isolation of bacteriophages obtained from limestones cave soils … · 2018-09-11 · their...

ISOLATION OF BACTERIOPHAGES FROM

LIMESTONE CAVE SOILS AND EVALUATION OF

THEIR POTENTIAL APPLICATION AS BIOCONTROL

AGENTS OF PSEUDOMONAS AERUGINOSA

By

HASINA MOHAMMED MKWATA

A thesis presented in fulfillment of the requirements for

the degree of Master of Science (Research)

School of Chemical Engineering and Science, Faculty of

Engineering, Computing and Science

SWINBURNE UNIVERSITY OF TECHNOLOGY

2018

i

ABSTRACT

The emergence and frequent occurrence of multidrug-resistant and extremely drug-

resistant bacteria have raised a major concern because infections caused by these

bacteria are often associated with high mortality rates, prolonged hospitalization, and

high treatment costs. This situation is predicted to worsen in the future due to a massive

decline in the development of new antibiotics in recent years. Bacteriophages and their

derivatives have long been exploited as powerful and promising alternative antibacterial

agents in phage therapy and biocontrol applications. Limestone caves remain relatively

unexplored as a source of novel lytic bacteriophages compared with other

environments, despite being one of the most propitious sources for the discovery of

novel antimicrobial compounds. This research presents, for the first time, the screening

and isolation of lytic bacteriophages targeting different pathogenic bacteria from

limestone caves of Sarawak, and evaluation of their potential application as biological

disinfectants to control P. aeruginosa infections. A total of 33 lytic bacteriophages were

isolated from samples obtained from FCNR and WCNR targeting bacterial strains

Pseudomonas aeruginosa, Staphylococcus aureus, Klebsiella pneumoniae, Streptococcus

pneumoniae, Escherichia coli and Vibrio parahaemolyticus, using enrichment culture

method. Phage amplification was performed, and lysates were obtained, and spot

tested on lawns of various bacteria strains to assess their lysis spectrum. The result

revealed that P. aeruginosa and V. parahaemolyticus infecting phage isolates showed

the broadest host range among all the phage isolates. An interesting feature observed,

was the ability of some phage isolates to exhibit trans-subdomain infectivity between

gram positive and gram-negative bacterial hosts. Phage bacteriolytic activity was

investigated in an in-vitro co-culture assay with P. aeruginosa PAO1 strain using five

multiplicity of infection (MOI) ratios. Viable P. aeruginosa PAO1 cells that survived phage

infection were enumerated at 6 hrs and 24 hrs post-infection and the counts were

compared with those of untreated control. Bacteriophages FCPA3 (MOI 105),

WCSS4PA (MOI 105) and Cocktail (MOI 104) showed the highest bacterial inactivation

among all the tested phages at the end of 6 hrs of incubation. The highest bacterial log10

CFU/mL reduction was 11.82 equivalent to 100% reduction in bacteria observed in

cultures treated with phage cocktail (Cocktail, MOI 104) at the end of 6 hrs of

ii

incubation. Similarly, surviving bacterial counts assessed 24 hrs post-infection showed

that phages WCSS4PA (MOI 104) and Cocktail (MOI 102) had the highest bacterial log10

CFU/mL reduction of 10.5 and 10.86 respectively, equivalent to 100% reduction in

bacterial cells. Some of the phages did not show any reduction in bacterial cells at 6 hrs

and 24 hrs post-infection, instead the cells rebounded and surpassed those of the

untreated control.

Assessment of phage’s ability to be utilized as a biological disinfectant was performed

on P. aeruginosa PAO1 contaminated sand samples. The sand samples served as a

simulant of any environmental surface exposed to contamination with P. aeruginosa.

Surviving bacterial cells following treatment with bacteriophages FCPA3, WCSS4PA

and Cocktail were enumerated at 0 hr, 6 hrs, 24 hrs and 48 hrs post-treatment, and the

counts were compared with those of untreated control. Over 99% reduction in bacterial

cells were observed on all phage treated sand samples harvested at 6 hrs post-

treatment. No reduction in bacterial cells was observed in sand samples harvested at 24

hrs and 48 hrs post-treatment despite phage recharge, instead, the cells rebounded and

surpassed those of untreated controls. The ‘Bacterial rebound’ phenomenon mentioned

in this study indicates that the bacteria evolved resistance against the infecting phage.

This study suggests that P. aeruginosa bacteriophages obtained from Sarawak limestone

caves (FCNR and WCNR) may present potentials to be developed into biological

disinfectants to control P. aeruginosa infections, upon further exploration.

iii

ACKNOWLEDGEMENT

Foremost, I am deeply grateful to my coordinating supervisor, Assoc. Prof Dr Peter

Morin Nissom for his valuable advice, critics, challenges, encouragement, and directions

throughout my research study. Special thanks goes to my co-supervisor, Dr Lee Tung Tan

for accepting to co-supervise my MSc research project. I extend my appreciation to

Sarawak Biodiversity Centre (SBC) and Sarawak Forestry Department (SFD) for issuing

the permits (SBC-RA-0110-PMN) which enabled me to have access to soil samples from

Fairy Cave and Wind Cave Nature Reserves, located in Bau, Kuching Division, Sarawak,

Malaysia. I also wish to thank the science laboratory officers, Nurul Arina Salleh,

Cinderella Sio and Marciana Jane Richard, as well as Chua JiaNi, biosafety officer, for

their assistance with regards to the provision of the materials and apparatus throughout

the course of my research. My heartfelt thanks extend to my boyfriend, Armstrong

Ighodalo Omoregie for his unwavering support and encouragement throughout my

research journey. I am sincerely grateful to my parents, Mohammed Mkwata and Fatma

Mwenda for their unconditional love, care, advice and encouragement throughout my

studies. I am profoundly thankful to my Dad, for his financial support used to partially

fund my research. I am grateful to Almighty God, whose blessings have enabled me to

successfully accomplish my research study.

iv

DECLARATION

I hereby declare that this research entitled “Isolation of bacteriophages obtained from

limestones cave soils and evaluation of their potential application as biocontrol agents

of Pseudomonas aeruginosa” is original and contains no material which has been

accepted for the award to the candidate of any other degree or diploma, except where

due reference is made in the text of the examinable outcome; to the best of my

knowledge contains no material previously published or written by another person

except where due reference is made in the text of the examinable outcome; and where

work is based on joint research or publications, discloses the relative contributions of

the respective workers or authors.

(HASINA MOHAMMED MKWATA)

DATE: August 15, 2018

In my capacity as the Principal Coordinating Supervisor of the candidate’s thesis, I

hereby certify that the above statements are true to the best of my knowledge.

(ASSOCIATE PROFESSOR DR. PETER MORIN NISSOM)

DATE: August 15, 2018

v

PUBLICATIONS

Mkwata, HM, Tan, LT, & Nissom, PM, 2016, ‘Assessing the Diversity of Viruses in Soils

Obtained from Limestone Caves,’ 15th International Peat Congress (IPC 2016),

International Peatland Society, pp. 156-160.

Mkwata, H.M., Omoregie, A. I., Musa, I. B., Suyuh, J., Yee, P.H., Sing, L.W., Tan, L. T. &

Nissom, P. M. (2018), “A Laboratory Practicum on Screening for Lytic Bacteriophages

from Soil Samples”, Transactions on Science and Technology, (6 pages), ISSN: 2289-8786,

published by e-VIBS, Faculty of Science and Natural Resources, Universiti Malaysia

Sabah. (Accepted).

Omoregie, A. I., Siah, J., Pei, B. C. S., Yie, S. P. J., Weissmann, L. S., Enn, T. G., Rafi, R., Zoe,

T. H. Y., Mkwata, H. M., Sio, C. A. & Nissom, P. M. (2018), “Integrating Biotechnology

into Geotechnical Engineering: A Laboratory Exercise”, Transactions on Science and

Technology, 13 pages), Volume 5, No. 2, ISSN: 2289-8786, published by e-VIBS, Faculty

of Science and Natural Resources, Universiti Malaysia Sabah.

CONFERENCE AND PRESENTATIONS

Poster presenter, Assessing the Diversity of Viruses in Soils Obtained from Limestone

Caves, 15th International Peat Congress (IPC 2016), International Peatland Society, 15-

19 August 2016, Kuching, Sarawak, Malaysia.

Oral presenter, Phage Therapy: The forgotten cure, Three Minute Thesis (3MT)

Competition, 17 June 2015, Swinburne University of Technology, Sarawak campus,

Kuching, Sarawak, Malaysia.

Participant, International Congress of the Malaysian Society for Microbiology, 7-10

December 2015, Batu Ferringhi, Penang, Malaysia.

Participant, Asian Congress on Biotechnology, 15-19 November 2015, Kuala Lumpur,

Selangor, Malaysia.

vi

TABLE OF CONTENTS

CONTENTS PAGE

ABSTRACT ................................................................................................................. i

ACKNOWLEDGEMENT ............................................................................................. iii

DECLARATION ......................................................................................................... iv

PUBLICATIONS ......................................................................................................... v

CONFERENCE AND PRESENTATIONS ......................................................................... v

TABLE OF CONTENTS ............................................................................................... vi

LIST OF FIGURES...................................................................................................... ix

LIST OF TABLES ....................................................................................................... xi

LIST OF ABBREVIATIONS ........................................................................................ xiii

INTRODUCTION AND LITERATURE ..............................................................................

1.1 Background ......................................................................................................... 1

1.2 Antibiotic-resistant pathogens: A global threat .................................................... 3

1.3 Antibiotic-resistant mechanisms .......................................................................... 5

1.3.1 Decreased drug permeability....................................................................................... 6

1.3.2 Active efflux ................................................................................................................. 6

1.3.3 Alteration or bypass of the drug target ....................................................................... 7

1.3.4 Production of antibiotic modifying enzymes ............................................................... 9

1.3.5 Antibiotic inactivation by transfer of a chemical group ............................................ 10

1.4 Bacteriophages ................................................................................................. 11

1.4.1 Bacteriophage structure and classification ............................................................... 12

1.4.2 Bacteriophage replication cycles ............................................................................... 14

1.5 Phage therapy, biocontrol, and its advantages ................................................... 18

1.5.1 Bactericidal capacity .................................................................................................. 18

1.5.2 Self-replicating pharmaceuticals................................................................................ 18

1.5.3 Specificity ................................................................................................................... 19

1.5.4 Narrow potential for inducing bacterial resistance ................................................... 19

1.5.5 Rapid discovery .......................................................................................................... 20

1.5.6 Safety and immunogenicity ....................................................................................... 20

1.5.7 Single dose potential.................................................................................................. 21

1.5.8 Minimal environmental impact and relatively low cost ............................................ 21

1.5.9 Biofilm clearance........................................................................................................ 22

1.6 History of phage therapy ................................................................................... 22

1.7 Early therapeutic applications of phages ............................................................ 23

1.8 Recent applications of phages in biocontrol and therapeutics ............................ 26

1.8.1 Human pathogens treatment .................................................................................... 26

1.8.2 Sanitation ................................................................................................................... 29

vii

1.8.3 Probiotics ................................................................................................................... 30

1.8.4 Food safety ................................................................................................................. 31

1.8.5 Water treatment ........................................................................................................ 32

1.9 Limestone Caves: A potential source for novel lytic phages ................................ 33

1.10 Exploring Sarawak’s limestone caves for potential lytic phages .......................... 36

1.11 Significance of the study .................................................................................... 39

1.12 Hypothesis ........................................................................................................ 39

1.13 Aims and objectives of the study ....................................................................... 39

1.14 Thesis Outline ................................................................................................... 40

MATERIALS AND METHODS .......................................................................................

2.1. Isolation of lytic bacteriophages targeting bacterial pathogens .......................... 41

2.1.1 Sampling site and sample collection .......................................................................... 41

2.1.2 Biological material ..................................................................................................... 41

2.1.3 Growth medium and sterilization .............................................................................. 42

2.1.4 Growth profiles of the bacterial hosts ....................................................................... 42

2.1.5 Maintenance and storage of bacterial hosts ............................................................. 43

2.1.6 Screening for lytic bacteriophages ............................................................................ 43

2.1.7 Phage isolation and amplification .............................................................................. 44

2.1.8 Screening and isolation of multiphages ..................................................................... 44

2.1.9 Determination of phage titer ..................................................................................... 45

2.1.10 Storage of lytic bacteriophages ............................................................................... 45

2.1.11 Revival of cryo-preserved lytic bacteriophages ....................................................... 46

2.1.12 Host range assay ...................................................................................................... 46

2.2. Phage in-vitro bacteriolytic activity and a small-scale treatment of experimentally

contaminated sand samples .......................................................................................... 47

2.2.1 Preparation of bacterial culture................................................................................. 47

2.2.2 Preparation of phage stocks ...................................................................................... 47

2.2.3 Phage in-vitro bacteriolytic activity ........................................................................... 48

2.2.4 Analysis of bacteria survival from phage treated cultures ........................................ 48

2.2.5 Preparation of sand samples ..................................................................................... 48

2.2.6 Phage preparation in spray bottles............................................................................ 49

2.2.7 Treatment of contaminated sand samples with phage ............................................. 49

2.2.8 Analysis of bacterial survival following phage treatment ......................................... 50

2.2.9 Statistical analysis ...................................................................................................... 50

RESULTS AND DISCUSSION .........................................................................................

3.1 Introduction ...................................................................................................... 51

3.2 Results .............................................................................................................. 52

3.2.1 Isolation of lytic bacteriophages targeting bacterial pathogens ............................... 52

3.2.2 In-vitro studies on phage bacteriolytic activity and assessment of bacterial survival

following phage treatment. ................................................................................................ 64

3.3 Discussion ....................................................................................................... 102

3.4 Conclusion ...................................................................................................... 117

GENERAL CONCLUSION AND FUTURE PERSPECTIVE ....................................................

4.1 General Conclusion ......................................................................................... 119

viii

4.1.1 Aim of the thesis ...................................................................................................... 119

4.1.2 Summary of the findings .......................................................................................... 120

4.2 Future Perspectives and Recommendation ...................................................... 123

4.2.1 Morphological and Molecular characterization of the phage isolates .................... 123

4.2.2 Broadening applications of the phage isolates ........................................................ 124

4.2.3 Assessment of phage stability ................................................................................. 124

4.2.4 Assessment of phage-resistant mutants ................................................................. 124

4.2.5 Investigative studies on the expansion of host range ............................................. 125

REFERENCES ........................................................................................................ 126

APPENDICES ........................................................................................................ 165

Appendix I: Plaque appearance of bacteriophages infecting (A) K. pneumoniae, (B) P.

aeruginosa, (C) E. coli and (D) V. parahaemolyticus, following an overnight incubation at

37oC. .................................................................................................................................. 165

Appendix II: Multiplicity of infection (MOI) ...................................................................... 166

Appendix III: Optical density (OD) values of phage FCPA1 obtained during an assessment

of phage bacteriolytic activity at varied MOI ratios ......................................................... 168

Appendix IV: Optical density (OD) values of phage FCPA2 obtained during an assessment


Appendix V: Optical density (OD) values of phage FCPA3 obtained during an assessment


Appendix VI: Optical density (OD) values of phage FCPA4 obtained during an assessment


Appendix VII: Optical density (OD) values of phage FCPA5 obtained during an assessment


Appendix VIII: Optical density (OD) values of phage FCPA6 obtained during an assessment


Appendix IX: Optical density (OD) values of phage WCSS4PA obtained during an

assessment of phage bacteriolytic activity at varied MOI ratios...................................... 174

Appendix X: Optical density (OD) values of phage WCSS5PA obtained during an

assessment of phage bacteriolytic activity at varied MOI ratios...................................... 175

Appendix XI: Optical density (OD) values of phage Cocktail obtained during an assessment


ix

LIST OF FIGURES

Figures Page

1.1 Death attributable to antimicrobial-resistance every year by 2050 4

1.2 Antibiotic targets and mechanisms of resistance 5

1.3 A generalized structure of a tailed phage (left) and electron micrograph

images of the three families of tailed dsDNA phages that infect bacteria

(right)

13

1.4 Schematic diagram showing lytic, lysogenic and pseudolysogenic cycles

of a bacteriophage

15

1.5 Phage bacteriolytic life cycle 17

1.6 Ancient phage preparations 25

1.7 Bacteriophage drugs produced by Eliava Biopreparations 29

1.8 Borneo Island’s map showing the geographical divisions

and features of Brunei Darussalam, Indonesia (Kalimantan) and

East Malaysia (Sarawak and Sabah)

37

3.1 Fairy Cave (FC) Bau, Sarawak, Malaysia 53

3.2 Wind Cave (WC) Bau, Sarawak, Malaysia 53

3.3 Growth profile of the bacterial host cultures 55

3.4 Plaque appearance of bacteriophages infecting (A) S. aureus [left] and

S. pneumoniae (B) [right]

57

3.5 Phage titer determination of FCPA3 by double-layer plaque assay 57

3.6 Re-confirmation of P. aeruginosa bacteriophage lytic ability by spot test

assay

61

3.7 In-vitro bacteriolytic activity of FCPA1 at different MOI ratios 67






3.13 In-vitro bacteriolytic activity of WCSS4PA at different MOI ratios 73

x

3.14 In-vitro bacteriolytic activity of WCSS5PA at different MOI ratios 74

3.15 In-vitro bacteriolytic activity cocktail at different MOI ratios 75

3.16 Survival of P. aeruginosa PAO1 cells after an in-vitro treatment with

FCPA1 at different MOI ratios

90



91



92



93



94



95


WCSS4PA at different MOI ratios

96


WCSS5PA at different MOI ratios

97


Cocktail at different MOI ratios

98

3.25 Survival of P. aeruginosa PAO1 cells on sand samples after treatment

with bacteriophages

101

xi

LIST OF TABLES

Table Page

2.1 Description of bacterial strains used in this study 42

3.1 Description of soil samples collected at FCNR and WCNR 52

3.2 Growth kinetics of bacterial hosts grown in batch cultures 56

3.3 Morphological characteristics of bacteriophages isolated from

FCNR and WCNR

59-60

3.4 Assessment of bacteriophage host range by spot test assay 62-63

3.5 Assessment of phage bacteriolytic activity at the end of 6 hrs of

incubation

65-66

3.6 Analysis of variance (ANOVA) results for the recovery of bacteria

following an in-vitro treatment with FCPA1

77



78



80



81



83



84


following an in-vitro treatment with WCSS4PA

86


following an in-vitro treatment with WCSS5PA.

87


following an in-vitro treatment with Cocktail.

89

xii

3.15 Analysis of variance (ANOVA) results for surviving bacterial cells

recovered from sand treated samples at 0 hr using different

phage samples.

99


recovered from sand treated samples at 6 hrs using different

phage samples.

100



phage samples.

100



phage samples.

101

xiii

LIST OF ABBREVIATIONS

ANOVA Analysis of variance

ASEAN Association of South East Asian Nations

ARB Antibiotic-resistant bacteria

BHI Brain heart infusion

CRE Carbapenem-resistant Enterobacteriaceae

CFU Colony forming unit

DNA Deoxyribonucleic acid

dsDNA Double-stranded DNA

dsRNA Double-stranded RNA

EIBMV Eliava Institute of Bacteriophages, Morphology, and Virology

ESBLs Extended-spectrum -lactamases

EPS Extracellular polymeric substances

XDR Extended-spectrum -lactamases

FCNR Fairy cave nature reserve

FDA Food and Drug Administration

GI Gastrointestinal

GRAS Generally Recognized as Safe

HIIET Hirszfeld Institute of Immunology and Experimental Therapy

ICTV International Committee for Taxonomy of Viruses

KPC- KP Klebsiella pneumoniae carbapenemase producing Klebsiella

pneumoniae

LPS Lipopolysaccharide

Log Logarithm

MRSA Methicillin-resistant Staphylococcus aureus

MRAB Multidrug Acinetobacter baumannii

MDR Multidrug-resistant

MOI Multiplicity of Infection

NDM1 New Delhi metallo-β- lactamase 1

OD Optical density

PB Phage buffer

xiv

PGHs Peptidoglycan hydrolases

PBS Phosphate buffered saline

PFU Plaque forming unit

PAGE Polyacrylamide gel electrophoresis

PFGE Pulsed-Field Gel Electrophoresis

RH Relative humidity

RFLP Restriction Fragment Length Polymorphism

RNA Ribonucleic acid

rRNA Ribosomal ribonucleic acid

ssDNA Single-stranded DNA

ssRNA Single-stranded RNA

SDS Sodium dodecyl sulfate

TEM Transmission Electron Microscopy

TTC 2,3,5-Triphenyltetrazolium chloride

TSA Tryptic soy agar

TSB Tryptic soy broth

WCNR Wind cave nature reserve

WHO World health organization

Chapter 1 INTRODUCTION AND LITERATURE

1

1.1 Background

Bacterial infectious diseases are known to be one of the biggest threats to health and

food security worldwide (Costelloe, et al., 2010, Prevention, 2013, Van Boeckel, et al.,

2014). Multidrug-resistant (MDR) and extremely drug resistant (XDR) bacterial

pathogens (Arora, et al., 2017) have recently emerged as a serious world threat (Bush,

2010, Michael, et al., 2014). For instance, ESKAPE bacterial pathogens (Enterococcus

faecium, Staphylococcus aureus, Klebsiella pneumoniae, Acinetobacter baumannii,

Pseudomonas aeruginosa, and Enterobacter species) which are increasingly associated

with nosocomial infections pose a serious challenge in medicine due to extreme

resistance towards multiple antimicrobial agents (Moellering Jr, 2010, Rice, 2010). One

of the most worrisome resistant pathogens undergoing pandemic dissemination is K.

pneumoniae producing KPC-type carbapenemases (KPC-KP) which very frequently show

an MDR or even an XDR phenotype, including last resort molecules such as colistin

(Cantón, et al., 2012, Lee, et al., 2016, Tzouvelekis, et al., 2012). The growing number of

antimicrobial-resistant pathogens, highlights a substantial liability on the healthcare

systems, leading to a worldwide economic expense. For instance, morbidity and death

rates, high treatment costs, diagnostic uncertainties, and lack of trust in traditional

medicine (Santajit and Indrawattana, 2016). Factors such as globalization and increasing

international mobility, abuse of antibiotics, horizontal gene transfer and evolution of

bacteria have facilitated the spread of antibiotic-resistant pathogens (D’Andrea, et al.,

2017, Lu and Koeris, 2011, Walsh, 2003). Nevertheless, shortage of new drug

development by the pharmaceutical industry due to reduced incentives and challenging

regulatory requirements (Jassim and Limoges, 2017), have also precipitated the

emergence of antibiotic-resistant crisis (Gould and Bal, 2013, Prevention, 2013,

Spellberg, 2014). This has resulted in a revived interest in unconventional antimicrobial

treatments such as bacteriophages (Lu and Koeris, 2011). Bacteriophages (phages) are

bacteria-specific viruses which infect and lyse their respective hosts (Sulakvelidze, et al.,

2001). Bacteriophages are the most abundant viruses in the ocean (Hambly and Suttle,

2005), with numbers estimated at 1027 phage particles whereas the entire viriosphere is

estimated to contain 1031 phage particles (Suttle, 2005, Wilhelm and Suttle, 1999). Due

to their widespread in the environment, phages can be obtained from any sample that

support bacteria proliferation (Jennifer, 2006).

2

Bacteriophages have shown great advancement since discovery and have been

recognized as potentially powerful tools for eliminating bacterial infections (Liu, 2014).

In addition, phages have facilitated the progress of modern biology, especially mastery

of biological processes at a molecular level which has been pivotal in the establishment

of modern biological sciences (Cairns, et al., 1968, Summers, 1999).

For a century, bacteriophages have been exploited as natural antibacterial agents in

phage therapy (Roach and Debarbieux, 2017). In the early years of discovery, phage

therapy resulted in mixed success, in large part due to poor understanding of the viruses

themselves, as well as how they infect and kill bacteria (Abedon, et al., 2011). With the

discovery of penicillin, phage therapies were largely superseded with the advent of the

antibiotic era. Nevertheless, the rise of multidrug-resistant (MDR) bacterial infections,

have renewed interest in the use of bacteriophages for treatment of human infections

as well as in agriculture, veterinary science, industry, and food safety (McCarville, et al.,

2016, Sulakvelidze, et al., 2001). Phages have shown an extensive application in

biocontrol of food pathogens as opposed to other systems. For instance, in biocontrol

of Listeria monocytogenes in food processing (Bai, et al., 2016), Salmonella enterica

serovars typhirium in food animals (Wong, et al., 2014) and E.coli O157: H7 in inanimate

surfaces (Viazis, et al., 2011). The effectiveness of phage therapy can be increased by

creating a combination of phages commonly referred to ‘phage cocktails’ to target a

wider variety of bacterial strains. Phage cocktails are well known to confer a broader

spectrum of activity against infectious bacteria and prevent rapid development of

phage-resistant mutants (Goodridge, 2010, Tanji, et al., 2005). Bacteriophages offers a

sustainable approach against bacterial pathogens due to several factors such as, the

ability to be easily isolated from the environment enriched with targeted bacteria, they

are relatively inexpensive to produce, capability of infecting their hosts specifically and

efficiently and development of phage products is relatively faster and more cost-

effective than conventional drugs (Nagel, et al., 2016, Semler, et al., 2011, Yu, et al.,

2017).

3

1.2 Antibiotic-resistant pathogens: A global threat

Human medicine and food production are heavily dependent on the effective use of

antibiotics (Baker, 2015). Antibiotics are arguably the most successful form of

chemotherapy developed in the 20th century and perhaps over the entire history of

medicine (Banin, et al., 2017). Since their discovery over 70 years ago, antibiotics have

been one of the most significant advances in the modern medicine (Medina and Pieper,

2016). These drugs have saved millions of lives not only by treating infections but also

as prophylactic measures in individuals with a weakened immune system such as those

undergoing chemotherapeutic treatments against cancer or after organ transplantation

(Medina and Pieper, 2016). In addition, antibiotics have found extensive applications in

animal husbandry and aquaculture for growth promotion, feed efficiency, prophylaxis,

as well as in the treatment of infections (Lekshmi, et al., 2017). Misuse of antibiotics

have been reported in every environment where they have found applications, from

small-scale clinical use by physicians (through unnecessary, indiscriminate or incorrect

prescribing) and by patients (through incorrect dosing and the course of durations) to

large-scale agricultural practice for disease treatment and prophylaxis or growth

promotion in animal husbandry and food production.

These actions have provoked the emergence of antibiotic-resistant pathogens and

present optimal environments for the dissemination and selection of resistance

determinants (Lekshmi, et al., 2017, Pendleton, et al., 2013). Globally, 10 million people

are expected to die by the year 2050 due to antimicrobial-resistance (Figure 1.1) (O’Neill,

2014). The advent of antibiotic-resistance reflects the ability of bacteria to evolve

resistance mechanisms by which bacterial cell can escape the lethal action of antibiotics.

Recent studies on metagenomics and functional genomics have provided an enthralling

evidence that antibiotic resistance genes are ubiquitous and the natural reservoir of

possible antibiotic resistance genes comprise of multiple ecosystems such as in

agriculture (e.g. animal manure, soil, water, wastewater lagoons), the gut of humans

and food animals, and even ancient soils (Lin, et al., 2015). The soil is an ideal forum for

genetic exchange, easily resulting in the movement of resistance determinants from

environmental or zoonotic bacteria to human pathogens (Pendleton, et al., 2013).

Various novel antibiotic resistance genes present in the soil could be available to

4

clinically relevant bacteria and facilitate the existence of antibiotic-resistant pathogens

(Lin, et al., 2015).

Bacteria have established mechanisms to fight back the noxious impact of antimicrobial

agents as an adaptive trait to their survival. The biological pressure inflicted by the

constant exposure to diverse antibiotics during clinical application has resulted to a

collective possession of resistant traits in major human pathogens, culminating in

multidrug-resistant (MDR) bacteria, which are almost impossible to treat. For instance,

methicillin-resistant Staphylococcus aureus (MRSA) is among some preeminent

reported example. Other reported MDR bacteria are -lactamase-producing (ESBL)

Klebsiella pneumoniae and Escherichia coli (Blair, et al., 2015), carbapenem-resistant

Enterobacteriaceae (CRE) and multidrug Acinetobacter baumannii (MRAB) (Medina and

Pieper, 2016).

Figure 1.1: Death attributable to antimicrobial-resistance every year by 2050. Over 4 million

deaths are predicted to occur from antimicrobial resistance in different regions situated in

Africa and Asia (Review on Antimicrobial Resistance, 2014).

5

1.3 Antibiotic-resistant mechanisms

Pathogenic microbes are able to resist specific antibiotics and they do so through

mutations in chromosomal genes and by horizontal gene transfer (Blair, et al., 2015).

The fundamental resistance of a bacterial species to a certain antibiotic is due to its

ability to withstand the effect of that antibiotic as a result of built-in structural or

functional characteristics (Blair, et al., 2015). Antibiotic resistance occurs through

different molecular mechanisms such as decreased drug permeability, active efflux,

alteration or bypass of the drug target, production of antibiotic-modifying enzymes and

physiological states such as biofilms that are less susceptible to antibiotic activity (Figure

1.2) (Wright, 2016). By using established high-throughput screens of high-density

genome mutant libraries constructed by targeted insertion or random transposons

mutagenesis in bacteria such as Staphylococcus aureus, Escherichia coli, and

Pseudomonas aeruginosa, many genes encoding for intrinsic resistance to antibiotics of

different classes have been determined (Blair, et al., 2015). Below are some major

important mechanisms responsible for antibiotic resistance in bacteria.

Figure 1.2: Antibiotic targets and mechanisms of resistance. Target modification,

efflux, immunity and bypass, and production of antibiotic modifying enzymes are the

major mechanisms of antibiotic-resistance (Wright, 2010).

6

1.3.1 Decreased drug permeability

Gram-negative bacteria are well known to be intrinsically less permeable to numerous

antibiotics unlike Gram-positive species as their outer membrane forms a permeability

hindrance (Kojima and Nikaido, 2013, Vargiu and Nikaido, 2012). Hydrophilic antibiotics

traverse the outer membrane by diffusing through outer-membrane porin proteins

(Blair, et al., 2015). Permeability reduction of the outer membrane which restricts the

antibiotic entrance into the bacterial cell is attained by the downregulation of porins or

by substitution of porins with more-selective channels (Tamber and Hancock, 2003).

Several reports have shown that reduction in porin expressions by Enterobacteriaceae,

Pseudomonas spp. and Acinetobacter spp. has remarkably contributed towards

resistance to newer drugs such as carbapenems and cephalosporins, to which resistance

is normally facilitated by enzymatic degradation (Baroud, et al., 2013, Lavigne, et al.,

2013, Tamber and Hancock, 2003). For instance, relevant clinical resistance to

carbapenems in Enterobacteriaceae can take place due to unavailability of

carbapenemase production if mutations reduce porin production or mutant porin alleles

are available (Baroud, et al., 2013, Poulou, et al., 2013, Wozniak, et al., 2012). Moreover,

several reports have highlighted that K. pneumoniae isolates that demonstrate porin

alternates have been linked with clonal lineages that have resulted in global epidemics

of infections (Novais, et al., 2012, Papagiannitsis, et al., 2013, Poulou, et al., 2013).

1.3.2 Active efflux

Bacterial efflux pumps which function by transferring many antibiotics out of the

bacterial cell are known to facilitate intrinsic resistance exhibited by Gram-negative

bacteria to numerous drugs often used to treat Gram-positive bacterial infections.

Following overexpression, these efflux pumps can present high levels of resistance to

antecedent clinically valuable antibiotics (Blair, et al., 2015). Bacteria that are known to

overexpress efflux pumps, such as Enterobacteriaceae, P. aeruginosa, and S. aureus

have been isolated from patients for over 20 years (Everett, et al., 1996, Kosmidis, et al.,

2012, Pumbwe and Piddock, 2000). Some efflux pumps such as the Tet pumps possess

limited substrate specificity (Blair, et al., 2015). However, the majority of efflux pumps

transport a broad range of structurally different substrates, thus commonly referred to

7

as multidrug resistance (MDR) efflux pumps (Blair, et al., 2015). For instance, MdeA in

Streptococcus mutants, FuaABC in Stenotrophomonas maltophilia, KexD in Klebsiella

pneumoniae and LmrS in Staphylococcus aureus (Floyd, et al., 2010, Hu, et al., 2012,

Ogawa, et al., 2012). While all bacteria bear several genes that encode MDR efflux

pumps on their chromosomes, some have been organized onto plasmids that can

transfer between bacteria (Blair, et al., 2015). For example, genes coding for a unique

tripartite resistance nodulation division (RND) pump have been uncovered on an IncH1

plasmid that was isolated from a Citrobacter freundii strain that also carried the gene

for the antibiotic-target enzyme New Delhi metallo-β- lactamase 1 (NDM1) (Blair, et al.,

2015). This is a fretting situation because it shows that, resistance mechanism is

transmissible and could be quickly spread to other clinically relevant pathogens (Blair,

et al., 2015). The RND family of MDR efflux pumps exist in Gram-negative bacteria and

is the most characterized of the clinically relevant MDR efflux transporters. Upon

overexpression, RND pumps result in clinically relevant levels of MDR and export an

exceptional range of substrates (Piddock, 2006). For instance, multidrug efflux pump

AcrB in E.coli and MexB in P.aeruginosa are some of the rigorously studied examples

(Blair, et al., 2015).

1.3.3 Alteration or bypass of the drug target

Alteration of the target structure that results in inefficient antibiotic binding, but that

still allows the target to proceed with its normal function can result in resistance (Blair,

et al., 2015). During an infection, a single point mutation in the gene encoding an

antibiotic target can confer resistance to antibiotics, thus pathogenic bacterial strains

possessing this mutation can grow rapidly (Blair, et al., 2015). Studies have reported that

genes that encode drug targets of certain antibiotics usually occur in multiple copies.

For instance, linezolid, a novel oxazolidinone antibiotic which has been used for more

than 10 years since its introduction into the market targets 23S rRNA ribosomal subunit

of Gram-positive bacteria, which is encoded by multiple, identical copies of its gene.

Clinical use of linezolid has resulted in resistance in S. pneumonieae and S. aureus by

mutations in one of these copies, followed by recombination at high frequency between

homologous alleles, which rapidly results in a population carrying the mutant allele

(Billal, et al., 2011, Gao, et al., 2010, Leclercq, 2002). Another example of a target change

8

involves possession of gene homologous to the original target, such as in methicillin-

resistant S. aureus (MRSA) in which methicillin resistance is acquired by the acquisition

of the staphylococcal cassette chromosome mec (SCCmec) element. This carries the

mecA gene, which encodes the β-lactam-insensitive protein PBP2a. This protein permits

cell wall biosynthesis despite inhibition of the native PBP by the presence of antibiotic

(Katayama, et al., 2000). Numerous SCCmec elements have been located in different

Staphylococcus species, and there is proof that the mecA allele has been mobilized

several times (Shore, et al., 2011).

Lately, targets protection has been recognized as a clinically relevant mechanism of

resistance exhibited by many major antibiotics. For example, the erythromycin ribosome

methylase (erm) family of genes methylate 16S rRNA can change the drug-binding site,

thus inhibiting the binding of macrolides, lincosamines, and streptogramins (Kumar, et

al., 2014). A recent spotted example is a chloramphenicol-florfenicol resistance (cfr)

methyltransferase, which methylates A2503 in the 23S rRNA thus conferring resistance

to a broad range of drugs with targets near the site such as phenicols, pleuromutilins,

streptogramins, lincosamides, and oxazolidonones, including linezolid (Long, et al.,

2006). The emr and cfr genes are carried on plasmids and operate as vectors to drive

their broad propagation (Leclercq, 2002, Zhang, et al., 2013).

Resistance to aminoglycosides can prevail due to alteration of the target ribosome by

methylation. This was not previously perceived as a clinically relevant mechanism of

resistance, however, in recent years the enzyme responsible for this type of mechanism

have been spotted in varieties of bacterial pathogens. For instance, clinical isolates of

Enterobacteriaceae obtained throughout North America, Europe and India, have been

found to carry the armA gene, which encodes a methyltransferase, likewise clinical

isolates obtained in North America, Central and South America and India have also been

found to carry the armA gene which encodes another methyltransferase (Fritsche, et al.,

2008, Hidalgo, et al., 2013).

In recent years, shortage of effective antibiotics has resulted to extensive use of last-

resort antibiotics such as colistin for treatment of infections caused by multidrug-

resistant P. aeruginosa, Accinetobacter spp. and Enterobacteriaceae, thus, leading to the

development of polymyxin (colistin) resistance (Blair, et al., 2015). Polymyxin antibiotics

9

are cyclic antimicrobial peptides characterized with long hydrophobic tails comprising

of polymyxin B and polymyxin E and target Gram-negative bacteria (Cai, et al., 2012, Lim,

et al., 2010). Antibacterial activity is conferred by the hydrophobic chain which distorts

both cell membranes (Kumar, et al., 2014, Wang, et al., 2013). This activity is frequently

associated with changes in the expression affecting Lipopolysaccharide (LPS)

production, which leads to target alterations thus, reducing the ability of the drug to

bind to the target (Blair, et al., 2015). Additionally, colistin resistance can occur due to

mutations in the genes encoding the PhoPO two-component system or its regulators

through increased expression of the PmrAB system (Cannatelli, et al., 2013, Miller, et

al., 2011). This type of resistance mechanism is very common in K. pneumoniae

(Cannatelli, et al., 2014).

1.3.4 Production of antibiotic modifying enzymes

Production of antibiotic modifying enzymes is a leading mechanism of antibiotic

resistance that has been pertinent since the emergence of antibiotics such as

penicillinase (a β-lactamase) in 1940 (Abraham and Chain, 1940). Since then, thousands

of enzymes capable of degrading and modifying antibiotics of different classes, including

-lactams, aminoglycosides, phenicols, and macrolides have been discovered.

Nevertheless, certain subclasses of enzymes capable of degrading various antibiotics

within the same class have been identified. For instance, the -lactam antibiotics, such

as penicillins, cephalosporins, clavams, carbapenems and monobactams, are hydrolyzed

by a diverse range of -lactamases (Livermore, 2008, Nordmann, et al., 2011, Woodford,

et al., 2011). Antibiotic classes expansion that target inclusion of derivatives of enhanced

properties has given rise to the emergence of hydrolytic enzymes with altered spectra

of activity. For example, expansion of early -lactamases which were effective against

first-generation -lactams resulted in the existence of extended-spectrum -lactamases

(ESBLs) with activity against-cephalosporins (Johnson and Woodford, 2013). Gram-

negative bacteria such as K. pneumoniae, E. coli, P. aeruginosa and A. baumannii, may

carry diverse ESBLs and carbapenemases, such as the IMP (imipenemase), VIM (Verena

integrin encoded metallo -lactamase), K. pneumoniae carbapenemase (KPC), OXA

(oxacillinase) and NDM enzymes which perpetuate the existence of -lactam antibiotics

resistant isolates (Blair, et al., 2015). This poses a serious consequence in the treatment

10

of hospitalized patients suffering from severe infections (Johnson and Woodford, 2013,

Lynch III, et al., 2013, Voulgari, et al., 2013).

The CTX-M14 and CTX-M15 enzymes represent one of the most widely isolated ESBLs

worldwide especially in cephalosporin-resistant E. coli and K. pneumoniae isolates. CTX-

M15-producing K. pneumoniae isolates and CTX-M15-producing E. coli strains are

predominantly nosocomial and community-acquired diseases respectively (Dhanji, et

al., 2010, Poirel, et al., 2012, Zhao and Hu, 2013). The use of carbapenem antibiotics in

clinical settings has grown over the past decade because of increased numbers of

bacteria carrying ESBL genes (Blair, et al., 2015). This has, in turn, resulted in increased

numbers of clinical isolates carrying -lactamases with carbapenem-hydrolyzing activity

(Queenan and Bush, 2007, Queenan, et al., 2010, Tzouvelekis, et al., 2012). Although it

was first detected on the chromosomes of single species, carbapenemase resistances

are now plasmid-mediated and have been reported in bacteria such as

Enterobacteriaceae, P. aeruginosa and A. baumannii (Tzouvelekis, et al., 2012).

Carbapenemases dissemination has occurred through various ways as demonstrated by

the kpc and ndm genes. For instance, serine carbapenemase KPC has been reported in

several Enterobacteriaceae since it was first reported in K. pneumoniae in 1996

(Deshpande, et al., 2006, Yigit, et al., 2001). The kpc gene is plasmid-borne and is linked

to a dominant clone of KPC-producing K. pneumoniae, ST258, which is found worldwide

(Qi, et al., 2010). Since it was first reported in India in 2009, the NDM carbapenemase

has grown to be one of the most extensive carbapenemases existing in Gram-negative

pathogens such as A. baumannii, K. pneumoniae and E. coli throughout the world

(Kumarasamy, et al., 2010). The ndm genes frequently occur on broad-host-range

conjugative plasmids present in many incompatibility or replicon types, including IncA,

IncC, IncF, IncHI1 and IncL-IncM (Giske, et al., 2012, Kumarasamy and Kalyanasundaram,

2011, Walsh, et al., 2011) and in concurrence with other antibiotic-resistance genes

(Nordmann, et al., 2011) and are reported to confer resistance to all -lactams except

aztreonam.

1.3.5 Antibiotic inactivation by transfer of a chemical group

The addition of a chemical group by bacterial enzymes to a vulnerable antibiotic

molecule can induce antibiotic resistance by inhibiting the antibiotic from binding to its

11

target protein because of steric hindrance (Blair, et al., 2015). This type of bacterial

resistance involves a large and diverse family of antibiotic-resistance enzymes which can

inactivate antibiotics by transfer of chemical groups such as acyl, phosphate, nucleotidyl,

and ribitoyl (Wright, 2005). Aminoglycoside antibiotics are exceptionally vulnerable to

modification because their molecules are large and have many exposed hydroxyl and

amide groups. Aminoglycoside-modifying enzymes confer high resistance levels to

antibiotics they modify (Blair, et al., 2015). For instance, a recent worrying incidence is

the discovery of a novel genomic island in Campylobacter coli isolated from broiler

chickens in China. This genomic island encodes six aminoglycoside-modifying enzymes,

comprising members of all three classes and confers resistance to diverse

aminoglycoside antibiotics often used to treat Campylobacter infections including

gentamicin (Qin, et al., 2012).

Mechanistic and structural understanding of bacterial resistance offers much better

opportunities for tackling antibiotic resistance problem because it allows the origin of

the problem to be addressed rather than simply generating additional resistance in the

future (Chellat, et al., 2016). Owing to the discovery gap during the last decades for novel

antibiotics chemotherapies in the pharmaceutical industry and the occurrence of

bacterial strains resistant to the current antibiotics, public health is running out of

treatment options for dealing with infectious diseases. To respond to this emerging

crisis, global organizations such as The World Health Organisation (WHO) have urged

the scientific community to search for new approaches to combat antibiotic resistance.

A lot of the research for new antibiotics is still focused on developing improved versions

of existing molecules. Screening for novel antibiotics from natural sources enables to

broaden the possibilities for treating infections. However, this approach does not

eliminate the intrinsic risk for the initiation of resistance to these novel antibiotics

(Chellat, et al., 2016).

1.4 Bacteriophages

Viruses are the most widely distributed biological entities on earth (Suttle, 2005), with

an estimate of 1031 virus-like particles in the biosphere most of which are bacteriophages

12

(McAuliffe, et al., 2007). Environmental viruses are indisputably the largest genetic

diversity pool on the planet (Hambly and Suttle, 2005). Virus particles are ecologically

significant as they shape microbial communities, cause the lysis of a large part of the

ocean biomass on a daily basis, transfer genetic material among host organisms, and

shunt nutrients between particulate and dissolved phases (Hambly and Suttle, 2005).

Bacteriophages, widely known as phages (Greek “Phagein” meaning “to eat”) are viruses

that specifically infect and lyse bacteria (Matsuzaki, et al., 2005, Sharma, et al., 2017).

Bacteriophages can be obtained in all environments on earth, ranging from soil,

sediments, water (both river and sea water) and in/on living or dead plants and animals

(Elbreki, et al., 2014). In fact, they can be isolated from any material that sustains

bacterial growth. For instance, many terrestrial ecosystem have been reported to

contain 107 bacteriophages per gram of soil (Parisien, et al., 2008, Pedulla, et al., 2003)

whereas, sewage is widely known to contain 108-1010 phage per millilitre (Dabrowska,

et al., 2005, Dublanchet and Bourne, 2007). Phages just like all viruses are absolute

parasites. Even though they carry all the information necessary in directing their own

reproduction in a suitable bacterial host, they lack machinery for energy and protein

production (Goldman and Green, 2015).

1.4.1 Bacteriophage structure and classification

Bacteriophages (Figure 1.3a) are small viruses of about 20-200 nm in size and may differ

greatly in size, shape, capsid symmetry and structure (Criscuolo, et al., 2017). A phage is

made up of a nucleic acid genome encapsulated by a protein coat (capsid) and may

contain lipids in the particle wall or in the envelope (Ackermann, 2006). Phage capsids

may exhibit different morphologies extending from small hexagonal structures to

filaments, or highly complex structures comprising a head and a tail (Melo, et al., 2017).

Bacteriophage genome may vary from as few as 5 kb (e.g. Phage phiX174) to as many as

500 kb such as in Bacillus Bacteriophage G, the phage presenting the biggest known

genome. Phages are regarded as metabolically inert particles due to the absence of

necessary machinery for energy production or ribosomes for protein synthesis. Thus,

phages depend on their hosts to produce progeny and their genome is devoted to

directing the host for that function (Guttman, et al., 2005).

13

Figure 1.3: A generalized structure of a tailed phage (left) and electron micrograph images

of the three families of tailed dsDNA phages that infect bacteria (right). a Myoviruses

generally isolated from natural marine viral communities. They have distinguishing

contractile tails, are typically lytic and frequently exhibit broad host ranges. b, Podoviruses

possess a non-contractile tail, are typically lytic, have narrow host ranges and are less

frequently isolated from seawater. c, Siphoviruses possess long non-contractile tails, they are

often isolated from seawater, in most cases exhibit broad host range, and most of them can

integrate into the host genome (Elbreki, et al., 2014, Suttle, 2005).

In most cases phages genetic material is carried in double-stranded DNA (dsDNA) and

sometimes as single-stranded DNA (ssDNA), single-stranded RNA (ssRNA) or rarely as

double-stranded RNA (dsRNA) (Melo, et al., 2017). Over the years, sophisticated phage

classification system has been drawn up by the International Committee on Taxonomy

of Viruses (ICTV) to account for the diversity. The ICTV has classified phages as one major

order, 13 families, and 31 genera based on nucleic acid content, morphology, and

genomic data. It is estimated that about 96% of all studied phages have tailed

morphology and belong to three families, the Myoviridae (tail contractile), Siphoviridae

(tail long and non-contractile) and Podoviridae (tail short) as shown in Figure 1.3b

(Ackermann and Prangishvili, 2012, Klumpp, et al., 2010). These families comprise the

order Caudovirales (Maniloff and Ackermann, 1998). Studies have reported most

therapeutic phages to be tailed. However, some cubic phages (X174 and Q)

(Bernhardt, et al., 2000, Bernhardt, et al., 2001) or filamentous phages (M13 and Pf3)

(Matsuzaki, et al., 2005) have also been used. The remaining 4% comprises of tailless

phages with varying structures: polyhedral (with either icosahedral or cubic symmetry),

14

pleomorphic (asymmetric e.g. shaped like a lemon or a droplet) and filamentous with a

long and a thin morphology (Ackermann, 2003, Maniloff and Ackermann, 1998).

1.4.2 Bacteriophage replication cycles

Bacteriophages are obligate parasites that can sustain two separate life cycles, lytic or

lysogenic and can be defined by their genetics and interaction with the bacterial host

(Feiner, et al., 2015, Ptashne, 2004). Distinctive receptors such as lipopolysaccharides,

teichoic acids, proteins and flagella present on the top of the host bacteria are essential

for the phage to infect bacteria (Sharma, et al., 2017). Owing to this specificity, phages

can only infect specific hosts (Sharma, et al., 2017). Phage genome excision and

integration are crucial steps in the onsite of the lytic and lysogenic cycles respectively.

These events are mediated by phage-encoded DNA recombinases, such as integrases

and excisionases, and take place at a specific attachment site in the bacterial genome

(attB) which is identical to attachment site (attP) in the phage genome (Feiner, et al.,

2015). Although the sequences select the phage specificity to the bacterial genome,

secondary sites can also be utilised in the absence of original attB site. Additionally,

some phages integrate randomly (e.g. phage Mu) within their host genome thus

expanding variation and possible mutations within the bacterial population (Harshey,

2012). The first contact between a phage and its host occurs by random collision, given

the cell carries specific receptors on its surface. This usually occurs between the receptor

molecules of the host (e.g. teichoic acid in Gram-positive or lipopolysaccharide in Gram-

negatives) and distinctive phage proteins located at the tip of the tail fibre, or at one

end of a filamentous phage (Elbreki, et al., 2014). Phage attachment on the bacteria-

host surface is influenced by different factors such as bacteria type (Gram-negative and

Gram-positive), growth conditions, and virulence (Rakhuba, et al., 2010). Following

adsorption, phage DNA is injected into the bacterial cytoplasm after a phage has firmly

and irreversibly adsorbed to the cell surface (Lengeler, et al., 1999). Due to their

propagation cycle, most phages can be broadly branched into two major groups: virulent

and temperate (Figure 1.4).

15

Figure 1.4. Schematic diagram showing lytic, lysogenic and pseudolysogenic cycles of a

bacteriophage. (a) Lytic phages enter a productive cycle, whereby the phage genome is

replicated, and phage capsid and tail proteins are manufactured by utilizing bacterial cell

machinery. This is followed by packaging of the phage genome into progeny phage particles

which are released through bacterial lysis (b) Temperate phages enter a lysogenic cycle, in

which the phage genome is integrated into the bacterial chromosome (prophage). Prophages

can either get replicated together with the bacterial host chromosome during host cell

replication or switch into lytic production due to DNA damage. (c) Pseudolysogeny is an

unstable state whereby the phage genome fails to replicate or become established as a

prophage due to nutrient-deprived circumstances. In this state, the phage genome exists as

a non-integrated preprophage for a considerable time, resembling an episome, until the

nutritional status is re-established and the phage can then enter an either a lysogenic or lytic

life cycle (Feiner, et al., 2015).

16

Virulent phages straightaway redirect the host metabolism into the production of new

phage virions which are released upon cell death within several minutes to hours

following the initial phage attachment process (Elbreki, et al., 2014). This is often

referred to as lytic cycle (Figure 1.5). Briefly, following phage DNA injection into the

bacterial host, the DNA is replicated, and multiple copies of synthesized DNA are taken

into the capsid, which is constructed de novo during the late stage of phage infection.

Progeny phage particles are finalized by the attachment of a tail to the DNA-filled head

(Matsuzaki, et al., 2005). The progeny phages are eventually liberated by the

coordinated action of two proteins, holing and endolysin (Lysin) coded by the phage

genome. Lysin is a peptidoglycan-degrading enzyme (peptidoglycan hydrolase). Holin

proteins form a “hole” in the cell membrane, allowing lysin to reach the outer

peptidoglycan layers (Wang, et al., 2000). The released descendant phages infect

neighboring bacteria in a speedy manner. Virulent phage infection results in clear

plaques on the lawns of the respective bacterial host (Elbreki, et al., 2014).

By contrast, temperate phages enter a lysogenic cycle, in which the phage genome is

integrated into the bacterial chromosome (forming a prophage) and remain in a state

called latent or dormant which does not promote cell death or production of phage

particles (Figure 1.4) (Feiner, et al., 2015). Some prophages, however, remain as a low

copy number plasmids and do not integrate into the bacterial chromosome (Edlin, et al.,

1977, Ravin, et al., 2000). Prophages are replicated together with the bacterial host

chromosome, and this lysogenic condition is sustained by the repression of the phage

lytic genes. A switch to lytic production begins when stressful conditions such as DNA

damage prompt the excision of the phage genome, which is followed by the expression

of lytic genes that promote DNA replication, phage assembly, DNA packaging and

bacterial lysis (Feiner, et al., 2015). Another documented but largely unexplored phage

life cycle is pseudolysogeny (Feiner, et al., 2015). Pseudolysogeny is a phase of stalled

development of a bacteriophage in the host cell, in which neither multiplication of the

phage genome nor replication synchronized with the cell cycle and stable maintenance

in the cell line, which proceeds with no viral genome degradation thus allowing the

subsequent restart and resumption of virus development (Łoś and Węgrzyn, 2012).

17

Figure 1.5: Phage bacteriolytic life cycle. Scanning electron microscopy of Acinetobacter

baumannii bacterial cell (false color) being lysed by phage vB-GEC_Ab-M-G7 during an

infection. Cell lysis can take place within minutes to hours depending on each phage and

metabolic status of the bacterium (Roach and Debarbieux, 2017).

Generally, pseudolysogeny itself is a nonreproductive stage. In this stage, the viral

genome may be maintained for a potentially long period of time and is sometimes called

“preprophage” (Miller and Ripp, 2002). This phenomenon occurs because of

unfavorable growth conditions such as starvation occurring to the host cell and is often

terminated with the instigation of either true lysogenization or lytic growth when the

growth conditions are restored (Łoś and Węgrzyn, 2012). Pseudolysogeny has been

postulated to play a crucial role in phage-bacteria interaction in water environments due

to lower concentrations of nutrients and seasonal variability. Additionally, Ripp and

Miller (1997) have demonstrated the importance of pseudolysogeny in maintaining the

presence of phages for a prolonged time in natural ecosystems.

18

1.5 Phage therapy, biocontrol, and its advantages

Phage therapy refers to the application of bacteria-specific viruses with the aim of

minimizing or eradicating pathogenic bacteria (Kutter, et al., 2010). Phage biocontrol

refers to the non-therapeutic antibacterial application of phages. More broadly, phages

have been employed as biocontrol agents, reducing bacterial loads in foods, e.g., such

as of Listeria monocytogenes in food processing (Bai, et al., 2016), of zoonotic pathogens

in food animals (Atterbury, 2009) or in the treatment of plant pathogenic bacteria

(Jones, et al., 2007). While phage therapy has become a predominantly pertinent

technology especially in veterinary, agriculture, and food microbiology applications, it is

for the treatment or prophylaxis of human infections that phage therapy first captured

the world’s attention (Kutter, et al., 2010). There has been a compelling need for new,

safe, effective and selectively non-toxic antibacterial agents, especially in the face of the

antibiotic resistance crisis (Aminov, 2010). Phages and their products thus present one

of the largest untapped resources of antibacterial agents (Abedon, et al., 2017). Phages

have several characteristics that make them attractive therapeutic and biocontrol

agents (Jassim and Limoges, 2017). Advantages of phages as therapeutic and biocontrol

agents can be drafted based on their properties as listed below:

1.5.1 Bactericidal capacity

Bacteriophages in contrast to antibiotics are bactericidal because after successfully

infecting bacterial cells, they are incapable of gaining their viability. In contrast to this,

antibiotics such as tetracycline are termed bacteriostatic because they can readily allow

bacterial evolution towards resistance (Loc-Carrillo and Abedon, 2011).

1.5.2 Self-replicating pharmaceuticals

Unlike antibiotics, phages increase in number specifically where their hosts are present

during the bacteria-killing process. However, limitations such as their relatively high

dependence on bacterial concentration may occur (Loc-Carrillo and Abedon, 2011). This

means bacteriophages low number can be pragmatic, they will amplify and persist until

the infection is eradicated or minimized to a point that the host immune system can

clear the infection (Clark, 2015). The kinetics of phage action is advantageous when

compared with antibiotics because the effects can be achieved with small doses (Payne

19

and Jansen, 2001). The downside of this is that the pharmacokinetics of phage therapy

varies from that of antibiotic, and is more complex than that of antibiotic (Clark, 2015).

1.5.3 Specificity

Bacteriophages are very specific to their bacterial hosts and frequently they target

strains or subtypes of bacteria (Hyman and Abedon, 2010). This phenomenon is

advantageous because phages can eliminate specific undesired bacterial strains while

leaving the rest of the microflora undisturbed (Skurnik, et al., 2007). As a result, phages

have been suggested as probiotic supplements particularly targeting bacteria which

cause an imbalance in the gut such as Clostridium difficile, at the same time sparing the

normal gut microflora (Rea, et al., 2013). However, in many real-world situations, phage

specificity is a disadvantage because, in most human infections, the agent causing

disease is not known (Clark, 2015). This is not an issue with relatively broad spectrum

small-molecule antibiotics but in the case of phages, it warrants the use of cocktails

which increases the complexity and cost of production (Clark, 2015, Kelly, et al., 2011).

1.5.4 Narrow potential for inducing bacterial resistance

The relatively strict host range displayed by most phages restricts the number of

bacterial types with which selection for specific phage-resistance mechanisms can arise

(Hyman and Abedon, 2010). This is the reverse when chemical antibiotics are employed,

as a considerable fraction of bacteria may be affected (Carlton, 1999). Moreover, some

mutations that arise due to resistance, negatively influence bacterial fitness or virulence

due to loss of pathogenicity-related receptors (Capparelli, et al., 2010, Skurnik and

Strauch, 2006). Nevertheless, phages lack cross-resistance with antibiotics.

Bacteriophages infect and kill their hosts using mechanisms dissimilar from those of

antibiotics (Carlton, 1999, Loc-Carrillo and Abedon, 2011), thus, specific antibiotic

resistant mechanisms cannot be transcribed into mechanisms of phage resistance

(Lobocka, et al., 2014, Weber-Dąbrowska, et al., 2014). Therefore, bacteriophages can

be utilized for curing antibiotic-resistant diseases such as those triggered by multi-drug

resistant Staphylococcus aureus (Gupta and Prasad, 2011, Mann, 2008).

20

1.5.5 Rapid discovery

This advantage comes from ubiquity and diversity of bacteriophages. Bacteriophages

targeting many pathogenic bacteria can be easily isolated from different sources such

as sewage or waste materials comprising high bacterial concentrations (Loc-Carrillo and

Abedon, 2011). Thus, although bacteria can easily mutate to phage resistance, the

natural environment can supply numerous phage substitutes differing in host range to

attack a variety of bacterial infections. These phages may be applied in cocktails so that

phage-resistant is challenged right from the start of the therapy (Goodridge and Abedon,

2003).

1.5.6 Safety and immunogenicity

One element that seems reliable throughout the history of phage therapy is their safety

as opposed to most antibiotics (Kutter, et al., 2010). It has been reported that phages

are normally highly tolerated by humans who are constantly exposed to immense

numbers of phages as part of their natural ecosystem (Clark, 2015). For instance,

Miedzybrodzki, et al. (2012) has reported on the therapeutic use of phages in 153

patients in a study which covered rigorous safety data. The only reported adverse effect

has been a relatively minor side effect, possibly due to endotoxin (and other super-

antigens) from lysed bacteria, either those delivered in the crude phage preparations

used or released in vivo by the destruction of the host phage replication (Clark, 2015).

The high specificity displayed by phages signifies that they do not actively interact with

human cells. However, phages do interact non-specifically with human cells, as the

immune system regards phages as inert virus-like particles (Merril, et al., 2003). Many

phages are reported to be immunogenic and can stimulate strong cellular (Keller and

Engley, 1958) and humoral (Clark and March 2006, Clark, et al., 2002) immune

responses. Although such immune responses do not affect the safety of phage products,

they can affect the effectiveness of the treatment. Phages administered systemically can

be cleared up by the immune response before any therapeutic effect occurs (Kutter,

2008). As a result, the first target for many phage applications is normally topical

(Abedon, et al., 2011, Górski, et al., 2009). However, it has been suggested that, if

conditions are optimized, phages can be applied systemically (Ryan, et al., 2011).

Additionally, phages can be administered orally, after which they traverse the gut barrier

21

and enter circulation. For instance, in a study reported by (Sarker, et al., 2012), a cocktail

of nine phages specific for E.coli (at up to 3 x 109 phages per dose ) was delivered to

healthy participants with no observed side effects.

1.5.7 Single dose potential

Single dose application relies on the phage’s ability to replicate, thus achieving an

‘active’ therapy. This situation is often regarded as phage amplification through auto

‘dosing” and culminate in substantial bacterial killing (Abedon and Thomas-Abedon,

2010, Capparelli, et al., 2010). Therefore, achieving efficacy following only a single dose,

or far less frequent dosing, is clearly unnecessary. However, in many or most occasions,

a single dose of phages is usually insufficient to achieve the desired efficacy (Capparelli,

et al., 2010). Nevertheless, the ability of phages to replicate in situ and increase in

density, given sufficient bacteria are present, could significantly minimize the treatment

cost by reducing phage dose sufficient to achieve efficacy (Loc-Carrillo and Abedon,

2011).

1.5.8 Minimal environmental impact and relatively low cost

Phages are predominantly composed of nucleic acids and proteins and usually exhibit

narrow host ranges (Abedon and Thomas-Abedon, 2010). Unlike broad-spectrum

antibiotics, discarded therapeutic phages will at worst affect only a small subset of

environmental bacteria (Ding and He, 2010, Hyman and Abedon, 2010). In addition,

phages that cannot tolerate degradative environmental factors such as sunlight,

desiccation, or extreme temperature will be rapidly inactivated (Loc-Carrillo and

Abedon, 2011). Phage production generally involves a combination of host growth and

subsequent purification (Gill and Hyman, 2010). The cost of growing bacterial hosts

differs depending on the species, whereas the cost of purifying the bacteria gets cheaper

as the technology advances (Kramberger, et al., 2010). Generally, the cost of production

per unit (Kutter, et al., 2010) are not out of line with the costs of pharmaceutical

products while the cost of discovery and characterization can be relatively low (Skurnik,

et al., 2007).

22

1.5.9 Biofilm clearance

Bacteria which persist as biofilms tend to be intrinsically more resistant to antibiotics as

the matrix physically restrict the entrance of the chemical to the target (Stewart and

Costerton, 2001), along with other factors, such as ‘persister’ cells, where phenotypic

drug tolerance occurs in a subpopulation of bacteria (Clark, 2015). Phages, unlike

antibiotics, can naturally disrupt bacterial biofilms through various mechanisms such as

through enzymes linked to the bacteriophage capsid, by carrying genes encoding biofilm

degrading enzymes in their genomes or by upregulating genes in the target bacteria that

make biofilm degrading enzymes (Abedon, 2011).

1.6 History of phage therapy

The history of bacteriophage discovery has been a matter of prolonged debates and

arguments over claims for priority for many decades (Sandeep, 2006, Skurnik and

Strauch, 2006, Sulakvelidze, et al., 2001). Ernest Hankin, a British bacteriologist,

reported in 1896 on the existence of antibacterial activity against Vibrio cholera in the

waters of the Ganges and Jumna rivers in India. He proposed that the unknown

substance capable of passing through the fine porcelain filters, and sensitive to heat was

responsible for this phenomenon and for regulating the dissemination of cholera

outbreaks (Sulakvelidze, et al., 2001). Two years later, Nikolay Fyodorovich Gamaleya

noticed the same phenomenon while working with Bacillus subtilis. In 1915, Frederick

Twort (a medically trained bacteriologist from England) re-introduced this matter

advancing the hypothesis that such antibacterial activity could be due to a virus

(Hermoso, et al., 2007). Due to several limitations encountered, such as financial

difficulties (Summers, 1999, Twort, 1915), Twort did not take up this discovery, and it

was another two years before bacteriophages were “officially” discovered by Felix

d’Herelle, a French-Canadian microbiologist at the Institut Pasteur in Paris (Sulakvelidze,

et al., 2001). Felix d’Herelle suggested this incident could have been by a virus

competent at parasitizing bacteria and named the virus ‘bacteriophage’, a word that is

derived from the fusion of ‘bacteria’ and ‘phagein’ (to eat in Greek) (Hermoso, et al.,

2007). The first trial involving the therapeutic use of phages was accomplished by

d’Herelle in 1919 where he used phages to tackle acute hemorrhagic dysentery despite

23

his phage phenomenon observation in 1910 while learning microbiologic means of

regulating an epizootic of locusts in Mexico (Hermoso, et al., 2007, Sulakvelidze, et al.,

2001). However, the first reported utilization of phages to treat infectious diseases in

humans came in 1921 from Richard Bruynoghe and Joseph Maisin, who employed

bacteriophages to treat staphylococcal skin disease. In 1930, various companies began

the commercialization of phages targeting various bacterial pathogens while at the same

time d’Herelle and other scientists continued advancing the study of phage therapy.

During this same time, d’Herelle established phage therapy centers in various countries

including the US, France, and Georgia (Hermoso, et al., 2007). During World War II the

German and Soviet armies utilized phages to treat dysentery, and the US army

conducted classified research on it (Hermoso, et al., 2007). Additionally, some

practitioners employed phages as therapeutic agents in the West, from the 1920s to the

early 1950s. This was considered as the ‘historic era’ for phage therapy. However, phage

therapy was widely deserted shortly after the establishment of antibiotics in the 1940s

and thus from 1950s to 1980s few data were published on this topic. Research focusing

of the therapeutic use of phages has been somewhat abandoned in the West ever since

until the past two decades when the growing incidence of antibiotic-resistant bacteria

revived the interest in phage therapy (Hermoso, et al., 2007).

1.7 Early therapeutic applications of phages

The first documented phage therapy research was a study conducted in Belgium by

Bruynoghe and Maisin in 1921, describing the treatment of staphylococcal skin furuncles

in human patients (Chhibber and Kumari, 2012). In this report, phages were

administered to six patients by injecting the phage preparation close to the base of

cutaneous boils (furuncles and carbuncles). This prompted recovery accompanied with

reduction of pain, swelling and fever within 48 hours of treatment. A substantial amount

of publications detailing phage treatment of typhoid fever, Shigella and Salmonella spp

related colitis, peritonitis, skin infections, surgical infections, septicaemia, urinary tract

infections and otolaryngology infections (Wittebole, et al., 2014) were also published in

the 1930s in the journal of La Médicine (Abedon, et al., 2011, Wittebole, et al., 2014).

Early, commercial production of phages was achieved by D’Herelle’s commercial

24

Laboratory in Paris, the Hirszfeld Institute of Immunology and Experimental Therapy

(HIIET), Poland (founded in 1952), the Eliava Institute (EIBMV) in Tbilisi, Georgia

(founded in 1923 by Giorgi Eliava and Felix d’Herelle) and companies such as Eli Lilly

Company (Indianapolis, Ind.) (Sulakvelidze, et al., 2001). D’Herelle’s commercial

laboratory in Paris produced at least five phage preparations such as Bacte-coli-phage,

Bacte-rhinophage, Bacte-intestine-phage, Bacte-pyo-phage, and Bacte-staphy- phage

targeting different bacterial infections. These phage preparations were marketed by the

famous large French company L’Ore´al (Sharma, et al., 2017). In the United States,

therapeutic phages were manufactured by the Eli Lilly Company (Indianapolis, Ind.) in

the 1940s. These preparations comprised of seven phage products for human use

against an array of pathogenic bacteria, such as staphylococci, streptococci and

Escherichia coli. The preparations included phage-lysed, bacteriologically sterile broth

cultures of the targeted bacteria or the same preparations in a water-soluble jelly base

and aimed at treating various infections including abscesses, festered wounds, vaginitis,

acute and chronic infections of the upper respiratory tract, and mastoid infections

(Sulakvelidze, et al., 2001). Nevertheless, the effectiveness of phage preparations was

contentious and with the emergence of antibiotics, commercial manufacturing of

therapeutic phages was discontinued in most parts of the Western world (Eaton and

Bayne-Jones, 1934, Krueger and Scribner, 1941).

The Eliava Institute (EIBMV) was regarded as one of the largest facilities involved in the

generation of therapeutic phage preparations in the world. This institute developed

phage preparations targeting a dozen of bacterial pathogens covering staphylococci,

pseudomonas, proteus and many enteric pathogens (Sulakvelidze, et al., 2001). The

Hirszfeld Institute of Immunology and Experimental Therapy (HIIET) phage laboratory

was actively involved in expansion and production of phages for the treatment of

septicaemia, furunculosis, and pulmonary and urinary tract infections and for the

prophylaxis or treatment of postoperative and post-traumatic infections (Sulakvelidze,

et al., 2001).

25

Figure 1.6. Ancient phage preparations. These preparations consisted of Monophages

(targeted Staphylococcal, Streptococcal, E. coli, Pseudomonas, Dysentrial, and Typhoid), Poly-

phages (Pyo and Intesti), Sera (targeted diphtheria, tetanus, gangrene, scarlet fever,

meningococcus) and for identification of Salmonella and Shigella (Kutateladze and Adamia,

2008).

One of the most extensive phage therapy studies was the one carried out in Tbilisi,

Georgia during 1963 and 1964 and focused on the application of therapeutic phages for

prevention of bacterial dysentery (Babalova, et al., 1968). Likewise, Smith and Huggins

revitalized phage therapy studies in the west in the 1980s. They reported successful

results on therapeutic use of phages against systemic infections and enteritis in mice,

calves, pigs and lambs (Smith and Huggins, 1983, Smith, et al., 1987, Smith, et al., 1987)

even demonstrating the superiority of phage therapy against antibiotics in a mouse

model of E. coli infection (Smith and Huggins, 1982). In recent years, the emergence of

multi-drug resistant pathogens combined with the discouragingly low-rate production

of clinically useful antibiotics has prompted a re-examination of bacteriophage therapy,

with work being carried out to modern regulatory standards. Furthermore, the

26

emergence of high-throughput sequencing technology has encouraged on-going

advancements in bacteriophage therapeutics and other uses (Monk, et al., 2010,

Rohwer and Edwards, 2002).

1.8 Recent applications of phages in biocontrol and therapeutics

Phages have proven extremely powerful at eradicating various bacterial diseases in

controlled animal studies, particularly as a biocontrol agent in the elimination of food-

borne diseases, owing to factors such as its target specificity, rapid bacterial killing and

self-replicating potential (Jassim and Limoges, 2014). Furthermore, the capability of

bacteriophage to reproduce at the infection site or whenever the bacterial host is

present and their nonexistence in sterile areas guarantee an optimal self-adjusting dose

of bacteriophages which is not common especially in non-biological modes of

antimicrobial agents (Mizoguchi, et al., 2003). These features have thus enabled phage

therapy and phage biocontrol grow into a predominantly applicable technology in

veterinary, agricultural, and food microbiology applications (Jassim and Limoges, 2014).

1.8.1 Human pathogens treatment

For many years since the introduction of phage therapy as a viable treatment of

pathogenic bacteria in Eastern Europe, numerous studies have been conducted to

evaluate the efficacy and safety of phage therapy including clinical experience

(Chanishvili, 2012, Kutter, et al., 2010, Sulakvelidze, et al., 2001). However, these trials

did not follow current Western rigorous standards (Barbu, et al., 2016). This rose many

questions regarding the safety of phage therapy. For this reason, phage therapy clinical

trials have focused on safety rather than efficacy, resolving some of these safety

concerns (Vandenheuvel, et al., 2015). The first reported double-blind, randomized,

placebo-controlled phase I trial to show the safety of phage treatment was performed

by Nestle Research Center (Lausanne, Switzerland) (Bruttin and Brüssow, 2005). In this

trial, it was reported that there were no significant side effects following administration

of phage. In addition, the results showed that oral administration of T4 did not disturb

the natural gut E. coli population. Subsequent studies were carried out to investigate

27

the metagenomic analysis of the entire anti-diarrheagenic phage collection to

determine the clinical risk of a subset of phages following oral administration in healthy

adults (Sarker, et al., 2012).

The first controlled phage therapy clinical trial occurred in 2009. This trial reported

efficacy and safety in chronic otitis caused by antibiotic-resistant Pseudomonas

aeruginosa following treatment with a therapeutic phage cocktail (Biophage-PA,

Biocontrol, UK) (Wright, et al., 2009). A year later, during an assessment of treatment of

chronic otitis infections in dogs using phages, the outcomes validated that topical

administration of the phage combination resulted to lysis of P. aeruginosa in the ear

without apparent toxicity and proved to be an appropriate and efficacious treatment

against P. aeruginosa otitis (Hawkins, et al., 2010, Jassim and Limoges, 2014). The first

FDA-approved phase I clinical phage trial was performed in 2007 at Southwest Reginal

Wound Care Center in Lubbock, Texas. This trial aimed at evaluating the local

administration of a small set of well-characterized phage in patients with chronic venous

leg ulcers (Rhoads, et al., 2009). The study revealed that topical phage administration

showed no safety concerns but at the same time did not affect wound healing. Although

efficacy was outside the scope of the clinical trial, no significant results were obtained.

The first fully regulated, placebo-controlled, double-blind, randomized phase II clinical

trial of the efficacy of a bacteriophage therapeutic was completed in 2007 and reported

a successful outcome against long-term infections with P. aeruginosa, despite using only

a single dose of input bacteriophages in the nanogram range (Wright, et al., 2009). This

trial supports the prediction that successful bacteriophage infection will lead to

therapeutically useful replication of the therapeutic agent if susceptible host bacteria

are present at the site of application (Monk, et al., 2010). Recently, Zhvania, et al. (2017),

have demonstrated a successful treatment of a 16-years old male with Netherton

syndrome (NS) (antibiotic resistant chronic Staphylococcus aureus skin infection)

accompanied with an allergy to multiple groups of antibiotics, with several anti-

staphylococcal phage preparations conducted at Eliava Phage Therapy Center. A

massive improvement and substantial changes in his symptoms and quality of life were

observed following a six months treatment.

28

Powerful supporting data on the potential for phage therapy has also been obtained

from animal models (Monk, et al., 2010). Many animal models of infections have been

utilized to study phages as prospective therapeutics, especially in the context of

antibiotic-resistant infections affecting humans. These animal models serve as an

essential bridge between in-vitro and clinical studies (Kusradze, et al., 2016). A

significant number of bacteriophage therapy studies in animals have concentrated on

respiratory infections, gastrointestinal infections and infections of skin and wounds

(Malik, et al., 2017). For example, Debarbieux, et al. (2010) utilized a mouse lung

infection model targeting P. aeruginosa. Both bacterial challenge and phage treatment

were performed via intranasal instillation. Reported phage doses were 108 per animal

treatment and 100-fold lower phage doses were found to be insufficient in preventing

death. Following bacterial densities measurement via bioluminescence, treatment

success in preventing lethality was found to diminish from 100% survival at 72 hr given

a 2-hr delay in phage treatment to 75% survival given 4-hr delays and then to 25%

survival given 6-hr delays in phage installations. Pre-treatment with phages 24-hr prior

to bacteria challenge resulted in 100% survival. Phage therapy studies with animals has

shown that in certain instances, it may help in reducing the densities of the infecting

bacterial populations to levels that may allow the immune response to mount a

successful defence to clear the infection (Alemayehu, et al., 2012, Debarbieux, et al.,

2010, Smith and Huggins, 1982).

Despite the long history of successful use, phage therapy has not yet managed to re-

enter Western medicine as a viable available treatment option due to a lack of

randomized controlled trials, quintessential in the age of evidence-based medicine

(Zhvania, et al., 2017). Safety concerns about the use of phages in human medicine have

also been a major hurdle to the phage therapy development in the Western world,

despite the fact that phage preparations have been commercially available in Russia and

Georgia for decades (Vandenheuvel, et al., 2015). An example of a successful

commercially available product is Pyo Bacteriophagum (Figure 1.7), a phage cocktail

developed by the Eliava Institute in Georgia, which targets bacteria strains such as

Staphylococcus aureus, Escherichia coli, Streptococcus spp., Pseudomonas aeruginosa,

Proteus spp. (Aminov, et al., 2017). In addition, clinical use of phage therapy is reported

29

to be faced with long product development and approval timelines in Western

regulatory frameworks. Due to this, many companies and researchers have instead

undertaken applications focusing on food safety, agricultural, industrial, and clinical

diagnostics (Lu and Koeris, 2011).

Figure 1.7. Bacteriophage drug produced by Eliava Biopreparations. A phage cocktail

targeting Staphylococcus aureus, Escherichia coli, Streptococcus spp., Pseudomonas

aeruginosa, Proteus spp. (Aminov, et al., 2017).

1.8.2 Sanitation

Phage application for disinfection has been conducted in Georgia to disinfect operating

rooms and medical apparatus as a preventive measure against nosocomial infections

(Kutter, 2008). A complementary approach suggested by Novolytics company involves

the use of a gel containing a phage cocktail targeting MRSA to treat nasal carriage of

MRSA, thus greatly minimizing the prevalence and spread of MRSA (Abedon, et al.,

2011). Elimination of S. aureus via experimental hand cleansing with phage-containing

Ringers solution has also been reported. In this study, roughly 100-fold reduction in

bacterial concentrations was detected after hand cleansing with a solution containing

108 phages/mL when compared with a phage-less control solution (O'flaherty, et al.,

30

2005). Different natural and man-made environments such as medical devices, dental

plaques, water pipes, industrial and food processing settings may be colonized with

microorganisms, causing microbial biofilm development (Kolter and Greenberg, 2006).

Biofilms refer to surface-related communities encased in hydrated extracellular

polymeric substances (EPS) matrix that is made up of polysaccharides, proteins, nucleic

acids, and lipids and assists in maintaining a complex heterogeneous structure (Lu and

Collins, 2007).

Biofilm-associated organisms especially those colonizing medical devices are resistant

to antimicrobial agents, can escape the host immune system, and can behave as a nidus

for infection (Donlan and Costerton, 2002). Hence, device-related infections, such as

catheter-associated bloodstream infections, culminate in high morbidity and mortality

among certain patients in populations (O'grady, et al., 2002). Bacteriophages among

other novel action plans have been suggested to counteract device-associated biofilms

either by reducing microbial attachment to the device or by targeting the biofilm

following its development (Fu, et al., 2010). For example, Curtin and Donlan (2006)

showed that a bacteriophage active against Staphylococcus epidermidis could be

integrated into a hydrogel coating on a catheter result in significant reduction in biofilm

formation by this bacteria in an in-vitro model system. Additionally, Sillankorva, et al.

(2004), showed that, phage S1 was capable of reducing Pseudomonas fluorescens

biofilm biomass by 85%. The biofilms tackled with phage S1 proved more efficient at

controlling the bacteria in comparison to traditional chemical biocides.

1.8.3 Probiotics

Due to high specificity, bacteriophages are regarded as unique tools for manipulating

the bacteria microflora composition of the gastrointestinal (GI) tract in a clearly defined

manner as opposed to other probiotic organisms or other antibacterial agents.

Additionally, bacteriophages deliver a novel, safe and effective method for controlling

the GI tract’s microflora (Abedon, et al., 2011). While probiotic bacterial formulations

introduce non-pathogenic bacteria to disrupt the ability of pathogenic bacteria to

colonize the GI tract, phage-based probiotics aid the GI tract balance by targeting

specific pathogenic bacteria (Abedon, et al., 2011). Phage probiotics have been

31

suggested as the most effective method to effectively control bacterial pathogens such

as Salmonella spp., Clostridium difficile, diarrheagenic E. coli and other bacteria with oral

entryway and demand short or long-term colonization of the GI tract to generate a

disease (Abedon, et al., 2011). The George Eliava Institute of Bacteriophages,

Microbiology, and Virology have formulated ‘Instestiphage’, a potential phage probiotic

among other phage therapies for human consumption (Nicastro, et al., 2016).

Additionally, IntraLytix company is currently developing ‘ShigActive’, a phage probiotic

that targets Shigella species in the gastrointestinal tract.

1.8.4 Food safety

Bacterial pathogens control present on fresh fruits, vegetables and ready to eat foods is

of utmost concern because these foods do not always go through further processing or

cooking that would destroy bacterial pathogens prior to consumption (O'Flaherty, et al.,

2009). The continuous rise in food-borne diseases due to pathogens such as Salmonella,

Campylobacter, Escherichia coli and Listeria (Chibeu, 2013) which are associated with

grave gastrointestinal infections has prompted interest to seek for alternative and

effective technologies aiming at inactivating bacteria in food (Endersen, et al., 2014,

Team, 2012). A necessity that needs to be presented by any of these new approaches is

that it should be safe for humans, animals, and the environment while maintaining the

nutritional value and the organoleptic properties of the final product (Rodríguez-Rubio,

et al., 2016). Nonetheless, containing bacteria can also gain access to food throughout

different stages of production such as slaughtering, milking, fermentation, processing,

storage, and packaging. Thus, these new alternative technologies need to be employed

throughout the entire food chain (farm to fork) (Rodríguez-Rubio, et al., 2016).

Phages have been suggested as natural substitutes for antibiotics in animal health, as

biopreservatives in food and as tools for detecting pathogenic bacteria throughout the

food chain (Garcia, et al., 2008). Magnone, et al. (2013), utilized EcoShield, SalmoFresh,

and ShigActive to control E. coli O157: H7, Salmonella and Shigella spp. on fresh fruits

and vegetables. EcoShield (ECP-100) is an FDA approved commercial phage cocktail

composed of phages ECML-4, ECML-117 and ECML-134 and is employed to eradicate or

minimize food contamination caused by E. coli O157: H7 (Carter, et al., 2012, Ferguson,

et al., 2013). Spraying EcoShield (1 Å~ 106 to 5 Å~ 106 PFU/g) reduced E.coli numbers

32

by 94% and 87% in beef and lettuce with an E.coli contamination of about 103 CFU/g,

respectively, throughout the 5-minute contact time (Carter, et al., 2012).

Many studies have also demonstrated the effectiveness of phage utilization on

contaminated working surfaces used during food processing. For instance, a

bacteriophage cocktail designated as BEC8 was examined for its ability to reduce

enterohemorrhagic E. coli (EHEC) 0157: H7 strains applied on materials typically

employed during food processing surfaces such as sterile stainless-steel chips, ceramic

tile chips and high-density polyethylene chips (Viazis, et al., 2011). Bacterial cultures of

EHEC O157: H7 strains were spot inoculated (106, 105 and 104 CFU/chip) on the surfaces

which were followed by phage treatment to achieve a multiplicity of infection (MOI)

ratio of 1,10 and 100. The results obtained showed that the phage cocktail was very

effective within an hour against low levels of the EHEC bacterial cocktail at above room

temperature in all three hard surfaces (Viazis, et al., 2011). Phage lytic enzymes such as

endolysins have also been applied in food preservation. Endolysins are peptidoglycan

hydrolases (PGHs) encoded by phage and applied to enzymatically disrupt the host cell

wall during the final phase of reproduction (Schmelcher, et al., 2012). For instance,

researchers have demonstrated that staphylococcal phage lysin LysH5 can eradicate S.

aureus bacteria present in pasteurized milk and does this synergistically with

bacteriocins (García, et al., 2010, Obeso, et al., 2008). Similarly, phage lysins designated

as Ply118, Ply511 and Ply500 have been utilized as an antibacterial agent on iceberg

lettuce (Schmeler, et al., 2011).

1.8.5 Water treatment

Water resources are becoming limited due to contamination caused by various life-

threatening bacterial pathogens and toxic chemicals. Existing issues facing water supply

are such as contamination with chemical compounds (e.g. pharmaceutical and personal

care products), and biological agents which can result in increased antimicrobial

resistance in bacteria (Hartmann, et al., 2014, Larsson, et al., 2007, Pruden, et al., 2013).

The use of these chemicals has increased to a point that propagation of antimicrobial

resistance has become unavoidable (Pruden, et al., 2006). Many enteric bacteria with

multiple resistance (MDR) such as Escherichia, Enterobacter, Klebsiella, Salmonella and

33

Shigella species, have been detected in drinking and recreational water resources

(Kumar, et al., 2013). Antibiotic-resistant Pseudomonas species have also been isolated

from drinking water (Vaz-Moreira, et al., 2012).

Traditional water purification methods such as chlorination, radiation, and filtration are

used for the reduction of pathogenic bacteria in water systems and have many

disadvantages (Ahiwale, et al., 2012). It has been reported that human exposure to

disinfection by-products (DBPs) such as chlorine in water can result in eye, nose,

stomach problems, and sinus irritation. Besides that, pathogenic bacteria residing in

water bodies are reported to confer resistance to chemical disinfectants (Ahiwale, et al.,

2012). Bacteriophages have been employed as a potential disinfectant in the natural

waterbodies alone or in combination with physical and chemical processes (Ahiwale, et

al., 2012). For instance, McLaughlin and Brooks (2008) demonstrated high inactivation

rate of Salmonella enterica subsp. enteric serovar Typhimurium (ATCC 14028) in

experimentally contaminated wells using mono phages and cocktail combinations.

Bacterial biofilms especially those formed by P. aeruginosa are known to obstruct filters

at drinking water plants and usually require chlorine and costly flushing procedures to

clean (Jassim, et al., 2016). Zhang and Hu (2013), have isolated P. aeruginosa phages

from sewage and evaluated the results in comparison to the standard treatment using

chlorine to destroy P. aeruginosa biofilms. The results showed 40% removal of P.

aeruginosa biofilms using chlorine as opposed to 89% removal using phages at a titer of

107 PFU/mL. Moreover, the addition of lower concentration (105 PFU/mL) of phages

followed by chlorine eradicated 96% of the biofilms. These studies demonstrate that a

combination of phages and chlorine is a propitious approach to control bacterial biofilms

in water systems.

1.9 Limestone Caves: A potential source for novel lytic phages

Caves are diagnostic dissolution features in karst landscapes underlain by soluble rock

such as limestone and dolomite, where surface water sinks into the subsurface and flows

in a network of self-evolving underground steam passages (Ford and Williams, 2013).

These features are sheltered from the atmospheric disturbances and represent an

34

ecosystem in which many environmental variables remain relatively constant.

Connectivity usually occurs through entrances, skylights and rock cracks, the latter

acting like narrow channels (Riquelme, et al., 2015). Caves constitute of an oligotrophic

ecosystem (less than 2mg of total organic carbon (TOC) per liter), characterized by

complete darkness or low level of light, low steady temperature and high humidity

(Tomczyk-Żak and Zielenkiewicz, 2016). Despite the oligotrophic nature of the caves, the

average number of microorganisms thriving in cave ecosystems is 106 cells/gram of rock

(Barton and Jurado, 2007). Photosynthetic activity is limited to places with access to

light, normally at the entryway to the cave, but also in the cave interior owing to the

presence of artificial lights installed for the public. Light absence hinders the production

of the primary organic matter by photosynthetic microorganisms (Tomczyk-Żak and

Zielenkiewicz, 2016). Other methods of carbon assimilation are related to

chemoautotrophy. In such conditions, energy is derived from binding chemical elements

such as hydrogen and nitrogen, or volatile organic compounds, and also from the

oxidation of reduced metal ions such as manganese and iron present on the rocks (Gadd,

2010, Northup and Lavoie, 2001).

The presence of organic matter in the caves permits the development of heterotrophs

(Groth, et al., 1999). Due to the oligotrophic environment of the caves, existence and

functioning of species are limited to those adapted to the oligotrophic conditions (Wu,

et al., 2015). This is explained by the domination of chemoautotrophic microorganisms

in a certain cave (Chen, et al., 2009, Sarbu, et al., 1996), which fixes carbon and imports

energy into food web (Wu, et al., 2015).

Studies on the microbial composition dominating oligotrophic cave settings, have

disclosed a surprisingly high degree of diversity and abundance within the domains of

bacteria and archaea in diverse cave habitats such as soils, sediments, stream waters,

and rock surfaces (Barton and Jurado, 2007, Engel, et al., 2004, Tomczyk-Żak and

Zielenkiewicz, 2016). Numerous and familiar bacterial phyla have been uncovered in

cave environments by sequencing of 16S rRNA genes, thus greatly advancing the

knowledge of bacterial diversity since its establishment in microbial ecology (Roesch, et

al., 2007). The dominant taxa on cave walls are largely associated with a few phyla such

as Proteobacteria, Acidobacteria, and Actinobacteria (Barton and Jurado, 2007, Cuezva,

35

et al., 2012, Pašić, et al., 2009). Bacterial abundance in cave sediments could be

proportionate to that in overlying soils, however, the rock surfaces are mostly colonized

by the lowest diversity natural microbial communities (Macalady, et al., 2007, Yang, et

al., 2011).

The study of virus diversity and specifically bacteriophages using samples of water, soil,

or sediments from caves remains undocumented until to date (Ghosh, et al., 2016).

However, studies of the viral communities of other extreme habitats which are

characterized by low level of nutrients have revealed the presence of viral families and

bacteriophages (Ghosh, et al., 2016). For instance, a metagenomic analysis conducted

on the viral diversity of Antarctica freshwater ecosystems revealed a high number of

viral families. Pyrosequencing of 89,347 sequences exhibited no similarity to the

available gene bank databases. Furthermore, a transition in genetic structures from

single-stranded (ssDNA) to double-stranded (dsDNA) was observed among assemblages

from an ice-covered lake in spring to an open water late in summer (López-Bueno, et al.,

2009). Studies have highlighted that microbial populations found in caves are regulated

by existing viral communities. For instance, the viral-mediated killing of algal blooms is

essential for microbial population regulation in the ocean, thereby influencing food-web

interactions and affecting geochemical cycles (Fuhrman, 1999, Suttle, 2007).

Thus, cave microbial consortia may also contain massive viral communities that require

assessment (Jurado, et al., 2014). This information is essential in understanding cave

microbial interactions and population dynamics. Nevertheless, cave viruses could also

serve as therapeutic agents because of their potential lytic properties (Tan, et al., 2008).

The emergency and widespread of antibiotic-resistant and multi-drug resistant bacterial

pathogens (superbugs) and the stalled novel antibiotic discovery are the main driving

forces towards the search for novel antimicrobial compounds from extreme

environments (Maria de Lurdes, 2013). Extreme environments are considered one of

the most propitious sources of beneficial compounds (Cheeptham, 2012). Several

studies have reported on secondary metabolites produced by microorganisms that

colonize extreme environments as possible sources of useful compounds such as

extremozymes (Singh, et al., 2011), exopolysaccharides (Nicolaus, et al., 2010),

biosurfactants (Banat, et al., 2010), antitumoral (Chang, et al., 2011), radiation-

36

protective drugs (Singh and Gabani, 2011), antibiotics , immunosuppressants, and

statins (Harvey, 2000). Bioactive compounds such as Cervimycins A-D and xiakemycin A

are some of the novel antibiotics generated by cave-dwelling bacteria (Herold, et al.,

2005, Jiang, et al., 2015). These antibiotics have shown activity against methicillin-

resistant Staphylococcus aureus and vancomycin-resistant Enterococcus faecalis.

Xiakemycin A is also efficacious against methicillin-resistant Staphylococcus epidermidis

and vancomycin-resistant Enterococcus faecalis. In addition, it demonstrates antifungal

and cytotoxic effects against cancer cells. To date, cervimycin C is the most studied

antimicrobial compound obtained from caves and its resistance in Bacillus subtilis, as

well as its biosynthesis, have been thoroughly investigated (Bretschneider, et al., 2012,

Herold, et al., 2004, Krügel, et al., 2010).

1.10 Exploring Sarawak’s limestone caves for potential lytic phages

Borneo is the third largest island in the world and is notable for its high level of

biodiversity (Myers, et al., 2000, Slik, et al., 2010). This island (Figure 1.8) has a total

landmass of 740,000 square kilometers and consist of the independent Sultanate of

Brunei Darussalam, the Indonesian territory of Kalimantan, and the Malaysian states of

Sarawak and Sabah (Rautner, et al., 2005, Sulaiman and Mayden, 2012). Borneo’s forests

are home to the highest level of plants and mammal species in Southeast Asia (Bellard,

et al., 2014), including 581 species of birds and 240 species of mammals, and the island

is regarded as a major evolutionary hotspot (De Bruyn, et al., 2014). Extensive

development has led to a significant land cover change on the island, with 389,566 km2,

approximately 53% of the total area of the island, remaining under natural forest cover

(Gaveau, et al., 2014). In 2007, the countries situated in Borneo Island made a

declaration to protect 220,000 square kilometers of pristine rainforest habitats (also

known as the “Heart of Borneo”) to prevent deforestation and develop plantation

activities for the sake of the island’s biodiversity (Sulaiman and Mayden, 2012).

Malaysia is ranked 14th on the list of the 17 global mega-diverse countries on earth

(Keong, 2015). Its forests sustain varieties of unique flora and fauna species of

extraordinarily abundance and very high rates of endemism and uniqueness (Keong,

37

2015). Furthermore, Malaysia ranks 14th in the world for its vascular plants (15,500

species as recorded in 2004) densities. It is a home to 336 species of mammals,

approximately 750 species of birds with a high level of endemism and 212 species of

amphibians. Thus, Malaysia’s ecosystem is regarded as one of the globally significant

and distinctive ecosystems with high conservation preference (Keong, 2015). Different

laws have been enacted to protect the environment and natural biodiversity. For

instance, Wildlife protection ordinance, Sarawak (1998) and Environmental protection

enactment, Sabah (2002, amended 2004) (Keong, 2015). At the regional level, the

ASEAN Centre for Biodiversity has been set up to strengthen coordination for the

purpose of conservation and sustainable utilization of biodiversity (Keong, 2015).

Figure 1.8: Borneo Island’s map showing the geographical divisions and features of Brunei

Darussalam, Indonesia (Kalimantan) and East Malaysia (Sarawak and Sabah). This island is

known as the world's third largest island and one of the twelve mega-biodiversity regions

(Lateef, et al., 2014, Tan, et al., 2009).

38

Sarawak is the largest state in Malaysia, located along the northwest coast of Borneo

island and covering 124,500 square kilometers (Rautner, et al., 2005). This state

comprises of 512, 387.47 hectares of the protected area constituting 18 National parks,

four wildlife sanctuaries, five nature reserves and the largest peatland area in Malaysia

(Forest Department Sarawak, 2013, Van der Meer, et al., 2013). This rich biodiversity

has attracted the attention of scientists within and outside Malaysia. So far, existing

scientific studies have focused on peat soils, plants, corals, microbes in aquatic and

forest environments (Cole, et al., 2015, Kuek, et al., 2015, Lateef, et al., 2014, Miyashita,

et al., 2013, Sa'don, et al., 2015). Malaysia is a home to abundant limestone caves

located in places such as Langkawi Island, Kedah-Perlis, Kinta Valley, Perak, Selangor,

Gua Musang, and Kelantan Bakhshipouri, et al. (2009).

Sarawak’s limestone forest is one of the nine main types of forests reported in Sarawak,

covering about 520 m2 or 0.4% of the total area (Banda, et al., 2004, Julaihi, 2004). This

forest constitutes of several limestone caves which have become the focal point of

investigating the varieties of bats indigenous to the Wind and Niah Caves (Mohd, et al.,

2011, Rahman, et al., 2010, Rahman, et al., 2010). Analyses such as the evolution of

limestone formation, biological influence on the formation of stalagmite, investigation

of trace metal ratios and carbon isotopic composition have also been performed in

Sarawak’s Niah and Mulu caves (Cucchi, et al., 2009, Dodge-Wan and Mi, 2013, Moseley,

et al., 2013). Many south-east Asia’s limestone outcrops which have been historically

free from agricultural practices due to their rugged terrain (Clements, et al., 2006), may

operate as biodiversity pool that restocks degraded environments during ecosystem

reassembly (Schilthuizen, 2004). Recently, studies have been reported on the presence

of microorganisms isolated from Fairy Cave and Wind Cave Nature Reserves, Sarawak

Malaysia, capable of producing urease enzyme and inducing calcium carbonate mineral

for the biocement application (Omoregie, 2016).To date, there has been no study on

phage diversity conducted in Sarawak limestone caves, utilizing samples such as water,

soil, or sediments despite the reported biodiversity and species endemism. This research

gap has initiated the relevance of screening for lytic phages from Fairy cave and Wind

cave nature reserves located in Bau, Sarawak.

39

1.11 Significance of the study

This current study explores the prospects of isolating lytic phages from Sarawak

limestone caves capable of infecting pathogenic bacteria strains. Phages infecting P.

aeruginosa were further studied for their biocontrol efficiency on P. aeruginosa PAO1

contaminated sand samples individually and in a cocktail. Sarawak limestone caves

represent one of Malaysia’s biodiversity reservoir that has not yet been explored for

potential therapeutic microbes including bacteriophages. Furthermore, the study of

virus diversity and specifically bacteriophages from limestone caves and their potential

applications have not been reported elsewhere in the literature. This research gap

initiated the relevance of the current study. The phages reported in this study present

potentials to be developed into biological disinfectants to control P. aeruginosa

infections.

1.12 Hypothesis

The hypotheses of the present study are listed below;

i. The reported abundance of bacteria in oligotrophic environments such as

limestone caves suggest the presence of phages capable of infecting them. Since

microbial populations in caves are regulated by existing viral communities,

hence, it is possible that lytic phages will be present and abundant and can be

isolated using standard phage isolation methods.

ii. There is a strong correlation between multiplicity of infection (MOI) ratio and the

bacterial inactivation during phage biocontrol studies.

iii. Phage cocktails and multiphages are more effective in inactivating bacterial

pathogens than monophages.

1.13 Aims and objectives of the study

This research sought to investigate the diversity of bacteriophages in limestone caves

and evaluate their potential application as biological disinfectants to control infections

caused by P. aeruginosa bacteria. To fulfill the general purpose of this research, three

independent studies were designed with the following objectives:

i. To screen and isolate lytic bacteriophages from limestone cave soil samples.

40

ii. To investigate the phage bacteriolytic activity in in-vitro.

iii. To treat sand samples contaminated with P. aeruginosa using the isolated

phages.

1.14 Thesis Outline

This thesis is divided into four chapters: Introduction and Literature Review (Chapter 1),

Materials and Methods (Chapter 2), Results and Discussion (Chapter 3) and General

Conclusion and Recommendations (Chapter 4). Concluding remark is shown at the end

of Chapter 3 to summarise the contents of this chapter.

Chapter 1, provides a brief introductory background of the study and a broad review of

the literature on phage therapy and biocontrol which has been reported by other

researchers. This chapter also introduces the prospect of screening for bacteriophages

from Sarawak’s limestone caves. The significance, hypothesis, and aim of the research

are also mentioned. Chapter 2, gives a detailed description of the materials and methods

undertaken to fulfill the main objective of the current study. This chapter provides a

detail description of the methods used to screen and isolate lytic phages from Sarawak

limestone caves. Phage bacteriolytic activities were investigated on all isolated P.

aeruginosa phages at varied multiplicity of infection (MOI) ratios. Assessment of phage

ability to disinfect P. aeruginosa contaminated sand samples was carried out using the

best P. aeruginosa phage candidates (FCPA3, WCSS4PA, Cocktail), selected based on

their high efficiency in inactivating the bacteria. Chapter 3, presents the results and

discusses the findings of the current study with relevant statistical analysis. The

exploration and biodiversity of lytic phages from Sarawak limestone caves and the

application of these phages in biocontrol of P. aeruginosa. Chapter 4, presents a concise

overview of the most significant findings extracted from the work presented in this

thesis. The scope for further research within this field is also presented as future

directions.

Chapter 2 MATERIALS AND METHODS

41

2.1. Isolation of lytic bacteriophages targeting bacterial pathogens

2.1.1 Sampling site and sample collection

Soil sampling was conducted at Fairy Cave (N 01°22’53.39” E 110°07’02.70”) and Wind

Cave (N 01°24’54.20” E 110°08’06.94”) Nature Reserves located in Bau, Kuching Division,

Sarawak, East Malaysia. Samples were collected upon authorized permission from

Sarawak Forest Department and Sarawak Biodiversity Centre (SBC-RA-0110-PMN). A

total of seven soil samples were collected at a depth of 0-25 cm, six of which were

obtained from regions surrounded by rocks and vegetation and one soil sample mixed

with bat Guano which was obtained from the cave floor of the Fairy Cave Nature Reserve

(FCNR). Temperature and percentage relative humidity of the sampling sites were

measured by using traceable digital hygrometer/thermometer (Thermo Fisher

Scientific). Each sample was collected using sterile tools, placed in sterile polystyrene

containers, sealed and stored in an ice box (at the sampling site) before being

transported to Swinburne University of Technology, Sarawak campus for further

microbiological analysis. In the laboratory, the soil samples were temporarily stored in

the refrigerator at 4oC prior to the commencement of the phage screening experiments.

2.1.2 Biological material

Bacterial strains used in this study are presented in Table 2.1. These strains were

purchased from American Type Culture Collection (ATCC) Manassas, Virginia, United

States of America except V. parahaemolyticus which was acquired from the Swinburne

University of Technology Sarawak Campus (SUTS) microbiology strain collections. All the

bacteria except V. parahaemolyticus were aseptically grown on Petri plates containing

Tryptic soy agar (TSA) (40.0 g. L-1, HiMedia Laboratories Pvt. Ltd). To grow V.

parahaemolyticus, Petri plates containing TSA supplemented with 1.5% NaCl (w/v)

(Sigma-Aldrich (M) Sdn Bhd) was used. The plates were incubated (Incucell, MMM

Medcenter Einrichtungen GmnH) at 37oC under aerobic conditions for up to 24 hrs and

then stored in the fridge at 4oC prior to use.

42

Table 2.1: Description of bacterial strains used in this study

Bacterial Strain Genotype Source Source Designation

E. coli MG 1655 ATCC 47076

K. pneumoniae K6 ATCC 700603

P. aeruginosa PAO1 ATCC 15692

S. aureus PS 88 ATCC 33742

S. pneumoniae R6 ATCC BAA-255

S. typhi TA 1537 ATCC 29630

V. parahaemolyticus Wild strain SUTS NA

2.1.3 Growth medium and sterilization

Brain-heart infusion (BHI) broth (HiMedia, Mumbai, India) served as growth media for

screening and amplification of bacteriophages from soil samples. Nutrient broth (Oxoid,

Basingstroke, UK), Nutrient agar (Oxoid, Basingstroke, UK), Tryptic soy broth (HiMedia,

Laboratories Pvt. Ltd) and Tryptic soy agar (HiMedia, Laboratories Pvt. Ltd) were utilised

as routine growth media for cultivation of all the bacterial hosts except V.

parahaemolyticus where 3% NaCl (w/v) (Sigma-Aldrich (M) Sdn Bhd) was supplemented

into the growth media. The growth media were prepared in accordance with their

respective manufacturer’s instructions. Sterilisation of growth media, chemicals and

glassware were performed with the use of an autoclave machine (Hirayama-HVE-110)

at 121oC, 103.42 kPa for 20 minutes.

2.1.4 Growth profiles of the bacterial hosts

A colony of a bacteria strains grown on TSA (40.0 g. L-1 HiMedia, Laboratories Pvt. Ltd)

was inoculated into the universal bottle (20 mL capacity) containing 10 mL of Brain heart

infusion (BHI) broth (37.0 g. L-1Oxoid, Basingstoke, UK) and then incubated overnight at

37oC and 150 rpm. Batch cultures were prepared by inoculating 2.5 mL from an

overnight grown bacterial culture into BHI broth prepared in a 250mL capacity conical

flask. The contents were grown at 37oC and 150 rpm for up to 6 hrs. Aliquots (3 mL) were

withdrawn from the culture flask every 30 minutes and optical density was measured at

600 nm wavelength using a spectrophotometer (Genesys TM 20- Thermo Scientific).

43

2.1.5 Maintenance and storage of bacterial hosts

Glycerol stock method was used for both short and long-term storage of the bacterial

hosts following a modified procedure of Fortier and Moineau (2009). For short-term

bacteria preservation, 500 µL of bacterial culture was transferred into a sterile 1 mL

cryotube. About 500µL of 50% glycerol (Sigma-Aldrich (M) Sdn Bhd) was added into the

cryotube to obtain a final glycerol concentration of 25% (v/v). The contents were mixed

gently by inverting the tube a few times and stored at -20oC. For long-term preservation

of bacteria, the same approach was used but the tubes were stored at -80oC. For the case

of reviving stored cells, sterile toothpick or inoculation loop was used to scrap off the

splinters of solid ice (Omoregie, 2016). The resulting culture was then streaked out on

either TSA or BHI agar plate which served as a stock plate for culture preparation. In the

case of revival of V. parahaemolyticus, the culture was streaked out on either BHI agar or

NA supplemented with 3% NaCl. Stock plates were replaced every three to four weeks or

sooner where necessary.

2.1.6 Screening for lytic bacteriophages

Phage enrichment and isolation were carried out by inoculating 1 g of soil sample into

100 mL of sterile BHI broth (37.0 g. L-1Oxoid, Basingstoke, UK). Four milliliters (4 mL) of

sterile 10 mM CaCl2 (Sigma-Aldrich (M) Sdn Bhd) was added into the same broth and the

contents were incubated for 1 hr at 37oC. About 5 mL of bacteria-host culture grown to

its mid-exponential phase was subsequently added to the soil sample broth and the

contents were incubated aerobically overnight with shaking at 37oC and 150 rpm.

Thereafter, 1 mL of 1% TTC (2,3,5-triphenyl tetrazolium chloride) (HiMedia, Laboratories

Pvt. Ltd) and 1.2 mL of sterile 10 mM CaCl2 solutions were added into 100 mL liquefied

nutrient agar (28.0 g. L-1, 45oC, Oxoid, HiMedia, Laboratories Pvt. Ltd) prepared in a

Schott bottle (250 mL). About 5 mL of previously cultured broth was then inoculated

into the Schott bottle and the contents were gently mixed and poured out into sterile

Petri dishes, avoiding the formation of any bubbles. The plates were left to air dry

inside a biological safety cabinet (Class II, type A2, Thermo ScientificTM) for 15 min and

then incubated (Incucell, MMM Medcenter Einrichtungen GmnH) without inversion

(Sambrook and Russell, 2001) under aerobic conditions for 24 hrs at 37oC. Incubation of

the plates without inversion was performed so as to encourage sweating of the fluid

44

onto the surface of the dish allowing bacteriophages to spread easily (Sambrook and

Russell, 2001).

2.1.7 Phage isolation and amplification

After a careful examination of the plates, plaques were identified and characterized

based on the size, shape, clarity, presence or absence of a halo as per Basra, et al. (2014).

A double-layered agar plate technique was performed on these plaque isolates for at

least three times to obtain a homogeneous plaque formation. Following this, a sterile

straw was used to excavate the agar part containing the plaque and this was amplified

in 4 mL of BHI broth (supplemented with 100 μL of 10 mM CaCl2 solution) containing 1

mL of bacterial culture grown to its mid-exponential phase. The contents were

incubated aerobically overnight at 37oC and 150 rpm. The bacteria were expected to

lyse in 6-8 hrs and become slightly turbid due to cell debris. The contents were then

centrifuged (Eppendorf®, 5424R) at 8000 g for 5 min and the supernatant containing

phage particles was filtered through a 0.22 μm syringe filter. A drop (approximately 50

L) of chloroform was added into the recovered phage lysates and the tubes were

stored temporarily in the fridge at 4oC.

2.1.8 Screening and isolation of multiphages

Multiphages were screened following the same procedure as indicated in subsection

2.1.6. After a careful examination of the plates following an overnight incubation was

performed to identify and record unique plaque morphologies, plaque isolation

followed by amplification was not performed as this method selects for lytic

monophages. Instead, phage amplification was carried out by inoculating 1 mL of filter

sterilized soil sample bacterial broth from which plaques were present in 9 mL of sterile

BHI broth. The contents were then incubated aerobically overnight at 37oC and 150 rpm.

After bacterial lysis was observed, the contents were centrifuged (Eppendorf®, 5424R)

at 8000 g for 5 min and the supernatant containing phage particles was filtered through

a 0.22 μm syringe filter. A drop of chloroform (approximately 50 L) was added to the

recovered phage lysates and the tubes were stored temporarily in the fridge at 4oC.

45

2.1.9 Determination of phage titer

Phage particles were enumerated using the double-layered agar plate technique

following a modified method of Merabishvili, et al. (2009). A serial dilution of the

bacteriophage lysate in microfuge tubes was performed using phage buffer (PB) (10 mM

Tris [pH 7.5], 10mM MgCl2 and 68 mM NaCl) (Sigma-Aldrich (M) Sdn Bhd) supplemented

with a 10 mM CaCl2 solution. About 0.1 mL of this dilution was inoculated into another

microfuge tube containing 0.5 mL of log-phase bacterial culture, and the tubes were

incubated at 37oC for 10 min to allow phage adsorption to occur. Each of the cell-phage

content was poured into a sterile 15 mL centrifuge tube containing 3 mL of top agar.

This top agar was prepared by adding 0.7% agar (w/v) in 37.0 g. L-1 BHI broth (Oxoid,

Basingstoke, UK), and the temperature maintained at 45oC in a water bath. The mixture

was then plated onto pre-warmed TSA plates. The plates were left to cool for

approximately 15 min inside a laminar flow biosafety cabinet and then incubated at 37oC

for up to 24 hrs. This experiment was performed in triplicates for each phage dilution.

To estimate the original bacteriophage concentration, plates with 30-300 (Kropinski, et

al., 2009, Sutton, 2011) distinguishable homogeneous plaques were enumerated and

the phage titer (PFU/mL) was calculated as shown in the formula below (eqn. 1):

𝑃𝑙𝑎𝑞𝑢𝑒 𝑓𝑜𝑟𝑚𝑖𝑛𝑔 𝑢𝑛𝑖𝑡 (𝑃𝐹𝑈

𝑚𝐿) =

(No of plaques)(dillution factor)

Volume plated (mL) (eqn. 1)

2.1.10 Storage of lytic bacteriophages

Short term storage of phage isolates was performed by transferring 100 μL of phage

lysate into a sterile cryotube containing a drop of chloroform. The contents were mixed

gently by inverting the tube a few times and then stored at 4oC in a fridge. For the long-

term storage of phage isolates, 100 μL of phage lysate was added into a sterile cryotube

containing 200 μL of 75% (v/v) glycerol in phage buffer (PB) (Fortier and Moineau, 2009,

Pardon, et al., 2014) and the contents were gently mixed by inverting the tube a few

times and the vials were frozen at -80oC. Cryoprotectant solution [75% (v/v) glycerol in

phage buffer] was prepared by adding 75% of glycerol (Thermo Fisher Scientific) in a

Schott bottle containing 25% of phage buffer (10 mM Tris [pH 7.5], 10mM MgCl2 and 68

mM NaCl) (Sigma-Aldrich (M) Sdn Bhd). The contents were thoroughly mixed by

46

inverting the bottle a few times and then sterilized by autoclaving at 121oC, 103.42 kPa

for 20 min.

2.1.11 Revival of cryo-preserved lytic bacteriophages

To revive cryo-preserved phages, an overnight culture of the host strain was prepared

and about 0.5 mL of it was transferred into a test tube containing 2.0 mL of TSB or BHI

broth. With the use of an inoculating loop, frozen top part of the phage solution was

scrapped off and added into the broth. The contents were incubated aerobically

overnight at 37oC and 150 rpm. The next morning, the amplified phage culture was

centrifuged (Eppendorf®, 5424R) for 10 min at 8000 g and the supernatant was filter

sterilized using 0.45 µm syringe filter and stored at 4oC.

2.1.12 Host range assay

Bacteria strains used for host range assay were grown to their mid-log phase in BHI

broth. About 0.3 mL of bacteria-host culture was then inoculated into a sterile 15 mL

centrifuge tube containing 3 mL molten top agar supplemented with 100 µL of 10 mM

CaCl2 solution. The top agar was prepared by adding 0.7% agar (w/v) in 37.0 g. L-1 BHI

broth (Oxoid, Basingstoke, UK) and the temperature maintained at 45oC in a water bath.

The contents were gently mixed and quickly poured onto pre-warmed TSA agar plate

and left to air dry in the laminar flow biosafety cabinet for 15 min. Phage host range was

determined by spotting 10 μL of phage lysate preparation (approximately 1015 PFU/mL)

three times onto different host plates. For control purpose, each bacteria strain was

mock infected with sterile phage buffer. The Petri plates were incubated at 37oC for up

to 36 hrs under aerobic conditions. A successful phage infection was scored based on

plaque formation on a susceptible bacterial lawn.

47

2.2. Phage in-vitro bacteriolytic activity and a small-scale treatment of

experimentally contaminated sand samples

2.2.1 Preparation of bacterial culture

Bacteria strain P. aeruginosa PAO1 used in sand decontamination studies was prepared

by inoculating a bacteria colony from a stock plate into 10 mL of sterile Brain-heart

infusion (BHI) broth (37.0 g. L-1, Oxoid Thermo Scientific Microbiology). The contents

were incubated at 37oC and 150 rpm for 24 hrs in an incubator shaker (CERTOMAT® CT

plus–Sartorius) under aerobic conditions. Prior to decontamination experiments,

aliquots of 100 μL were transferred into pre-sterilised universal bottles containing 9 mL

of sterile BHI broth and the contents grown at 37oC and 150 rpm to mid-exponential

phase.

2.2.2 Preparation of phage stocks

This study utilized six highly lytic single plaque phages designated as FCPA1, FCPA2,

FCPA3, FCPA4, FCPA5 and FCPA6, and two multi-phages designated as WCSS4PA

and WCSS5PA specific for P. aeruginosa PAO1 bacteria. A phage cocktail (Cocktail)

was prepared by combining equal volumes of phage lysates (approximately 1015

PFU/mL) obtained from the six single-plaque bacteriophages (FCPA1, FCPA2, FCPA3,

FCPA4, FCPA5 and FCPA6,) following a modified procedure of Viazis, et al. (2011). To

obtain high titer phage stocks for the experiments, a modified procedure by Fortier and

Moineau (2009) was adopted. Briefly, P. aeruginosa host strain was grown in 10 mL of

BHI broth to its early log phase (OD6000.1). About 100 μL of 10 mM CaCl2 solution and

100 μL of phage lysate were added into the bacterial culture. The contents were

incubated for 8 hrs at 37oC and 150 rpm to allow amplification of the phage. Afterward,

the contents were centrifuged at 8000 g for 10 min and the supernatant containing

phage particles was filtered through 0.22 μm syringe filter. Each time bacteriophage

stocks were grown or amplified, their titer was determined using the double agar

overlay technique following a modified protocol of Merabishvili, et al. (2009). Phage

lysates were stored in the fridge at 4oC prior to use.

48

2.2.3 Phage in-vitro bacteriolytic activity

The bacteriolytic activity of phages was performed as suggested by Wang, et al. (2016)

with minor modifications. To begin with, an overnight P. aeruginosa PAO1 culture was

diluted 1:100 (v/v) in sterile BHI broth and incubated at 37oC and 150 rpm until a mid-

exponential phase was attained (7.76 x 1010 CFU/mL). This culture was then diluted using

Phosphate buffer saline (PBS) solution to obtain a concentration of 1.0 x 1010 CFU/mL.

About 25 mL aliquots of the culture were dispensed into 100-mL capacity conical flasks

and equal volumes (25 mL) of bacteriophage lysates were added to obtain different

multiplicity of infection (MOI) ratios (101,102,103,104 and 105). The titers of the phages

were diluted to desired concentrations using phage buffer (PB). The contents were then

incubated at 37oC and 150 rpm for up to 6 hrs. P. aeruginosa PAO1 bacterial culture with

an equal volume of phage buffer (PB); (10 mM Tris [pH 7.5], 10mM MgCl2 and 68 mM

NaCl) (Sigma-Aldrich (M) Sdn Bhd) was used as a control. The phage bacteriolytic activity

was determined by monitoring the cell absorbance of the culture solution (OD600) for 6

hrs with 30 minutes interval. Incubation was continued for up to 24 hrs and viable counts

(CFU/mL) of the recovered bacteria were determined at 6th and 24th hrs post-incubation.

Optical density measured at a wavelength of 600 nm and viable bacterial counts

(CFU/mL) were recorded as an average of three independent biological repeats.

2.2.4 Analysis of bacteria survival from phage treated cultures

To determine the CFU/mL counts of the recovered bacteria from phage treated cultures,

1 mL sample was withdrawn at a specific time and centrifuged at 8000 g for 10 min in a

1.5 mL capacity micro-centrifuge tube. The supernatant was discarded, and the pellet

was washed twice using Phosphate-buffered saline (PBS); 137 mM NaCl, 2.7 mM KCl,10

mM Na2HPO4, 1.8 mM KH2PO4 pH 7.4 (Sigma-Aldrich (M) Sdn Bhd) before being

resuspended in the same solution. Serial dilution was performed in PBS and plating was

done on Tryptic soy agar (40.0 g. L-1, HiMedia Laboratories Pvt. Ltd). Plates were

incubated at 37oC overnight and viable bacteria cells (CFU/ml) were enumerated.

2.2.5 Preparation of sand samples

Evaluation of phage’s ability to be utilized as a biological disinfectant to control

infections caused by P. aeruginosa was performed on P. aeruginosa PAO1 contaminated

49

sand samples. The sand samples served as a simulant of any environmental surface

exposed to contamination with P. aeruginosa. Sand decontamination studies were

performed on Petri plates (Surface area=56.75cm2) containing 20 g of sterile sand. Sand

was obtained from Swinburne University of Technology Sarawak Concrete Laboratory

(E002) and was thoroughly cleaned by washing it several times under running water,

followed by rinsing it with deionized water at least three times. The sand was then dried

overnight in an oven set to 100oC. About 20 g of sand was dispensed into clean and dry

universal bottles and sterilized by autoclaving at 121oC, 103.42 kPa for 30 minutes prior

to use.

2.2.6 Phage preparation in spray bottles

Plastic spray bottles of 50 mL capacity were purchased from a local supermarket and

sterilization was done by soaking 10% (v/v) of bleach inside the bottles overnight,

followed by rinsing the bottles at least three times with sterile deionized water. Spray

bottles were then further sterilized by exposing them to ultraviolet (UV) radiation inside

a laminar flow hood for 45 min. About 25 mL of phage lysates (approximately 1015

PFU/mL) for bacteriophages FCPA3, WCSS4PA and Cocktail were dispensed into the

bottles and samples were temporarily stored in the fridge at 4oC prior to use.

2.2.7 Treatment of contaminated sand samples with phage

The ability of bacteriophage isolates to decontaminate P. aeruginosa PAO1 immobilized

sand samples were assessed as follows; to begin with, 4 mL of freshly grown mid-

exponential phase P. aeruginosa PAO1 culture (OD600=0.5) having a concentration of

7.76 x 1010 CFU/mL was uniformly mixed with 20 g of sterile sand in a Petri dish using a

sterile spatula. This sand was compacted to form a matrix of approximately 3 mm thick.

Using a spray bottle, phage lysate (approximately 1015 PFU/mL) which was prepared as

demonstrated in subsection 2.2.2, was sprayed on the surface of the sand (thirty times)

delivering a volume of 2.55 mL. A negative control sample was prepared following the

same procedure as per section 2.2.7, but no phage was sprayed on it. This experiment

was performed in triplicate and the samples were incubated at 37oC for 6 hrs, 24 hrs,

and 48 hrs. Exactly twenty-four hours post-incubation, samples were sprayed with

phage for the second time (2.55 mL) and incubation was continued until 48 hrs. Phage

50

recharge was performed to investigate the effect of an additional dose of phage at

preventing regrowth of bacteria which was evident during the phage in-vitro

bacteriolytic activity studies presented in (Subsection 2.2.3).

2.2.8 Analysis of bacterial survival following phage treatment

About 1 g from phage treated and non-treated (control) sand samples were collected at

different time intervals during the treatment process (t= 0 hr, t=6 hrs, t=24 hrs and t=48

hrs) and placed into sterile test tubes containing 10 mL of PBS solution (pH 7.4). The

tubes were vortexed for 1 min and a serial dilution in PBS solution was conducted.

Plating was done on TSA Petri dishes and incubation was performed for up to 24 hrs at

37oC. Viable bacterial cell reductions (CFU/mL) were calculated by subtracting treated

sand sample cell counts from negative control cell counts (Tomat, et al., 2014).

2.2.9 Statistical analysis

The data obtained in this study was presented as mean SE (standard deviation) for

three independent replicates. The rate of bacterial inactivation by phages (FCPA1,

FCPA2, FCPA3, FCPA4, FCPA5, FCPA6, WCSS4PA, WCSS5PA and the Cocktail)

in comparison to untreated control at different MOIs was evaluated and analyzed using

GraphPad Prism software (version 7.0d). A one-way analysis of variance (ANOVA) and

Tukey-Kramer’s post hoc analysis was performed using StatPlus program (version 6.0)

to indicate any significant difference between groups. The value of p<0.05 was

considered as significant. Logarithmic values in terms of log10 CFU/mL for viable bacterial

count were used in order to normalize the data. The logarithmic mean, mean log10

CFU/mL was calculated by averaging the individual log10 CFU/mL values. The mean log

reduction (LR) in CFU/mL was calculated by subtracting the mean log10 CFU/mL of

negative control from mean log10 CFU/mL of test samples. Mean LR CFU/mL ≥1 was

considered as significant. Percentage bacterial load reduction was calculated as shown

in following formula below (eqn. 1):

𝑃𝑒𝑟𝑐𝑒𝑛𝑡𝑎𝑔𝑒 𝑟𝑒𝑑𝑢𝑐𝑡𝑖𝑜𝑛 = (A−B) x 100

𝐴 (eqn. 2)

Where,

A is the number of viable bacteria before treatment

B is the number of viable bacteria after treatment

Chapter 3 RESULTS AND DISCUSSION

51

3.1 Introduction

The emergence and spread of multi-drug resistant (MDR) bacteria are alarming and have

prompted interests in the search for fresh alternative schemes to tackle the problem

(Parmar, et al., 2017). One example is P. aeruginosa, an opportunistic pathogen

commonly isolated in clinical samples (Yu, et al., 2017). Phage biocontrol has received

an increasing level of interest by many researchers to mitigate the propagation of

antibiotic-resistance bacteria (Viertel, et al., 2014). Virulent bacteriophages (phages)

represent a viable antibacterial scheme that could be particularly beneficial to control

pathogenic bacteria with little impact on the rest of microbial community (Loc-Carrillo

and Abedon, 2011). One rising application of lytic phages is disinfection of surfaces and

materials commonly used in hospitals and food processing industries. The disinfection

of hard surfaces faces considerable challenges due to an increase in bacterial resistance

to traditional chemical sanitizers including hypochlorous acid and benzalkonium

chloride (Abuladze, et al., 2008). The study in this chapter explores the prospect of

isolating lytic bacteriophages from limestone caves with potentials to be utilised as

biological disinfectants to control infections caused by P. aeruginosa bacteria. Studies

on isolation and application of lytic bacteriophages obtained from limestone cave

environment have not been reported in preceding literature. However, the potential of

using cave microorganisms as a source of antimicrobial agents and drug discovery has

been recently reviewed (Ghosh, et al., 2016). This research gap forms the basis of the

current study. This chapter discusses the outcome of the experiments conducted to

isolate lytic bacteriophages from Sarawak limestone cave soils targeting various

pathogenic bacteria. Investigative studies on assessment of lytic abilities of P.

aeruginosa phages in an in-vitro co-culture assay at a varied MOI ratio and their

potentials to treat sand samples contaminated with P. aeruginosa PAO1 cells are also

reported. The results presented in this chapter shows presence and diversity of

bacteriophages in limestone cave environment with potentials to be further explored

and developed into biological disinfectants of P. aeruginosa.

52

3.2 Results

3.2.1 Isolation of lytic bacteriophages targeting bacterial pathogens

3.2.1.1 Soil collection

A total of seven samples (Table 3.1) were collected in January 2016 from FC (also known

as Gua Pari) as shown in Figure 3.1 and WC (also known as Lubang Angin) as shown in

Figure 3.2. These caves are about 5-7 km south-west of Bau and 30 km from Kuching,

Sarawak (Mohd, et al., 2011). The caves are part of the nature reserves protected by

environmental laws that preserve the forest, national parks, and nature reserve

(Omoregie, 2016). They cover 56 and 6.16 hectares respectively and are largely

surrounded by forests (Sarawak Forest Department, 1992).

Table 3.1: Description of soil samples collected at FCNR and WCNR

WC= Wind cave; FC= Fairy cave; oC= Temperature; (%) RH = Relative humidity.

Sample Code ID

Sample collected

Colour Texture oC (%) RH

WC1 Soil mixed with Guano

Yellowish-brown

Fine 30.6 94

WC2 Soil Brown Fine 29.7 90

WC3 Soil Black Coarse 28.7 84

FC1 Soil Brown Fine 24.8 76

FC2 Soil Black Coarse 26.5 73

FC2 Soil Brown Fine 28.1 80

FC4 Soil Brown Clay 30.6 79

53

Figure 3.1: Fairy Cave (FC) Bau, Sarawak, Malaysia. [A] Entrance view of the cave

(left). [B] View of cave chamber (right). Four samples were taken from inside the

cave chamber.

Figure 3.2: Wind Cave (WC) Bau, Sarawak, Malaysia. [A] Entrance to the cave (left). [B]

View of cave chamber (right). Three samples were taken from inside the cave chamber.

54

3.2.1.2 Bacterial host growth profile

Optical density (OD) at a wavelength of 600 nm, an indicator of bacterial growth, was

studied for up to 6 hr under aerobic batch conditions in a sterile Brain-heart infusion

(BHI) broth as presented in Figure 3.3. It was observed from the graph that, the growth

curve of the bacterial host increased in response to time and all the tested bacteria had

similar growth patterns for the total duration of the incubation. As indicated in Figure

3.3, bacterial cultures continued to have a progressive cell growth, hence, stationary

phase or death phase was not observed. The lag phase of all the bacterial hosts was

brief, noticeably and lasted for 0.5 hrs. The lag phase is usually characterized by no

immediate increase in cell numbers, as the bacteria are synthesizing new components.

During this stage, the cells may be old and depleted of ATP, essential, cofactors, and

ribosomes, thus these must be synthesized before growth can begin (Willey, et al.,

2009). This was followed by the log (exponential) phase marked by constant bacterial

growth rate and cell doubling in number at regular intervals. As seen in Figure 3.3 all the

bacterial hosts entered exponential phase after 1 hr of incubation and this phase lasted

for up to 3 hrs. Table 3.2 summarises the results of the growth kinetics of the bacterial

hosts during the batch culturing. The growth rate (specific growth rate) refers to the

change in a number of cells per minute, which can be estimated as the change in OD per

minute. Ideally, bacterial cultures grow exponentially mimicking a first-order chemical

reaction and the OD increases as a function of ln (OD), not OD itself (Hall, et al., 2013).

In this study, specific growth rate, (eqn. 3), at the different times of sampling was

estimated from the OD600 growth curve using five consecutive OD600 measurements

as described by Berney, et al. (2006) in the formula below. In practice, specific growth

rate, , is equal to the slope of ln OD versus time (t).

Specific growth rate, =lnOD600

t , where t is time (eqn. 3)

Doubling time (td) =𝐥𝐧𝟐

, where is the specific growth rate (eqn. 4)

Doubling time (eqn. 4) or generation time refers to the time it takes for cell division to

occur, with shorter time implying a more rapid bacterial growth (Maier, et al., 2009).

Doubling time can be calculated from a linear portion of a semilog plot of growth versus

55

time. The mathematical expression for this portion of the growth curve can be

rearranged and solved to calculate doubling time as shown in equation 4 above. Analysis

of the growth kinetics of the bacterial hosts showed that the highest specific growth rate

() (0.644 h-1) was exhibited by S. aureus whereas the lowest specific growth rate was

0.359 h-1 exhibited by P. aeruginosa. On the other hand, analysis of doubling time of the

bacteria revealed that P. aeruginosa had the shortest doubling time (td) of 0.249 and S.

aureus had the longest doubling time of 2.726. Maximum optical density (OD600) which

was studied for up to 6 hrs was achieved by P. aeruginosa (1.496), whereas, V.

parahaemolyticus (1.277) had the lowest maximum optical density among all the

bacteria strains.

Figure 3.3: Growth profile of the bacterial host cultures. The bacteria host were

grown in sterile brain-heart infusion broth (37.0 g. L-1Oxoid, Basingstoke, UK) at

37oC and 150 rpm for 6 hrs under aerobic conditions. Error bars represent standard

error of the mean.

0 2 4 60.0

0.5

1.0

1.5

2.0

Time (hrs)

Op

tical d

en

sit

y (

OD

600)

V. parahaemolyticus

P. aeruginosa

S. aureus

E. coli S. pneumoniae

K. pneumoniae

56

Table 3.2: Growth kinetics of bacterial hosts grown in batch cultures

3.2.1.3 Enrichment culturing and bacteriophage isolation

Using the methods presented in subsection 2.1.6, 2.1.7 and 2.1.8, a total of thirty-three

bacteriophages targeting different bacterial strains, and having distinct morphological

plaque formation were isolated from Sarawak limestone cave samples. Soil samples

obtained from Fairy and Wind Caves, were cultured in Brain-heart infusion (BHI) broth

containing 10 mM CaCl2 solution and a respective bacteria host, to screen for lytic

bacteriophages. About 5 mL of this soil-bacteria culture was inoculated into liquefied

nutrient agar supplemented with 1% (v/v) TTC solution (2,3,5-triphenyl tetrazolium

chloride) and 10 mM CaCl2 solution, and the contents plated out on pre-warmed Petri

dishes. Following an overnight incubation, plaques were formed in the areas where

phages destroyed bacteria cells. Uninfected viable bacteria cells developed into a

smooth lawn of confluent bacteria growth, which reduced TTC to red formazan turning

the agar red. Tetrazolium chloride (TTC) which was incorporated into the agar served as

a motility assay. The metabolic activity of viable active cells can break down TTC to TPF

(1,3,5-triphenyl formazan), a red colored compound (Kumar, et al., 2011). Lytic

bacteriophages were isolated based on their ability to form clear plaques on their

respective bacteria lawns. Figure 3.4 shows plaque formation due to lysis of the bacterial

host S. aureus and S. pneumoniae after 24 hrs incubation at 37oC. About 79% of the

phage isolates were obtained from FCNR soil samples whereas the remaining 21%

represented phages obtained from WCNR. In Figure 3.5, the isolated phage particles

were enumerated using double-layer plaque assay and their titer recorded as PFU/mL.

Bacterial host cultures

Specific growth rate

, [h-1]

Doubling time, td [g]

Maximum optical density (OD600) of bacteria

V. parahaemolyticus 0.623 0.432 1.277

P. aeruginosa 0.359 0.249 1.496

S. aureus 0.644 0.446 1.476

E. coli 0.406 0.282 1.357

S. pneumoniae 0.453 0.314 1.333

K. pneumoniae 0.418 0.290 1.476

57

Figure 3.4: Plaque appearance of bacteriophages infecting (A) S. aureus [left] and S.

pneumoniae (B) [right]. The incubated Petri dishes contained nutrient agar supplemented

with 10 mM CaCl2 and 1% TTC (2,3,5-triphenyltetrazolium chloride).

Figure 3.5: Phage titer determination of FCPA3 by double-layer plaque assay. Petri plates

from top-left to top-right shows lower dilution of viral titer, the Petri plates from the lower-

left to lower-right shows higher dilution of viral titer.

58

Table 3.3 outlines the sample description and phage plaque characteristics of the

isolates obtained from Sarawak limestone cave (FCNR and WCNR) soil samples. Majority

of the phage isolates exhibited distinctive features such as mixed plaque morphology

suggesting the presence of different phage traits infecting the same bacterial host

(Gallet, et al., 2011). Several turbid phages were discarded because it was likely they

were formed by temperate phages which are not fit for phage therapy studies. After a

careful examination of plaque morphology, distinctive phage plaques were amplified in

BHI broth enriched with their respective bacteria host as explained in section 2.1.7.

Amplified phage lysates were subjected to phage titer assay using the double-layer

plaque technique so as to determine the concentration of the phage particles. Amongst

all the isolates, P. aeruginosa infecting bacteriophages designated as FCPA4,

WCSS4PA and WCSS5PA showed the highest phage titer (1015 PFU/mL). Figure 3.5,

shows phage titer assay, determined by double-layer plaque assay for phage FCPA3

after 24 hrs of incubation at 37oC. Titrated phages were preserved in sterile cryotube

containing 200 μL of 75% (v/v) glycerol in phage buffer (PB) using modified methods

adapted from Fortier and Moineau (2009) and (Pardon, et al., 2014) as explained in

subsection 2.1.10 and the cryotube was then stored in a freezer at -80oC.

59

Table 3.3: Morphological characteristics of bacteriophages isolated from FCNR and WCNR

Sampling Origin Designated phage

ID Bacteria host

Plaque

Size

(mm)

Plaque description Phage titer

(PFU/mL)

FCNR FCVP1 V. parahaemolyticus 6 Clear, round 1.19 x 109






WCNR WCVP3 V. parahaemolyticus Nil Turbid Nil



FCNR FCSA1 S. aureus 4 Clear, round 1.89 x 107




FCNR FCKP1 K. pneumoniae 3 Clear, round 2.26 x 1012

WCNR WCKP1 K. pneumoniae 3 Clear, round 6.1 x 1015

WCNR WCKP4 K. pneumoniae 2 Clear, round 1.51 x 1014

FCNR FCEC1 E. coli 6 Clear, round 4.3 x 106


60


FCNR FCEC7 E. coli Nil Turbid Nil

FCNR FCPA1 P. aeruginosa 3 Clear, round 2.28 x 1013






WCNR WCSS4PA P. aeruginosa Nil Clear, web-pattern 1.25 x 1015

WCNR WCSS5PA P. aeruginosa Nil Clear, web-pattern 4.5 x 1015

FCNR FCSP1 S. pneumoniae 5 Clear, round 1.50 x 108





61

3.2.1.4 Bacteriophage host range analysis

One of the goals of this study was to determine the phage specificity with the

expectation that some phages would have broader host range than others due to the

presence or absence of phage receptor molecules or intracellular restriction

mechanisms (Jensen, et al., 2015). Spot tests were performed on TSA Petri plates

containing lawns of various bacteria as described in subsection 2.1.12. The Petri plates

were then assessed for the presence of plaques on the lawns of the bacteria (Figure 3.6).

Based on spot test results as shown in Table 3.4, the majority of phage isolates were

capable of infecting E. coli (72.7%), P. aeruginosa (66.7%) and K. pneumoniae

(48.5%) bacterial strains. The broadest host range was exhibited by a P. aeruginosa

phage designated as FCPA3 which was capable of lysing S. aureus, K. pneumonia,

E. coli and S. typhimurium bacterial strains. This phage exhibited high virulence on

S. aureus and K. pneumoniae bacteria lawns. Generally, broad host range was seen

in V. parahaemolyticus and P. aeruginosa phage isolates. For example, V.

parahaemolyticus phages (FCVP1, FCVP2, FCVP3) were able to lyse bacteria

stains S. aureus, P. aeruginosa and E. coli, while P. aeruginosa phages (FCPA1,

FCPA2, FCPA4, FCPA5 and FCPA6) were capable of lysing bacteria strains S.

aureus, E. coli and S. typhirium. In addition, bacterial strains S. pneumoniae and S.

typhirium were lysed by the least number of phages among all the bacteria tested.

Figure 3.6: Re-confirmation of P. aeruginosa bacteriophage lytic ability by spot

test assay. Bacteriophages were spot-tested on Tryptic soy agar (supplemented

with 10 mM CaCl2) containing lawn of P. aeruginosa.

62

Table 3.4: Assessment bacteriophage host range by spot test assay

Designated

Phage ID

V. parahaem

olyticus

S. aureus P.

aeruginosa

S. pneumon

are

K. pneumo

niae

E. coli S.

typhirium

FCVP1 + ++ ++ - - + -

FCVP2 ++ ++ + - - ++ -

FCVP3 ++ ++ ++ - - ++ -

FCVP4 ++ ++ ++ - - ++ -

FCVP5 ++ - ++ - - ++ -

FCVP6 ++ - ++ - - ++ -

WCVP3 - - - - + ++ -

WCVP4 - - ++ - + ++ -

WCVP5 - - ++ - - + -

FCSA1 - ++ - - ++ + -

FCSA3 - ++ - - ++ + -

FCSA4 - - - - ++ + -

FCSA6 - - - - ++ + -

FCKP1 - - - ++ ++ - -

WCKP1 - - - - ++ - -

WCKP4 - - ++ - ++ - -

FCEC1 - - - - ++ ++ -

FCEC3 - - - - ++ ++ -

WCEC6 - - - - ++ - -

WCEC7 - - - - ++ - -

FCPA1 - + ++ - - + +

FCPA2 - + ++ - - + +

63

Lysis pattern was measured qualitatively as (++) for full lysis, (+) for partial lysis and (-) for no lysis.

FCPA3 - ++ ++ - ++ + +

FCPA4 - + ++ - - + +

FCPA5 - + ++ - - + +

FCPA6 - + ++ - - + +

WCSS4PA - - ++ - ++ - -

WCSS5PA - - ++ - ++ - -

FCSP1 - - ++ ++ - ++ -

FCSP2 - - ++ ++ - ++ -

FCSP3 - - ++ ++ - ++ -

FCSP4 - - ++ - - - -

FCSP5 - + ++ - - - -

64

3.2.2 In-vitro studies on phage bacteriolytic activity and assessment of bacterial

survival following phage treatment.

3.2.2.1 Phage bacteriolytic activity

The lytic abilities of bacteriophages against P. aeruginosa PAO1 cells were evaluated in

an in-vitro co-culture assay for up to 6 hrs at varied MOI ratios as shown in Figure 3.7 to

Figure 3.15. The results showed that the growth of P. aeruginosa PAO1 was inactivated

when co-cultured with phage in a concentration-dependent manner, with OD values

declining more quickly at higher MOI (104 and 105) than at lower MOI (103, 102 or 101).

The OD values decrease very quickly, just 30 minutes after phage addition except for

phages FCPA4 and FCPA6, suggesting the occurrence of bacterial lysis. However, it

was possible to differentiate these phages into three clusters, the first consisting of

FCPA1, FCPA2, FCPA4 and FCPA6, for which OD values decreased slowly and

steadily until the end of the incubation time when compared with uninfected control.

This group was not very effective at inactivating the bacteria in all the tested MOI ratios

except FCPA2 (MOI 105). The second cluster consisted of phages FCPA5, WCSS4PA

and WCSS5PA. In this group, the OD values decreased slowly for a period of time and

then rose slowly again until the end of the incubation time. For instance, in FCPA5, the

OD decreased steadily for up to 4 hrs and then slowly started rising until the end of the

6 hrs of incubation. In phage WCSS5PA, the OD values decreased steadily for the first

3 hrs and then took a sharp rise until the end of the 6 hrs of incubation. In phage

WCSS4PA, OD values were seen to decrease sharply in the first 2 hrs and then rose

again slowly and steadily until the end of the 6 hrs of incubation, except for the highest

MOI (105) where the lower OD values were maintained throughout the incubation

period. The last cluster consisted of only the phage cocktail (Cocktail). In phage cocktail

(Cocktail), the OD decreased sharply for the first 2 hrs then maintained this lower OD

until the end of the 6 hrs of incubation. Table 3.5 shows absorbance readings obtained

from the phage treated cultures at the end of the 6 hrs of incubation. From this result,

bacteriophages FCPA3 (MOI 105), WCSS4PA (MOI 105) and Cocktail (MOI 104)

showed the highest bacterial inactivation with OD values decreasing to 0.200, 0.319 and

0.288 when compared with the OD of untreated controls i.e. 1.370, 1.533, 1.557

respectively.

65

Table 3.5: Assessment of phage bacteriolytic activity at the end of 6 hrs of incubation

Phage ID Tested MOI ratio Samples absorbance

(OD600)

FCPA1 101 1.377

102 1.362

103 1.461

104 1.283

105 1.155

Uninfected control 1.370

FCPA2 101 1.418

102 1.171

103 1.043

104 1.069

105 0.703


FCPA3 101 0.766

102 0.566

103 0.588

104 0.383

105 0.200


FCPA4 101 1.431

102 1.441

103 1.493

104 1.527

105 1.495


66

FCPA5

101 1.407

102 1.129

103 0.941

104 0.919

105 0.900


FCPA6 101 1.188

102 1.266

103 1.309

104 1.317

105 1.329


WCSS4PA 101 1.229

102 1.189

103 1.168

104 0.897

105 0.319


WCSS5PA 101 1.136

102 1.076

103 1.105

104 1.096

105 1.146


Cocktail 101 0.945

102 0.616

103 0.349

104 0.288

105 0.405


67

Figure 3.7: In-vitro bacteriolytic activity of FCPA1 at different MOI ratios. Mid–

exponential cultures of P. aeruginosa PAO1 were co-cultured with bacteriophage

FCPA1 at different multiplicity of infection (MOI) ratios. Error bars represent

standard error of the mean

0 2 4 60.0

0.5

1.0

1.5

Time (hrs)

Op

tical d

en

sit

y (

OD

600)

Control

ΦFCPA1 (MOI 105)ΦFCPA1 (MOI 104)ΦFCPA1 (MOI 103)

ΦFCPA1 (MOI 102)ΦFCPA1 (MOI 101)

68




standard error of the mean

0 2 4 60.0

0.5

1.0

1.5

Time (hrs)

Op

tical d

en

sit

y (

OD

600)

Control



69


exponential cultures of P. aeruginosa PAO1 were co-cultured with phage FCPA3

at different multiplicity of infection (MOI) ratios. Error bars represent standard

error of the mean.

0 2 4 60.0

0.5

1.0

1.5

2.0

Time (hrs)

Op

tical d

en

sit

y (

OD

600)

Control



70




standard error of the mean.

0 2 4 60.0

0.5

1.0

1.5

2.0

Time (hrs)

Op

tical d

en

sit

y (

OD

600)

Control



71





0 2 4 60.0

0.5

1.0

1.5

2.0

Time (hrs)

Op

tical d

en

sit

y (

OD

600)

Control



72





0 2 4 60.0

0.5

1.0

1.5

2.0

Time (hrs)

Op

tical d

en

sit

y (

OD

600)

Control



73

Figure 3.13: In-vitro bacteriolytic activity of WCSS4PA at different MOI ratios.

Mid–exponential cultures of P. aeruginosa PAO1 were co-cultured with

bacteriophage WCSS4PA at different multiplicity of infection (MOI) ratios. Error

bars represent standard error of the mean.

0 2 4 60.0

0.5

1.0

1.5

2.0

Time (hrs)

Op

tical d

en

sit

y (

OD

600)

Control

ΦWCSS4PA (MOI 105)ΦWCSS4PA (MOI 104)ΦWCSS4PA (MOI 103)

ΦWCSS4PA (MOI 102)WCSS4PA (MOI 101)

74

Figure 3.14: In-vitro bacteriolytic activity of WCSS5PA at different MOI ratios.

Mid–exponential cultures of P. aeruginosa PAO1 were co-cultured with

bacteriophage WCSS5PA at different multiplicity of infection (MOI) ratios. Error

bars represent standard error of the mean.

0 2 4 60.0

0.5

1.0

1.5

2.0

Time (hrs)

Op

tical d

en

sit

y (

OD

600)

Control

ΦWCSS5PA (MOI 105)ΦWCSS5PA (MOI 104)ΦWCSS5PA (MOI 103)

ΦWCSS5PA (MOI 102)ΦWCSS5PA (MOI 101)

75

Figure 3.15: In-vitro bacteriolytic activity of Cocktail at different MOI ratios. Mid–

exponential cultures of P. aeruginosa PAO1 were co-cultured with phage cocktail

(Cocktail) at different multiplicity of infection (MOI) ratios. Error bars represent


3.2.2.2 Assessment of bacterial survival following phage in-vitro bacteriolytic activity.

To evaluate the efficiency of the phage isolates to inhibit or eradicate P. aeruginosa

PAO1 cells, the in-vitro susceptibility of P. aeruginosa PAO1 bacteria cells to

bacteriophages at varied MOI ratios were assessed at 6 hrs and 24 hrs post-infection as

explained in subsection 2.2.4. The surviving bacterial cells were expressed as log10

CFU/mL and were compared with those of uninfected control. Surviving P. aeruginosa

PAO1 cells following an in-vitro treatment with bacteriophages FCPA1, FCPA2,

FCPA3, FCPA4, FCPA5, FCPA6, WCSS4PA, WCSS5PA and Cocktail at varied MOI

ratios are presented in Figure 3.16 to 3.24.

0 2 4 60.0

0.5

1.0

1.5

2.0

Time (hrs)

Op

tical d

en

sit

y (

OD

600)

Control

ΦCocktail (MOI 105)ΦCocktail (MOI 104)Φ(MOI 103)

Φ(MOI 102)Φ(MOI 101)

76

The in-vitro susceptibility results of P. aeruginosa PAO1 cells to phage FCPA1 as

presented in Figure 3.16, showed significant bacterial log reductions at both 6 hrs and

24 hrs post-infection. The lowest bacterial recovery was achieved at MOI of 105. Exactly

6 hrs post-infection, the recovered bacteria cells at MOI of 105 were 10.6 log10 CFU/mL

and those of uninfected control were 16.9 log10 CFU/mL. This resulted in a 6.3 log

reduction in bacteria cells. Even at lower MOI ratios (101, 102, 103, 104), FCPA1

managed to reduce P. aeruginosa PAO1 cells by 4.2, 4.7, 5.5 and 5.3 logs respectively.

At 24 hrs post-infection, recovered bacteria cells at MOI of 105 were 13.7 log10 CFU/mL

and those of uninfected control were 19.2 log10 CFU/mL, resulting in a 5.5 log reduction

in bacteria cells. The results obtained from ANOVA (Table 3.6) and Tukey-Kramer’s post

hoc analysis revealed that there were significant differences among all the group means

except when the mean of MOI 102 at 6 hrs was compared with the means of MOI 103 at

24 hrs and MOI 104 at 24 hrs, when the mean of MOI 103 at 6 hrs was compared with

the means of MOI 104 at 6 hrs and MOI 105 at 6 hrs, when the mean of MOI 104 at 6 hrs

was compared with the means of MOI 105 at 6 hrs, when the mean of MOI 101 at 24 hrs

was compared with the means of MOI 102 at 24 hrs and MOI 104 at 24 hrs and when

MOI 105 at 24 hrs, when the mean of MOI 102 at 24 hrs was compared with the means

of MOI 105 at 24 hrs, when the mean of MOI 104 at 24 hrs was compared with the means

of MOI 103 at 24 hrs and MOI 105 at 24 hrs, and when the uninfected control at 6 hrs

was compared with the means of MOI 101 at 24 hrs, MOI 102 at 24 hrs and MOI 105 at

24 hrs, respectively.

77

Table 3.6: Analysis of variance (ANOVA) results for the recovery of bacteria following an

in-vitro treatment with FCPA1

Group Sum Mean Sample variance

Standard deviation

MOI 101 (6 hrs) 38.048 12.683 0.359 0.3461

MOI 101 (24 hrs) 52.858 17.619 0.288 0.3098

MOI 102 (6 hrs) 36.504 12.168 0.279 0.3052

MOI 102 (24 hrs) 45.087 15.029 0.000351

0.0106

MOI 103 (6 hrs) 34.354 11.451 0.287 0.3092

MOI 103 (24 hrs) 55.312 18.437 0.098 0.1808

MOI 104 (6 hrs) 34.718 11.573 0.397 0.3639

MOI 104 (24 hrs) 42.918 14.306 0.0000792

0.0051

MOI 105 (6 hrs) 31.794 10.598 0.23 0.2768

MOI 105 (24 hrs) 41.078 13.693 0.171 0.2389

Control (6 hrs) 50.713 16.9043 0.2801 0.3056

Control (24 hrs) 57.498 19.166 0.000218

0.0085

sample size mean =3; degrees of freedom =11,24; sum of squares =286.728, 4.768; mean square =26.066, 0.199 and F statistic =130.803.

The results of phage FCPA2 (Figure 3.17) revealed significant bacterial log reductions

in higher MOI ratios (103, 104, 105) than lower MOI ratios (101 and 102) at 6 hrs post-

infection. MOI ratio of 104 had the highest bacterial log reduction when compared with

the rest MOI ratios. Recovered bacteria cells were 7.7 log10 CFU/mL and those of the

uninfected control were 16.9 log10 CFU/mL, resulting in a 9.2 log reduction in bacterial

cells. On the other hand, MOI ratio of 103 showed the highest bacterial log reduction

when compared with the rest tested MOI ratios at 24 hrs post-infection. At this MOI,

the recovered bacteria cells were 13.7 log10 CFU/mL whereas those of uninfected control

were 19.2 log10 CFU/mL resulting in a 5.5 log reduction in bacteria cells. The results

obtained from ANOVA (Table 3.7) and Tukey-Kramer’s post hoc analysis revealed that

78

there were significant differences among all the group means except when the mean of

MOI 102 at 6 hrs was compared with the means of MOI 103 at 24 hrs and MOI 104 at 24

hrs, when the mean of MOI 103 at 6 hrs was compared with the means of MOI 104 at 6

hrs and MOI 105 at 6 hrs, when the mean of MOI 104 at 6 hrs was compared with the

means of MOI 105 at 6 hrs, when the mean of MOI 101 at 24 hrs was compared with the

means of MOI 102 at 24 hrs and MOI 104 at 24 hrs and when MOI 105 at 24 hrs, when

the mean of MOI 102 at 24 hrs was compared with the means of MOI 105 at 24 hrs, when

the mean of MOI 104 at 24 hrs was compared with the means of MOI 103 at 24 hrs and

MOI 105 at 24 hrs, and when the uninfected control at 6 hrs was compared with the

means of MOI 101 at 24 hrs, MOI 102 at 24 hrs and MOI 105 at 24 hrs, respectively.



Group Sum Mean Sample

variance

Standard

deviation

MOI 101 (6 hrs) 27.985 9.328 0.0000166 1.039

MOI 101 (24 hrs) 47.617 15.872 0.309 0.042

MOI 102 (6 hrs) 23.812 7.937 0.048 0.133

MOI 102 (24 hrs) 44.154 14.718 0.202 0.568

MOI 103 (6 hrs) 26.828 8.943 0.000227 0.015

MOI 103 (24 hrs) 41.185 13.728 0.222 0.471

MOI 104 (6 hrs) 41.015 13.672 0.018 0.218

MOI 104 (24 hrs) 50.945 16.982 0.322 0.45

MOI 105 (6 hrs) 36.049 12.016 1.08 0.004

MOI 105 (24 hrs) 46.876 15.625 0.002 0.556

Control (6 hrs) 50.713 16.9043 0.2801 0.3056

Control (24 hrs) 57.498 19.166 0.000218 0.0085

sample size mean =3; degrees of freedom =11,24; sum of squares =471.440, 4.405; mean square =42.858, 0.184, and F statistic =233.513.

79

The results of phage FCPA3, as shown in Figure 3.18 revealed presence of significant

bacterial log reductions in samples infected at MOI of 104 and 105 when compared with

the rest tested MOIs at 6 hrs post-infection. Recovered bacteria cells were 7.7 log10

CFU/mL and 8.2 log10 CFU/mL respectively, whereas those of an uninfected control was

16.9 log10 CFU/mL. This resulted in a 9.2 and 8.7 log reduction in bacterial cells

respectively. At 24 hrs post-infection, MOI of 105 had the highest bacteria log reduction

when compared with the rest MOIs. At this MOI, recovered bacteria cells were 10.9 log10

CFU/mL and those of uninfected control were 19.2 log10 CFU/mL, resulting in an 8.3 log

reduction in bacteria cells. The results obtained from ANOVA (Table 3.8) and Tukey-

Kramer’s post hoc analysis revealed that there were significant differences among all

the group means except when the mean of MOI 101 at 6 hrs was compared with the

means of MOI 104 at 24 hrs and MOI 105 at 24 hrs, when MOI 102 at 6 hrs was compared

with the means of MOI 103 at 6 hrs, MOI 104 at 24 hrs and MOI 105 at 24 hrs, when the

means of MOI 103 at 6 hrs was compared with the means of MOI 104 at 6 hrs, MOI 103

at 24 hrs, MOI 104 at 24 hrs and MOI 105 at 24 hrs, when the means of MOI 104 at 6 hrs

was compared with the means of MOI 103 at 24 hrs and MOI 104 at 24 hrs, when the

means of MOI 101 at 24 hrs was compared with the means of MOI 102 at 24 hrs, when

the means of MOI 103 at 24 hrs was compared with the means of MOI 104 at 24 hrs and

MOI 105 at 24 hrs, when the means of MOI 104 at 24 hrs was compared with the means

of MOI 105 at 24 hrs, and the mean of uninfected control at 6 hrs was compared with

the means of MOI 101 at 24 hrs and MOI 102 at 24 hrs respectively.

80




variance

Standard

deviation

MOI 101 (6 hrs) 35.221 11.74 0.000250 0.0091

MOI 101 (24 hrs) 44.981 14.994 0.328 0.3306

MOI 102 (6 hrs) 33.813 11.271 0.0000880 0.0054

MOI 102 (24 hrs) 44.252 14.751 0.0770 0.1605

MOI 103 (6 hrs) 30.751 10.25 0.0000238 0.0028

MOI 103 (24 hrs) 42.771 14.257 0.0000249 0.0029

MOI 104 (6 hrs) 22.952 7.651 0.155 0.2274

MOI 104 (24 hrs) 41.725 13.908 0.000115 0.0062

MOI 105 (6 hrs) 24.742 8.247 0.159 0.2302

MOI 105 (24 hrs) 32.596 10.865 0.293 0.3123

Control (6 hrs) 50.713 16.9043 0.2801 0.3056

Control (24 hrs) 57.498 19.166 0.000218 0.0085


The results of phage FCPA4 as shown in Figure 3.19 revealed presence of significant

bacterial log reductions in samples infected at MOI of 105 when compared with the rest

MOIs (101, 102, 103, 104) at 6 hrs post-infection. Recovered bacteria cells at MOI 105

were 8.6 log10 CFU/mL whereas those of uninfected control were 16.9 log10 CFU/mL,

resulting in an 8.3 log reduction of bacterial cells. At 24 hrs post-infection, MOI of 103

showed the highest bacteria log reduction when compared with the rest MOIs. At this

MOI, recovered bacteria cells were 12.2 log10 CFU/mL and those of uninfected control

were 19.2 log10 CFU/mL, resulting in a 7.0 log reduction in bacterial cells. The results

obtained from ANOVA (Table 3.9) and Tukey-Kramer’s post hoc analysis revealed that

there were significant differences among all the group means except when the mean of

81

MOI 101 at 6 hrs was compared with the means of MOI 104 at 24 hrs and MOI 105 at 24

hrs, when MOI 102 at 6 hrs was compared with the means of MOI 103 at 6 hrs, MOI 104

at 24 hrs and MOI 105 at 24 hrs, when the means of MOI 103 at 6 hrs was compared with

the means of MOI 104 at 6 hrs, MOI 103 at 24 hrs, MOI 104 at 24 hrs and MOI 105 at 24

hrs, when the means of MOI 104 at 6 hrs was compared with the means of MOI 103 at

24 hrs and MOI 104 at 24 hrs, when the means of MOI 101 at 24 hrs was compared with

the means of MOI 102 at 24 hrs, when the means of MOI 103 at 24 hrs was compared

with the means of MOI 104 at 24 hrs and MOI 105 at 24 hrs, when the means of MOI 104

at 24 hrs was compared with the means of MOI 105 at 24 hrs, and the mean of uninfected

control at 6 hrs was compared with the means of MOI 101 at 24 hrs and MOI 102 at 24

hrs respectively.

Table 3.9: Analysis of variance (ANOVA) results for recovery of the bacteria following an



variance

Standard

deviation

MOI 101 (6 hrs) 41.103 13.701 0.0009 0.0174

MOI 101 (24 hrs) 49.181 16.394 0.2594 0.294

MOI 102 (6 hrs) 40.505 13.502 0.1629 0.233

MOI 102 (24 hrs) 49.702 16.567 0.2465 0.2867

MOI 103 (6 hrs) 37.311 12.437 0.0626 0.1445

MOI 103 (24 hrs) 36.683 12.228 0 0.0036

MOI 104 (6 hrs) 35.299 11.766 0.3647 0.3487

MOI 104 (24 hrs) 38.334 12.778 0 0.0015

MOI 105 (6 hrs) 25.787 8.596 0.0001 0.0042

MOI 105 (24 hrs) 39.238 13.079 0 0.0021

Control (6 hrs) 50.713 16.9043 0.2801 0.3056

Control (24 hrs) 57.498 19.166 0.000218 0.0085


82

The results of phage FCPA5 (Figure 3.20) revealed the presence of significant bacterial

log reduction in samples infected at higher MOI (103, 104, 105) than lower MOI (101 and

102) at 6 hrs post-infection. The highest bacterial log reduction was observed from

recovered bacterial cells which were infected at MOI of 105. At this MOI, recovered

bacterial cells were 7.5 log10 CFU/mL and those of uninfected control were 16.9 log10

CFU/mL, resulting in a 9.4 log reduction in bacterial cells. On the other hand, recovered

bacterial cells harvested at 24 hrs post-infection revealed a significant log reduction at

MOI of 104 when compared with the rest MOIs. At this MOI, recovered bacteria cells

were 13.3 log10 CFU/mL whereas those of uninfected control were 19.2 log10 CFU/mL

resulting in a 5.9 log reduction in bacterial cells. The results obtained from ANOVA (Table

3.10) and Tukey-Kramer’s post hoc analysis revealed that there were significant

differences among all the group means except when the mean of MOI 101 at 6 hrs was

compared with the means of MOI 102 at 6 hrs and MOI 104 at 24 hrs, when MOI 103 at

6 hrs was compared with the means of MOI 104 at 6 hrs and MOI 105 at 6 hrs, when MOI

101 at 24 hrs was compared with the means of MOI 102 at 24 hrs and MOI 103 at 24 hrs,

when MOI 103 at 24 hrs was compared with the means of MOI 101 at 24 hrs and MOI

105 at 24 hrs, and when the mean of uninfected control at 6 hrs was compared with the

mean of MOI 101 at 24 hrs, MOI 102 at 24 hrs and MOI 103 at 24 hrs, respectively.

83

Table 3.10: Analysis of variance (ANOVA) results for the recovery of bacteria following

an in-vitro treatment with FCPA5


variance

Standard

deviation

MOI 101 (6 hrs) 38.189 12.73 0.000154 0.0071

MOI 101 (24 hrs) 50.209 16.736 0.297 0.3147

MOI 102 (6 hrs) 34.592 11.531 0.001 0.02

MOI 102 (24 hrs) 50.532 16.844 0.271 0.3005

MOI 103 (6 hrs) 28.928 9.643 0.341 0.3373

MOI 103 (24 hrs) 47.148 15.716 0.199 0.2573

MOI 104 (6 hrs) 26.334 8.778 0.000360 0.0109

MOI 104 (24 hrs) 39.935 13.312 0.0000184 0.0026

MOI 105 (6 hrs) 22.544 7.515 0.186 0.2489

MOI 105 (24 hrs) 44.632 14.877 0.304 0.3183

Control (6 hrs) 50.713 16.9043 0.2801 0.3056

Control (24 hrs) 57.498 19.166 0.000218 0.0085


The results of phage FCPA6, as shown in Figure 3.21, revealed significant log reduction

in bacteria at MOI of 103 and 104 compared with the rest MOIs at 6 hrs post-infection.

The highest bacterial log reduction was observed in samples infected at MOI of 104. At

this MOI ratio, recovered bacterial cells were 7.3 log10 CFU/mL whereas those of

uninfected control were 16.9 log10CFU.mL-1, resulting in 9.6 log reduction in bacterial

cells. On the other hand, bacterial cells recovered at 24 hrs post-infection revealed a

significant bacterial log reduction at MOI of 105 when compared with the rest MOI

ratios. At this MOI, recovered bacteria cells were 13.0 log10 CFU/mL whereas those of

uninfected control were 19.2 log10 CFU/mL, resulting in a 6.2 log reduction in bacterial

cells. The results obtained from ANOVA (Table 3.11) and Tukey-Kramer’s post hoc

analysis revealed that there were significant differences among all the group means

84

except when the mean of MOI 101 at 6 hrs was compared with the means of MOI 102 at

6 hrs and MOI 105 at 6 hrs, when the mean of MOI 105 at 6 hrs was compared with the

means of MOI 102 at 6 hrs and MOI 101 at 6 hrs, when the mean of MOI 101 at 24 hrs

was compared with the means of MOI 102 at 24 hrs and MOI 103 at 24 hrs, and when

the mean of uninfected control at 6 hrs was compared with the mean of MOI 102 at 24

hrs and MOI 103 at 24 hrs, respectively.


an in-vitro treatment with FCPA6


variance

Standard

deviation

MOI 101 (6 hrs) 50.713 16.904 0.280 0.1921

MOI 101 (24 hrs) 57.498 19.166 0.000219 0.0102

MOI 102 (6 hrs) 50.713 16.904 0.28 0.3482

MOI 102 (24 hrs) 57.498 19.166 0.000219 0.0073

MOI 103 (6 hrs) 50.713 16.904 0.28 0.3896

MOI 103 (24 hrs) 57.498 19.166 0.000219 0.2938

MOI 104 (6 hrs) 50.713 16.904 0.280 0.1479

MOI 104 (24 hrs) 57.498 19.166 0.000219 0.0062

MOI 105 (6 hrs) 50.713 16.904 0.28 0.3025

MOI 105 (24 hrs) 57.498 19.166 0.000219 0.0023

Control (6 hrs) 50.713 16.9043 0.2801 0.3056

Control (24 hrs) 57.498 19.166 0.000218 0.0085

sample size mean =3; degrees of freedom =11,24; sum of squares (46.036, 3.64) and mean square (4.185, 0.140), F statistic (229.857).

85

The results of phage WCSS4PA as seen in Figure 3.22, showed significant log reductions

in bacteria cells at 6 hrs post-infection in all the MOI ratios. However, the highest

bacterial log reduction was observed in samples infected at MOI of 105. At this MOI ratio,

the recovered bacterial cells were 6.6 log10 CFU/mL and those of uninfected control

were 16.9 log10 CFU/mL, resulting in a 10.3 log reduction in bacterial cells. Similarly, the

recovered bacterial cells at 24 hrs post-infection revealed significant bacterial log

reductions in all the MOI ratios. However, the highest log reduction was observed in

bacterial cultures infected at MOI of 104. At this MOI, recovered bacteria cells were 8.7

log10 CFU/mL whereas those of uninfected control were 19.2 log10 CFU/mL, resulting in

a 10.5 log reduction in bacterial cells. The results obtained from ANOVA (Table 3.12) and

Tukey-Kramer’s post hoc analysis revealed that there were significant differences

among all the group means except when the mean of MOI 101 at 6 hrs was compared

with the means of MOI 102 at 6 hrs, MOI 103 at 6 hrs, MOI 104 at 6 hrs, MOI 101 at 24

hrs, MOI 102 at 24 hrs, MOI 103 at 24 hrs and MOI 105 at 24 hrs, the mean of MOI 102 at

6 hrs was compared with the means of MOI 103 at 6 hrs, MOI 104 at 6 hrs, MOI 101 at 24

hrs, MOI 102 at 24 hrs, MOI 103 at 24 hrs, MOI 104 at 24 hrs and MOI 105 at 24 hrs, the

mean of MOI 103 at 6 hrs was compared with the means of MOI 101 at 24 hrs, MOI 103

at 24 hrs and MOI 104 at 24 hrs, the mean of MOI 104 at 6 hrs was compared with the

means of MOI 101 at 24 hrs, MOI 102 at 24 hrs and MOI 103 at 24 hrs, the mean of MOI

101 at 24 hrs was compared with the means of MOI 102 at 24 hrs, MOI 103 at 24 hrs and

MOI 105 at 24 hrs, the mean of MOI 102 at 24 hrs was compared with the means of MOI

103 at 24 hrs, and MOI 105 at 24 hrs and the mean of MOI 103 at 24 hrs was compared

with the means of MOI 104 at 24 hrs, and MOI 105 at 24 hrs, respectively.

86


an in-vitro treatment with WCSS4PA


variance

Standard

deviation

MOI 101 (6 hrs) 31.276 10.425 0.0710 0.1535

MOI 101 (24 hrs) 31.311 10.437 0.247 0.287

MOI 102 (6 hrs) 30.204 10.068 0.24 0.2828

MOI 102 (24 hrs) 32.645 10.882 0.263 0.2958

MOI 103 (6 hrs) 27.255 9.085 0.0000298 0.0032

MOI 103 (24 hrs) 28.894 9.631 0.258 0.293

MOI 104 (6 hrs) 31.956 10.652 0.0190 0.0805

MOI 104 (24 hrs) 26.149 8.716 0.241 0.2836

MOI 105 (6 hrs) 19.876 6.625 0.166 0.235

MOI 105 (24 hrs) 31.995 10.665 0.151 0.2242

Control (6 hrs) 50.713 16.9043 0.2801 0.3056

Control (24 hrs) 57.498 19.166 0.000218 0.0085


The results of phage WCSS5PA as seen in Figure 3.23, revealed the lowest bacterial log

reduction in samples infected at MOI of 104 at 6 hrs post-infection when compared with

the rest MOI ratios. At this MOI, the recovered bacterial cells were 12.1 log10 CFU/mL

whereas those of uninfected control were 16.9 log10 CFU/mL, resulting in a 4.8 log

reduction in bacterial cells. At 24 hrs post-infection, the highest bacteria log reduction

was observed in cultures infected at MOI of 105. At this MOI, recovered bacterial cells

were 12.2 log10 CFU/mL whereas those of uninfected control were 19.2 log10 CFU/mL,

resulting in a 7.0 log reduction in bacterial cells. The results obtained from ANOVA (Table

3.13) and Tukey-Kramer’s post hoc analysis revealed that there were significant

87

differences among all the group means except when the mean of MOI 101 at 6 hrs was

compared with the means of MOI 102 at 6 hrs, MOI 103 at 6 hrs, MOI 105 at 6 hrs and

MOI 101 at 24 hrs, when the mean of MOI 102 at 6 hrs was compared with the means of

MOI 103 at 6 hrs, MOI 104 at 6 hrs, MOI 105 at 6 hrs, MOI 104 at 24 hrs and MOI 105 at

24 hrs, when the mean of MOI 103 at 6 hrs was compared with the means of MOI 105 at

6 hrs and MOI 104 at 24 hrs, when the mean of MOI 104 at 6 hrs was compared with the

means of MOI 104 at 24 hrs, and when the mean of MOI 101 at 24 hrs was compared

with the means MOI 102 at 24, respectively.


an in-vitro treatment with WCSS5PA.


variance

Standard

deviation

MOI 101 (6 hrs) 40.725 13.575 0.0550 0.135

MOI 101 (24 hrs) 47.243 15.748 0.116 0.1968

MOI 102 (6 hrs) 37.934 12.645 0.182 0.246

MOI 102 (24 hrs) 54.215 18.072 0.000388 0.0114

MOI 103 (6 hrs) 39.462 13.154 0.00200 0.0251

MOI 103 (24 hrs) 44.928 14.976 0.0000494 0.0041

MOI 104 (6 hrs) 36.186 12.062 0.0000330 0.0033

MOI 104 (24 hrs) 39.785 13.262 0.0000132 0.0021

MOI 105 (6 hrs) 40.468 13.489 0.007 0.0495

MOI 105 (24 hrs) 36.557 12.186 0.244 0.2852

Control (6 hrs) 50.713 16.9043 0.2801 0.3056

Control (24 hrs) 57.498 19.166 0.000218 0.0085


88

The results of phage cocktail (Cocktail) as shown in Figure 3.24, revealed significant

bacterial log reductions in all the MOI ratios at both 6 hrs and 24 hrs post-infection. At

6 hrs post-infection, the lowest bacterial recovery was 5.08 log10 CFU/mL obtained from

samples treated at MOI 104. The uninfected control count was 16.90 log10 CFU/mL,

resulting in an 11.82 log reduction in bacterial cells. At 24 hrs post-infection, MOI of 102

had the highest bacterial log reduction when compared with the rest MOI ratios. At this

MOI, recovered bacterial cells were 8.34 log10 CFU/mL whereas those of uninfected

control were 19.2 log10 CFU/mL, resulting in a 10.86 log reduction in bacteria cells. The

results obtained from ANOVA (Table 3.14) and Tukey-Kramer’s post hoc analysis

revealed that there were significant differences among all the group means except when

the mean of MOI 101 at 6 hrs was compared with the means of MOI 102 at 24 hrs, MOI

103 at 24 hrs, MOI 104 at 24 hrs and MOI 105 at 24 hrs, when the mean of MOI 102 at 6

hrs was compared with the means of MOI 102 at 24 hrs, when the mean of MOI 103 at 6

hrs was compared with the means of MOI 104 at 6 hrs and MOI 105 at 6 hrs, when the

mean of MOI 104 at 6 hrs was compared with the means of MOI 105 at 6 hrs, when the

mean MOI 102 at 24 hrs was compared with the means of MOI 104 at 24 hrs, when the

mean MOI 103 at 24 hrs was compared with the means of MOI 104 at 24 hrs and MOI

105 at 24 hrs, and when the mean MOI 104 at 24 hrs was compared with the means of

MOI 105 at 24 hrs, respectively.

89


an in-vitro treatment with Cocktail.


variance

Standard

deviation

MOI 101 (6 hrs) 26.691 8.897 0.000648 0.0147

MOI 101 (24 hrs) 39.284 13.095 0.00002.84 0.0031

MOI 102 (6 hrs) 22.766 7.589 0.26 0.0156

MOI 102 (24 hrs) 25.023 8.341 0.148 0.2222

MOI 103 (6 hrs) 17.608 5.869 0.0000345 0.0034

MOI 103 (24 hrs) 28.279 9.426 0.244 0.285

MOI 104 (6 hrs) 15.241 5.08 0.0000303 0.0032

MOI 104 (24 hrs) 27.447 9.149 0.175 0.2415

MOI 105 (6 hrs) 17.04 5.68 0.002 0.0234

MOI 105 (24 hrs) 29.169 9.723 0.00200 0.0245

Control (6 hrs) 26.691 8.897 0.000648 0.0147

Control (24 hrs) 39.284 13.095 0.0000284 0.0031

sample size mean =3; degrees of freedom =11,24; sum of squares (636.020, 1.2.223) and mean square (57.820, 0.093), F statistic (624.098)

90

Figure 3.16: Survival of P. aeruginosa PAO1 cells after an in-vitro treatment with

FCPA1 at different MOI ratios. Bacteria recovery was performed at 6 hrs and 24

hrs post-incubation. Error bars represent standard error of the mean.

6 hr

s

24 h

rs0

5

10

15

20

25

Time

Lo

g10 C

FU

/mL

Control

MOI 101

MOI 102

MOI 103

MOI 104

MOI 105

91




6 hr

s

24 h

rs0

5

10

15

20

25

Time

Lo

g10 C

FU

/mL

Control

MOI 101

MOI 102

MOI 103

MOI 104

MOI 105

92




6 hr

s

24 h

rs

0

5

10

15

20

25

Time

Lo

g10 C

FU

/mL

Control

MOI 101

MOI 102

MOI 103

MOI 104

MOI 105

93




6 hr

s

24 h

rs0

5

10

15

20

25

Time

Lo

g10 C

FU

/mL

Control

MOI 101

MOI 102

MOI 103

MOI 104

MOI 105

94




6 hr

s

24 h

rs

0

5

10

15

20

25

Time

Lo

g10 C

FU

/mL

Control

MOI 101

MOI 102

MOI 103

MOI 104

MOI 105

95




6 hr

s

24 h

rs0

5

10

15

20

25

Time

Lo

g10 C

FU

/mL

Control

MOI 101

MOI 102

MOI 103

MOI 104

MOI 105

96


WCSS4PA at different MOI ratios. Bacteria recovery was performed at 6 hrs and

24 hrs post-incubation. Error bars represent standard error of the mean.

6 hrs

24 h

rs

0

5

10

15

20

25

Time

Lo

g10 C

FU

/mL

Control

MOI 101

MOI 102

MOI 103

MOI 104

MOI 105

97


WCSS5PA at different MOI ratios. Bacteria recovery was performed at 6 hrs and

24 hrs post-incubation. Error bars represent standard error of the mean.

6 hr

s

24 h

rs

0

5

10

15

20

25

Time

Lo

g10 C

FU

/mL

Control

MOI 101

MOI 102

MOI 103

MOI 104

MOI 105

98


Cocktail at different MOI ratios. Bacteria recovery was performed at 6 hrs and 24


3.2.2.3 Survival of bacterial cells on sand treated samples

The ability of phage isolates to inhibit or eradicate P. aeruginosa PAO1 bacterial cells in

experimentally contaminated sand samples were assessed at 0 hr, 6 hrs, 24 hrs and 48

hrs post-treatment. The surviving bacterial cells following treatment with phages

(FCPA3, WCSS4PA and Cocktail) were expressed as log10 CFU/mL and were compared

with those of uninfected control as shown in Figure 3.25. The highest bacteria log

reduction was seen exactly 6 hrs after treatment with phages (FCPA3, WCSS4PA and

Cocktail). Recovered bacterial cells were 7.04 log10 CFU/mL, 6.61 log10 CFU/mL and 5.55

log10 CFU/mL respectively.

6 hr

s

24 h

rs0

5

10

15

20

25

Time

Lo

g10 C

FU

/mL

Control

MOI 101

MOI 102

MOI 103

MOI 104

MOI 105

99

This resulted in 4.6, 5.03 and 6.09 logs reduction respectively when compared with the

uninfected control (11.64 log10 CFU/mL). The highest bacterial log reduction was 6.09

log10 CFU/mL obtained from sand samples treated with a phage cocktail (Cocktail). At

24 hrs and 48 hrs post-treatment, surviving bacterial counts showed no significant

difference when compared with that of the untreated control. The ANOVA data

presented in Table 3.15-3.18 and post hoc analysis using the Tukey-Kramer’s procedure

(α=0.05) further revealed that at 6 hrs post-treatment, the mean of WCSS4PA (M=

6.608; SD= 0.182), and cocktail (M= 5.554; SD= 0.205) were significantly different from

the mean of FCPA3 (M= 7.043; SD= 0.281), and untreated control (M= 11.645; SD=

0.643).

Table 3.15: Analysis of variance (ANOVA) results for surviving bacterial cells recovered

from sand treated samples at 0 hr using different phage samples.


variance

Standard

deviation

⏀FCPA3 30.5747 10.19 0.019 0.079

⏀WCSS4PA 29.0346 9.678 0.269 0.3

⏀Phage

Cocktail

29.5304 9.844 0.177 0.243

⏀Control 29.3091 9.77 0.303 0.318

sample size mean =3; degrees of freedom =3, 8; sum of squares = 0.4529, 1.5357; mean square =0.1510, 0.1920 and F statistic =0.7864.

100


from sand treated samples at 6 hrs using different phage samples.


variance

Standard

deviation

⏀FCPA3 21.13 7.043 0.079 0.162

⏀WCSS4PA 19.82 6.608 0.033 0.104

⏀Phage Cocktail 16.66 5.554 0.042 0.118

⏀Control 34.93 11.64 0.414 0.371





variance

Standard

deviation

⏀FCPA3 24.1513 8.05 0.0001 0.006

⏀WCSS4PA 23.1924 7.731 0.172 0.24

⏀Phage Cocktail 22.2131 7.404 0.142 0.218

⏀Control 21.5348 7.178 0.183 0.247


101



Group Sum Mean Sample variance

Standard deviation

⏀FCPA3 23.1346 7.712 0.166 0.235

⏀WCSS4PA 21.1268 7.042 0.02 0.081 ⏀Phage Cocktail 23.2816 7.761 0.343 0.338

⏀Control 22.666 7.555 0.094 0.177

sample size mean =3; degrees of freedom =3, 8; sum of squares = 0.9719, 1.2446; mean square =0.3240. 0.1556 and F statistic =2.0834.

Figure 3.25: Survival of P. aeruginosa PAO1 cells on sand samples after treatment

with bacteriophages. Error bars represent standard error of the mean.

0 hrs

6 hrs

24 h

rs

48 h

rs0

5

10

15

Time

Lo

g10 C

FU

/mL

Control (uninfected)

ΦWCSS5PA

ΦFCPA3

ΦCocktail

102

3.3 Discussion

This study reports on isolation, host range analysis, and assessment of in-vitro

bacteriolytic activities of lytic phages isolated from Sarawak limestone caves (FCNR and

WCNR). Investigative studies designed to test the potentials of selected P. aeruginosa

phages to decontaminate sand samples are also reported. The study of viral diversity

from limestone caves is very limited despite the abundance of caves all over the world.

Most studies on phage isolation have utilized sewage as an optimal resource (Lobocka,

et al., 2014). Although caves are considered as an extreme environment to life due to

lack of organic carbon input from photosynthesis and absence of light and various

physicochemical micro-gradients, studies have reported on the presence of vast

microbial communities with unexpected biodiversity (Northup and Lavoie, 2001,

Tomczyk-Żak and U., 2015). Nevertheless, caves have been reported to harbor

microorganisms that display variable enzymatic and antimicrobial activities which are

different from those observed in other extreme environments. This further explains that

caves are rich reservoir of potential antimicrobials and suggests that investigating cave

microbiota opens new frontiers for drug discovery (Ghosh, et al., 2016, Lamprinou, et

al., 2015, Nimaichand, et al., 2015).

It is crucial to understand bacteriophages and their interactions with bacterial hosts as

this provides insights into the molecular biology and may result to an improved

understanding of treatment methods against bacterial infection (Bolger-Munro, et al.,

2013). The study of bacterial growth kinetics is specifically significant in determining the

number of bacterial cells present in the liquid medium (Mohammed, 2013). Previous

studies have reported that metabolic state of bacterial cells influences their

susceptibility to phage infection, phage latent period and burst size and hence the

success of new phage isolation attempts (Weinbauer, 2004). In most cases,

exponentially growing cells are the most susceptible and can support the fastest and the

most efficient phage production (Wommack and Colwell, 2000). It has been reported

that Gram-negative bacteria such as E. coli tend to minimize their overall rate of protein

synthesis while upgrading the expression of certain groups of proteins recognized as

stationary phage specific . These proteins safeguard cells from oxidative damage and

allow cells to stay viable during stationary phase (Braun, et al., 2006). Furthermore, it

has also been urged that due to nutrient starvation, stationary phase cells decrease in

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size and become more spherical rather than rod-like shape thus, providing a very small

surface area to enable proper binding of the phage (Abedon, 1990, Ingraham, et al.,

1983, Lange and Hengge-Aronis, 1991). Since stationary phase bacterial cells are

smaller, the number of collision between phage and bacterial cells is significantly

lowered because the cells tend to absorb less phage resulting to the production of

insufficient progeny (Braun, et al., 2006). Nevertheless, factors such as cell metabolic

poisons or free phage poisons released by cells in the media can reduce bacterial

susceptibility to infection and phage progeny production (Abedon, 1990). Therefore,

due to dramatic effects associated with the use of stationary phase bacteria during

phage therapy studies, it's recommended to use log phase cultures for optimal viral

progeny production. Thus, the growth profiles of the selected phage host i.e. V.

parahaemolyticus, S. aureus, K. pneumoniae, E. coli, P. aeruginosa and S. pneumoniae

were performed as shown in Figure 3.3 and Table 3.2, and mid-exponential phase

bacterial cultures were employed during phage isolation and decontamination

experiments.

About 33 lytic bacteriophages were isolated from FCNR and WCNR soil samples

following an enrichment technique. Among these isolates, were two multiphages

designated as WCSS4PA and WCSS5PA. Majority of these phage isolates were

obtained from FCNR soil samples (79%). Despite phage abundance in the environment,

less than 1% of phage species present in the environmental samples can be detected by

plaque assay with cultivable hosts (Ashelford, et al., 2003, Williamson, et al., 2003). The

possibility to visualize bacterial host lysis due to phage attack, in the form of plaques on

the lawns of bacterial cells enables detection and isolation of most of the environmental

phages against cultivable hosts, if only they are present (Kropinski, et al., 2009,

Mazzocco, et al., 2009). Numerous ways have been proposed to enhance phage plaque

visibility on a bacterial lawn. A feasible method may involve the use of compounds such

as 2,3,5-triphenyltetrazolium chloride (TTC), ferric ammonium citrate and sodium

thiosulfate. These compounds facilitate visualization or detection of plaques that are

too small or too turbid to be easily seen (McLaughlin and Balaa, 2006). Sublethal doses

of antibiotics such as 2.5–3.5 μg/mL of ampicillin can be incorporated into the top agar

to improve phage plaque contrast. This method enables the formation of plaques with

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increased diameter and visibility in standard conditions such as in the case of E. coli

phages (Łoś, et al., 2008). Phage isolation reported in this study utilized pour plate

method in which 1% TTC (2,3,5-triphenyl tetrazolium chloride) solution was added. This

enabled improved resolution of phage plaques as seen in Figure 3.4. The use of TTC

(2,3,5-triphenyl tetrazolium chloride) to improve phage plaque resolution was first

reported by Pattee (1966) who worked on Phage 83, which was employed in the genetic

analysis of Staphylococcus aureus. In his study, phage isolation was carried out using the

agar-layer technique and the plates were incubated at 37oC for 8 hrs or until the plaques

were sufficiently developed to be scored. The assay plates containing fully developed

plaques were then flooded with 10 mL of TSB containing 0.1% TTC (2,3,5-triphenyl

tetrazolium chloride) solution and plates were incubated for 20 minutes at 37oC. Each

plaque appeared as sharp and clear area against the intense red background produced

by the reduction of TTC to the insoluble formazan by the indicator cells.

Elimination of temperate phages from the collection of newly obtained isolates is one

of the most important early tasks in the selection of phages for therapeutic use, as

temperate phages are estimated to comprise about 50% of environmental isolates

(Ackermann, 2005). A commonly accepted criterion to distinguish obligately lytic and

temperate phages is the ability to form clear plaques by the former (Guttman, et al.,

2005). In most cases, plaque clarity is a good indicator of phage propagation strategy.

However, the preliminary classification of a phage as obligatory lytic or temperate based

on plaque observation should be treated with caution. For instance, Lobocka, et al.

(2014) postulated that obligately lytic phages that form clear plaques can be more or

less overgrown by resistant or infection escaping bacteria on cell layers of certain other

strains and make the impression of being turbid. Following isolation, distinctive plaques

were amplified in BHI broth containing respective bacterial host. The contents were

filter sterilized and the titer of lysates was determined as seen in Figure 3.5 and Table

3.3.

Bacteriophage host-range analysis is a crucial factor in assessing phage diversity in the

environment (Malki, et al., 2015). Broad host range phages are rendered suitable in

phage therapy or phage biocontrol applications. In phage therapy, a broad host range

phage that can kill multiple species of bacteria is equivalent to a broad spectrum

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antibiotic (Ross, et al., 2016). Thus, a smaller number of broad host range is more useful

than many narrow host range phages. Another key advantage of host range analysis in

phage therapy studies is the specificity of the phage-host range which spares non-

pathogenic bacteria from being killed during the treatment. Contrarily, this same

specificity may limit the ability and usage of a specific phage to a small set of potential

pathogens requiring more precise diagnosis (Mapes, et al., 2016, Nilsson, 2014, Ross, et

al., 2016). In the present study, P. aeruginosa and V. parahaemolyticus infecting

bacteriophages exhibited the broadest host range among all the isolates. Interestingly,

this broad lysis spectrum extended beyond a single bacteria phylum. For instance, V.

parahaemolyticus phages designated as FCVP2, FCVP3 and FCVP4 were capable of

infecting S. aureus bacteria which belongs to a completely different phylum i.e.

Eubacteria. Another important feature observed during the host range analysis was the

ability of some phage isolates to display trans-subdomain infectivity between gram

positive and gram-negative bacterial hosts. For instance, S. pneumoniae phage isolates

(FCSP1, FCSP2 and FCSP3) were able to infect and lyse P. aeruginosa and E. coli

bacteria. This has been previously argued that such results might be caused not by

the added bacteriophage but by the temperate bacteriophage which was originally

in the host bacteria tested (Khan, et al., 2002). Previous studies have assigned

classification as a “generalist” when a phage demonstrates capacity to infect more than

one species of a bacterial genus, while some restricting the definition further to include

strains of a specific species (Bono, et al., 2013, Czajkowski, et al., 2014, Merabishvili, et

al., 2014). Similar findings have been reported by Malki, et al. (2015) where

bacteriophages isolated from lake Michigan was capable of infecting several bacteria

phyla and it was proposed that such a broad-host-range was likely related to the

oligotrophic nature of the lake and the competitive benefit this characteristic may have

contributed to phages in nature. These results imply that bacteriophage host-range is

not always genera-restricted, and the oligotrophic environment of Fairy cave might have

in one way or another contributed to the broad lytic spectrum exhibited by V.

parahaemolyticus infecting phage isolates. This scenario has also been highlighted by

(Nilsson, 2014) where it was urged that phages with the ability to use more universal

surface receptors for adsorption, exhibit broader host range and are usually found in

environments with poor nutrients.

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The host range assay results also revealed that a significant number of the phage

isolates failed to infect some bacteria strains such as V. parahaemolyticus, S.

pneumoniae, and S. typhimurium. The resistance of bacteria on a bacteriophage may

be due to several mechanisms. These include modification of phage attachment or

adsorption sites (receptors), restriction-modification systems and CRISPER-Cas

mechanisms or Clustered regularly interspaced short palindromic repeats (CRISPRs) and

the CRISPR-associated (cas) genes (Seed, 2015, Sharma, et al., 2017). Alteration of phage

adsorption sites (receptors) appears to be the most common mechanism by which

bacteria evade phage infection and become resistant to phage (Bohannan and Lenski,

2000, Hyman and Abedon, 2010). Additionally, bacteria may synthesize

exopolysaccharide (EPS) or masking proteins e.g. protein A of S. aureus to mask the

phage receptor and thus conferring resistance to phage attack. To encounter the effect

of EPS or masking proteins, bacteriophages may conquer the barrier by cleaving the EPS

layer using polysaccharide lyase or a polysaccharide hydrolase (Labrie, et al., 2010,

Örmälä and Jalasvuori, 2013). Bacteria may also possess restriction-modification

systems that defend hosts from exogenous DNA. In this case, bacteria system can

recognize and modify the phage DNA. In most cases, the phage DNA is cleaved by

restriction endonucleases upon entering the bacterial cell through the restriction-

modification system and thus protecting bacterial cell from phage DNA attack (Pleška,

et al., 2016, Sharma, et al., 2017). To encounter this, phages may adopt an anti-

restriction strategy to avoid recognition by endonuclease enzyme. For example, T4

phage evades restriction endonuclease attack because it possesses

hydroxymethylcytosine (HMC) instead of cytosine. Even so, some bacteria may modify

their system to recognize hydroxymethylcytosine (HMC) and destroy phage DNA (Bickle

and Kruger, 1993, Borgaro and Zhu, 2013). Interestingly, anti-restriction strategy in

Staphylococcus phage K possesses 5′ GATC-3′ cleavage site which confers DNA

protection from restriction endonucleases (Bryson, et al., 2015, Tock and Dryden,

2005).

CRISPER-Cas mechanism presents a novel strategy by which prokaryotes acquire

resistance against viruses (Sharma, et al., 2017). The main function of CRISPER-Cas is

to provide immunity against foreign DNA such as phage genomic DNA and plasmid

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DNA (Gasiunas, et al., 2014). Clustered regularly interspaced short palindromic repeats

(CRISPR) loci are arrays of short repeats separated by equally short “spacer” sequences

(Biswas, et al., 2016). Along with the CRISPR-associated (cas) genes, encode an

adaptive immune system of archaea and bacteria that protects the cell against viral

infection (Barrangou, et al., 2007, Marraffini and Sontheimer, 2008). This system

functions by inserting a short piece of an infecting viral genome as a spacer in the

CRISPR array. The spacer sequence is transcribed and processed to generate a small

antisense RNA (the CRISPR RNA or crRNA) that is used as a guide for the recognition

and destruction of the invader in subsequent infections (Brouns, et al., 2008, Carte, et

al., 2008, Deltcheva, et al., 2011). Thus, spacer acquisition immunizes the bacterium

and its progeny against the virus from which it was taken. Barrangou, et al. (2007) has

highlighted an example of antiphage activity of CRISPER-Cas mechanism in

Streptococcus thermophiles where exposure to virulent phage gave rise to the phage-

resistant mutants due to insertion of additional 30 bp spacer resembling protospacer

of infecting phage. The event of acquiring immunity against phage can be explained

briefly in the following steps like adaptation or spacer attainment, transcription of

acquired spacer (small CRISPER RNAs (crRNAs), on recurrent phage attack this crRNAs

form a complex with Cas protein), and immunity against phage (crRNAs-Cas complex

direct nuclease to trace and chop the invading phage DNA (Marraffini, 2015).

Another key observation seen in this study was the inability of some phage isolates (e.g.

WCVP3, WCVP4, WCVP5, FCSA4, FCSA6, FCSP4 and FCSP5) to lyse the

bacteria hosts from which they were first isolated during the host range analysis

experiments. Although plaque formation was not observed, this does not necessarily

mean that infection did not occur. It is possible that the phage-infected the host

bacteria and resulted in a lysogenic relationship. But regardless of whether

bacteriophage and the host started lysogenic interactions or not, the observed

phenomena certainly indicate that infectivity of the bacteriophages changed or the

resistance of host bacteria to the bacteriophages varied during the course of the study.

The possibility that these bacteriophages could not re-infect their bacterial hosts might

be due to the occurrence of spontaneous single mutation in the host bacteria which

caused it to gain resistance to the phage (Khan, et al., 2002). According to Ross, et al.

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(2016), bacteriophage host range is not a fixed property of each species of

bacteriophage. Rather, it is one that can evolve over time and can show unexpected

plasticity. Modifying procedures and growth conditions can favor the isolation of novel

phages with broader host ranges. By mixing a few well differentiated phage host

bacterial strains in a cultivation flask which is inoculated with an original source of

phages (typically a filtrate of water, sewage, soil suspension etc.), usually suffices for

the isolation of polyvalent phages assuming they are present in the tested sample

(Carvalho, et al., 2010, Lobocka, et al., 2014, Van Twest and Kropinski, 2009). For

instance, Mapes, et al. (2016) developed a host range expansion protocol that aimed at

broadening the host range of P. aeruginosa-specific bacteriophages. Their study

reported culturing a mixture of four phages with a mix of 16 different host strains and

isolated individual phage strains by plaque isolation after multiple passages of phage

mix onto the fresh mix. Over the course of 30 cycles, host range was expanded following

spot test assay on both the 16 host strains and an additional 10 P. aeruginosa strains.

Assessment of phage in-vitro bacteriolytic activity and decontamination of sand samples

utilized P. aeruginosa crude phage lysates, selected based on their broad host range,

high titer and virulence. It is recommended that bacteriophage preparations especially

those targeting Gram-negative bacteria, be purified in order to remove endotoxins or

lipopolysaccharides, cell debris and other contaminating substances prior to

applications (Van Belleghem, et al., 2017). However, the degree of purification of phage

preparation is largely a function of the type of application the phage will be used for.

For instance, phages which are to be administered in a medical (human or animal)

setting must be thoroughly purified to remove bacterial endotoxins as these elicit a wide

variety of pathophysiological effects in the body due to their immunogenic, pro-

inflammatory and pyrogenic effects (Aderem and Ulevitch, 2000, Bonilla, et al., 2016).

Excessive or systemic exposure to endotoxins may prompt a systemic inflammatory

reaction associated with multiple pathophysiological effects such as endotoxin shock,

tissue injury and death (Anspach, 2001, Erridge, et al., 2002, Ogikubo, et al., 2004). Thus,

it is important to remove endotoxins from phage preparations as these may affect

efficacy and safety of the administration during phage therapy (Van Belleghem, et al.,

2017). On the other hand, extensive purification of phage preparations is of less

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importance especially if the phage preparations are to be used for disinfection or as

foliar sprays in plant agriculture, as the later will be washed away well before any animal

or human consumes the plant (Balogh, et al., 2010, Gutiérrez, et al., 2016). In such cases,

simply removing the live bacterial cells along with larger cell debris via filtration may be

sufficient (Gill and Hyman, 2010).

Phage bacteriolytic activities were investigated in an in-vitro co-culture assay as

presented in Figure 3.7 to 3.15. Five MOI ratios were tested to optimize the phage dose

required to inhibit or completely eradicate the bacteria, and also to provide a basis on

which to select the most appropriate phages for subsequent sand decontamination

experiments. High MOI dose was intended to examine whether P. aeruginosa PAO1

bacterial cells could be reduced by passive inundation (Payne and Jansen, 2001). Passive

inundation refers to a scenario where numbers of bacterial cells are depleted by

attachment of overwhelming numbers of phage but without productive replication of

the phage (Carrillo, et al., 2005). The use of lower phage doses was anticipated to initiate

active proliferation of the phage and bacteria, with the phage eventually overwhelming

their host (Carrillo, et al., 2005). Additionally, the use of higher MOI dose was considered

as an appropriate strategy for inhibiting or eliminating P. aeruginosa PAO1 bacterial cells

with the purpose of minimizing the likelihood of acquired host resistance to phages over

time (Carrillo, et al., 2005). Generally, the growth of P. aeruginosa PAO1 was inactivated

when co-cultured with phage in a concentration-dependent manner, with OD values

declining more quickly at higher MOI (104 and 105) than at lower MOI (103, 102 or 101).

The uninfected P. aeruginosa PAO1 cells showed steady growth, with OD600 values

increasing at different time points as expected. The highest bacterial inactivation at the

end of the 6 hrs of incubation was seen in cultures infected with bacteriophages FCPA2,

FCPA3, WCSS4PA and Cocktail. For instance, FCPA2 at MOI 105, FCPA3 at MOI 105,

WCSS4PA at MOI 105 and Cocktail at MOI 104 managed to decrease the absorbance

(OD600) values of the infected cultures to 0.702, 0.200, 0.319 and 0.288 respectively

when compared with the uninfected control cultures. The absorbance (OD600) readings

of uninfected control cultures for phages FCPA2, FCPA3, WCSS4PA and Cocktail

were 1.370, 1.370, 1.533 and 1.557 respectively. The phage bacteriolytic activity curves

of the named phages followed a very similar pattern where an initial rise in turbidity was

110

observed which was then followed by a decrease in turbidity which stabilized to the end

of the incubation period. This result implies that the phages continued to grow during

the initial infection and lysis occurred as the infection proceeded. However, cell lysis was

observed much earlier in cultures infected with phage at high MOI ratio than at low MOI

ratio. For example, in bacterial cultures infected with phage FCPA3 at MOI of 105, cell

lysis began 30 minutes post-infection whereas at lower MOI ratios (104, 103,102,101) cell

lysis began 4 hrs post-infection. This observation could be due to increased stress on the

host cells which resulted in bacterial lysis (Brewster, et al., 2012). Another possible

reason to this could be each phage causes lysis at different rates with phages FCPA3,

WCSS4PA and Cocktail being the fastest when compared with the rest phages. In

addition, this phenomenon could be due to due to the speed at which the phages

attaches to the host cell receptors, enters the cell, replicates, assembles or lyses the cell

(Young, et al., 2003). The phage bacteriolytic activities of FCPA4, FCPA5 FCPA6,

WCSS5PA were marked by a slow decrease in absorbance (OD600) which lasted for a

period of time, followed by a slow rise in absorbance which lasted to the end of 6 hrs

incubation. The increase in OD may be due to the presence of phage-resistant bacterial

cells (Tan, et al., 2014).

Development of phage resistant bacteria is often correlated with a concomitant loss of

virulence (Laanto, et al., 2012). This arises mainly due to the cell surface components

such as lipopolysaccharides (LPS) and proteins that act as receptors for phage

adsorption, which can also act as virulence factors. Furthermore, mutations occurring

on these receptors which causes bacteriophage resistance results in a reduction in

pathogenicity (Silva, et al., 2014). Therefore, bacteria regrowth after phage therapy will

result too few or no big consequences in terms of virulence (Capparelli, et al., 2010,

Filippov, et al., 2011, Wagner and Waldor, 2002). It is recommended to carry out further

studies to detect mutations in the outer bacterial molecules of resistant bacteria

following phage therapy, as these can act as phage receptors and possibly at the same

time as virulence factors (Silva, et al., 2014). Bacteriophages have several features that

make them potential therapeutic agents against infectious bacteria. One of these

features is the highly specific and effective lysis of the targeted pathogenic bacteria

(Zhang, et al., 2015). Generally, the in-vitro bacteriolytic activity results obtained in this

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study demonstrated that phage isolates possessed strong bacteriolytic activity which is

important for use in phage therapy. The highest bacteriolytic activity was observed in P.

aeruginosa bacterial cultures infected with phages FCPA3 (MOI 105) and Cocktail

(MOI 104). Various studies have revealed that two or more phages with different host

ranges in a single suspension (a phage cocktail) acts more effective than a use of a single

phage alone (Chan, et al., 2013, Gu, et al., 2012, Jaiswal, et al., 2013). The use of

multiphage therapy has also been reported to be more efficient in reducing the bacterial

density when compared with monophage therapy. For instance, Hall, et al. (2012)

investigated the effect of using one, two or four phages either sequentially or

simultaneously against P. aeruginosa PAO1 planktonic cultures and the results showed

that simultaneous application of phages was consistently equal or superior to the

sequential application with respect to efficacy. This study reports similar observation

where a multiphage designated as WCSS4PA showed efficient bacterial reduction

when compared with monophage isolates. One possible limitation of the study

conducted by Hall, et al. (2012) was the use of optical density measurements to estimate

bacterial population density. It was claimed that the relationship between population

size and optical density can be altered by the evolution of phage resistance. For

example, resistant genotypes may overproduce alginate or extracellular polymeric

substances (EPS) that results in OD measurements inflation.

Viable P. aeruginosa PAO1 cells that survived in-vitro treatment with bacteriophages

were enumerated at 6 hrs and 24 hrs post-infection as shown in Figure 3.16 to 3.24. The

number of surviving bacteria cells assessed 6 hrs post-infection were compared with

bacterial counts assessed prior to phage infection (10.89 log10 CFU/mL) and also at 6 hrs

post-infection (16.90 log10 CFU/mL). Significant (p0.05) log10 CFU/mL reductions were

observed when surviving bacteria cells harvested 6 hrs post-infection were compared

with those of untreated control, also harvested at 6 hrs post-infection. All the phages

showed over 99.99 % reduction in bacteria following the treatment. The highest

bacterial log10 CFU/mL reduction was 11.82 equivalent to 100% reduction in bacteria

observed in cultures treated with phage cocktail (Cocktail) at MOI of 104. Moreover,

bacteriophages designated as FCPA3, FCPA5, FCPA6 and WCSS4PA also showed

higher bacterial log10 CFU/mL reductions of 9.2, 9.4, 9.6 and 10.3 respectively,

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equivalent to over 99.99% reduction in bacteria. The lowest bacterial log10 CFU/mL

reduction was observed in cultures infected with phage WCSS5PA (4.8 log10 CFU/mL).

However, when surviving bacteria cells harvested 6 hrs post-infection were compared

with bacterial counts harvested prior to phage infection, not all phages were effective

at reducing P. aeruginosa PAO1 bacterial cells. The highest bacterial log10 CFU/mL

reduction was 5.8 equivalent to 99.99% bacterial reduction which was observed in

cultures treated with phage cocktail (Cocktail). Following this was a multiphage

(WCSS4PA) which showed a log10 CFU/mL reduction of 4.26 equivalent to 99.99%

bacterial reduction. Bacteriophage WCSS5PA did not show any reduction in bacterial

cells but instead the bacterial cells rebounded and surpassed those of the untreated

control. Phage FCPA1 showed the lowest bacterial log10 CFU/mL reduction (0.16)

equivalent to 30 % reduction in bacterial cells.

Another comparison was performed between surviving bacterial cells harvested 24 hrs

post-infection and that of untreated control harvested before any infection was initiated

(10.89 log10 CFU/mL). The results showed significant bacterial log reductions in cultures

treated with phages WCSS4PA and Cocktail only. Phages WCSS4PA and Cocktail

showed a bacterial log10 CFU/mL reduction of 2.17 and 1.74 equivalent to 99.33% and

98.18% bacterial load reduction respectively. Again, the rest bacteriophages did not

show any reductions in bacterial cells but instead, the cells rebounded and surpassed

those of the untreated control. Surviving bacteria cells harvested 24 hrs post-infection

were then compared with uninfected control (19.17 log10 CFU/mL) harvested at the

same time. The results showed significant bacterial log reduction amongst all the tested

phages. All the phages achieved greater than 99.99% reduction in bacterial load when

compared with uninfected control. The highest bacterial log reduction was observed in

bacteriophages WCSS4PA (MOI 104) and Cocktail (MOI 102). These phages showed a

bacterial log10 CFU/mL reduction of 10.5 and 10.86 respectively, equivalent to 100%

reduction in bacterial cells. Nevertheless, bacteriophage FCPA3 (MOI 105) showed the

highest bacterial log10 CFU/mL reduction of 8.3 equivalent to over 99.99% reduction in

bacterial cells amongst all the tested monophages. Surprisingly, multiphage WCSS5PA

showed higher bacterial log10 CFU/mL reduction (7.0) equivalent to over 99.99%

bacterial reduction after 24 hrs post-infection. Previous results indicated that, phage

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WCSS5PA achieved the lowest bacterial log10 CFU/mL reduction (4.8) among all the

tested phages at 6 hrs post-infection when compared with uninfected control assessed

at the same time. Additionally, phage WCSS5PA showed no reduction in bacterial cells

following assessment at 6 hrs and 24 hrs post-infection when compared with uninfected

control assessed before the infection was initiated. Instead, the cells rebounded and

surpassed those of uninfected control.

Treatment of experimentally contaminated sand samples utilized crude phage lysates

obtained from phages FCPA3, WCSS4PA and Cocktail, owing to their efficient

bacteriolytic activities. Surviving bacterial cells following phage treatment were

enumerated at 0 hr, 6 hrs, 24 hrs and 48 hrs as shown in Figure 3.25. When surviving

bacterial cells at 0 hrs (assessed just after phage addition and before incubation) were

compared with viable P. aeruginosa PAO1 counts obtained from untreated sand

samples (9.77 log10 CFU/mL), harvested at the same time, no significant bacterial

reduction was observed as expected. However, a bacterial log10 CFU/mL reduction of

0.09 equivalent to 19.18% reduction in bacteria cells was observed in sand samples

sprayed with phage WCSS4PA. Surviving bacterial cells assessed at 6 hrs, 24 hrs, and

48 hrs post-treatment were compared with bacterial counts obtained from untreated

sand samples (9.77 log10 CFU/mL) assessed prior to phage treatment. The results

showed significant bacterial log10 CFU/mL reduction in all the three phages. However,

the highest bacterial log10 CFU/mL reduction was 4.2 equivalent to 99.99% bacterial load

reduction observed in sand samples sprayed with a phage cocktail (Cocktail). Phages

FCPA3 and WCSS4PA showed bacterial log10 CFU/mL reductions of 2.73 and 3.16

equivalent to 99.8% and 99.9% bacterial load reduction. Surviving bacterial cells

assessed 24 hrs post-treatment also showed the highest bacterial log10 CFU/mL

reduction in phage cocktail (Cocktail), followed by WCSS4PA and FCPA3. Bacterial

log10 CFU/mL reductions observed in these samples were 2.36, 2.04 and 1.72 equivalent

to 99.57%, 99.09% and 98.09% bacterial load reductions respectively. At 48 hrs post-

treatment, the highest bacterial log10 CFU/mL reduction was 2.73 equivalent to 99.81%

bacterial load reduction observed in WCSS4PA followed by 2.05 equivalent to 99.13%

and 2.01 equivalent to 99.2% observed in FCPA3 and Cocktail respectively. The

number of surviving bacteria cells assessed 6 hrs post-treatment were compared with

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those of untreated sand samples (11.64 log10 CFU/mL) assessed at the same time. All the

three phages i.e. FCPA3, WCSS4PA and Cocktail achieved high bacterial log10

CFU/mL reductions of 4.60, 5.03 and 6.09 respectively, equivalent to over 99.99%

reduction in bacterial cells.

It was also observed that P. aeruginosa PAO1 cells in the untreated control sand samples

showed favorable growth at 6 hrs post-treatment despite limited nutrient supply.

Various studies have reported that P. aeruginosa exhibits extensive metabolic diversity

which enables it to thrive in different ecological niches such as soils, plants, water and

animals (LaBauve and Wargo, 2012, Orlandi, et al., 2015). It is this metabolic flexibility

that allows P. aeruginosa to succeed as an opportunistic pathogen causing both

community-acquired and hospital-acquired infections which can be life-threatening

(LaBauve and Wargo, 2012). Studies have reported that beach sands can serve as a

vehicle for exposure of humans to pathogens, resulting in increased health risks. For

instance, analysis of Israel beaches revealed various levels of P. aeruginosa, with higher

counts found on beach sands than in the seawater samples (Ghinsberg, et al., 1994). In

an interesting report by Velonakis, et al. (2014), factors affecting the survival of

pathogenic bacteria such as P. aeruginosa in beach sands were examined. The results

indicated greater survival and proliferation of P. aeruginosa along with S. aureus in

sterile beach sands than seawater. In the current study, it was possible for P. aeruginosa

cells to grow in sand samples at 6 hrs post-incubation because the cells were still at a

mid-exponential phase when introduced into the sand samples. At this phase bacteria,

cells divide rapidly and double in number at regular intervals. The growth kinetics result

presented in Table 3.2 showed that P. aeruginosa had a rapid growth with a very short

doubling time (td) of 0.249. In addition, the prospect of P. aeruginosa cells to replicate

in sand samples at 6 hrs post-incubation could be due to the presence of nutrients

supplied by Brain-heart infusion (BHI) media used to grow the bacteria cells.

When the number of surviving, bacterial cells assessed at 24 hrs post-treatment were

compared with those obtained from untreated sand samples assessed at the same time

(7.18 log10 CFU/mL), no reduction in bacterial cells was observed in all the three phages.

In fact, all the three phages showed an increase in bacterial cells which surpassed that

of the untreated control. Similarly, when the number of surviving bacterial cells assessed

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at 48 hrs post-treatment was compared with those of untreated sand samples assessed

at the same time (7.56 log10 CFU/mL), no significant reduction in bacterial cells was

observed in all the three phages. However, bacteriophage WCSS4PA showed bacterial

log10 CFU/mL reduction of 0.51 equivalent to 69.36% reduction in bacterial cells. Sand

samples treated with bacteriophages FCPA3 and Cocktail showed an increase in

bacteria cells which surpassed those of the untreated control. This study demonstrates

the usefulness of virulent bacteriophages isolated from Sarawak limestone caves in

inactivating the growth of P. aeruginosa PAO1 cells. The effectiveness of the phages

against P. aeruginosa PAO1 bacterial target varied significantly when compared with the

control and among each other. The results attained in this study showed that effective

phage infection and subsequent destruction of the host cells is strongly determined by

multiplicity of infection (MOI) ratio. The multiplicity of infection (MOI) refers to the ratio

of phages to host cells (Bigwood, et al., 2009). Higher MOI ratio resulted in significant

growth suppression of the bacterial host. For example, all phages showed better

inactivation when MOI ratio of at least 103 was used. This agrees with the findings

reported in other analyses that showed that application of higher MOI ratio resulted in

higher bacterial inactivation. For instance, O'flynn, et al. (2004) achieved efficient

inactivation of Escherichia coli O157: H7 on beef using MOI of 106 in which 2 x 108 PFU

of phages were applied to pieces of meat inoculated with 2 x 102 CFU of the pathogen.

Similarly, Atterbury, et al. (2007) reported better efficacy with phage application at MOI

of 106 during in-vitro studies against Salmonella serovars.

Based on the results obtained in this study, a phage cocktail (Cocktail) was the most

efficient in reducing colonialization of P. aeruginosa bacteria during in-vitro bacteriolytic

activity and sand decontamination studies when compared with monophages. The use

of combination of two or more phages with different host ranges in a single suspension

(a phage cocktail) has been reported to be more effective than the use of a single phage

alone (Chan, et al., 2013, Gu, et al., 2012, Hall, et al., 2012). For instance, Fu, et al. (2010)

reported a significant reduction of biofilm formation by P. aeruginosa M4 on catheters

using a cocktail of five best phages as opposed to treatment with a single phage. As

clearly explained by Schmerer, et al. (2014), one advantage of using a phage cocktail is

the presence of a large collective host range that may obviate the need to characterize

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phage sensitivities of the infecting pathogenic bacteria. Another advantage relies on

thwarting resistance when multiple phages target the same bacterium. In this case, the

evolution of resistance to all such phages may be required before treatment fails. A third

possible mechanism is dynamical: two phages may collectively kill the bacterial

population more rapidly or more completely than either phage alone. This latter process

of ‘synergy’ between phages is relatively unexplored, perhaps because its

demonstration requires a quantitative assessment of bacterial densities during

treatment (Schmerer, et al., 2014). Although the results reported in this study showed

efficient growth suppression of P. aeruginosa bacteria in both the phage in-vitro

bacteriolytic activity and sand treatment experiments, a regrowth of bacteria at 24 hrs

and 48 hrs post-treatment (in the case of sand treatment) was noticed. As highlighted

in literature, surviving bacteria may be due to a reduced probability of viruses to find

host bacteria (Bull, et al., 2002, Levin and Bull, 2004), a non-replicating condition of

surviving bacteria that is physiologically refractory to phage infection (Bull, et al., 2002),

lysogenic conversion (Skurnik and Strauch, 2006) and due to the development of phage

resistance by the bacterial host (Levin and Bull, 2004). The assumption of bacteria

regrowth due to the low probability of an encounter between viruses and the bacteria-

host is not likely because an increase in MOI (MOI 105) did not increase the efficiency of

phage therapy. Furthermore, the assumption of non-replicating bacteria to be

physiologically refractory to phage infection is also unlikely because following the peak

of bacterial inactivation by the phages, the remaining bacteria grew at a high rate and

reached densities similar to those observed in the controls (Silva, et al., 2014).

The occurrence of lysogeny, which can also render the bacterium immune to not only

the original phages buy also to related phages might be one of the reasons for bacterial

regrowth after phage treatment. However, it is essential to evaluate the occurrence of

lysogenic conversions following rigorous testing to exclude this possibility (Silva, et al.,

2014). The hypothesis of bacteria re-growth due to the presence of phage-resistant

bacterial mutants may be possible. The resistance of bacteria towards phage may be

due factors such as mutations that affect phage adsorption, restriction modification or

the mechanisms of abortive infection such as the presence of clustered regularly

interspaced short palindromic repeats (CRISPRs) in the bacterial genome as discussed

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earlier in this chapter (Allison and Klaenhammer, 1998, Barrangou, et al., 2007, Donlan,

2009). When pre-treated sand samples were treated a second time with phage

(recharged) at 24 hrs, P. aeruginosa bacterial densities were not significantly different

from those assessed at 48 hrs post-treatment. The observation that recharged

treatment of sand samples did not allow bacteria growth suppression explains that

surviving bacteria during the first treatment may be phage-resistant mutants and an

increase in phage dose did not render bacterial cells susceptible to phage. These findings

support the idea that, applying phages in a single dose takes advantage of the phage

potential to replicate and thereby achieve ‘active’ therapy, i.e., significant phage

amplification via auto “dosing” that results in greater bacterial killing (Abedon and

Thomas-Abedon, 2010, Capparelli, et al., 2010). Achieving efficacy following only a single

dose, or far less frequent dosing, is an obvious convenience, though in many or most

instances a single dosage of phages should not be expected, a priori, to be sufficient to

achieve desired efficacy (Capparelli, et al., 2010). Another reason that might have

contributed to the re-growth of bacteria even after phage recharge could be due to

impaired diffusion of bacteriophages depending on the structure and composition of

the matrices (Marcó, et al., 2010). It is assumed that, in solid media, the diffusion of

bacteriophages could be limited, thus reducing phage adsorption on bacteria and

consequently the phage infection capacity. For example in a study reported by

Guenther, et al. (2009) it was shown that the use of bacteriophages was limited by their

diffusion in solid food matrices such as hot dogs, smoked salmon, and seafood.

3.4 Conclusion

The study in this chapter reports, the isolation of lytic bacteriophages from soil samples

collected at Sarawak limestone caves (FCNR and WCNR), targeting different pathogenic

bacteria. Phage lysates were spot tested on various bacterial strains to determine their

lysis spectrum. Analysis of phage bacteriolytic activity was performed in an in- vitro co-

culture assay with P. aeruginosa PAO1 using different multiplicity of infection (MOI)

ratios. Surviving P. aeruginosa PAO1 cells following an in-vitro treatment with phage

were enumerated at 6 hrs and 24 hrs post-infection and the counts were compared with

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those of uninfected P. aeruginosa PAO1 cultures (Controls). The final part of this work

explored the applicability of the phage isolates as biological disinfectants to control

infections caused by P. aeruginosa. Decontamination experiments were conducted by

spraying selected phages (a monophage FCPA3, a multiphage WCSS4PA and a phage

cocktail Cocktail) individually onto sand samples immobilized with P. aeruginosa PAO1

cells, followed by incubation for up to 48 hrs. About 33 lytic phages were isolated from

limestone cave soil samples with P. aeruginosa and V. parahaemolyticus phage isolates

displaying the broadest host range. The highest bacterial inactivation was seen in

cultures infected with phages FCPA3, WCSS4PA and Cocktail when compared with

uninfected P. aeruginosa PAO1 cultures (Controls). Plate count results performed to

assess bacterial survival following in-vitro treatment with phage reveled that phage

cocktail (Cocktail, MOI 104) had the highest bacterial log reduction of 11.82 log10

CFU/mL equivalent to 100% reduction in bacterial cells at the end of 6 hrs of incubation.

Similarly, phages WCSS4PA (MOI 104) and Cocktail (MOI 102) also reported the highest

bacterial log reduction of 10.86 log10 CFU/mL and 10.5 log10 CFU/mL respectively,

equivalent to 100% reduction in bacterial cells at the end of 24 hrs of incubation. Some

of the phages failed to show any bacterial reduction at 6 hrs and 24 hrs post-infection,

instead the cells rebounded and surpassed those of the untreated control. Sand

decontamination experiments reported over 99% reduction in P. aeruginosa PAO1

bacterial cells in all three tested phages (FCPA3, WCSS4PA and Cocktail) when

compared with untreated control at 6 hrs post-treatment. The highest bacterial log

reduction was 4.2 log10 CFU/mL equivalent to 99.99% reduction in bacterial cells

achieved by sand samples sprayed with a phage cocktail (Cocktail). However, no

significant bacterial reduction was seen in sand samples harvested at 24 hrs and 48 hrs

post-treatment despite phage recharge at 24 hrs, instead, the cells rebounded and

surpassed those of untreated control. The results presented in this chapter

demonstrates the presence of lytic phages from Sarawak limestone cave soils capable

of inactivating P. aeruginosa PAO1 bacterial cells. However, further studies are

warranted especially on the emergence of phage-resistant mutants, assumed to be the

cause of bacterial regrowth during phage bacteriolytic and small-scale sand

decontamination studies reported in this study.

Chapter 4 GENERAL CONCLUSION AND FUTURE PERSPECTIVE

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4.1 General Conclusion

4.1.1 Aim of the thesis

The emergence and widespread of multi-drug resistant bacteria, accompanied by a slow

progress in the development of new antibiotics, have built up interests in the search for

alternative and natural antimicrobial agents (Gorski, et al., 2016, Jassim and Limoges,

2014). Virulent bacteriophages represent a viable antibacterial technology that has

proven effective to control multi-drug resistant bacterial pathogens (Nagel, et al., 2016).

The large variation of increasingly multi-drug resistant bacteria causing infections,

demands exploration of previously untapped biological niche such as limestone caves

for potential novel lytic bacteriophages. Preceding studies have reported on the

presence of novel bioactive compounds from caves with antimicrobial properties. Novel

antibiotics such as Cervimycins A-D and xiakemycin A have been successfully isolated

from cave bacteria (Herold, et al., 2005, Jiang, et al., 2015). These antibiotics have shown

activity against methicillin-resistant Staphylococcus aureus and vancomycin-resistant

Enterococcus faecalis. Xiakemycin A has been reported to extend its activity to include

Staphylococcus epidermidis and vancomycin-resistant Enterococcus faecium. In

addition, it has demonstrated additional antifungal and cytotoxic effects against cancer

cells (Bretschneider, et al., 2012, Herold, et al., 2004). These findings indicate that caves

are a rich reservoir of potential and novel antimicrobials which can open new frontiers

for drug discovery. This thesis is thus, the first report on the isolation of lytic

bacteriophages from Sarawak limestone caves (FCNR and WCNR) with the potential to

be developed into biological disinfection agents to control infections caused by P.

aeruginosa bacteria.

The preceding chapters (Chapter 2 and Chapter 3) described the studies undertaken to

fulfill the major aims of the thesis, which were:

i. To screen and isolate lytic phages from limestone cave soil samples.

ii. To investigate the phage bacteriolytic activity in in-vitro.

iii. To treat sand samples contaminated with P. aeruginosa using isolated phages.

This chapter provides an overview of the major findings of this research study as well as

identifying the scope for further research.

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4.1.2 Summary of the findings

4.1.2.1 Bacteriophage isolation and host range analysis

There are no documented studies on presence and diversity of bacteriophages

inhabiting limestone caves despite the fact that caves are known to harbor novel

antimicrobial compounds, active against many multi-drug resistant bacteria. This study

broadens our knowledge about the presence and diversity of bacteriophages inhabiting

Sarawak limestone caves (FCNR and WCNR). A total of 33 lytic bacteriophages were

isolated from Sarawak limestone cave samples targeting bacterial strains V.

parahaemolyticus, S. aureus, K. pneumoniae, E. coli, P. aeruginosa and S. pneumoniae

following enrichment method. The phage isolates were tested against strains of a well-

defined bacterial collection (host range assay), a strategy used to screen for suitable

biocontrol candidates.

In phage therapy and biocontrol studies, a broad host range phage capable of killing

multiple species of bacteria is equivalent to a broad spectrum antibiotic (Ross, et al.,

2016). In this study phage isolates, V. parahaemolyticus and P. aeruginosa phages

showed the broadest host range. A fascinating observation was the ability of V.

parahaemolyticus phage isolates to infect bacterial strain S. aureus which belongs to a

completely different phylum i.e. Eubacteria. Nevertheless, another observation seen

was the ability of some phage isolates to display trans-subdomain infectivity between

gram positive and gram-negative bacterial hosts. For instance, S. pneumoniae phage

isolates (FCSP1, FCSP2 and FCSP3) were capable of infecting P. aeruginosa and

E. coli bacteria. This phenomenon is attributed to the oligotrophic nature of the cave

environment from which the samples were collected (Malki, et al., 2015).

Furthermore, phages with the ability to use more universal receptors for adsorption

exhibit broader host range and are usually found in environments with poor

nutrients (Nilsson, 2014).

4.1.2.2 Assessment of phage in-vitro bacteriolytic activity

The most important criteria for selecting phages for therapeutic or biocontrol

applications are specificity and effective lysis of the targeted bacteria (Zhang, et al.,

2015). This study assessed the ability of selected P. aeruginosa phage isolates

individually and, in a cocktail, to inhibit or completely inactivate the bacterial host in an

121

in-vitro co-culture assay for up to 6 hrs. The spectrophotometric method which involves

the use of optical density measurement was utilized in the assessment of phage lytic

activity because it is rapid and non-destructive to the cells (Sutton, 2011). Five MOI

ratios were tested to optimize the phage dose required to inhibit or completely

inactivate the bacterial host and to provide a basis on which to select the most suitable

phages for subsequent sand decontamination experiments. The results showed that the

growth of P. aeruginosa PAO1 was inactivated when co-cultured with phage in a

concentration-dependent manner, with OD values declining more quickly at higher MOI

(104 and 105) than at lower MOI (103, 102 or 101). Overall, bacteriophages designated as

FCPA3, WCSS4PA and Cocktail showed the highest bacterial inactivation when

compared with the rest phages. One possible drawback in the use of optical density

measurement to estimate bacterial population density during phage bacteriolytic

studies is the evolution of phage resistance. Resistant genotypes may overproduce

alginate or extracellular polymeric substances (EPS) as mentioned earlier in this thesis

(Section 3.1), which may result in OD measurements inflation Hall, et al. (2012). Hence,

it was imperative that bacterial cell concentration by plate count needed to be

performed for accurate results. Survival of P. aeruginosa PAO1 bacterial cells following

an in-vitro co-culture with phage were determined using plate count at 6 hrs and 24 hrs

post-infection and the results were expressed as log10 CFU/mL. Uninfected bacterial

cultures were used as the controls of the experiment. The results revealed that phage

cocktail (Cocktail, MOI 104) had the highest bacterial log10 CFU/mL reduction of 11.82

equivalent to 100% reduction in bacterial cells at the end of 6 hrs of incubation.

However, surviving bacterial counts assessed 24 hrs post-infection showed that a

multiphage (WCSS4PA, MOI 104) and a phage cocktail (Cocktail, MOI 102) had the

highest bacterial log10 CFU/mL reduction of 10.5 and 10.86 respectively, equivalent to

100% reduction in bacterial cells. These findings support the notion that, when two or

more phages in the cocktail attack the same bacterium, the combination results in a

better killing than the application of a single phage. Some of the phages did not show

any reduction in bacterial cells at 6 hrs and 24 hrs post-infection, but instead, the cells

rebounded and surpassed those of the untreated control. This phenomenon has been

attributed to presence and development of phage-resistant mutants.

122

4.1.2.3 Evaluation of phage isolates as biological disinfectants against P. aeruginosa

In order to evaluate the applicability of the phage isolates to be employed as biological

disinfectants against P. aeruginosa PAO1 bacteria, a small-scale decontamination

experiment was designed, which utilized experimentally contaminated sand samples.

Decontamination process was performed by spraying selected phages (a monophage

FCPA3, a multiphage WCSS4PA and a phage cocktail Cocktail) individually onto the

sand samples followed by incubation for up to 48 hrs. Untreated samples were used as

the controls of the experiment. Over 99% reduction in P. aeruginosa PAO1 bacterial cells

were observed on all phage treated sand samples harvested at 6 hrs post-treatment.

However, the highest bacterial log10 CFU/mL reduction was 4.2 equivalent to 99.99%

reduction in bacteria achieved by sand samples sprayed with a phage cocktail

(Cocktail). No significant reduction in bacterial cells was observed in sand samples

harvested at 24 hrs and 48 hrs post-treatment despite phage recharge at 24 hrs, but

instead, the cells rebounded and surpassed those of untreated controls. Phage recharge

was performed to investigate the effect of an additional dose of phage at preventing

regrowth of bacteria and possibly reduce bacterial colonialization on the sand samples.

The occurrence of bacterial regrowth despite phage recharge could be due to the timing

of the additional dose, as this has been shown to be crucial in effective infection control

(Hall, et al., 2012, Torres-Barceló, et al., 2014). The results attained in this study

demonstrates that phage cocktail (Cocktail) was the most efficient at reducing P.

aeruginosa PAO1 colonialization when compared with a monophage (FCPA3). This is in

line with preceding literature that has reported the use of phage cocktail to be more

effective than the use of a single phage alone (Chan, et al., 2013, Gu, et al., 2012, Hall,

et al., 2012).

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4.2 Future Perspectives and Recommendation

The work presented in this thesis intended to screen and isolate lytic bacteriophages

from limestone cave environment and demonstrate their applicability as biological

disinfectants to control infections caused by P. aeruginosa. The results attained in this

study suggests that the phage isolates designated as FCPA3, WCSS4PA and Cocktail,

are promising biological disinfectant candidates against colonialization of P. aeruginosa.

Although this work focused on disinfection of contaminated sand samples, phage

application can be extended to include other materials such as hospital equipment or

indwelling devices that would benefit from an additional method of sterilization. This

will, in turn, reduce the use of antibiotics in the treatment of human diseases as well as

minimizing the number of chemicals and detergents needed to decontaminate such

surfaces. However, this can only be achieved by carrying out further investigative studies

on the phage isolates reported in this study. Additionally, this work can be used as a

starting point for several other lines of investigations to attain a deeper understanding

of the application of phages in biocontrol of bacterial pathogens.

4.2.1 Morphological and Molecular characterization of the phage isolates

Future studies should consider the preliminary characterization of the phage isolates by

transmission electron microscopy (TEM). This will not only permit phage classification

to one of the morphologically distinguishable families but will allow its inclusion to a

group of phages of similar size and morphology within a family (Lobocka, et al., 2014).

Phage characterization focusing on growth kinetics and DNA analysis-based methods of

phage grouping such as Pulsed-field gel electrophoresis (PFGE) and Restriction fragment

length polymorphisms (RFLP) should be performed. Phage growth kinetics will

determine the latent period and the burst size of the isolates. PFGE will allow grouping

or identification of phages based on the size of their virion DNA. The DNA of phages

within each genome size group can be further differentiated based on digestion profiles

with restriction endonucleases. The optimal set of enzymes for digestion needs to be

selected either based on an in silico analysis of genomes of known phages that infect

bacteria of the same or related species as those infected by the tested phage isolates or

empirically (Lobocka, et al., 2014). RFLP allows assessment of bacteriophage genome

diversity (Clokie, et al., 2011). Proteomic approaches to identify viral structural proteins

124

by SDS-PAGE can also be performed. Whole genome sequencing of the phage isolates

will allow a thorough investigation of the phage’s obligate lytic nature and estimate its

safety, especially the risk of participating in the horizontal transfer of bacterial, plasmid

and temperate phage’s genes. It is postulated that a distinctive feature of temperate

phages that infect bacterial pathogens are genes encoding pathogenicity-related traits.

Thus, identification of such genes should exclude a phage for studies involving phage

therapy (Lobocka, et al., 2014).

4.2.2 Broadening applications of the phage isolates

It will be interesting to further analyze phage specificity, infection process, adsorption

potentials and biocontrol efficiency of the phage isolates using a wide collection of

clinical strains. Testing the potential of phage isolates obtained in this study on clinical

strains will expand applications of the phages as biocontrol agents of pathogenic

bacteria. Future studies should also examine the influence of non-target host cells on

sand samples to be decontaminated. There is a possibility that, a non-target host

bacteria may affect the ability of the phages to adsorb to the intended host as

highlighted by Wilkinson (2001).

4.2.3 Assessment of phage stability

A sensible investigation step should evaluate phage stability under storage conditions

and formulations. Phages are composed of protein structures which may render them

unstable in solution formulations (Vandenheuvel, et al., 2015). Phage storage should

ensure the stability of phage particles in the form and conditions in which preparation

is stored, but the form of application should also protect phage particles against losing

their activity (Weber-Dąbrowska, et al., 2016). Stability of the phage can be monitored

based on parameters such as temperature and pH. Furthermore, it will be worthwhile

to investigate the synergistic effect of the best bacteriophage candidates reported in

this study (FCPA3, WCSS4PA and Cocktail) and commercial chemical disinfectants in

decontamination studies.

4.2.4 Assessment of phage-resistant mutants

Regarding bacterial regrowth observed during phage in-vitro bacteriolytic activity

studies, more specific tests such as searching for modifications in bacterial phage

125

receptors should be performed to evaluate the development of resistance and explain

the occurrence of viable P. aeruginosa PAO1 bacteria during phage biocontrol studies.

Investigations focusing on potential phage receptors identification to determine

whether there is any correlation between phage susceptibility and phage receptor types

can be conducted. Additionally, the frequency of the occurrence and the stability of

bacteriophage resistant mutants (BRM) can also be analyzed.

4.2.5 Investigative studies on the expansion of host range

The use of a phage cocktail (Cocktail) and a multiphage (WCSS4PA) proved to be more

effective at reducing bacterial colonialization during both in-vitro bacteriolytic activity

and sand treatment studies reported in this work. The use of phage cocktail is claimed

to eliminate cross-resistance, and based on this fact, a bacterium which is resistant to

one phage may remain sensitive to another. Additionally, cocktails that are composed

of different receptors for binding to bacteria may be a better solution for eliminating

the development of resistance in bacteria (Gill and Hyman, 2010). Future work should

look at modifying isolation procedures and growth conditions that favor the isolation of

phages with broader host ranges. Recent advances in sequencing technologies and

genetic engineering have made it possible to design phages with more predictable and

domesticated therapeutic properties. For instance, recombinant phages with hybrid tail

fibers can be created to broaden the bacterial host ranges (Lin, et al., 2012).

126

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APPENDICES

Appendix I: Plaque appearance of bacteriophages infecting (A) K. pneumoniae, (B) P.

aeruginosa, (C) E. coli and (D) V. parahaemolyticus, following an overnight incubation at

37oC.

A B

C D

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Appendix II: Multiplicity of infection (MOI)

Multiplicity of infection (MOI) refers to the ratio of infectious virions to the cells in a

culture (Shabram and Aguilar-Cordova, 2000). Multiplicity of infection (MOI) can be

calculated by dividing the number of phage added (mL added x PFU/mL) by the number

of bacteria added (mL added x cells/mL) as shown below:

MOI= (PFU/mL x Volume added) (CFU/mL x Volume added)

In order to calculate the MOI, it is recommended to determine the number of cells you

are infecting and the titer of the virus inoculated on them. In this study P. aeruginosa

bacterial cells grown to their mid-exponential phase (7.76 x 1010 CFU/mL) were treated

with phages having titers ranging from 1.25 x 1015 PFU/mL and 5.83 x 1015 PFU/mL.

Dilution formula (C1V1=C2V2) was used to prepared desired concentrations of both

bacteriophage and bacteria. Desired phage concentration was 1.0 x 1015 PFU/mL and

that of bacteria was 1.0 x 1010 CFU/mL.

For instance, to make 50 mL of 1.0 x 1015 PFU/mL phage lysate from a stock solution of

5.83 x 1015 PFU/mL, the following calculation was performed.

C1V1=C2V2

(5.83 x 1015 PFU/mL) V1= (1.0 x 1015 PFU/mL) x 50 mL

V1= 8.58mL

Thus, 8.58 mL of 5.83 x 1015 PFU/mL phage lysate stock was added into (50 mL - 8.58

mL) = 41.42 mL of a diluent (PB).

Likewise, to make 50 mL of (1.0 x 1010 CFU/mL) bacteria solution from a stock solution

of 7.76 x 1010 CFU/mL, the following calculation was performed.

C1V1=C2V2

(7.76 x 1010 CFU/mL) V1= (1.0 x 1010 CFU/mL) x 50 mL

V1=6.44 mL

Thus, 6.44 mL of 7.76 x 1010 CFU/mL bacteria stock solution was added into (50 mL –

6.44 mL) = 43.56 mL of a diluent (PBS).

Since the aim of this study was to test infection frequencies at different MOI ratios, it

was necessary that conditions were identical between experiments. Thus, equal

volumes of phage and bacteria were used, with the total volume set to 50 mL. Five MOI

ratios were selected (101, 102, 103, 104, 105). The reason to why high MOI ratios were

selected, was to investigate whether the use of high MOI dose could inhibit or eliminate

167

P. aeruginosa bacterial cells by passive inundation and possibly minimise the occurrence

of phage resistant mutants. The use of similar MOI range has also been reported by

Atterbury, et al. (2007) where application of phage 10 at MOI of 106 reduced S. enterica

serotype Typhimurium counts to below the limit of detection in 24 hrs.

In order to obtain these selected MOI ratios (101, 102, 103, 104, 105), phage stock with a

titer of 1.0 x 1015 PFU/mL was diluted using the dilution formula (C1V1=C2V2) as described

earlier, to obtain phage lysates with titer ranging from (1x 1011, 1x 1012, 1x 1013, 1x 1014

and 1x 1015) PFU/mL. About 25 mL of the desired phage concentration was added into

25 mL of P. aeruginosa bacterial culture with a concentration of 1.0 x 1010 CFU/mL. The

MOI ratios were calculated as follows:

MOI=𝑃𝑓𝑢/𝑚𝐿

𝐶𝑓𝑢/𝑚𝐿

MOI 10 or 101= (1x 1011 PFU/mL) (1x 1010 CFU/mL)





168

Appendix III: Optical density (OD) values of phage FCPA1 obtained during an assessment of phage bacteriolytic activity at varied MOI ratios

169

Appendix IV: Optical density (OD) values of phage FCPA2 obtained during an assessment of phage bacteriolytic activity at varied MOI

ratios

170

Appendix V: Optical density (OD) values of phage FCPA3 obtained during an assessment of phage bacteriolytic activity at varied MOI

ratios

171

Appendix VI: Optical density (OD) values of phage FCPA4 obtained during an assessment of phage bacteriolytic activity at varied MOI

ratios

172

Appendix VII: Optical density (OD) values of phage FCPA5 obtained during an assessment of phage bacteriolytic activity at varied MOI

ratios

173

Appendix VIII: Optical density (OD) values of phage FCPA6 obtained during an assessment of phage bacteriolytic activity at varied MOI

ratios

174

Appendix IX: Optical density (OD) values of phage WCSS4PA obtained during an assessment of phage bacteriolytic activity at varied

MOI ratios

175

Appendix X: Optical density (OD) values of phage WCSS5PA obtained during an assessment of phage bacteriolytic activity at varied

MOI ratios

176

Appendix XI: Optical density (OD) values of phage Cocktail obtained during an assessment of phage bacteriolytic activity at varied MOI

ratios

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