage - SA38 27 of 81% and 69% of adherent S. aureus and P. aeruginosa respectively, 24 while FU-3 displayed reductions of 74% and 52% for S. aureus and P. 21 Dead aeruginosa respectively. 18 bacteria F Br 15 Several strategies have been explored in the past to immobilize Br Live 12 bacteria O O Br QS inhibiting compounds on the surfaces to inhibit biofilm formation N 9 O O N H H [31,40,41]. For example, a furanone derivative has been physically 6 adsorbed on various polymer surfaces commonly used for medical Bacterial coverage (%) coverage Bacterial 3 0 devices [40]. However, such a non-uniform coating is highly prone to APTS ABA FU-1 FU-2 FU-3 DHP-1 DHP-2 DHP-3 leaching and gradual loss of the active compound. In another approach, FUs and DHPs were coated on surfaces via specific attachment B strategies [30,31,41]. Although this attachment strategy overcomes the limitations of uneven coating and leaching, it requires extensive modification of the compound for surface attachment. Any structural Bacterial coverage - PA01 14 change or modification of the active compound may also result in 12 decrease in activity. The non-specific attachment strategy employed in Br this study does not require structural modification or functionalization 10 Dead bacteria of the compound. Also, unlike the previous attachment strategies, the 8 Br O O Br O N azide reaction described in this study is much faster, making it easier 6 H Live bacteria and more convenient to implement. This study is the first to investigate 4 the antimicrobial activity of photo-immobilized DHPs on surfaces. 2 Bacterial coverage (%) coverage Bacterial DHPs act by interfering with the bacterial QS system. In particular, 0 DHPs are able to disrupt the N-acyl homoserine lactones (AHL) APTS ABA FU-1 FU-2 FU-3 DHP-1 DHP-2 DHP-3 regulated QS system in Gram-negative bacteria [29]. The mechanism through which DHPs inhibit QS is postulated to be similar to that Figure 3: Percentage bacterial coverage of live and dead bacteria for (A) S. aureus and (B) P. aeruginosa (mean ± standard error of the mean); *indicates of FUs, that is, via displacing the AHL signal from the receptor site p<0.001 compared to APTS and ABA control; × indicates p<0.05 compared without affecting bacterial growth [19,39,42,43]. Surface immobilized to FU surfaces; ^indicates p<0.05 compared to DHP surfaces. DHPs were capable of interfering with the AHL regulated las QS system in P. aeruginosa, thereby inhibiting biofilm formation [31].

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In the current study, about 97% of adherent bacteria on the coated Sustained antibiotic release from an intraocular lens-hydrogel assembly for surfaces were alive, supporting the previous data that FUs and DHPs cataract surgery. Investig Ophthalmol Vis Sci 52: 6109–6116. act without killing the bacteria and exerting no selective pressure on 9. Raad I, Darouiche R, Dupuis J, Abi-Said D, Gabrielli A, et al. (1997) Central the bacteria to develop resistance [24,30]. Therefore, it is likely that venous catheters coated with minocycline and rifampin for the prevention of catheter-related colonization and bloodstream infections. A randomized, the FU and DHP surfaces generated in this study act through the double-blind trial. The Texas Medical Center Catheter Study Group. Ann Intern same mechanism of action for Gram-negative bacteria. On the other Med 127: 267-274. hand, for Gram-positive bacteria, the mode of action of FU and 10. Leung D, Spratt DA, Pratten J, Gulabivala K, Mordan NJ, et al. (2005) DHP is still not fully understood. Research investigating the effect of Chlorhexidine-releasing methacrylate dental composite materials. Biomaterials 26: 7145-7153. the AHL, N-(3-oxododecanoyl)-L-homoserine lactone (3-oxo-C12-

HSL), on S. aureus showed that the mode of action of 3-oxo-C12-HSL 11. Fazly Bazzaz BS, Khameneh B, Jalili-Behabadi MM, Malaekeh-Nikouei B, involves inhibition of the agr-dependent QS system by binding to the Mohajeri SA (2014) Preparation, characterization and antimicrobial study of cytoplasmic membrane of S. aureus [44]. Similarly, two new classes a hydrogel (soft contact lens) material impregnated with silver nanoparticles. Cont Lens Anterior Eye 37: 149-152. of compounds recently identified, one derived from 3-oxo AHLs and other from 3-acyl tetronic acids, have displayed agr QS inhibitory 12. Wang R, Neoh KG, Kang ET, Tambyah PA, Chiong E (2015) Antifouling coating with controllable and sustained silver release for long-term inhibition activity in S. aureus [45]. In another study, the mechanism of action of infection and encrustation in urinary catheters. 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Antonelli M, De Pascale G, Ranieri VM, Pelaia P, Tufano R, et al. (2012) antibacterial coatings. All the FU and DHP coated surfaces were able Comparison of triple-lumen central venous catheters impregnated with silver to reduce adhesion of S. aureus and P. aeruginosa, the most common nanoparticles (AgTive®) vs. conventional catheters in intensive care unit pathogens associated with biomaterial infections. This suggests patients. J Hosp Infect 82: 101-107. that the non-specific attachment of FUs and DHPs to the highly 16. de Nys R, Wright AD, König GM, Sticher O (1993) New halogenated furanones reactive azide groups does not impair the antibacterial activity of the from the marine alga Delisea pulchra (cf. fimbriata). Tetrahedron 49: 11213– compounds, indicating that the compounds retain their activity even 11220. after attachment. Since prior functionalization of compounds was not 17. de Nys R, Steinberg PD, Willemsen P, Dworjanyn SA, Gabelish CL, et al. needed, it is a fast and easy technique for developing novel coatings for (1995) Broad spectrum effects of secondary metabolites from the red alga Delisea pulchra in antifouling assays. Biofouling 8: 259–271. prevention of infections of biomedical devices. 18. Kjelleberg S, Steinberg P, Givskov M, Gram L, Manefield M, et al. (1997) Do Acknowledgement marine natural products interfere with prokaryotic AHL regulatory systems? Aquat Microb Ecol 13: 85–93. This work was supported by a Discovery Project from Australian Research Council grant (DP 140102195). AT was supported by an Australian Postgraduate 19. Manefield M, de Nys R, Kumar N, Read R, Givskov M, et al. (1999) Evidence award. 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