Isolation of bacteriophages obtained from limestones cave soils … · 2018-09-11 · THEIR...
Transcript of 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
of phage bacteriolytic activity at varied MOI ratios ......................................................... 169
Appendix V: Optical density (OD) values of phage FCPA3 obtained during an assessment
of phage bacteriolytic activity at varied MOI ratios ......................................................... 170
Appendix VI: Optical density (OD) values of phage FCPA4 obtained during an assessment
of phage bacteriolytic activity at varied MOI ratios ......................................................... 171
Appendix VII: Optical density (OD) values of phage FCPA5 obtained during an assessment
of phage bacteriolytic activity at varied MOI ratios ......................................................... 172
Appendix VIII: Optical density (OD) values of phage FCPA6 obtained during an assessment
of phage bacteriolytic activity at varied MOI ratios ......................................................... 173
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
of phage bacteriolytic activity at varied MOI ratios ......................................................... 176
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
3.8 In-vitro bacteriolytic activity of FCPA2 at different MOI ratios 68
3.9 In-vitro bacteriolytic activity of FCPA3 at different MOI ratios 69
3.10 In-vitro bacteriolytic activity of FCPA4 at different MOI ratios 70
3.11 In-vitro bacteriolytic activity of FCPA5 at different MOI ratios 71
3.12 In-vitro bacteriolytic activity of FCPA6 at different MOI ratios 72
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
3.17 Survival of P. aeruginosa PAO1 cells after an in-vitro treatment with
FCPA2 at different MOI ratios
91
3.18 Survival of P. aeruginosa PAO1 cells after an in-vitro treatment with
FCPA3 at different MOI ratios
92
3.19 Survival of P. aeruginosa PAO1 cells after an in-vitro treatment with
FCPA4 at different MOI ratios
93
3.20 Survival of P. aeruginosa PAO1 cells after an in-vitro treatment with
FCPA5 at different MOI ratios
94
3.21 Survival of P. aeruginosa PAO1 cells after an in-vitro treatment with
FCPA6 at different MOI ratios
95
3.22 Survival of P. aeruginosa PAO1 cells after an in-vitro treatment with
WCSS4PA at different MOI ratios
96
3.23 Survival of P. aeruginosa PAO1 cells after an in-vitro treatment with
WCSS5PA at different MOI ratios
97
3.24 Survival of P. aeruginosa PAO1 cells after an in-vitro treatment with
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
3.7 Analysis of variance (ANOVA) results for the recovery of bacteria
following an in-vitro treatment with FCPA2
78
3.8 Analysis of variance (ANOVA) results for the recovery of bacteria
following an in-vitro treatment with FCPA3
80
3.9 Analysis of variance (ANOVA) results for the recovery of bacteria
following an in-vitro treatment with FCPA4
81
3.10 Analysis of variance (ANOVA) results for the recovery of bacteria
following an in-vitro treatment with FCPA5
83
3.11 Analysis of variance (ANOVA) results for the recovery of bacteria
following an in-vitro treatment with FCPA6
84
3.12 Analysis of variance (ANOVA) results for the recovery of bacteria
following an in-vitro treatment with WCSS4PA
86
3.13 Analysis of variance (ANOVA) results for the recovery of bacteria
following an in-vitro treatment with WCSS5PA.
87
3.14 Analysis of variance (ANOVA) results for the recovery of bacteria
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
3.16 Analysis of variance (ANOVA) results for surviving bacterial cells
recovered from sand treated samples at 6 hrs using different
phage samples.
100
3.17 Analysis of variance (ANOVA) results for surviving bacterial cells
recovered from sand treated samples at 24 hrs using different
phage samples.
100
3.18 Analysis of variance (ANOVA) results for surviving bacterial cells
recovered from sand treated samples at 48 hrs using different
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
FCNR FCVP2 V. parahaemolyticus 5 Clear, round 6.3 x 108
FCNR FCVP3 V. parahaemolyticus 4 Clear, round 1.1 x 107
FCNR FCVP4 V. parahaemolyticus 4 Clear, round 8.5 x 107
FCNR FCVP5 V. parahaemolyticus 6 Clear, round 8.0 x 106
FCNR FCVP6 V. parahaemolyticus 4 Clear, round 1.18 x 109
WCNR WCVP3 V. parahaemolyticus Nil Turbid Nil
WCNR WCVP4 V. parahaemolyticus Nil Turbid Nil
WCNR WCVP5 V. parahaemolyticus Nil Turbid Nil
FCNR FCSA1 S. aureus 4 Clear, round 1.89 x 107
FCNR FCSA3 S. aureus 3 Clear, round 1.20 x 107
FCNR FCSA4 S. aureus 3 Clear, round 1.36 x 108
FCNR FCSA6 S. aureus 2 Clear, round 1.22 x 106
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
FCNR FCEC3 E. coli 5 Clear, round 1.01 x 108
60
FCNR FCEC6 E. coli 5 Clear, round 4.2 x 107
FCNR FCEC7 E. coli Nil Turbid Nil
FCNR FCPA1 P. aeruginosa 3 Clear, round 2.28 x 1013
FCNR FCPA2 P. aeruginosa 3 Clear, round 1.37 x 1013
FCNR FCPA3 P. aeruginosa 5 Clear, round 3.01 x 1014
FCNR FCPA4 P. aeruginosa 3 Clear, round 1.52 x 1015
FCNR FCPA5 P. aeruginosa 3 Clear, round 2.16 x 1013
FCNR FCPA6 P. aeruginosa 4 Clear, round 9.4 x 108
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
FCNR FCSP2 S. pneumoniae 4 Clear, round 2.29 x 107
FCNR FCSP3 S. pneumoniae 4 Clear, round 1.19 x 108
FCNR FCSP4 S. pneumoniae 3 Clear, round 1.25 x 108
FCNR FCSP5 S. pneumoniae 3 Clear, round 1.87 x 107
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
Uninfected control 1.370
FCPA3 101 0.766
102 0.566
103 0.588
104 0.383
105 0.200
Uninfected control 1.370
FCPA4 101 1.431
102 1.441
103 1.493
104 1.527
105 1.495
Uninfected control 1.563
66
FCPA5
101 1.407
102 1.129
103 0.941
104 0.919
105 0.900
Uninfected control 1.471
FCPA6 101 1.188
102 1.266
103 1.309
104 1.317
105 1.329
Uninfected control 1.563
WCSS4PA 101 1.229
102 1.189
103 1.168
104 0.897
105 0.319
Uninfected control 1.533
WCSS5PA 101 1.136
102 1.076
103 1.105
104 1.096
105 1.146
Uninfected control 1.533
Cocktail 101 0.945
102 0.616
103 0.349
104 0.288
105 0.405
Uninfected control 1.557
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
Figure 3.8: In-vitro bacteriolytic activity of FCPA2 at different MOI ratios. Mid–
exponential cultures of P. aeruginosa PAO1 were co-cultured with bacteriophage
FCPA2 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
ΦFCPA2 (MOI 105)ΦFCPA2 (MOI 104)ΦFCPA2 (MOI 103)
ΦFCPA2 (MOI 102)ΦFCPA2 (MOI 101)
69
Figure 3.9: In-vitro bacteriolytic activity of FCPA3 at different MOI ratios. Mid–
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
ΦFCPA3 (MOI 105)ΦFCPA3 (MOI 104)ΦFCPA3 (MOI 103)
ΦFCPA3 (MOI 102)ΦFCPA3 (MOI 101)
70
Figure 3.10: In-vitro bacteriolytic activity of FCPA4 at different MOI ratios. Mid–
exponential cultures of P. aeruginosa PAO1 were co-cultured with bacteriophage
FCPA4 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
ΦFCPA4 (MOI 105)ΦFCPA4 (MOI 104)ΦFCPA4 (MOI 103)
ΦFCPA4 (MOI 102)ΦFCPA4 (MOI 101)
71
Figure 3.11: In-vitro bacteriolytic activity of FCPA5 at different MOI ratios. Mid–
exponential cultures of P. aeruginosa PAO1 were co-cultured with bacteriophage
FCPA5 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
ΦFCPA5 (MOI 105)ΦFCPA5 (MOI 104)ΦFCPA5 (MOI 103)
ΦFCPA5 (MOI 102)ΦFCPA5 (MOI 101)
72
Figure 3.12: In-vitro bacteriolytic activity of FCPA6 at different MOI ratios. Mid–
exponential cultures of P. aeruginosa PAO1 were co-cultured with bacteriophage
FCPA6 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
ΦFCPA6 (MOI 105)ΦFCPA6 (MOI 104)ΦFCPA6 (MOI 103)
ΦFCPA6 (MOI 102)ΦFCPA6 (MOI 101)
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
standard error of the mean.
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.
Table 3.7: Analysis of variance (ANOVA) results for the recovery of bacteria following an
in-vitro treatment with FCPA2
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
Table 3.8: Analysis of variance (ANOVA) results for the recovery of bacteria following an
in-vitro treatment with FCPA3
Group Sum Mean Sample
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
sample size mean =3; degrees of freedom =11,24; sum of squares =368.556, 2.642; mean square =33.505, 0.110, and F statistic =304.227.
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
in-vitro treatment with FCPA4
Group Sum Mean Sample
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
sample size mean =3; degrees of freedom =11,24; sum of squares =15.5883, 0.1097; mean square =33.505, 0.110, and F statistic =142.0855.
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
Group Sum Mean Sample
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
sample size mean =3; degrees of freedom =11,24; sum of squares =448.240, 3.760; mean square =40.749, 0.157, and F statistic =260.117.
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.
Table 3.11: Analysis of variance (ANOVA) results for the recovery of bacteria following
an in-vitro treatment with FCPA6
Group Sum Mean Sample
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
Table 3.12: Analysis of variance (ANOVA) results for the recovery of bacteria following
an in-vitro treatment with WCSS4PA
Group Sum Mean Sample
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
sample size mean =3; degrees of freedom =11,24; sum of squares (399.208, 3.871) and mean square (36.292, 0.161), F statistic (225.014).
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.
Table 3.13: Analysis of variance (ANOVA) results for the recovery of bacteria following
an in-vitro treatment with WCSS5PA.
Group Sum Mean Sample
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
sample size mean =3; degrees of freedom =11,24; sum of squares (185.790, 1.773) and mean square (16.890, 0.074), F statistic (228.652).
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
Table 3.14: Analysis of variance (ANOVA) results for the recovery of bacteria following
an in-vitro treatment with Cocktail.
Group Sum Mean Sample
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
Figure 3.17: Survival of P. aeruginosa PAO1 cells after an in-vitro treatment with
FCPA2 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
92
Figure 3.18: Survival of P. aeruginosa PAO1 cells after an in-vitro treatment with
FCPA3 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
93
Figure 3.19: Survival of P. aeruginosa PAO1 cells after an in-vitro treatment with
FCPA4 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
94
Figure 3.20: Survival of P. aeruginosa PAO1 cells after an in-vitro treatment with
FCPA5 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
95
Figure 3.21: Survival of P. aeruginosa PAO1 cells after an in-vitro treatment with
FCPA6 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
96
Figure 3.22: Survival of P. aeruginosa PAO1 cells after an in-vitro treatment with
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
Figure 3.23: Survival of P. aeruginosa PAO1 cells after an in-vitro treatment with
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
Figure 3.24: Survival of P. aeruginosa PAO1 cells after an in-vitro treatment with
Cocktail at different MOI ratios. Bacteria recovery was performed at 6 hrs and 24
hrs post-incubation. Error bars represent standard error of the mean.
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.
Group Sum Mean Sample
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
Table 3.16: Analysis of variance (ANOVA) results for surviving bacterial cells recovered
from sand treated samples at 6 hrs using different phage samples.
Group Sum Mean Sample
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
sample size mean =3; degrees of freedom =3, 8; sum of squares = 65.3752, 1.1349; mean square =21.7917, 0.1419 and F statistic =153.6141.
Table 3.17: Analysis of variance (ANOVA) results for surviving bacterial cells recovered
from sand treated samples at 24 hrs using different phage samples.
Group Sum Mean Sample
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
sample size mean =3; degrees of freedom =3, 8; sum of squares = 1.3074, 0.9961; mean square =0.4358, 0.1245 and F statistic =3.4998.
101
Table 3.18: Analysis of variance (ANOVA) results for surviving bacterial cells recovered
from sand treated samples at 48 hrs using different phage samples.
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
103
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
104
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
105
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.
106
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
107
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.
108
(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
109
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
111
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,
112
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
113
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
114
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
115
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
116
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
117
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
118
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
119
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.
120
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).
123
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
REFERENCES
Abedon, S. T. 2011. Bacteriophages and biofilms, p. 1-58. In W. C. Bailey (ed.), Biofilms:
Formation, development and properties Nova Science Publishers, Inc.
Abedon, S. T. 1990. The ecology of bacteriophage t4. University of Arizona Univesity of
Arizona (UA) Campus Repository.
Abedon, S. T., García, P., Mullany, P., and Aminov, R. 2017. Phage therapy: Past, present
and future. Frontiers in Microbiology. 8, 1-7.
Abedon, S. T., Kuhl, S. J., Blasdel, B. G., and Kutter, E. M. 2011. Phage treatment of
human infections. Bacteriophage. 1, 66-85.
Abedon, S. T. and Thomas-Abedon, C. 2010. Phage therapy pharmacology. Current
Pharmaceutical Biotechnology. 11, 28-47.
Abraham, E. P. and Chain, E. 1940. An enzyme from bacteria able to destroy penicillin.
Nature. 146, 837.
Abuladze, T., Li, M., Menetrez, M. Y., Dean, T., Senecal, A., and Sulakvelidze, A. 2008.
Bacteriophages reduce experimental contamination of hard surfaces, tomato,
spinach, broccoli, and ground beef by escherichia coli o157: H7. Applied and
Environmental Microbiology. 74, 6230-6238.
Ackermann, H.-W. 2005. Bacteriophage classification, p. 67-89. In E. Kutter and A.
Sulakvelidze (eds.), Bacteriophages: Biology and applications, CRC Press.
Ackermann, H.-W. 2003. Bacteriophage observations and evolution. Research in
Microbiology. 154, 245-251.
Ackermann, H.-W. 2006. Classification of bacteriophages, p. 8-16. In R. Calendar (ed.),
The bacteriophages, Oxford University Press, USA.
Ackermann, H.-W. and Prangishvili, D. 2012. Prokaryote viruses studied by electron
microscopy. Archives of Virology. 157, 1843-1849.
Aderem, A. and Ulevitch, R. J. 2000. Toll-like receptors in the induction of the innate
immune response. Nature. 406, 782-787.
Ahiwale, S., Koparde, P., Deore, P., Gunale, V., and Kapadnis, B. P. 2012. Bacteriophage
based technology for disinfection of different water systems, p. 289-313.
Microorganisms in environmental management, Springer.
127
Alemayehu, D., Casey, P. G., McAuliffe, O., Guinane, C. M., Martin, J. G., Shanahan, F.,
Coffey, A., Ross, R. P., and Hill, C. 2012. Bacteriophages phimr299-2 and phinh-4
can eliminate pseudomonas aeruginosa in the murine lung and on cystic fibrosis
lung airway cells. MBio. 3, 1-9.
Allison, G. E. and Klaenhammer, T. R. 1998. Phage resistance mechanisms in lactic acid
bacteria. International Dairy Journal. 8, 207-226.
Aminov, R., Caplin, J., Chanishvili, N., Coffey, A., Cooper, I., De Vos, D., Doskar, J., Friman,
V.-P., Kurtboke, I., and Pantucek, R. 2017. Application of bacteriophages.
Microbiology Australia. 63-66.
Aminov, R. I. 2010. A brief history of the antibiotic era: Lessons learned and challenges
for the future. Frontiers in Microbiology. 1, 1-7.
Anspach, F. B. 2001. Endotoxin removal by affinity sorbents. Journal of Biochemical and
Biophysical Methods. 49, 665-681.
Arora, G., Sajid, A., and Kalia, V. C. 2017. Drug resistance in bacteria, fungi, malaria, and
cancer, p. Pages. Springer, Cham, Switzerland.
Ashelford, K. E., Day, M. J., and Fry, J. C. 2003. Elevated abundance of bacteriophage
infecting bacteria in soil. Applied and Environmental Microbiology. 69, 285-289.
Atterbury, R. J. 2009. Bacteriophage biocontrol in animals and meat products. Microbial
Biotechnology. 2, 601-612.
Atterbury, R. J., Van Bergen, M. A. P., Ortiz, F., Lovell, M. A., Harris, J. A., De Boer, A.,
Wagenaar, J. A., Allen, V. M., and Barrow, P. A. 2007. Bacteriophage therapy to
reduce salmonella colonization of broiler chickens. Applied and Environmental
Microbiology. 73, 4543-4549.
Babalova, E. G., Katsitadze, K. T., Sakvarelidze, L. A., Imnaishvili, N., Sharashidze, T. G.,
Badashvili, V. A., Kiknadze, G. P., Meĭpariani, A. N., Gendzekhadze, N. D., and
Machavariani, E. V. 1968. Preventive value of dried dysentery bacteriophage.
Zhurnal Mikrobiologii, Epidemiologii, i Immunobiologii 45, 143-145.
Bai, J., Kim, Y.-T., Ryu, S., and Lee, J.-H. 2016. Biocontrol and rapid detection of food-
borne pathogens using bacteriophages and endolysins. Frontiers in
Microbiology. 7, 1-15.
128
Bai, J., Kim, Y.-T., Ryu, S., and Lee, J.-H. 2016. Biocontrol and rapid detection of food-
borne pathogens using bacteriophages and endolysins. Frontiers in microbiology.
7.
Baker, S. 2015. A return to the pre-antimicrobial era? Science. 347, 1064-1066.
Bakhshipouri, Z., Omar, H., Yousof, Z., and Ghiasi, V. 2009. An overview of subsurface
karst features associated with geological studies in malaysia. Electronic Journal
of Geotechnical Engineering. 14, 1-15.
Balogh, B., Jones, J. B., Iriarte, F. B., and Momol, M. T. 2010. Phage therapy for plant
disease control. Current Pharmaceutical Biotechnology. 11, 48-57.
Banat, I. M., Franzetti, A., Gandolfi, I., Bestetti, G., Martinotti, M. G., Fracchia, L., Smyth,
T. J., and Marchant, R. 2010. Microbial biosurfactants production, applications
and future potential. Applied Microbiology and Biotechnology. 87, 427-444.
Banda, R. M., Gendang, R., and Ambun, A. V. 2004. Geology and geochemistry of
limestone in sarawak. The Sarawak Museum Journal. 6, 41-61.
Banin, E., Hughes, D., and Kuipers, O. P. 2017. Bacterial pathogens, antibiotics and
antibiotic resistance. FEMS Microbiology Reviews. 41, 450-452.
Barbu, E. M., Cady, K. C., and Hubby, B. 2016. Phage therapy in the era of synthetic
biology. Cold Spring Harbor Perspectives in Biology. 8, 1-16.
Baroud, á., Dandache, I., Araj, G. F., Wakim, R., Kanj, S., Kanafani, Z., Khairallah, M.,
Sabra, A., Shehab, M., and Dbaibo, G. 2013. Underlying mechanisms of
carbapenem resistance in extended-spectrum β-lactamase-producing klebsiella
pneumoniae and escherichia coli isolates at a tertiary care centre in lebanon:
Role of oxa-48 and ndm-1 carbapenemases. International Journal of
Antimicrobial Agents. 41, 75-79.
Barrangou, R., Fremaux, C., Deveau, H., Richards, M., Boyaval, P., Moineau, S., Romero,
D. A., and Horvath, P. 2007. Crispr provides acquired resistance against viruses
in prokaryotes. Science. 315, 1709-1712.
Barton, H. A. and Jurado, V. 2007. What's up down there? Microbial diversity in caves.
Microbe-American Society for Microbiology. 2, 132.
Basra, S., Anany, H., Brovko, L., Kropinski, A. M., and Griffiths, M. W. 2014. Isolation and
characterization of a novel bacteriophage against mycobacterium avium
subspecies paratuberculosis. Archives of Virology. 159, 2659-2674.
129
Bellard, C., Leclerc, C., Leroy, B., Bakkenes, M., Veloz, S., Thuiller, W., and Courchamp,
F. 2014. Vulnerability of biodiversity hotspots to global change. Global Ecology
and Biogeography. 23, 1376-1386.
Berney, M., Weilenmann, H.-U., Ihssen, J., Bassin, C., and Egli, T. 2006. Specific growth
rate determines the sensitivity of escherichia coli to thermal, uva, and solar
disinfection. Applied and Environmental Microbiology. 72, 2586-2593.
Bernhardt, T. G., Roof, W. D., and Young, R. 2000. Genetic evidence that the
bacteriophage φx174 lysis protein inhibits cell wall synthesis. Proceedings of the
National Academy of Sciences. 97, 4297-4302.
Bernhardt, T. G., Wang, N., Struck, D. K., and Young, R. 2001. A protein antibiotic in the
phage qβ virion: Diversity in lysis targets. Science. 292, 2326-2329.
Bickle, T. A. and Kruger, D. H. 1993. Biology of DNA restriction. Microbiology and
Molecular Biology Reviews. 57, 434-450.
Bigwood, T., Hudson, J. A., and Billington, C. 2009. Influence of host and bacteriophage
concentrations on the inactivation of food-borne pathogenic bacteria by two
phages. FEMS Microbiology Letters. 291, 59-64.
Billal, D. S., Feng, J., Leprohon, P., Légaré, D., and Ouellette, M. 2011. Whole genome
analysis of linezolid resistance in streptococcus pneumoniae reveals resistance
and compensatory mutations. BMC Genomics. 12, 1-10.
Biswas, A., Staals, R. H. J., Morales, S. E., Fineran, P. C., and Brown, C. M. 2016.
Crisprdetect: A flexible algorithm to define crispr arrays. BMC Genomics. 17, 1-
14.
Blair, J. M., Webber, M. A., Baylay, A. J., Ogbolu, D. O., and Piddock, L. J. 2015. Molecular
mechanisms of antibiotic resistance. Nature reviews. Microbiology. 13, 42.
Blair, J. M. A., Webber, M. A., Baylay, A. J., Ogbolu, D. O., and Piddock, L. J. V. 2015.
Molecular mechanisms of antibiotic resistance. Nature Reviews Microbiology.
13, 42-51.
Bohannan, B. J. M. and Lenski, R. E. 2000. Linking genetic change to community
evolution: Insights from studies of bacteria and bacteriophage. Ecology Letters.
3, 362-377.
130
Bolger-Munro, M., Cheung, K., Fang, A., and Wang, L. 2013. T4 bacteriophage average
burst size varies with escherichia coli b23 cell culture age. Journal of Experimental
Microbiology and Immunology (JEMI). 17, 115-119.
Bonilla, N., Rojas, M. I., Cruz, G. N. F., Hung, S.-H., Rohwer, F., and Barr, J. J. 2016. Phage
on tap–a quick and efficient protocol for the preparation of bacteriophage
laboratory stocks. PeerJ. 4, 1-18.
Bono, L. M., Gensel, C. L., Pfennig, D. W., and Burch, C. L. 2013. Competition and the
origins of novelty: Experimental evolution of niche-width expansion in a virus.
Biol Lett. 9, 2012-2016.
Borgaro, J. G. and Zhu, Z. 2013. Characterization of the 5-hydroxymethylcytosine-
specific DNA restriction endonucleases. Nucleic acids research. 41, 4198-4206.
Braun, A., Brugger, H., Gandham, L., and Leinweber, K. 2006. The effect of culture age
and extrinsic factors on t4 bacteriophage progeny production in recovered
stationary phase escherichia coli zk126. Journal of Experikental Microbiology and
Immunology (JEMI). 9, 44-50.
Braun, A., Brugger, H., Gandham, L., and Leinweber, K. 2006. The effect of culture age
and extrinsic factors on t4 bacteriophage progeny production in recovered
stationary phase escherichia coli zk126.
Bretschneider, T., Zocher, G., Unger, M., Scherlach, K., Stehle, T., and Hertweck, C. 2012.
A ketosynthase homolog uses malonyl units to form esters in cervimycin
biosynthesis. Nature Chemical Biology. 8, 154-161.
Brewster, L., Langley, M., and Twa, D. 2012. Co-infection of c3000 escherichia coli with
bacteriophages ms2 and, t7 or φx-174 results in differential cell lysis patterns.
Journal of Experimental Microbiology and Immunology (JEMI) 16, 139-143.
Brouns, S. J. J., Jore, M. M., Lundgren, M., Westra, E. R., Slijkhuis, R. J. H., Snijders, A. P.
L., Dickman, M. J., Makarova, K. S., Koonin, E. V., and Van Der Oost, J. 2008. Small
crispr rnas guide antiviral defense in prokaryotes. Science. 321, 960-964.
Bruttin, A. and Brüssow, H. 2005. Human volunteers receiving escherichia coli phage t4
orally: A safety test of phage therapy. Antimicrobial Agents and Chemotherapy.
49, 2874-2878.
131
Bryson, A. L., Hwang, Y., Sherrill-Mix, S., Wu, G. D., Lewis, J. D., Black, L., Clark, T. A., and
Bushman, F. D. 2015. Covalent modification of bacteriophage t4 DNA inhibits
crispr-cas9. MBio. 6, 1-9.
Bull, J. J., Levin, B. R., DeRouin, T., Walker, N., and Bloch, C. A. 2002. Dynamics of success
and failure in phage and antibiotic therapy in experimental infections. BMC
Microbiology. 2, 1-10.
Bush, K. 2010. Alarming β-lactamase-mediated resistance in multidrug-resistant
enterobacteriaceae. Current Opinion in Microbiology. 13, 558-564.
Cai, Y., Chai, D., Wang, R., Liang, B., and Bai, N. 2012. Colistin resistance of acinetobacter
baumannii: Clinical reports, mechanisms and antimicrobial strategies. Journal of
Antimicrobial Chemotherapy. 67, 1607-1615.
Cairns, J., Stent, G. S., and Watson, J. D. 1968. Phage and the origins of molecular biology.
Journal of the History of Biology. 1, 155-161.
Cannatelli, A., D'Andrea, M. M., Giani, T., Di Pilato, V., Arena, F., Ambretti, S., Gaibani,
P., and Rossolini, G. M. 2013. In vivo emergence of colistin resistance in klebsiella
pneumoniae producing kpc-type carbapenemases mediated by insertional
inactivation of the phoq/phop mgrb regulator. Antimicrobial Agents and
Chemotherapy. 57, 5521-5526.
Cannatelli, A., Giani, T., D'Andrea, M. M., Di Pilato, V., Arena, F., Conte, V.,
Tryfinopoulou, K., Vatopoulos, A., and Rossolini, G. M. 2014. Mgrb inactivation is
a common mechanism of colistin resistance in kpc carbapenemase-producing
klebsiella pneumoniae of clinical origin. Antimicrobial Agents and Chemotherapy.
1-26.
Cantón, R., Akóva, M., Carmeli, Y., Giske, C. G., Glupczynski, Y., Gniadkowski, M.,
Livermore, D. M., Miriagou, V., Naas, T., and Rossolini, G. M. 2012. Rapid
evolution and spread of carbapenemases among enterobacteriaceae in europe.
Clinical Microbiology and Infection. 18, 413-431.
Capparelli, R., Nocerino, N., Iannaccone, M., Ercolini, D., Parlato, M., Chiara, M., and
Iannelli, D. 2010. Bacteriophage therapy of salmonella enterica: A fresh appraisal
of bacteriophage therapy. The Journal of infectious diseases. 201, 52-61.
Capparelli, R., Nocerino, N., Lanzetta, R., Silipo, A., Amoresano, A., Giangrande, C.,
Becker, K., Blaiotta, G., Evidente, A., and Cimmino, A. 2010. Bacteriophage-
132
resistant staphylococcus aureus mutant confers broad immunity against
staphylococcal infection in mice. PLoS ONE. 5, e11720.
Carlton, R. M. 1999. Phage therapy: Past history and future prospects. Archivum
Immunologiae et Therapiae Experimentalis. 47, 267-274.
Carrillo, C. L., Atterbury, R., El-Shibiny, A., Connerton, P., Dillon, E., Scott, A., and
Connerton, I. 2005. Bacteriophage therapy to reduce campylobacter jejuni
colonization of broiler chickens. Applied and environmental microbiology. 71,
6554-6563.
Carrillo, C. L., Atterbury, R. J., El-Shibiny, A., Connerton, P. L., Dillon, E., Scott, A., and
Connerton, I. F. 2005. Bacteriophage therapy to reduce campylobacter jejuni
colonization of broiler chickens. Applied and Environmental Microbiology. 71,
6554-6563.
Carte, J., Wang, R., Li, H., Terns, R. M., and Terns, M. P. 2008. Cas6 is an
endoribonuclease that generates guide rnas for invader defense in prokaryotes.
Genes & Development. 22, 3489-3496.
Carter, C. D., Parks, A., Abuladze, T., Li, M., Woolston, J., Magnone, J., Senecal, A.,
Kropinski, A. M., and Sulakvelidze, A. 2012. Bacteriophage cocktail significantly
reduces escherichia coli o157: H7 contamination of lettuce and beef, but does
not protect against recontamination. Bacteriophage. 2, 178-185.
Carvalho, C., Susano, M., Fernandes, E., Santos, S., Gannon, B., Nicolau, A., Gibbs, P.,
Teixeira, P., and Azeredo, J. 2010. Method for bacteriophage isolation against
target campylobacter strains. Letters in Applied Microbiology. 50, 192-197.
Chan, B. K., Abedon, S. T., and Loc-Carrillo, C. 2013. Phage cocktails and the future of
phage therapy. Future Microbiology. 8, 769-783.
Chang, C.-C., Chen, W.-C., Ho, T.-F., Wu, H.-S., and Wei, Y.-H. 2011. Development of
natural anti-tumor drugs by microorganisms. Journal of Bioscience and
Bioengineering. 111, 501-511.
Chanishvili, N. 2012. Phage therapy—history from twort and d’herelle through soviet
experience to current approaches. Bacteriophages. 83, 3-40.
Cheeptham, N. 2012. Cave microbiomes: A novel resource for drug discovery, p. Pages.
Springer Science & Business Media.
133
Chellat, M. F., Raguž, L., and Riedl, R. 2016. Targeting antibiotic resistance. Angewandte
Chemie International Edition. 55, 6600-6626.
Chen, Y., Wu, L., Boden, R., Hillebrand, A., Kumaresan, D., Moussard, H., Baciu, M., Lu,
Y., and Murrell, J. C. 2009. Life without light: Microbial diversity and evidence of
sulfur-and ammonium-based chemolithotrophy in movile cave. The ISME
Journal. 3, 1093-1104.
Chhibber, S. and Kumari, S. 2012. Application of therapeutic phages in medicine, p. 140-
158. In I. Kurtboke (ed.), Bacteriophages, InTech.
Chibeu, A. 2013. Bacteriophages in food safety. Microbial pathogens and strategies for
combating them: science, technology and education 1041-1052.
Clark, J. R. 2015. Bacteriophage therapy: History and future prospects. Future Virology.
10, 449-461.
Clark, J. R. and March, J. B. 2006. Bacteriophages and biotechnology: Vaccines, gene
therapy and antibacterials. Trends in Biotechnology. 24, 212-218.
Clark, L., Greenbaum, C., Jiang, J., Lernmark, Å., and Ochs, H. 2002. The antibody
response to bacteriophage is linked to the lymphopenia gene in congenic
biobreeding rats. FEMS Immunology & Medical Microbiology. 32, 205-209.
Clements, R., Sodhi, N. S., Schilthuizen, M., and Ng, P. K. L. 2006. Limestone karsts of
southeast asia: Imperiled arks of biodiversity. BioScience. 56, 733-742.
Clokie, M. R. J., Millard, A. D., Letarov, A. V., and Heaphy, S. 2011. Phages in nature.
Bacteriophage. 1, 31-45.
Cole, L. E., Bhagwat, S. A., and Willis, K. J. 2015. Long-term disturbance dynamics and
resilience of tropical peat swamp forests. Journal of Ecolology. 103, 16-30.
Costelloe, C., Metcalfe, C., Lovering, A., Mant, D., and Hay, A. D. 2010. Effect of antibiotic
prescribing in primary care on antimicrobial resistance in individual patients:
Systematic review and meta-analysis. BMJ. 340, 1-11.
Criscuolo, E., Spadini, S., Lamanna, J., Ferro, M., and Burioni, R. 2017. Bacteriophages
and their immunological applications against infectious threats. Journal of
Immunology Research. 2017, 1-14.
Cucchi, T., Fujita, M., and Dobney, K. 2009. New insights into pig taxonomy,
domestication and human dispersal in island south east asia: Molar shape
134
analysis of sus remains from niah caves, sarawak. International Journal of
Osteoarchaeology 19, 508-530.
Cuezva, S., Fernandez-Cortes, A., Porca, E., Pašić, L., Jurado, V., Hernandez-Marine, M.,
Serrano-Ortiz, P., Hermosin, B., Cañaveras, J. C., and Sanchez-Moral, S. 2012. The
biogeochemical role of actinobacteria in altamira cave, spain. FEMS
Microbiology Ecology. 81, 281-290.
Curtin, J. J. and Donlan, R. M. 2006. Using bacteriophages to reduce formation of
catheter-associated biofilms by staphylococcus epidermidis. Antimicrobial
Agents and Chemotherapy. 50, 1268-1275.
Czajkowski, R., Ozymko, Z., Zwirowski, S., and Lojkowska, E. 2014. Complete genome
sequence of a broad-host-range lytic dickeya spp. Bacteriophage varphid5. Arch
Virol. 159, 3153-3155.
D’Andrea, M. M., Marmo, P., De Angelis, L. H., Palmieri, M., Ciacci, N., Di Lallo, G.,
Demattè, E., Vannuccini, E., Lupetti, P., and Rossolini, G. M. 2017. Φbo1e, a
newly discovered lytic bacteriophage targeting carbapenemase-producing
klebsiella pneumoniae of the pandemic clonal group 258 clade ii lineage.
Scientific Reports. 7, 1-8.
Dabrowska, K., Switała‐Jelen, K., Opolski, A., Weber‐Dabrowska, B., and Gorski, A. 2005.
Bacteriophage penetration in vertebrates. Journal of Applied Microbiology. 98,
7-13.
De Bruyn, M., Stelbrink, B., Morley, R. J., Hall, R., Carvalho, G. R., Cannon, C. H., Van Den
Bergh, G., Meijaard, E., Metcalfe, I., and Boitani, L. 2014. Borneo and indochina
are major evolutionary hotspots for southeast asian biodiversity. Systematic
Biology. 63, 879-901.
Debarbieux, L., Leduc, D., Maura, D., Morello, E., Criscuolo, A., Grossi, O., Balloy, V., and
Touqui, L. 2010. Bacteriophages can treat and prevent pseudomonas aeruginosa
lung infections. The Journal of infectious diseases. 201, 1096-1104.
Deltcheva, E., Chylinski, K., Sharma, C. M., Gonzales, K., Chao, Y., Pirzada, Z. A., Eckert,
M. R., Vogel, J., and Charpentier, E. 2011. Crispr rna maturation by trans-encoded
small rna and host factor rnase iii. Nature. 471, 602-607.
Deshpande, L. M., Jones, R. N., Fritsche, T. R., and Sader, H. S. 2006. Occurrence and
characterization of carbapenemase-producing enterobacteriaceae: Report from
135
the sentry antimicrobial surveillance program (2000–2004). Microbial Drug
Resistance. 12, 223-230.
Dhanji, H., Doumith, M., Rooney, P. J., O'leary, M. C., Loughrey, A. C., Hope, R.,
Woodford, N., and Livermore, D. M. 2010. Molecular epidemiology of
fluoroquinolone-resistant st131 escherichia coli producing ctx-m extended-
spectrum β-lactamases in nursing homes in belfast, uk. Journal of Antimicrobial
Chemotherapy. 66, 297-303.
Ding, C. and He, J. 2010. Effect of antibiotics in the environment on microbial
populations. Applied Microbiology and Biotechnology. 87, 925-941.
Dodge-Wan, D. and Mi, A. D. H. 2013. Biologically influenced stalagmites in niah and
mulu caves (sarawak, malaysia). Acta Carsologica. 42, 155-163.
Donlan, R. M. 2009. Preventing biofilms of clinically relevant organisms using
bacteriophage. Trends in Microbiology. 17, 66-72.
Donlan, R. M. and Costerton, J. W. 2002. Biofilms: Survival mechanisms of clinically
relevant microorganisms. Clinical Microbiology Reviews. 15, 167-193.
Dublanchet, A. and Bourne, S. 2007. The epic of phage therapy. Canadian Journal of
Infectious Diseases and Medical Microbiology. 18, 15-18.
Eaton, M. D. and Bayne-Jones, S. 1934. Bacteriophage therapy: Review of the principles
and results of the use of bacteriophage in the treatment of infections. Journal of
the American Medical Association. 103, 1769-1776.
Edlin, G., Lin, L., and Bitner, R. 1977. Reproductive fitness of p1, p2, and mu lysogens of
escherichia coli. Journal of Virology. 21, 560-564.
Elbreki, M., Ross, R. P., Hill, C., O'Mahony, J., McAuliffe, O., and Coffey, A. 2014.
Bacteriophages and their derivatives as biotherapeutic agents in disease
prevention and treatment. Journal of Viruses. 2014, 1-20.
Elbreki, M., Ross, R. P., Hill, C., O'Mahony, J., McAuliffe, O., and Coffey, A. 2014.
Bacteriophages and their derivatives as biotherapeutic agents in disease
prevention and treatment. Journal of Viruses. 2014.
Endersen, L., O'Mahony, J., Hill, C., Ross, R. P., McAuliffe, O., and Coffey, A. 2014. Phage
therapy in the food industry. Annual Review of Food Science and Technology. 5,
327-349.
136
Engel, A. S., Stern, L. A., and Bennett, P. C. 2004. Microbial contributions to cave
formation: New insights into sulfuric acid speleogenesis. Geology. 32, 369-372.
Erridge, C., Bennett-Guerrero, E., and Poxton, I. R. 2002. Structure and function of
lipopolysaccharides. Microbes and Infection. 4, 837-851.
Everett, M. J., Jin, Y. F., Ricci, V., and Piddock, L. J. 1996. Contributions of individual
mechanisms to fluoroquinolone resistance in 36 escherichia coli strains isolated
from humans and animals. Antimicrobial Agents and Chemotherapy. 40, 2380-
2386.
Feiner, R., Argov, T., Rabinovich, L., Sigal, N., Borovok, I., and Herskovits, A. A. 2015. A
new perspective on lysogeny: Prophages as active regulatory switches of
bacteria. Nature Reviews Microbiology. 13, 641-650.
Feiner, R., Argov, T., Rabinovich, L., Sigal, N., Borovok, I., and Herskovits, A. A. 2015. A
new perspective on lysogeny: Prophages as active regulatory switches of
bacteria. Nature reviews. Microbiology. 13, 641.
Ferguson, S., Roberts, C., Handy, E., and Sharma, M. 2013. Lytic bacteriophages reduce
escherichia coli o157: H7 on fresh cut lettuce introduced through cross-
contamination. Bacteriophage. 3, 1-7.
Filippov, A. A., Sergueev, K. V., He, Y., Huang, X.-Z., Gnade, B. T., Mueller, A. J.,
Fernandez-Prada, C. M., and Nikolich, M. P. 2011. Bacteriophage-resistant
mutants in yersinia pestis: Identification of phage receptors and attenuation for
mice. PLoS ONE. 6, e25486.
Floyd, J. L., Smith, K. P., Kumar, S. H., Floyd, J. T., and Varela, M. F. 2010. Lmrs is a
multidrug efflux pump of the major facilitator superfamily from staphylococcus
aureus. Antimicrobial Agents and Chemotherapy. 54, 5406-5412.
Ford, D. and Williams, P. D. 2013. Karst hydrogeology and geomorphology, p. Pages.
John Wiley & Sons.
Forest Department Sarawak. 2013. Annual report. Kuching, Sarawak Malaysia.
Fortier, L.-C. and Moineau, S. 2009. Phage production and maintenance of stocks,
including expected stock lifetimes, p. 203-219. Bacteriophages, Springer.
Fritsche, T. R., Castanheira, M., Miller, G. H., Jones, R. N., and Armstrong, E. S. 2008.
Detection of methyltransferases conferring high-level resistance to
137
aminoglycosides in enterobacteriaceae from europe, north america, and latin
america. Antimicrobial Agents and Chemotherapy. 52, 1843-1845.
Fu, W., Forster, T., Mayer, O., Curtin, J. J., Lehman, S. M., and Donlan, R. M. 2010.
Bacteriophage cocktail for the prevention of biofilm formation by pseudomonas
aeruginosa on catheters in an in-vitro model system. Antimicrobial Agents and
Chemotherapy. 54, 397-404.
Fuhrman, J. A. 1999. Marine viruses and their biogeochemical and ecological effects.
Nature. 399, 541-548.
Gadd, G. M. 2010. Metals, minerals and microbes: Geomicrobiology and
bioremediation. Microbiology. 156, 609-643.
Gallet, R., Kannoly, S., and Wang, I. N. 2011. Effects of bacteriophage traits on plaque
formation. BMC Microbiol. 11, 1-16.
Gao, W., Chua, K., Davies, J. K., Newton, H. J., Seemann, T., Harrison, P. F., Holmes, N.
E., Rhee, H.-W., Hong, J.-I., and Hartland, E. L. 2010. Two novel point mutations
in clinical staphylococcus aureus reduce linezolid susceptibility and switch on the
stringent response to promote persistent infection. PLoS Pathogens. 6,
e1000944.
García, M. S., De la Torre, M. Á., Morales, G., Peláez, B., Tolón, M. J., Domingo, S., Candel,
F. J., Andrade, R., Arribi, A., and García, N. 2010. Clinical outbreak of linezolid-
resistant staphylococcus aureus in an intensive care unit. JAMA. 303, 2260-2264.
Garcia, P., Martinez, B., Obeso, J. M., and Rodriguez, A. 2008. Bacteriophages and their
application in food safety. Letters in Applied Microbiology. 47, 479-485.
Gasiunas, G., Sinkunas, T., and Siksnys, V. 2014. Molecular mechanisms of crispr-
mediated microbial immunity. Cellular and Molecular Life Sciences. 71, 449-465.
Gaveau, D. L. A., Sloan, S., Molidena, E., Yaen, H., Sheil, D., Abram, N. K., Ancrenaz, M.,
Nasi, R., Quinones, M., and Wielaard, N. 2014. Four decades of forest
persistence, clearance and logging on borneo. PLoS ONE 9, 1-11.
Ghinsberg, R. C., Rogol, M., Sheinberg, Y., and Nitzan, Y. 1994. Monitoring of selected
bacteria and fungi in sand and sea water along the tel aviv coast. Microbios. 77,
29-40.
Ghosh, S., Kuisiene, N., and Cheeptham, N. 2016. The cave microbiome as a source for
drug discovery: Reality or pipe dream? Biochemical Pharmacology. 134, 18-34.
138
Gill, J. J. and Hyman, P. 2010. Phage choice, isolation, and preparation for phage therapy.
Current Pharmaceutical Biotechnology. 11, 2-14.
Gill, J. J. and Hyman, P. 2010. Phage choice, isolation, and preparation for phage therapy.
Current Pharmaceutical Biotechnology. 11, 2-14.
Giske, C. G., Fröding, I., Hasan, C. M., Turlej-Rogacka, A., Toleman, M., Livermore, D.,
Woodford, N., and Walsh, T. R. 2012. Diverse sequence types of klebsiella
pneumoniae contribute to the dissemination of blandm-1 in india, sweden, and
the united kingdom. Antimicrobial Agents and Chemotherapy. 56, 2735-2738.
Goldman, E. and Green, L. H. 2015. Practical handbook of microbiology, p. Pages. CRC
Press.
Goodridge, L. and Abedon, S. T. 2003. Bacteriophage biocontrol and bioprocessing:
Application of phage therapy to industry. SIM News. 53, 254-262.
Goodridge, L. D. 2010. Designing phage therapeutics. Current Pharmaceutical
Biotechnology. 11, 15-27.
Górski, A., Miedzybrodzki, R., Borysowski, J., Weber-Dabrowska, B., Lobocka, M.,
Fortuna, W., Letkiewicz, S., Zimecki, M., and Filby, G. 2009. Bacteriophage
therapy for the treatment of infections. Current Opinion in Investigational Drugs
(London, England: 2000). 10, 766-774.
Gorski, A., Międzybrodzki, R., Weber-Dąbrowska, B., Fortuna, W., Letkiewicz, S., Rogóż,
P., Jończyk-Matysiak, E., Dąbrowska, K., Majewska, J., and Borysowski, J. 2016.
Phage therapy: Combating infections with potential for evolving from merely a
treatment for complications to targeting diseases. Frontiers in Microbiology. 7,
1-9.
Gould, I. M. and Bal, A. M. 2013. New antibiotic agents in the pipeline and how they can
help overcome microbial resistance. Virulence. 4, 185-191.
Groth, I., Vettermann, R., Schuetze, B., Schumann, P., and Saiz-Jimenez, C. 1999.
Actinomycetes in karstic caves of northern spain (altamira and tito bustillo).
Journal of microbiological methods. 36, 115-122.
Gu, J., Liu, X., Li, Y., Han, W., Lei, L., Yang, Y., Zhao, H., Gao, Y., Song, J., and Lu, R. 2012.
A method for generation phage cocktail with great therapeutic potential. PLoS
One. 7, e31698.
139
Guenther, S., Huwyler, D., Richard, S., and Loessner, M. J. 2009. Virulent bacteriophage
for efficient biocontrol of listeria monocytogenes in ready-to-eat foods. Applied
and Environmental Microbiology. 75, 93-100.
Gupta, R. and Prasad, Y. 2011. Efficacy of polyvalent bacteriophage p-27/hp to control
multidrug resistant staphylococcus aureus associated with human infections.
Current Microbiology. 62, 255-260.
Gutiérrez, D., Rodríguez-Rubio, L., Martínez, B., Rodríguez, A., and García, P. 2016.
Bacteriophages as weapons against bacterial biofilms in the food industry.
Frontiers in Microbiology. 7, 1-15.
Guttman, B., Raya, R., and Kutter, E. 2005. Bacteriophages: Biology and applications, p.
Pages. CRC Press.
Guttman, B., Raya, R., and Kutter, E. 2005. Basic phage biology. Bacteriophages: Biology
and applications. 4.
Hall, A. R., De Vos, D., Friman, V.-P., Pirnay, J.-P., and Buckling, A. 2012. Effects of
sequential and simultaneous applications of bacteriophages on populations of
pseudomonas aeruginosa in-vitro and in wax moth larvae. Applied and
Environmental Microbiology. 78, 5646-5652.
Hall, B. G., Acar, H., Nandipati, A., and Barlow, M. 2013. Growth rates made easy.
Molecular Biology and Evolution. 31, 232-238.
Hambly, E. and Suttle, C. A. 2005. The viriosphere, diversity, and genetic exchange within
phage communities. Current opinion in microbiology. 8, 444-450.
Harshey, R. M. 2012. The mu story: How a maverick phage moved the field forward.
Mobile DNA. 3, 1-9.
Hartmann, J., Beyer, R., and Harm, S. 2014. Effective removal of estrogens from drinking
water and wastewater by adsorption technology. Environmental Processes. 1,
87-94.
Harvey, A. 2000. Strategies for discovering drugs from previously unexplored natural
products. Drug Discovery Today. 5, 294-300.
Hawkins, C., Harper, D., Burch, D., Änggård, E., and Soothill, J. 2010. Topical treatment
of pseudomonas aeruginosa otitis of dogs with a bacteriophage mixture: A
before/after clinical trial. Veterinary Microbiology. 146, 309-313.
140
Hermoso, J. A., García, J. L., and García, P. 2007. Taking aim on bacterial pathogens: From
phage therapy to enzybiotics. Current opinion in microbiology. 10, 461-472.
Herold, K., Gollmick, F. A., Groth, I., Roth, M., Menzel, K. D., Möllmann, U., Gräfe, U., and
Hertweck, C. 2005. Cervimycin a–d: A polyketide glycoside complex from a cave
bacterium can defeat vancomycin resistance. Chemistry-A European Journal. 11,
5523-5530.
Herold, K., Xu, Z., Gollmick, F. A., Gräfe, U., and Hertweck, C. 2004. Biosynthesis of
cervimycin c, an aromatic polyketide antibiotic bearing an unusual
dimethylmalonyl moiety. Organic & Biomolecular Chemistry. 2, 2411-2414.
Hidalgo, L., Hopkins, K. L., Gutierrez, B., Ovejero, C. M., Shukla, S., Douthwaite, S.,
Prasad, K. N., Woodford, N., and Gonzalez-Zorn, B. 2013. Association of the novel
aminoglycoside resistance determinant rmtf with ndm carbapenemase in
enterobacteriaceae isolated in india and the uk. Journal of Antimicrobial
Chemotherapy. 68, 1543-1550.
Hu, R.-M., Liao, S.-T., Huang, C.-C., Huang, Y.-W., and Yang, T.-C. 2012. An inducible
fusaric acid tripartite efflux pump contributes to the fusaric acid resistance in
stenotrophomonas maltophilia. PLoS ONE. 7, e51053.
Hyman, P. and Abedon, S. T. 2010. Bacteriophage host range and bacterial resistance.
Advances in Applied Microbiology. 70, 217-248.
Ingraham, J., L., Maaløe, O., and Neidhardt, F. C. 1983. Growth of the bacterial cell, p.
Pages. Sinauer Associates.
Jaiswal, A., Koley, H., Ghosh, A., Palit, A., and Sarkar, B. 2013. Efficacy of cocktail phage
therapy in treating vibrio cholerae infection in rabbit model. Microbes and
Infection. 15, 152-156.
Jassim, S. A. A. and Limoges, R. G. 2017. Bacteriophages: Practical applications for
nature's biocontrol, p. Pages. Springer, Cham, Switzerland.
Jassim, S. A. A. and Limoges, R. G. 2014. Natural solution to antibiotic resistance:
Bacteriophages ‘the living drugs’. World Journal of Microbiology and
Biotechnology. 30, 2153-2170.
Jassim, S. A. A., Limoges, R. G., and El-Cheikh, H. 2016. Bacteriophage biocontrol in
wastewater treatment. World Journal of Microbiology and Biotechnology. 32, 1-
10.
141
Jennifer, E. 2006. Bacteriophages of burkholderia pseudomallei: Friend or foe? James
Cook University.
Jensen, K. C., Hair, B. B., Wienclaw, T. M., Murdock, M. H., Hatch, J. B., Trent, A. T., White,
T. D., Haskell, K. J., and Berges, B. K. 2015. Isolation and host range of
bacteriophage with lytic activity against methicillin-resistant staphylococcus
aureus and potential use as a fomite decontaminant. PLOS ONE. 10, e0131714.
Jiang, Z.-k., Guo, L., Chen, C., Liu, S.-w., Zhang, L., Dai, S.-j., He, Q.-y., You, X.-f., Hu, X.-x.,
and Tuo, L. 2015. Xiakemycin a, a novel pyranonaphthoquinone antibiotic,
produced by the streptomyces sp. Cc8-201 from the soil of a karst cave. The
Journal of Antibiotics. 68, 771-774.
Jiang, Z.-k., Guo, L., Chen, C., Liu, S.-w., Zhang, L., Dai, S.-j., He, Q.-y., You, X.-f., Hu, X.-x.,
and Tuo, L. 2015. Xiakemycin a, a novel pyranonaphthoquinone antibiotic,
produced by the streptomyces sp. Cc8-201 from the soil of a karst cave. Journal
of Antibiotics. 68, 771.
Johnson, A. P. and Woodford, N. 2013. Global spread of antibiotic resistance: The
example of new delhi metallo-β-lactamase (ndm)-mediated carbapenem
resistance. Journal of medical microbiology. 62, 499-513.
Jones, J. B., Jackson, L. E., Balogh, B., Obradovic, A., Iriarte, F. B., and Momol, M. T. 2007.
Bacteriophages for plant disease control. Annual Review of Phytopathology. 45,
245-262.
Julaihi, L. C. J. 2004. Altitudinal analyses of limestone vegetation of gunung api, gunung
mulu national park, miri, sarawak. Universiti Malaysia Sarawak, Sarawak,
Malaysia.
Jurado, V., Novakova, A., Hernández Mariné, M., and Saiz-Jimenez, C. 2014. The
conservation of subterranean cultural heritage, p. Pages. CRC Press/Balkema,
Leiden, The Netherlands Sevilla, Spain
Katayama, Y., Ito, T., and Hiramatsu, K. 2000. A new class of genetic element,
staphylococcus cassette chromosome mec, encodes methicillin resistance in
staphylococcus aureus. Antimicrobial Agents and Chemotherapy. 44, 1549-1555.
Keller, R. and Engley, J. R. F. B. 1958. Fate of bacteriophage particles introduced into
mice by various routes. Proceedings of the Society for Experimental Biology and
Medicine. 98, 577-580.
142
Kelly, D., McAuliffe, O., O’Mahony, J., and Coffey, A. 2011. Development of a broad-host-
range phage cocktail for biocontrol. Bioengineered Bugs. 2, 31-37.
Keong, C. Y. 2015. Sustainable resource management and ecological conservation of
mega-biodiversity: The southeast asian big-3 reality. International Journal of
Environmental Science and Development. 6, 876.
Khan, M. A., Satoh, H., Katayama, H., Kurisu, F., and Mino, T. 2002. Bacteriophages
isolated from activated sludge processes and their polyvalency. Water research
36, 3364-3370.
Khan, M. A., Satoh, H., Katayama, H., Kurisu, F., and Mino, T. 2002. Bacteriophages
isolated from activated sludge processes and their polyvalency. Water Research.
36, 3364-3370.
Klumpp, J., Lavigne, R., Loessner, M. J., and Ackermann, H.-W. 2010. The spo1-related
bacteriophages. Archives of Virology. 155, 1547-1561.
Kojima, S. and Nikaido, H. 2013. Presented at the Proceedings of the National Academy
of Sciences of the United States of America.
Kolter, R. and Greenberg, E. P. 2006. Microbial sciences: The superficial life of microbes.
Nature. 441, 300-302.
Kosmidis, C., Schindler, B. D., Jacinto, P. L., Patel, D., Bains, K., Seo, S. M., and Kaatz, G.
W. 2012. Expression of multidrug resistance efflux pump genes in clinical and
environmental isolates of staphylococcus aureus. International Journal of
Antimicrobial Agents. 40, 204-209.
Kramberger, P., Honour, R. C., Herman, R. E., Smrekar, F., and Peterka, M. 2010.
Purification of the staphylococcus aureus bacteriophages vdx-10 on
methacrylate monoliths. Journal of Virological Methods. 166, 60-64.
Kropinski, A. M., Mazzocco, A., Waddell, T. E., Lingohr, E., and Johnson, R. P. 2009.
Enumeration of bacteriophages by double agar overlay plaque assay, p. 69-76.
In M. R. Clokie and A. M. Kropinski (eds.), Bacteriophages: Methods and
protocols, volume 1: Isolation, characterization, and interactions, Humana Press.
Kropinski, A. M., Mazzocco, A., Waddell, T. E., Lingohr, E., and Johnson, R. P. 2009.
Enumeration of bacteriophages by double agar overlay plaque assay.
Bacteriophages: Methods and Protocols, Volume 1: Isolation, Characterization,
and Interactions. 69-76.
143
Krueger, A. P. and Scribner, E. J. 1941. The bacteriophage: Its nature and its therapeutic
use. Journal of the American Medical Association. 116, 2269-2277.
Krügel, H., Licht, A., Biedermann, G., Petzold, A., Lassak, J., Hupfer, Y., Schlott, B.,
Hertweck, C., Platzer, M., and Brantl, S. 2010. Cervimycin c resistance in bacillus
subtilis is due to a promoter up-mutation and increased mrna stability of the
constitutive abc-transporter gene bmra. FEMS Microbiology Letters. 313, 155-
163.
Kuek, F. W. I., Lim, L.-F., Ngu, L.-H., Mujahid, A., Lim, P.-T., Leaw, C.-P., and Müller, M.
2015. The potential roles of bacterial communities in coral defence: A case study
at talang-talang reef. Ocean Science Journal. 50, 1-14.
Kumar, N., Radhakrishnan, A., Wright, C. C., Chou, T. H., Lei, H. T., Bolla, J. R., Tringides,
M. L., Rajashankar, K. R., Su, C. C., and Purdy, G. E. 2014. Crystal structure of the
transcriptional regulator rv1219c of mycobacterium tuberculosis. Protein
Science. 23, 423-432.
Kumar, P. B., Kannan, M. M., and Quine, S. D. 2011. Litsea deccanensis ameliorates
myocardial infarction in wistar rats: Evidence from biochemical histological
studies. Journal of Young Pharmacists. 3, 287-296.
Kumar, S., Tripathi, V. R., and Garg, S. K. 2013. Antibiotic resistance and genetic diversity
in water-borne enterobacteriaceae isolates from recreational and drinking water
sources. International Journal of Environmental Science and Technology. 10, 789-
798.
Kumarasamy, K. and Kalyanasundaram, A. 2011. Emergence of klebsiella pneumoniae
isolate co-producing ndm-1 with kpc-2 from india. Journal of Antimicrobial
Chemotherapy. 67, 243-244.
Kumarasamy, K. K., Toleman, M. A., Walsh, T. R., Bagaria, J., Butt, F., Balakrishnan, R.,
Chaudhary, U., Doumith, M., Giske, C. G., and Irfan, S. 2010. Emergence of a new
antibiotic resistance mechanism in india, pakistan, and the uk: A molecular,
biological, and epidemiological study. The Lancet Infectious Diseases. 10, 597-
602.
Kusradze, I., Karumidze, N., Rigvava, S., Dvalidze, T., Katsitadze, M., Amiranashvili, I., and
Goderdzishvili, M. 2016. Characterization and testing the efficiency of
144
acinetobacter baumannii phage vb-gec_ab-m-g7 as an antibacterial agent.
Frontiers in Microbiology. 7, 1-7.
Kutateladze, M. 2015. Experience of the eliava institute in bacteriophage therapy.
Virologica Sinica. 30, 80-81.
Kutateladze, M. and Adamia, R. 2008. Phage therapy experience at the eliava institute.
Médecine et maladies infectieuses. 38, 426-430.
Kutter, E. 2008. Phage therapy: Bacteriophages as naturally occurring antimicrobials.
Practical Handbook of Microbiology. 713-730.
Kutter, E., De Vos, D., Gvasalia, G., Alavidze, Z., Gogokhia, L., Kuhl, S., and Abedon, S. T.
2010. Phage therapy in clinical practice: Treatment of human infections. Current
Pharmaceutical Biotechnology. 11, 69-86.
Laanto, E., Bamford, J. K. H., Laakso, J., and Sundberg, L.-R. 2012. Phage-driven loss of
virulence in a fish pathogenic bacterium. PLoS ONE. 7, e53157.
LaBauve, A. E. and Wargo, M. J. 2012. Growth and laboratory maintenance of
pseudomonas aeruginosa. Current protocols in microbiology. 6E. 1.1-6E. 1.8.
Labrie, S. J., Samson, J. E., and Moineau, S. 2010. Bacteriophage resistance mechanisms.
Nature Reviews Microbiology. 8, 317-327.
Lamprinou, V., Tryfinopoulou, K., Velonakis, E. N., Vatopoulos, A., Antonopoulou, S.,
Fragopoulou, E., Pantazidou, A., and Economou-Amilli, A. 2015. Cave
cyanobacteria showing antibacterial activity. International Journal of Speleology.
44, 231-238.
Lange, R. and Hengge-Aronis, R. 1991. Growth phase-regulated expression of bola and
morphology of stationary-phase escherichia coli cells are controlled by the novel
sigma factor sigma s. Journal of Bacteriology. 173, 4474-4481.
Larsson, D. G. J., de Pedro, C., and Paxeus, N. 2007. Effluent from drug manufactures
contains extremely high levels of pharmaceuticals. Journal of Hazardous
Materials. 148, 751-755.
Lateef, A. A., Sepiah, M., and Bolhassan, M. H. 2014. Presented at the Proceedings of
International Conference on Beneficial Microbes (ICOBM 2014): Micobes for the
Benefit of Mankind, ParkRoyal Penang Resort, Penang, Malaysia.
145
Lateef, A. A., Sepiah, M., and Bolhassan, M. H. 2014. Presented at the Proceedings of
the International Conference on Beneficial Microbes ICOBM 2014: Microbes for
the Benefits of Mankind, Penang, Malaysia.
Lavigne, J.-P., Sotto, A., Nicolas-Chanoine, M.-H., Bouziges, N., Pagès, J.-M., and Davin-
Regli, A. 2013. An adaptive response of enterobacter aerogenes to imipenem:
Regulation of porin balance in clinical isolates. International Journal of
Antimicrobial Agents. 41, 130-136.
Leclercq, R. 2002. Mechanisms of resistance to macrolides and lincosamides: Nature of
the resistance elements and their clinical implications. Clinical Infectious
Diseases. 34, 482-492.
Lee, C.-R., Lee, J. H., Park, K. S., Kim, Y. B., Jeong, B. C., and Lee, S. H. 2016. Global
dissemination of carbapenemase-producing klebsiella pneumoniae:
Epidemiology, genetic context, treatment options, and detection methods.
Frontiers in Microbiology. 7, 1-30.
Lekshmi, M., Ammini, P., Kumar, S., and Varela, M. F. 2017. The food production
environment and the development of antimicrobial resistance in human
pathogens of animal origin. Microorganisms. 5, 11.
Lengeler, J. W., Drews, G., and Schlegel, H. G. 1999. Biology of the prokaryotes, p. Pages.
Georg Thieme Verlag.
Levin, B. R. and Bull, J. J. 2004. Population and evolutionary dynamics of phage therapy.
Nature Reviews Microbiology. 2, 166-173.
Lim, L. M., Ly, N., Anderson, D., Yang, J. C., Macander, L., Jarkowski, A., Forrest, A.,
Bulitta, J. B., and Tsuji, B. T. 2010. Resurgence of colistin: A review of resistance,
toxicity, pharmacodynamics, and dosing. Pharmacotherapy: The Journal of
Human Pharmacology and Drug Therapy. 30, 1279-1291.
Lin, J., Nishino, K., Roberts, M. C., Tolmasky, M., Aminov, R. I., and Zhang, L. 2015.
Mechanisms of antibiotic resistance. Frontiers in Microbiology. 6, 1-3.
Lin, T.-Y., Lo, Y.-H., Tseng, P.-W., Chang, S.-F., Lin, Y.-T., and Chen, T.-S. 2012. A t3 and t7
recombinant phage acquires efficient adsorption and a broader host range. PLoS
ONE. 7, e30954.
Liu, S. 2014. Presented at the Symposium: Student Journal of Science & Math
146
Livermore, D. M. 2008. Defining an extended‐spectrum β‐lactamase. Clinical
Microbiology and Infection. 14, 3-10.
Lobocka, M., Hejnowicz, M., Gagala, U., Weber-Dabrowska, B., Wegrzyn, G., and Dadlez,
M. 2014. The first step to bacteriophage therapy: How to choose the correct
phage. Phage Therapy: Current Research and Applications.
Lobocka, M., Hejnowicz, M. S., Gkagała, U., Weber-Dkabrowska, B., Wkegrzyn, G., and
Dadlez, M. 2014. The first step to bacteriophage therapy – how to choose the
correct phage, p. 378. In J. Borysowski, R. Mikedzybrodzki and A. Górski (eds.),
Phage therapy: Current research and applications, Caister Academic Press,
Norfolk, England.
Loc-Carrillo, C. and Abedon, S. T. 2011. Pros and cons of phage therapy. Bacteriophage.
1, 111-114.
Long, K. S., Poehlsgaard, J., Kehrenberg, C., Schwarz, S., and Vester, B. 2006. The cfr rrna
methyltransferase confers resistance to phenicols, lincosamides, oxazolidinones,
pleuromutilins, and streptogramin a antibiotics. Antimicrobial Agents and
Chemotherapy. 50, 2500-2505.
López-Bueno, A., Tamames, J., Velázquez, D., Moya, A., Quesada, A., and Alcamí, A.
2009. High diversity of the viral community from an antarctic lake. Science. 326,
858-861.
Łoś, J. M., Golec, P., Węgrzyn, G., Węgrzyn, A., and Łoś, M. 2008. Simple method for
plating escherichia coli bacteriophages forming very small plaques or no plaques
under standard conditions. Applied and Environmental Microbiology. 74, 5113-
5120.
Łoś, M. and Węgrzyn, G. 2012. Pseudolysogeny, p. Pages.
Lu, T. K. and Collins, J. J. 2007. Dispersing biofilms with engineered enzymatic
bacteriophage. Proceedings of the National Academy of Sciences. 104, 11197-
11202.
Lu, T. K. and Koeris, M. S. 2011. The next generation of bacteriophage therapy. Current
Opinion in Microbiology. 14, 524-531.
Lynch III, J. P., Clark, N. M., and Zhanel, G. G. 2013. Evolution of antimicrobial resistance
among enterobacteriaceae (focus on extended spectrum β-lactamases and
carbapenemases). Expert opinion on pharmacotherapy. 14, 199-210.
147
Macalady, J. L., Jones, D. S., and Lyon, E. H. 2007. Extremely acidic, pendulous cave wall
biofilms from the frasassi cave system, italy. Environmental Microbiology. 9,
1402-1414.
Magnone, J. P., Marek, P. J., Sulakvelidze, A., and Senecal, A. G. 2013. Additive approach
for inactivation of escherichia coli o157: H7, salmonella, and shigella spp. On
contaminated fresh fruits and vegetables using bacteriophage cocktail and
produce wash. Journal of Food Protection. 76, 1336-1341.
Maier, R. M., Pepper, I. L., and Gerba, C. P. 2009. Environmental microbiology, p. Pages.
Academic Press.
Malik, D. J., Sokolov, I. J., Vinner, G. V., Mancuso, F., Cinquerrui, S., Vladisavljevic, G. T.,
Clokie, M. R. J., Garton, N. J., Stapley, A. G. F., and Kirpichnikova, A. 2017.
Formulation, stabilisation and encapsulation of bacteriophage for phage
therapy. Advances in Colloid and Interface Science. 249, 100-133.
Malki, K., Kula, A., Bruder, K., Sible, E., Hatzopoulos, T., Steidel, S., Watkins, S. C., and
Putonti, C. 2015. Bacteriophages isolated from lake michigan demonstrate broad
host-range across several bacterial phyla. Virol J. 12, 1-5.
Maniloff, J. and Ackermann, H.-W. 1998. Taxonomy of bacterial viruses: Establishment
of tailed virus genera and the other caudovirales. Archives of Virology. 143, 2051-
2063.
Mann, N. H. 2008. The potential of phages to prevent mrsa infections. Research in
Microbiology. 159, 400-405.
Mapes, A. C., Trautner, B. W., Liao, K. S., and Ramig, R. F. 2016. Development of
expanded host range phage active on biofilms of multi-drug resistant
pseudomonas aeruginosa. Bacteriophage. 6.
Marcó, M. B., Reinheimer, J. A., and Quiberoni, A. 2010. Phage adsorption to
lactobacillus plantarum: Influence of physiological and environmental factors.
International Journal of Food Microbiology. 138, 270-275.
Maria de Lurdes, N. 2013. Cave biofilms and their potential for novel antibiotic
discovery, p. 35-45. Cave microbiomes: A novel resource for drug discovery,
Springer, New York, NY, SpringerBriefs in Microbiology.
Marraffini, L. A. 2015. Crispr-cas immunity in prokaryotes. Nature. 526, 55-61.
148
Marraffini, L. A. and Sontheimer, E. J. 2008. Crispr interference limits horizontal gene
transfer in staphylococci by targeting DNA. Science. 322, 1843-1845.
Matsuzaki, S., Rashel, M., Uchiyama, J., Sakurai, S., Ujihara, T., Kuroda, M., Ikeuchi, M.,
Tani, T., Fujieda, M., and Wakiguchi, H. 2005. Bacteriophage therapy: A
revitalized therapy against bacterial infectious diseases. Journal of Infection and
Chemotherapy. 11, 211-219.
Mazzocco, A., Waddell, T. E., Lingohr, E., and Johnson, R. P. 2009. Enumeration of
bacteriophages using the small drop plaque assay system, p. 81-85. In M. R.
Clokie and A. M. Kropinski (eds.), Bacteriophages: Methods and protocols,
volume 1: Isolation, characterization, and interactions, Humana Press.
McAuliffe, O., Ross, R. P., and Fitzgerald, G. F. 2007. The new phage biology: From
genomics to applications, p. Pages. Caister Academic Press, Norfolk.
McCarville, J. L., Caminero, A., and Verdu, E. F. 2016. Novel perspectives on therapeutic
modulation of the gut microbiota. Therapeutic Advances in Gastroenterology. 9,
580-593.
McLaughlin, M. and Balaa, M. 2006. Enhanced contrast of bacteriophage plaques in
salmonella with ferric ammonium citrate and sodium thiosulfate (facst) and
tetrazolium red (tzr). Journal of Microbiological Methods. 65, 318-323.
McLaughlin, M. R. and Brooks, J. P. 2008. Epa worst case water microcosms for testing
phage biocontrol of salmonella. Journal of Environmental Quality. 37, 266-271.
Medina, E. and Pieper, D. H. 2016. Tackling threats and future problems of multidrug-
resistant bacteria, p. Pages., Cham, Switzerland.
Medina, E. and Pieper, D. H. 2016. Tackling threats and future problems of multidrug-
resistant bacteria. How to Overcome the Antibiotic Crisis: Facts, Challenges,
Technologies and Future Perspectives. 3-33.
Melo, L. D. R., Oliveira, H., Santos, S. B., Sillankorva, S., and Azeredo, J. 2017. Phages
against infectious diseases, p. 269-294. In R. Paterson and N. Lima (eds.),
Bioprospecting: Success, potential and constraints, Springer, Cham, Germany.
Merabishvili, M., Pirnay, J.-P., Verbeken, G., Chanishvili, N., Tediashvili, M., Lashkhi, N.,
Glonti, T., Krylov, V., Mast, J., Van Parys, L., Lavigne, R., Volckaert, G., Mattheus,
W., Verween, G., De Corte, P., Rose, T., Jennes, S., Zizi, M., De Vos, D., and
Vaneechoutte, M. 2009. Quality-controlled small-scale production of a well-
149
defined bacteriophage cocktail for use in human clinical trials. PLoS ONE. 4,
e4944.
Merabishvili, M., Vandenheuvel, D., Kropinski, A. M., Mast, J., De Vos, D., Verbeken, G.,
Noben, J. P., Lavigne, R., Vaneechoutte, M., and Pirnay, J. P. 2014.
Characterization of newly isolated lytic bacteriophages active against
acinetobacter baumannii. PLoS One. 9, 1-11.
Merril, C. R., Scholl, D., and Adhya, S. L. 2003. The prospect for bacteriophage therapy
in western medicine. Nature Reviews Drug discovery. 2, 489-497.
Michael, C. A., Dominey-Howes, D., and Labbate, M. 2014. The antimicrobial resistance
crisis: Causes, consequences, and management. Frontiers in Public Health. 2, 1-
8.
Miedzybrodzki, R., Borysowski, J., Weber-Dabrowska, B., Fortuna, W., Letkiewicz, S.,
Szufnarowski, K., Pawelczyk, Z., Rogóz, P., Klak, M., and Wojtasik, E. 2012. Clinical
aspects of phage therapy, p. Pages.
Miller, A. K., Brannon, M. K., Stevens, L., Johansen, H. K., Selgrade, S. E., Miller, S. I.,
Høiby, N., and Moskowitz, S. M. 2011. Phoq mutations promote lipid a
modification and polymyxin resistance of pseudomonas aeruginosa found in
colistin-treated cystic fibrosis patients. Antimicrobial Agents and Chemotherapy.
55, 5761-5769.
Miller, R. V. and Ripp, S. A. 2002. Pseudolysogeny: A bacteriophage strategy for
increasing longevity in situ, p. 81-91. In M. Syvanen and C. I. Kado (eds.),
Horizontal gene transfer, Academic Press.
Miyashita, N. T., Iwanaga, H., Charles, S., Diway, B., Sabang, J., and Chong, L. 2013. Soil
bacterial community structure in five tropical forests in malaysia and one
temperate forest in japan revealed by pyrosequencing analyses of 16s rrna gene
sequence variation. Genes & Genetic Systems. 88, 93-103.
Mizoguchi, K., Morita, M., Fischer, C. R., Yoichi, M., Tanji, Y., and Unno, H. 2003.
Coevolution of bacteriophage pp01 and escherichia coli o157: H7 in continuous
culture. Applied and Environmental Microbiology. 69, 170-176.
Moellering Jr, R. C. 2010. Ndm-1—a cause for worldwide concern. New England Journal
of Medicine. 363, 2377-2379.
150
Mohammed, T. A. 2013. Isolation, characterization and application of calcite producing
bacteria from urea rich soils. Islamic University of Gaza, Gaza, Palestine.
Mohd, R. A. R., Roberta, C. T. T., Mohd, I. A., Noor, H. H., and Mohd, T. A. 2011. Bats of
the wind cave nature reserve, sarawak, malaysian borneo Tropical Natural
History 11, 159-175.
Monk, A. B., Rees, C. D., Barrow, P., Hagens, S., and Harper, D. R. 2010. Bacteriophage
applications: Where are we now? Letters in Applied Microbiology. 51, 363-369.
Moseley, G. E., Richards, D. A., Smith, C. J. M., Smart, P. L., Hoffmann, D. L., and Farrant,
A. R. 2013. U–th dating of speleothems to investigate the evolution of limestone
caves in the gunung mulu national park, sarawak, malaysia. Cave and Karst
Science. 40, 13-16.
Myers, N., Mittermeier, R. A., Mittermeier, C. G., Da Fonseca, G. A. B., and Kent, J. 2000.
Biodiversity hotspots for conservation priorities. Nature. 403, 853-858.
Nagel, T. E., Chan, B. K., De Vos, D., El-Shibiny, A., Kang'ethe, E. K., Makumi, A., and
Pirnay, J.-P. 2016. The developing world urgently needs phages to combat
pathogenic bacteria. Frontiers in Microbiology. 7, 1-4.
Nicastro, J., Wong, S., Khazaei, Z., Lam, P., Blay, J., and Slavcev, R. A. 2016. Bacteriophage
applications-historical perspective and future potential, p. Pages. Springer,
Cham, Switzerland.
Nicolaus, B., Kambourova, M., and Oner, E. T. 2010. Exopolysaccharides from
extremophiles: From fundamentals to biotechnology. Environmental
Technology. 31, 1145-1158.
Nilsson, A. S. 2014. Phage therapy—constraints and possibilities. Upsala Journal of
Medical Sciences. 119, 192-198.
Nimaichand, S., Devi, A. M., Tamreihao, K., Ningthoujam, D. S., and Li, W.-J. 2015.
Actinobacterial diversity in limestone deposit sites in hundung, manipur (india)
and their antimicrobial activities. Frontiers in Microbiology. 6, 1-10.
Nordmann, P., Poirel, L., Carrër, A., Toleman, M. A., and Walsh, T. R. 2011. How to detect
ndm-1 producers. Journal of Clinical Microbiology. 49, 718-721.
Nordmann, P., Poirel, L., Walsh, T. R., and Livermore, D. M. 2011. The emerging ndm
carbapenemases. Trends in Microbiology. 19, 588-595.
151
Northup, D. E. and Lavoie, K. H. 2001. Geomicrobiology of caves:A review.
Geomicrobiology Journal. 18, 199-222.
Novais, A., Rodrigues, C., Branquinho, R., Antunes, P., Grosso, F., Boaventura, L., Ribeiro,
G., and Peixe, L. 2012. Spread of an ompk36-modified st15 klebsiella pneumoniae
variant during an outbreak involving multiple carbapenem-resistant
enterobacteriaceae species and clones. European Journal of Clinical
Microbiology & Infectious Diseases. 31, 3057-3063.
O'Flaherty, S., Ross, R. P., and Coffey, A. 2009. Bacteriophage and their lysins for
elimination of infectious bacteria. FEMS Microbiology Reviews. 33, 801-819.
O'flaherty, S., Ross, R. P., Meaney, W., Fitzgerald, G. F., Elbreki, M. F., and Coffey, A.
2005. Potential of the polyvalent anti-staphylococcus bacteriophage k for control
of antibiotic-resistant staphylococci from hospitals. Applied and Environmental
Microbiology. 71, 1836-1842.
O'flynn, G., Ross, R. P., Fitzgerald, G. F., and Coffey, A. 2004. Evaluation of a cocktail of
three bacteriophages for biocontrol of escherichia coli o157: H7. Applied and
Environmental Microbiology. 70, 3417-3424.
O'grady, N. P., Alexander, M., Dellinger, E. P., Gerberding, J. L., Heard, S. O., Maki, D. G.,
Masur, H., McCormick, R. D., Mermel, L. A., and Pearson, M. L. 2002. Guidelines
for the prevention of intravascular catheter–related infections. Clinical Infectious
Diseases. 35, 1281-1307.
O’Neill, J. 2014. Antimicrobial resistance: Tackling a crisis for the health and wealth of
nations.
Obeso, J. M., Martínez, B., Rodríguez, A., and García, P. 2008. Lytic activity of the
recombinant staphylococcal bacteriophage φh5 endolysin active against
staphylococcus aureus in milk. International Journal of Food Microbiology. 128,
212-218.
Ogawa, W., Onishi, M., Ni, R., Tsuchiya, T., and Kuroda, T. 2012. Functional study of the
novel multidrug efflux pump kexd from klebsiella pneumoniae. Gene. 498, 177-
182.
Ogikubo, Y., Norimatsu, M., Noda, K., Takahashi, J., Inotsume, M., Tsuchiya, M., and
Tamura, Y. 2004. Evaluation of the bacterial endotoxin test for quantification of
endotoxin contamination of porcine vaccines. Biologicals. 32, 88-93.
152
Omoregie, A. I. 2016. Characterization of ureolytic bacteria isolated from limestone
caves of sarawak and evaluation of their efficiency in biocementation. Swinburne
University of Technology (Sarawak Campus), Malaysia.
Orlandi, V. T., Bolognese, F., Chiodaroli, L., Tolker-Nielsen, T., and Barbieri, P. 2015.
Pigments influence the tolerance of pseudomonas aeruginosa pao1 to
photodynamically induced oxidative stress. Microbiology. 161, 2298-2309.
Örmälä, A.-M. and Jalasvuori, M. 2013. Phage therapy: Should bacterial resistance to
phages be a concern, even in the long run? Bacteriophage. 3, e24219-24211-
e24219-24213.
Papagiannitsis, C. C., Giakkoupi, P., Kotsakis, S. D., Tzelepi, E., Tzouvelekis, L. S.,
Vatopoulos, A. C., and Miriagou, V. 2013. Ompk35 and ompk36 porin variants
associated with specific sequence types of klebsiella pneumoniae. Journal of
Chemotherapy. 25, 250-254.
Pardon, E., Laeremans, T., Triest, S., Rasmussen, S. G. F., Wohlkönig, A., Ruf, A.,
Muyldermans, S., Hol, W. G. J., Kobilka, B. K., and Steyaert, J. 2014. A general
protocol for the generation of nanobodies for structural biology. Nature
protocols. 9, 674-693.
Parisien, A., Allain, B., Zhang, J., Mandeville, R., and Lan, C. Q. 2008. Novel alternatives
to antibiotics: Bacteriophages, bacterial cell wall hydrolases, and antimicrobial
peptides. Journal of Applied Microbiology. 104, 1-13.
Parmar, K. M., Hathi, Z. J., and Dafale, N. A. 2017. Control of multidrug-resistant gene
flow in the environment through bacteriophage intervention. Applied
Biochemistry and Biotechnology. 181, 1007-1029.
Pašić, L., Kovče, B., Sket, B., and Herzog-Velikonja, B. 2009. Diversity of microbial
communities colonizing the walls of a karstic cave in slovenia. FEMS Microbiology
Ecology. 71, 50-60.
Pattee, P. A. 1966. Use of tetrazolium for improved resolution of bacteriophage plaques.
Journal of Bacteriology. 92, 787-788.
Payne, R. J. and Jansen, V. A. 2001. Understanding bacteriophage therapy as a density-
dependent kinetic process. Journal of Theoretical Biology. 208, 37-48.
Payne, R. J. H. and Jansen, V. A. A. 2001. Understanding bacteriophage therapy as a
density-dependent kinetic process. Journal of Theoretical Biology. 208, 37-48.
153
Pedulla, M. L., Ford, M. E., Houtz, J. M., Karthikeyan, T., Wadsworth, C., Lewis, J. A.,
Jacobs-Sera, D., Falbo, J., Gross, J., and Pannunzio, N. R. 2003. Origins of highly
mosaic mycobacteriophage genomes. Cell. 113, 171-182.
Pendleton, J. N., Gorman, S. P., and Gilmore, B. F. 2013. Clinical relevance of the eskape
pathogens. Expert Review of Anti-Infective Therapy. 11, 297-308.
Piddock, L. J. V. 2006. Clinically relevant chromosomally encoded multidrug resistance
efflux pumps in bacteria. Clinical Microbiology Reviews. 19, 382-402.
Pleška, M., Qian, L., Okura, R., Bergmiller, T., Wakamoto, Y., Kussell, E., and Guet, C. C.
2016. Bacterial autoimmunity due to a restriction-modification system. Current
Biology. 26, 404-409.
Poirel, L., Bonnin, R. A., and Nordmann, P. 2012. Genetic support and diversity of
acquired extended-spectrum β-lactamases in gram-negative rods. Infection,
Genetics and Evolution. 12, 883-893.
Poulou, A., Voulgari, E., Vrioni, G., Koumaki, V., Xidopoulos, G., Chatzipantazi, V.,
Markou, F., and Tsakris, A. 2013. Outbreak caused by an ertapenem-resistant,
ctx-m-15-producing klebsiella pneumoniae sequence type 101 clone carrying an
ompk36 porin variant. Journal of Clinical Microbiology. 51, 3176-3182.
Prevention, U. S. D. o. H. a. H. S. C. f. D. C. a. 2013. Antibiotic resistance threats in the
united states, 2013.
Pruden, A., Larsson, D. G. J., Amézquita, A., Collignon, P., Brandt, K. K., Graham, D. W.,
Lazorchak, J. M., Suzuki, S., Silley, P., and Snape, J. R. 2013. Management options
for reducing the release of antibiotics and antibiotic resistance genes to the
environment. Environmental Health Perspectives. 121, 878-885.
Pruden, A., Pei, R., Storteboom, H., and Carlson, K. H. 2006. Antibiotic resistance genes
as emerging contaminants: Studies in northern colorado. Environmental Science
& Technology. 40, 7445-7450.
Ptashne, M. 2004. A genetic switch: Phage lambda revisited, p. Pages. Cold Spring
Harbor Laboratory Press.
Pumbwe, L. and Piddock, L. J. V. 2000. Two efflux systems expressed simultaneously in
multidrug-resistant pseudomonas aeruginosa. Antimicrobial Agents and
Chemotherapy. 44, 2861-2864.
154
Qi, Y., Wei, Z., Ji, S., Du, X., Shen, P., and Yu, Y. 2010. St11, the dominant clone of kpc-
producing klebsiella pneumoniae in china. Journal of Antimicrobial
Chemotherapy. 66, 307-312.
Qin, S., Wang, Y., Zhang, Q., Chen, X., Shen, Z., Deng, F., Wu, C., and Shen, J. 2012.
Identification of a novel genomic island conferring resistance to multiple
aminoglycoside antibiotics in campylobacter coli. Antimicrobial Agents and
Chemotherapy. 56, 5332-5339.
Queenan, A. M. and Bush, K. 2007. Carbapenemases: The versatile β-lactamases. Clinical
Microbiology Reviews. 20, 440-458.
Queenan, A. M., Shang, W., Flamm, R., and Bush, K. 2010. Hydrolysis and inhibition
profiles of β-lactamases from molecular classes a to d with doripenem,
imipenem, and meropenem. Antimicrobial Agents and Chemotherapy. 54, 565-
569.
Rahman, M. R. A., Achadi, A. S., Tingga, R. C. T., and Hasan, N. H. 2010. A new
distributional record of the rare bat coelops robinsoni from sarawak,malaysian
borneo. Journal of tropical biology and conservation. 7, 87-92.
Rahman, M. R. A., Tingga, R. C. T., Noor Haliza, H., Sigit, W., Anang, S. A., Eileen, L., Besar,
K., Huzal, I. H., and Abdullah, M. T. 2010. Diversity of bats in two protected
limestone areas in sarawak, malaysia. Sarawak Museum Journal. 88, 209-246.
Rakhuba, D. V., Kolomiets, E. I., Dey, E. S., and Novik, G. I. 2010. Bacteriophages
receptors, mechanisms of phage adsorption and penetration into host cell.
Polish Journal of Microbiology. 59, 145-155.
Rautner, M., Hardiono, M., and Alfred, R. J. 2005. Borneo: Treasure island at risk.
Frankfurt: WWF Germany.
Rautner, M., Hardiono, M., and Alfred, R. J. 2005. Borneo: Treasure island at risk. Status
of forest, wildlife and related threats on the island of borneo. World Wildlife
Fund,
Frankfurt am Main, Germany.
Ravin, V., Ravin, N., Casjens, S., Ford, M. E., Hatfull, G. F., and Hendrix, R. W. 2000.
Genomic sequence and analysis of the atypical temperate bacteriophage n15.
Journal of Molecular Biology. 299, 53-73.
155
Rea, M. C., Alemayehu, D., Ross, R. P., and Hill, C. 2013. Gut solutions to a gut problem:
Bacteriocins, probiotics and bacteriophage for control of clostridium difficile
infection. Journal of Medical Microbiology. 62, 1369-1378.
Review on Antimicrobial Resistance. 2014. Antimicrobial resistance: Tackling a crisis for
the health and wealth of nations.
Rhoads, D. D., Wolcott, R. D., Kuskowski, M. A., Wolcott, B. M., Ward, L. S., and
Sulakvelidze, A. 2009. Bacteriophage therapy of venous leg ulcers in humans:
Results of a phase i safety trial. Journal of Wound Care. 18, 237-243.
Rice, L. B. 2010. Progress and challenges in implementing the research on eskape
pathogens. Infection Control & Hospital Epidemiology. 31, S7-S10.
Ripp, S. and Miller, R. V. 1997. The role of pseudolysogeny in bacteriophage-host
interactions in a natural freshwater environment. Microbiology. 143, 2065-2070.
Riquelme, C., Rigal, F., Hathaway, J. J. M., Northup, D. E., Spilde, M. N., Borges, P. A. V.,
Gabriel, R., Amorim, I. R., and Dapkevicius, M. d. L. N. E. 2015. Cave microbial
community composition in oceanic islands: Disentangling the effect of different
colored mats in diversity patterns of azorean lava caves. FEMS Microbiology
Ecology. 91, 1-12.
Roach, D. R. and Debarbieux, L. 2017. Phage therapy: Awakening a sleeping giant.
Emerging Topics in Life Sciences. 1, 93-103.
Rodríguez-Rubio, L., Gutiérrez, D., Donovan, D. M., Martínez, B., Rodríguez, A., and
García, P. 2016. Phage lytic proteins: Biotechnological applications beyond
clinical antimicrobials. Critical Reviews in Biotechnology. 36, 542-552.
Roesch, L. F. W., Fulthorpe, R. R., Riva, A., Casella, G., Hadwin, A. K. M., Kent, A. D.,
Daroub, S. H., Camargo, F. A. O., Farmerie, W. G., and Triplett, E. W. 2007.
Pyrosequencing enumerates and contrasts soil microbial diversity. The ISME
Journal. 1, 283-290.
Rohwer, F. and Edwards, R. 2002. The phage proteomic tree: A genome-based taxonomy
for phage. Journal of Bacteriology. 184, 4529-4535.
Ross, A., Ward, S., and Hyman, P. 2016. More is better: Selecting for broad host range
bacteriophages. Frontiers in Microbiology. 7, 1-6.
Ryan, E. M., Gorman, S. P., Donnelly, R. F., and Gilmore, B. F. 2011. Recent advances in
bacteriophage therapy: How delivery routes, formulation, concentration and
156
timing influence the success of phage therapy. Journal of Pharmacy and
Pharmacology. 63, 1253-1264.
Sa'don, N. M., Karim, A. R. A., Jaol, W., and Lili, W. H. W. 2015. Sarawak peat
characteristics and heat treatment. UNIMAS e-Journal of Civil Engineering. 5, 6-
12.
Sambrook, J. and Russell, D. W. 2001. Molecular cloning, p. Pages. Cold Spring Harbor
Laboratory Press, New York, USA.
Sandeep, K. 2006. Bacteriophage precision drug against bacterial infections. Current
Science. 90, 631-633.
Santajit, S. and Indrawattana, N. 2016. Mechanisms of antimicrobial resistance in eskape
pathogens. BioMed Research International. 2016, 1-8.
Sarawak Forest Department. 1992. Proposal to constitute wind cave and fairy cave as
nature reserve. In S. National Park Wildlife Office Forest Department (ed.).
Sarbu, S. M., Kane, T. C., and Kinkle, B. K. 1996. A chemoautotrophically based cave
ecosystem. Science. 272, 1953-1955.
Sarker, S. A., McCallin, S., Barretto, C., Berger, B., Pittet, A.-C., Sultana, S., Krause, L.,
Huq, S., Bibiloni, R., and Bruttin, A. 2012. Oral t4-like phage cocktail application
to healthy adult volunteers from bangladesh. Virology. 434, 222-232.
Schilthuizen, M. 2004. Land snail conservation in borneo: Limestone outcrops act as
arks. Journal of Conchology Special Publication. 3, 149-154.
Schmelcher, M., Donovan, D. M., and Loessner, M. J. 2012. Bacteriophage endolysins as
novel antimicrobials. Future Microbiology. 7, 1147-1171.
Schmeler, K. M., Frumovitz, M., and Ramirez, P. T. 2011. Conservative management of
early stage cervical cancer: Is there a role for less radical surgery? Gynecologic
Oncology. 120, 321-325.
Schmerer, M., Molineux, I. J., and Bull, J. J. 2014. Synergy as a rationale for phage therapy
using phage cocktails. PeerJ. 2, 1-19.
Seed, K. D. 2015. Battling phages: How bacteria defend against viral attack. PLOS
Pathogens. 11, 1-5.
Semler, D. D., Lynch, K. H., and Dennis, J. J. 2011. The promise of bacteriophage therapy
for burkholderia cepacia complex respiratory infections. Frontiers in Cellular and
Infection Microbiology.
157
Shabram, P. and Aguilar-Cordova, E. 2000. Multiplicity of infection/multiplicity of
confusion. Molecular Therapy. 2, 420-421.
Sharma, S., Chatterjee, S., Datta, S., Prasad, R., Dubey, D., Prasad, R. K., and Vairale, M.
G. 2017. Bacteriophages and its applications: An overview. Folia Microbiologica.
62, 17-55.
Shore, A. C., Deasy, E. C., Slickers, P., Brennan, G., O'Connell, B., Monecke, S., Ehricht,
R., and Coleman, D. C. 2011. Detection of staphylococcal cassette chromosome
mec type xi carrying highly divergent meca, meci, mecr1, blaz, and ccr genes in
human clinical isolates of clonal complex 130 methicillin-resistant
staphylococcus aureus. Antimicrobial Agents and Chemotherapy. 55, 3765-3773.
Sillankorva, S., Oliveira, R., Vieira, M. J., Sutherland, I., and Azeredo, J. 2004.
Bacteriophage φ s1 infection of pseudomonas fluorescens planktonic cells versus
biofilms. Biofouling. 20, 133-138.
Silva, Y. J., Costa, L., Pereira, C., Cunha, Â., Calado, R., Gomes, N., and Almeida, A. 2014.
Influence of environmental variables in the efficiency of phage therapy in
aquaculture. Microbial Biotechnology. 7, 401-413.
Silva, Y. J., Costa, L., Pereira, C., Mateus, C., Cunha, Â., Calado, R., Gomes, N. C. M., Pardo,
M. A., Hernandez, I., and Almeida, A. 2014. Phage therapy as an approach to
prevent vibrio anguillarum infections in fish larvae production. PLoS ONE. 9, 1-
23.
Silva, Y. J., Costa, L., Pereira, C., Mateus, C., Cunha, n., Calado, R., Gomes, N. C. M., Pardo,
M. A., Hernandez, I., and Almeida, A. 2014. Phage therapy as an approach to
prevent <italic>vibrio anguillarum</italic> infections in fish larvae production.
PLoS ONE. 9, e114197.
Singh, G., Bhalla, A., and Ralhan, P. K. 2011. Extremophiles and extremozyrnes:
Importance in current biotechnology. Extreme Life, Biospeology & Astrobiology.
3, 46-54.
Singh, O. V. and Gabani, P. 2011. Extremophiles: Radiation resistance microbial reserves
and therapeutic implications. Journal of Applied Microbiology. 110, 851-861.
Skurnik, M., Pajunen, M., and Kiljunen, S. 2007. Biotechnological challenges of phage
therapy. Biotechnology Letters. 29, 995-1003.
158
Skurnik, M. and Strauch, E. 2006. Phage therapy: Facts and fiction. International Journal
of Medical Microbiology. 296, 5-14.
Slik, J. W. F., Aiba, S. I., Brearley, F. Q., Cannon, C. H., Forshed, O., Kitayama, K.,
Nagamasu, H., Nilus, R., Payne, J., and Paoli, G. 2010. Environmental correlates
of tree biomass, basal area, wood specific gravity and stem density gradients in
borneo's tropical forests. Global Ecology and Biogeography. 19, 50-60.
Smith, H. W. and Huggins, M. B. 1983. Effectiveness of phages in treating experimental
escherichia coli diarrhoea in calves, piglets and lambs. Microbiology. 129, 2659-
2675.
Smith, H. W. and Huggins, M. B. 1982. Successful treatment of experimental escherichia
coli infections in mice using phage: Its general superiority over antibiotics.
Microbiology. 128, 307-318.
Smith, H. W., Huggins, M. B., and Shaw, K. M. 1987. The control of experimental
escherichia coli diarrhoea in calves by means of bacteriophages. Microbiology.
133, 1111-1126.
Smith, H. W., Huggins, M. B., and Shaw, K. M. 1987. Factors influencing the survival and
multiplication of bacteriophages in calves and in their environment.
Microbiology. 133, 1127-1135.
Spellberg, B. 2014. The future of antibiotics. Critical Care. 18, 1-7.
Stewart, P. S. and Costerton, J. W. 2001. Antibiotic resistance of bacteria in biofilms. The
Lancet. 358, 135-138.
Sulaiman, Z. H. and Mayden, R. L. 2012. Cypriniformes of borneo (actinopterygii,
otophysi): An extraordinary fauna for integrated studies on diversity,
systematics, evolution, ecology, and conservation. Zootaxa. 3586, 359-376.
Sulaiman, Z. H. and Mayden, R. L. 2012. Cypriniformes of borneo (actinopterygii,
otophysi): An extraordinary fauna for integrated studies on diversity,
systematics, evolution, ecology, and conservation. Zootaxa Journal. 3586, 359–
376.
Sulakvelidze, A., Alavidze, Z., and Morris, J. G. 2001. Bacteriophage therapy.
Antimicrobial Agents and Chemotherapy. 45, 649-659.
Sulakvelidze, A., Alavidze, Z., and Morris, J. G., Jr. 2001. Bacteriophage therapy.
Antimicrob Agents Chemother. 45, 649-659.
159
Summers, W. C. 1999. Felix d’herelle and the origins of molecular biology, p. Pages. Yale
University Press.
Suttle, C. A. 2007. Marine viruses--major players in the global ecosystem. Nature
Reviews Microbiology. 5, 801-812.
Suttle, C. A. 2005. Viruses in the sea. Nature. 437, 356-361.
Sutton, S. 2011. Accuracy of plate counts. Journal of validation technology. 17, 42.
Sutton, S. 2011. Measurement of microbial cells by optical density. Journal of Validation
technology. 17, 46.
Tamber, S. and Hancock, R. E. 2003. On the mechanism of solute uptake in
pseudomonas. Frontiers in Bioscience: a Journal and Virtual Library. 8, s472-483.
Tan, D., Gram, L., and Middelboe, M. 2014. Vibriophages and their interactions with the
fish pathogen vibrio anguillarum. Applied and Environmental Microbiology. 80,
3128-3140.
Tan, G. H., Nordin, M. S., and Napsiah, A. B. 2008. Isolation and characterization of lytic
bacteriophages from sewage water. Journal of Tropical Agriculture and Food
Science 36.
Tan, H. S., Nadiah, I., and Eryani, S. S. 2009. Tree flora of sabah and sarawak project–
progress and future activities. Blumea-Biodiversity, Evolution and Biogeography
of Plants. 54, 23-24.
Tanji, Y., Shimada, T., Fukudomi, H., Miyanaga, K., Nakai, Y., and Unno, H. 2005.
Therapeutic use of phage cocktail for controlling escherichia coli o157: H7 in
gastrointestinal tract of mice. Journal of Bioscience and Bioengineering. 100,
280-287.
Team, E. E. 2012. The european union summary report on trends and sources of
zoonoses, zoonotic agents and food-borne outbreaks in 2010. Eurosurveillance:
Europe Journal on Infectious Disease, Epidemiology, Prevention and Control 17.
Tock, M. R. and Dryden, D. T. F. 2005. The biology of restriction and anti-restriction.
Current Opinion in Microbiology. 8, 466-472.
Tomat, D., Quiberoni, A., Mercanti, D., and Balagué, C. 2014. Hard surfaces
decontamination of enteropathogenic and shiga toxin-producing escherichia coli
using bacteriophages. Food Research International. 57, 123-129.
160
Tomczyk-Żak, K. and U., Z. 2015. Microbial diversity in caves. Review of Geomicrobiology.
1-19.
Tomczyk-Żak, K. and Zielenkiewicz, U. 2016. Microbial diversity in caves.
Geomicrobiology Journal. 33, 20-38.
Torres-Barceló, C., Arias-Sánchez, F. I., Vasse, M., Ramsayer, J., Kaltz, O., and Hochberg,
M. E. 2014. A window of opportunity to control the bacterial pathogen
pseudomonas aeruginosa combining antibiotics and phages. PLoS ONE. 9, 1-7.
Twort, F. W. 1915. An investigation on the nature of ultra-microscopic viruses. The
Lancet. 186, 1241-1243.
Tzouvelekis, L. S., Markogiannakis, A., Psichogiou, M., Tassios, P. T., and Daikos, G. L.
2012. Carbapenemases in klebsiella pneumoniae and other enterobacteriaceae:
An evolving crisis of global dimensions. Clinical Microbiology Reviews. 25, 682-
707.
Van Belleghem, J. D., Merabishvili, M., Vergauwen, B., Lavigne, R., and Vaneechoutte,
M. 2017. A comparative study of different strategies for removal of endotoxins
from bacteriophage preparations. Journal of Microbiological Methods. 132, 153-
159.
Van Boeckel, T. P., Gandra, S., Ashok, A., Caudron, Q., Grenfell, B. T., Levin, S. A., and
Laxminarayan, R. 2014. Global antibiotic consumption 2000 to 2010: An analysis
of national pharmaceutical sales data. The Lancet Infectious Diseases. 14, 742-
750.
Van der Meer, P. J., Demies, M. A., Shabudin, M. S., Verwer, C., Beintema, A., Hilligers,
P. J. M., and Teo, S. P. 2013. Tropical peat swamp forests of sarawak, p. 1-39.
Sustainable use and biodiversity conservation in a changing environment.
Alterra, Wageningen UR, Netherland.
Van Twest, R. and Kropinski, A. M. 2009. Bacteriophage enrichment from water and soil,
p. 15-21. In M. R. J. Clokie and A. M. Kropinski (eds.), Bacteriophages: Methods
and protocols, volume 1: Isolation, characterization, and interactions, Humana
Press.
Vandenheuvel, D., Lavigne, R., and Brüssow, H. 2015. Bacteriophage therapy: Advances
in formulation strategies and human clinical trials. Annual Review of Virology. 2,
599-618.
161
Vargiu, A. V. and Nikaido, H. 2012. Multidrug binding properties of the acrb efflux pump
characterized by molecular dynamics simulations. Proceedings of the National
Academy of Sciences of the United States of America. 109, 20637-20642.
Vaz-Moreira, I., Nunes, O. C., and Manaia, C. M. 2012. Diversity and antibiotic resistance
in pseudomonas spp. From drinking water. Science of the Total Environment.
426, 366-374.
Velonakis, E., Dimitriadi, D., Papadogiannakis, E., and Vatopoulos, A. 2014. Present
status of effect of microorganisms from sand beach on public health. Journal of
Coastal Life Medicine. 2, 746-756.
Viazis, S., Akhtar, M., Feirtag, J., and Diez-Gonzalez, F. 2011. Reduction of escherichia
coli o157: H7 viability on hard surfaces by treatment with a bacteriophage
mixture. International journal of food microbiology. 145, 37-42.
Viertel, T. M., Ritter, K., and Horz, H.-P. 2014. Viruses versus bacteria—novel approaches
to phage therapy as a tool against multidrug-resistant pathogens. Journal of
Antimicrobial Chemotherapy. 69, 2326-2336.
Voulgari, E., Poulou, A., Koumaki, V., and Tsakris, A. 2013. Carbapenemase-producing
enterobacteriaceae: Now that the storm is finally here, how will timely detection
help us fight back? Future Microbiology. 8, 27-39.
Wagner, P. L. and Waldor, M. K. 2002. Bacteriophage control of bacterial virulence.
Infection and Immunity. 70, 3985-3993.
Walsh, C. 2003. Opinion--anti-infectives: Where will new antibiotics come from? Nature
Reviews Microbiology. 1, 65.
Walsh, T. R., Weeks, J., Livermore, D. M., and Toleman, M. A. 2011. Dissemination of
ndm-1 positive bacteria in the new delhi environment and its implications for
human health: An environmental point prevalence study. The Lancet Infectious
Diseases. 11, 355-362.
Wang, I.-N., Smith, D. L., and Young, R. 2000. Holins: The protein clocks of bacteriophage
infections. Annual Reviews in Microbiology. 54, 799-825.
Wang, K., Pei, H., Huang, B., Zhu, X., Zhang, J., Zhou, B., Zhu, L., Zhang, Y., and Zhou, F.-
F. 2013. The expression of abc efflux pump, rv1217c–rv1218c, and its association
with multidrug resistance of mycobacterium tuberculosis in china. Current
Microbiology. 66, 222-226.
162
Wang, Z., Zheng, P., Ji, W., Fu, Q., Wang, H., Yan, Y., and Sun, J. 2016. Slpw: A virulent
bacteriophage targeting methicillin-resistant staphylococcus aureus in- vitro and
in vivo. Frontiers in Microbiology. 7, 1-10.
Weber-Dąbrowska, B., Jończyk-Matysiak, E., Żaczek, M., Łobocka, M., Łusiak-
Szelachowska, M., and Górski, A. 2016. Bacteriophage procurement for
therapeutic purposes. Frontiers in Microbiology. 7, 1-14.
Weber-Dąbrowska, B., Żaczek, M., Dziedzic, B., Łusiak-Szelachowska, M., Kiejzik, M.,
Górski, A., Gworek, B., Wierzbicki, K., and Eymontt, A. 2014. Presented at the
Industrial, Medical and Environmental Applications of Microorganisms: Current
Status and Trends: Proceedings of the international conference on
environmental, industrial and applied microbiology, Madrid, Spain.
Weinbauer, M. G. 2004. Ecology of prokaryotic viruses. FEMS Microbiology Reviews. 28,
127-181.
Wilhelm, S. W. and Suttle, C. A. 1999. Viruses and nutrient cycles in the sea. BioScience.
49, 781-788.
Wilkinson, M. H. F. 2001. Predation in the presence of decoys: An inhibitory factor on
pathogen control by bacteriophages or bdellovibrios in dense and diverse
ecosystems. Journal of Theoretical Biology. 208, 27-36.
Willey, J. M., Sherwood, L., and Woolverton, C. J. 2009. Prescott's principles of
microbiology, p. Pages. McGraw-Hill Higher Education.
Williamson, K. E., Wommack, K. E., and Radosevich, M. 2003. Sampling natural viral
communities from soil for culture-independent analyses. Applied and
Environmental Microbiology. 69, 6628-6633.
Wittebole, X., De Roock, S., and Opal, S. M. 2014. A historical overview of bacteriophage
therapy as an alternative to antibiotics for the treatment of bacterial pathogens.
Virulence. 5, 226-235.
Wommack, K. E. and Colwell, R. R. 2000. Virioplankton: Viruses in aquatic ecosystems.
Microbiology and Molecular Biology Reviews. 64, 69-114.
Wong, C. L., Sieo, C. C., Tan, W. S., Abdullah, N., Hair-Bejo, M., Abu, J., and Ho, Y. W.
2014. Evaluation of a lytic bacteriophage, φ st1, for biocontrol of salmonella
enterica serovar typhimurium in chickens. International journal of food
microbiology. 172, 92-101.
163
Woodford, N., Turton, J. F., and Livermore, D. M. 2011. Multiresistant gram‐negative
bacteria: The role of high‐risk clones in the dissemination of antibiotic resistance.
FEMS Microbiology Reviews. 35, 736-755.
Wozniak, A., Villagra, N. A., Undabarrena, A., Gallardo, N., Keller, N., Moraga, M.,
Roman, J. C., Mora, G. C., and García, P. 2012. Porin alterations present in non-
carbapenemase-producing enterobacteriaceae with high and intermediate
levels of carbapenem resistance in chile. Journal of Medical Microbiology. 61,
1270-1279.
Wright, A., Hawkins, C. H., Änggård, E. E., and Harper, D. R. 2009. A controlled clinical
trial of a therapeutic bacteriophage preparation in chronic otitis due to
antibiotic‐resistant pseudomonas aeruginosa; a preliminary report of efficacy.
Clinical Otolaryngology. 34, 349-357.
Wright, G. D. 2016. Antibiotic adjuvants: Rescuing antibiotics from resistance. Trends in
Microbiology. 24, 862-871.
Wright, G. D. 2005. Bacterial resistance to antibiotics: Enzymatic degradation and
modification. Advanced Drug Delivery Reviews. 57, 1451-1470.
Wright, G. D. 2010. Q&a: Antibiotic resistance: Where does it come from and what can
we do about it? BMC biology. 8, 123.
Wu, Y., Tan, L., Liu, W., Wang, B., Wang, J., Cai, Y., and Lin, X. 2015. Profiling bacterial
diversity in a limestone cave of the western loess plateau of china. Frontiers in
Microbiology. 6, 1-10.
Yang, H., Ding, W., Zhang, C. L., Wu, X., Ma, X., He, G., Huang, J., and Xie, S. 2011.
Occurrence of tetraether lipids in stalagmites: Implications for sources and gdgt-
based proxies. Organic Geochemistry. 42, 108-115.
Yigit, H., Queenan, A. M., Anderson, G. J., Domenech-Sanchez, A., Biddle, J. W., Steward,
C. D., Alberti, S., Bush, K., and Tenover, F. C. 2001. Novel carbapenem-
hydrolyzing β-lactamase, kpc-1, from a carbapenem-resistant strain of klebsiella
pneumoniae. Antimicrobial Agents and Chemotherapy. 45, 1151-1161.
Young, D. F., Andrejeva, L., Livingstone, A., Goodbourn, S., Lamb, R. A., Collins, P. L.,
Elliott, R. M., and Randall, R. E. 2003. Virus replication in engineered human cells
that do not respond to interferons. Journal of Virology. 77, 2174-2181.
164
Yu, P., Mathieu, J., Lu, G. W., Gabiatti, N., and Alvarez, P. J. 2017. Control of antibiotic-
resistant bacteria in activated sludge using polyvalent phages in conjunction with
a production host. Environmental Science & Technology Letters. 4, 137-142.
Zhang, L., Bao, H., Wei, C., Zhang, H., Zhou, Y., and Wang, R. 2015. Characterization and
partial genomic analysis of a lytic myoviridae bacteriophage against
staphylococcus aureus isolated from dairy cows with mastitis in mid-east of
china. Virus Genes. 50, 111-117.
Zhang, W.-J., Xu, X.-R., Schwarz, S., Wang, X.-M., Dai, L., Zheng, H.-J., and Liu, S. 2013.
Characterization of the inca/c plasmid pscec2 from escherichia coli of swine
origin that harbours the multiresistance gene cfr. Journal of Antimicrobial
Chemotherapy. 69, 385-389.
Zhang, Y. and Hu, Z. 2013. Combined treatment of pseudomonas aeruginosa biofilms
with bacteriophages and chlorine. Biotechnology and Bioengineering. 110, 286-
295.
Zhao, W.-H. and Hu, Z.-Q. 2013. Epidemiology and genetics of ctx-m extended-spectrum
β-lactamases in gram-negative bacteria. Critical Reviews in Microbiology. 39, 79-
101.
Zhvania, P., Hoyle, N. S., Nadareishvili, L., Nizharadze, D., and Kutateladze, M. 2017.
Phage therapy in a 16-year-old boy with netherton syndrome. Frontiers in
Medicine. 4, 94-98.
165
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
166
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)
MOI 100 or 102= (1x 1012 PFU/mL) (1x 1010 CFU/mL)
MOI 1000 or 103= (1x 1013 PFU/mL) (1x 1010 CFU/mL)
MOI 10000 or 104= (1x 1014 PFU/mL) (1x 1010 CFU/mL)
MOI 100000 or 105= (1x 1015 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