Systematic Study of Bacterial Phenotypes Caused by Pseudomonas aeruginosa Prophage Adaptive Genes

by

Yu-Fan Tsao

A thesis submitted in conformity with the requirements for the degree of Master of Science

Department of Molecular Genetics University of Toronto

© Copyright by Yu-Fan Tsao 2015

ii

Systematic Study of Bacterial Phenotypes Caused by

Pseudomonas aeruginosa Prophage Adaptive Genes

Yu-Fan Tsao

Master of Science

Department of Molecular Genetics

University of Toronto

2015

Abstract

In addition to being the major predator of bacteria, some bacteriophages adapted a symbiotic

relationship with their bacterial hosts through the formation of prophages. Unique genes in

prophages known as Prophage Adaptive Genes (PrAGs) function to alter host biological

processes, and they contribute greatly to bacterial pathogenicity. I aimed to investigate the

phenotypic effects of PrAGs on Pseudomonas aeruginosa, an opportunistic human pathogen.

Individual PrAGs in P. aeruginosa PA14 or PAO1 strain background were screened for bacterial

phenotypes related to virulence and survivability, including phage resistance, motility,

production of biofilm and pyocyanin, and virulence in Drosophila melanogaster. I found that 11

out of 17 PrAGs screened modified the host phenotype in some manner. In addition, some of

them were observed to affect multiple phenotypes or cause bacterial strain-specific changes,

which suggests that PrAGs interact with host components to cause the phenotypes observed.

iii

Acknowledgments

My journey through Master’s study has been a wonderful one, and I want to thank all the people

who made this possible. Without any of you, I would not be standing where I am today. First of

all, I would like to express my gratitude to my supervisors Dr. Karen Maxwell and Dr. Alan

Davidson for allowing me to work in their labs and providing me with all the resources and

guidance for my graduate study. Their continual support, both academically and mentally, has

helped me pass numerous obstacles in the past few years and is the key to the successful

completion of this project.

I would also like to thank my committee members Dr. Trevor Moraes, Dr. Alex Ensminger and

Dr. John Parkinson for their helpful advice throughout my study. They guided me forward in my

project while keeping me on track. Every committee meeting was valuable to me as they always

clear away confusion and inspire me with new ideas. Special thanks to Dr. Trevor Moraes and

his lab members for their help with structural studies. I have learned a lot about protein structures

and X-ray crystallography through working with the Moraes lab. In addition, I would like to

thank Dr. Paul Sadowski for sharing his experiences in experimental techniques and other

helpful comments.

Thanks to all the past and present members of the Maxwell and Davidson labs for their input and

inspirations at the lab meetings and elsewhere. They made the lab environment and my

experience there really positive and enjoyable, and they have also filled my life outside of the lab

with laughter and happiness. Especially, I need to thank Diane Bona for guiding me through my

early days in the lab as well as assisting me with my experiments, and Joe Bondy-Denomy for

his work on prophages that led to my project and his valuable guidance. I also wish to thank

Nichole Cumby, Senjuti Saha, Smriti Kala and Kris Hon for answering thousands of my

questions, helping and mentoring me through my study, and being such great friends. My

graduate school life will be very different without them.

Finally, I would like to thank my parents Charles Tsao and Jennifer Fu, my brother Oliver Tsao,

my boyfriend Dan Cheng and all my friends for their love and encouragement through stressful

times which made my journey so much easier and wonderful.

iv

Table of Contents

Acknowledgments .......................................................................................................................... iii

Table of Contents ........................................................................................................................... iv

List of Tables ................................................................................................................................. vi

List of Figures ............................................................................................................................... vii

List of Abbreviations ................................................................................................................... viii

Chapter 1 Introduction .................................................................................................................... 1

1.1 Bacteriophages and their Life Cycle ................................................................................... 1

1.2 Bacteriophages and Human Health ..................................................................................... 2

1.3 The Importance of Bacteriophages in Bacterial Physiology ............................................... 3

1.4 Prophage Adaptive Genes (PrAGs) .................................................................................... 5

1.5 Pseudomonas aeruginosa – An Opportunistic Human Pathogen ....................................... 7

1.5.1 P. aeruginosa and Human Health ........................................................................... 7

1.5.2 Virulence Factors of P. aeruginosa ........................................................................ 8

1.6 P. aeruginosa Prophages and PrAGs ................................................................................ 10

1.7 Thesis Objectives .............................................................................................................. 10

Chapter 2 Materials and Methods ................................................................................................. 12

2.1 Construction of Expression Plasmids and Protein Test Expression in E. coli .................. 12

2.2 Transformation of plasmids into P. aeruginosa ............................................................... 12

2.3 PCR Reactions and Colony Screening .............................................................................. 13

2.4 Bacterial Growth ............................................................................................................... 13

2.5 Twitching Motility Assay ................................................................................................. 14

2.6 Swimming Motility Assay ................................................................................................ 14

2.7 Induction of P. aeruginosa Lysogens and Plaquing Assays ............................................. 15

v

2.8 Drug Resistance Assay ..................................................................................................... 15

2.9 Biofilm Production Assay ................................................................................................. 15

2.10 Virulence Assay ................................................................................................................ 16

Chapter 3 Results .......................................................................................................................... 17

3.1 Acknowledgement ............................................................................................................ 17

3.2 P. aeruginosa PrAGs possess diverse genomic and bioinformatics properties ................ 17

3.3 The PrAGs all express and make proteins in E. coli ......................................................... 19

3.4 Some PrAGs introduce growth defects in the bacterial host ............................................ 21

3.5 PrAGs increase bacterial survivability by protecting their hosts from phage infection ... 24

3.6 Bacterial motility is often modified by PrAGs ................................................................. 29

3.7 PrAGs are capable of altering the production of bacterial virulence factors .................... 33

3.8 Two PrAGs modify bacterial virulence in Drosophila melanogaster .............................. 36

Chapter 4 Discussion and Future Directions ................................................................................ 38

References ..................................................................................................................................... 45

Appendix ....................................................................................................................................... 53

vi

List of Tables

Table 1. Summary of PrAGs included in this study. .................................................................... 19

Table 2. Increase in phage resistance caused by PrAG expression in PA14 compared to no PrAG

expression. .................................................................................................................................... 25

Table 3. Fold increase in phage resistance caused by PrAG expression in PAO1 compared to no

PrAG expression. .......................................................................................................................... 26

Table 4. Difference in phage resistance caused by PrAG expression in PA14 and PAO1. .......... 28

Table 5. List of drugs tested in drug resistance assay. .................................................................. 29

Table 6. Summary of host phenotypic changes caused by PrAGs in P. aeruginosa PA14 and

PAO1 strains. ................................................................................................................................ 43

vii

List of Figures

Figure 1. The life cycle of a bacteriophage. .................................................................................... 2

Figure 2. Example of PrAGs identified in phage genomes. ........................................................... 6

Figure 3. Genome comparison between related phage strains and positions of PrAGs. .............. 18

Figure 4. Protein expression of PrAGs in E. coli. ......................................................................... 20

Figure 5. PrAG expression affect bacterial growth rate. .............................................................. 23

Figure 6. Some PrAGs affect twitching motility of P. aeruginosa. ............................................. 30

Figure 7. PrAGs could alter swimming motility of the bacteria. .................................................. 32

Figure 8. PrAGs cause changes in bacterial biofilm production. ................................................. 34

Figure 9. JBD5-4 alters the levels of pyocyanin produced by P. aeruginosa. ............................. 35

Figure 10. PrAGs have effects on bacterial virulence towards Drosophila malenogaster. .......... 37

Figure 11. PrAG expression affect various bacterial elements. .................................................... 44

Figure 12. Detail genome alignment for phages used in this study. ............................................. 53

viii

List of Abbreviations

ATPase adenosine triphosphatase

BLAST basic local alignment search tool

cm centimeter

C. botulinum Clostridium botulinum

C. diphtheria Corynebacterium diphtheriae

CF cystic fibrosis

CFTR cystic fibrosis transmembrane conductance regulator

d day

°C degrees Celsius

dNTP deoxynucleotide

DNA deoxyribonucleic acid

ssDNA single-stranded deoxyribonucleic acid

dsDNA double-stranded deoxyribonucleic acid

D. malenogaster Drosophila malenogaster

EV empty vector

E. coli Escherichia coli

EPS extracellular polymeric substance

gp gene product

g gram

ix

h hour

IPTG isopropyl β-D-1 thiogalactopyranosaide

kDa kilodalton

λ Enterobacterophage lambda

LB lysogeny broth

NaCl sodium chloride

µg microgram

µL microliter

μM micromolar

mL milliliter

mm millimeter

mM millimolar

NCBI National Center for Biotechnology Information

OD optical density

OD595 optical density measured at 595 nm

OD600 optical density measured at 600 nm

ORF open reading frame

PCR polymerase chain reaction

PrAG Prophage Adaptive Gene

P. aeruginosa Pseudomonas aeruginosa

x

ssRNA single-stranded ribonucleic acid

dsRNA double-stranded ribonucleic acid

RNAseq ribonucleic acid sequencing

S. enterica Salmonella enterica

SDS-PAGE sodium dodecyl sulfate polyacrylamide gel electrophoresis

S. aureus Staphylococcus aureus

S. mitis Streptococcus mitis

T4P Type IV pili

V. cholerae Vibrio cholera

V volt

w/v weight/volume

1

Chapter 1 Introduction

1.1 Bacteriophages and their Life Cycle

In the early 1900s, Twort and d’Herelle first discovered bacteriophages (phages), the viruses that

specifically infect bacteria. They are the most abundant biological entity on earth with an

estimated population of 1031, outnumbering their bacterial host by approximately 10 fold1. Their

population is widely spread as they can be identified in all places where their bacterial hosts are

found, including soil, water, plants, animals and all non-sterile sites in the human body2,3.

According to the International Committee on Taxonomy of Viruses, phages were classified into

different families based on their virion morphology and form of nucleic acid4. Phages virions can

be binary (tailed), cubic, helical or pleomorphic5. Most of the phages contain double-stranded

(ds) DNA as their genetic material, while some phages have single-stranded (ss) DNA, dsRNA

or ssRNA5. Phages, having narrow host range6 and high population in a given environment, are

actively involved in shaping the composition of the bio-community, genetic transfer, nutrient and

energy recycling and they are an important factor in bacterial evolution7.

Phages are suspected to have originated from bacterial gene clusters that escaped from cellular

control resulting in a lifeform that do not have the necessary machinery to replicate themselves.

In order for a phage to replicate its genome, it must infect a bacterial host. Upon infection,

phages inject their genetic material into the bacterial cell, where they can go into two different

lifecycles: the lytic cycle or the lysogenic cycle (Figure 1)5. According to how phages behave

during infections, they can be virulent or temperate5. Virulent phages only go through the lytic

cycle, while temperate phages are able to survive with either life cycle. During the lytic cycle,

the phage proteins are actively produced and assembled into phage virions inside the bacterial

cell. Eventually, the phage particles exit the host, usually through lysis which kills the host cells.

In the lysogenic life cycle, the phage genome is either maintained as a plasmid or gets integrated

into the bacterial genome, forming a prophage. In this state, most of the phage genes are silenced

by a repressor protein. The prophage is replicated passively when the bacterial cells divide.

Although prophages can remain in the bacterial genome for numerous generations, temperate

phages are able to switch from the lysogenic cycle to the lytic cycle when the bacterial cell is

2

under certain stress conditions, including DNA damage8, changes in temperature or pH9, and

exposure to heavy metal9 or oxidative radicals10.

Figure 1. The life cycle of a bacteriophage.

When a phage infects a bacterial cell it injects its genetic material into the host cell and it can

replicate itself through two different life cycles: the lysogenic cycle or the lytic cycle. In the

lysogenic cycle, the phage DNA gets incorporated into the bacterial genome, and it is replicated

passively with the host genome through host cell divisions. In the lytic cycle, the phage genome

is replicated and phage proteins are produced and assembled into phage virions. Eventually, the

mature phage particles exit the host cell via lysis. Phages can switch from lysogenic to lytic cycle

upon induction by cell stress, such as DNA damage.

1.2 Bacteriophages and Human Health

Although bacteriophages do not infect any other organisms except bacteria, they can exert

profound effects on mammalian health indirectly through their bacterial hosts. Bacteriophages

are found in any environments where bacteria reside, including inside human bodies. Up to 1015

bacteriophage particles are found in the human gut, and the phage populations in the body are

sometimes referred to as the phageome11. The relationship between bacteriophages, bacteria and

humans is very complex, and our knowledge on the impact of phages on human health is limited.

Phages are known to encode genes that modify various biological processes in the bacterial life

cycle12. Based on how phages infect and alter the physiology of their host, they are likely

3

involved significantly in the shaping of the number and diversity of bacteria in the human

microbiome. In one example, studies have shown that both the phage and bacteria population in

the gastrointestinal tract of healthy individual and Crohn’s disease patients were noticeably

different13-16. It is possible that differences in the phageome contribute to the difference in

microbiota and the onset of the disease, but further studies are required to confirm this

hypothesis.

In addition to the natural residence in human bodies, phages have been used to treat bacterial

infections as an alternative to antibiotics as early as 192117. Recent successful use of phage

therapy was reported in Pseudomonas aeruginosa and Staphylococcus aureus infection models18-

21, which suggests the potential of phage therapy becoming more common in the future,

especially against antibiotic-resistant bacteria. Phages have also been investigated as direct

treatments or vectors to deliver vaccines and anticancer agents in order to prevent or treat human

diseases22-24. There are also cases where phages were used in food safety, where they were added

during food production to reduce pathogens in the food chain25,26. Phages play important roles in

bacterial and human biology, and as potential disease treatments and clinical biomarkers, their

applications in human health will likely increase as our knowledge of them expands.

1.3 The Importance of Bacteriophages in Bacterial Physiology

As previously mentioned, some phages replicate themselves by infecting and lysing bacterial

cells, and the number of phage exceeds the bacteria by approximately 10 fold. As a result, phage

killing has been the major way of how phage affect bacterial physiology, and it is also one of the

most common factors limiting bacterial population levels.

While phages are seen as parasitic to bacteria in the traditional view, the prophage exists in a

symbiotic relationship with the bacterial host. There are numerous known cases where prophage

genes make the bacteria more fit by enhancing its survivability or virulence. The most common

way of phages contributing to bacterial virulence is the encoding of extracellular toxins within

prophage genomes. Prophages were found to encode various toxins produced by both Gram-

positive and Gram-negative bacteria, including C. botulinum27, C. diphtheria28,29, V. cholerae30

and E. coli31-33. The phage-encoded toxins aid the bacteria in invading their targets via various

mechanisms, such as inhibiting protein synthesis (e.g. shiga toxin34) or interfering with

mammalian immune-response (e.g. enterotoxins35), and they are often the main cause of

4

diseases. Prophages can also encode proteins which help their host during initial bacterial

infection, for example, S. mitis surface proteins PblA and PblB that are important for adhesion

onto target cells36,37 as well as the Group A Strepotococcal hyaluronidase that is essential for

invasion38 are both encoded by prophages. Once the bacteria is inside the target cell, the

prophage can protect the bacterial host from the human immune system via a variety of methods.

Some prophages encode enzymes that alter the bacterial antigens to help evade immune-

responses, such as the O-antigen acetylase (oac) encoded by Shigella flexneri Sf6 prophage,

which is able to convert the serotype of its bacterial host 39,40. Other prophages encode proteins

that interfere with components of the mammalian immune system. For example, the chemotaxis

inhibitory protein encoded by Staphylococci prophage binds to neutrophil receptors and

attenuates its activity41. The S. aureus prophage encodes another such protein known as Panton-

Valentine leucocidin, a cytotoxin that directly targets and attacks human phagocytes42. There are

also examples where a prophage protects bacterial hosts from oxidative stress generated by

mammalian immune system, such as the superoxide dismutase (SodC) encoded by S. enterica

Gifsy-2 prophage which neutralizes superoxide radicals in phagocytes43. Thus, prophages

contribute greatly to bacterial pathogenicity by a variety of mechanisms.

In addition to protecting their hosts during bacterial infections, prophages were also found to

increase the survivability of the host in different environments. Through increasing the fitness of

their hosts, the phages themselves gain benefit as more copies of the prophage are replicated

when the bacterial cells divide. Therefore, the most successful phages likely express genes that

enhance the fitness of their host. Prophages encode proteins that protect the bacteria from further

phage infections. For example, the superinfection exclusion protein gp15 in E. coli is encoded by

the HK97 prophage44. Since phage killing is one of the main factors limiting bacterial population

levels, blocking phage infections results in higher survivability for the bacterial host. It is

believed that phages may be involved in the transduction of antibiotic-resistance genes among

bacteria. Studies have shown that the rate of phage transduction of non-phage genetic elements is

much higher than previously known45, and gene sequences related to antibiotic-resistance were

found in viral population in human gut and lungs46. In addition, antibiotic treatment has been

observed to increase the number of antibiotic-resistance genes detected in the phage genome47.

These findings suggest that phages may contribute indirectly to the distribution of host resistance

genes against various antibiotics in both environmental and clinical bacterial strains.

5

Multiple prophage elements can be found integrated in a bacterial genome, averaging around 3-

10% of the host genome48. An extreme case involves the enterohemorragic E. coli 0157:H7,

which harbours a total of 18 prophages, making up 16% of its entire genome content49. While

single prophages can cause a significant effect on bacterial phenotypes, the interactions between

multiple prophages can lead to very diverse phenotypic outcomes. Therefore, in order to

understand the physiology of bacteria, it is essential to note the importance of prophages, as they

contribute greatly to bacterial diversity and evolution.

1.4 Prophage Adaptive Genes (PrAGs)

As previously mentioned, prophages cause a wide variety of phenotypic effects in their bacterial

hosts, and they are able to do so through a class of genes we call Prophage Adaptive Genes

(PrAGs). Some of these genes have also been referred to as “accessory genes” or “morons”12,50.

PrAGs are phage genes that are not essential for the lytic or lysogenic phage life cycle. Since

they are non-essential genes, the presence of PrAGs is not necessarily conserved among related

phage strains, and their positions in the genome can be variable. These genetic elements are

thought to be inserted into the genome from a foreign source via horizontal gene transfer, as

many PrAGs have homologs in the bacterial genome outside of the prophage-encoded regions.

For example, the Pseudomonas phage D3-encoded O-antigen acetylase (Oac) has several

bacterial homologs, including the Pseudomonas acetyltransferases (WbpC proteins)12,51. Since

related phage genomes tend to have conserved gene orders, PrAG elements can be easily spotted

through genome alignments as they are usually only present in one or a few genomes (Figure 2).

PrAGs have been identified in many phage genomes, and they can be diverse in sequence and

function.

6

Figure 2. Example of PrAGs identified in phage genomes.

Since the presence of PrAGs are not necessarily conserved, they can be easily spotted through

genome alignment of closely related phages that have conserved gene orders, such as P.

aeruginosa phages JBD30, JBD23, JBD5 and DMS3. Gene JBD30-30 (red arrow a) and DMS3-

25 (red arrow b) are PrAGs identical in sequence and are found between the phage small and

large terminase genes in JBD30 and DMS3, but they are not present in JBD23 or JBD5. Note

that the small terminase genes share more than 90% identity and the large terminase gene share

35-99% identity in the phage genomes examined.

When expressed from a prophage, PrAG elements may modify the physiological properties of

the host cell. In fact, many of the effects on bacterial virulence and survivability mentioned

previously that are caused by prophages arose from the expression of PrAGs. In some examples,

prophages protect their hosts from further phage infections by expressing PrAGs to inactive

receptors on bacterial cell surface required for phage entry (e.g. Cor proteins expressed by E. coli

phages HK022, N15 and ϕ80 inactivate FhuA receptors52), or through other mechanisms (e.g.

PrAG gp15 from E. coli phage HK97 was hypothesized to block the formation of the phage entry

complex in the bacterial cell membrane as it has been shown to be responsible for the

superinfection exclusion phenotype observed in the lysogen44,53). In other examples, PrAGs help

their bacterial host in invading mammalian cells (e.g. PrAG-expressed λ Lom protein in E.

coli54and SopE protein expressed by Salmonella phages55) or surviving the mammalian immune

system (e.g. Bor protein expressed from λ PrAG protect the host from serum killing56). The

previously described bacterial toxins encoded by prophages are also PrAGs, including cholera,

diphtheria, botulism and shiga toxins27-32, which directly enhance the virulence of the bacterial

hosts. Although a number of PrAGs have been shown to be of great importance in bacterial

biology, the biological roles of most PrAGs are still unknown.

7

1.5 Pseudomonas aeruginosa – An Opportunistic Human Pathogen

Pseudomonas aeruginosa is a ubiquitous, gram-negative bacterium that can be found in diverse

environments, including soil, water and inside animal hosts. P. aeruginosa and its phages are

great model organisms for the studies of prophages, PrAGs, and their effects in bacterial

pathogenesis, because this bacterium is well-studied with many fully-sequenced bacterial and

phage strains. P. aeruginosa is also known to produce a large number of virulence factors, most

of which are well-characterized with techniques and assays available for related studies. In

addition, P. aeruginosa infects a variety of hosts, including several common model organisms,

such as Drosophila melanogaster57,58 and Caenorhabditis elegans59, which can be used to

measure the virulence level of P. aeruginosa with infection models.

1.5.1 P. aeruginosa and Human Health

P. aeruginosa is one of the most common opportunistic human pathogens. It is able to infect

humans who are immuno-compromised or elderly, as well as patients with burns, lung diseases,

open fractures or other injuries. P. aeruginosa infections often lead to complication of disease

states or secondary fungal infections, and in some cases can result in the death of the patient.

This bacterium especially causes serious issues among cystic fibrosis patients. Cystic fibrosis

(CF) is a genetic disorder caused by mutations in the cystic fibrosis transmembrane conductance

regulator (CFTR) gene, resulting in non-functional chloride ion channel important for the

production of sweat, digestive fluids and mucus60. The disease mainly affects the lungs, but it

can also affect intestines, liver, kidneys and pancreas61. P. aeruginosa can worsen CF conditions,

since it is commonly found in the lungs of the patients, where it can cause both acute and chronic

infections in the respiratory systems62. P. aeruginosa infections among CF patients can be severe

and persistent as the bacterial strains often possess high resistance to multiple antibiotics. In

addition, the ability of the bacteria to grow in various conditions significantly increases the

occurrence of new infections and outbreaks in hospitals 63. Because of its infectious and highly

drug-resistant nature, P. aeruginosa is a huge concern for public health, and many studies have

been done in the hope of developing effective treatments against these infections64,65.

8

1.5.2 Virulence Factors of P. aeruginosa

P. aeruginosa expresses various virulence factors, some of which help the bacteria invade a wide

spectrum of hosts, including mammals, plants and insects, while others enable the bacteria to

survive environmental stresses, such as antibiotic treatment and mammalian immune response.

The expression of virulence factors in P. aeruginosa is controlled by the quorum sensing

system66. The quorum sensing system is an intercellular communication network that is

responsible for the regulation of bacterial virulence based on the cell density in the surrounding

environment66. This study focused on investigating the phenotypic effects of PrAGs on three P.

aeruginosa virulence factors: bacterial motility, production of biofilm and pyocyanin.

1.5.2.1 Bacterial Motility

In P. aeruginosa, motility is mediated by two cell surface structures known as the Type IV pilus

(T4P) and the flagellum. The T4P is responsible for twitching motility, which describes the

ability of the bacteria to move across solid surfaces. The main T4P structure is a polymer of

Type IV pilins (PilA), and it is connected to an ATPase motor (PilT, PilB and PilU) associated

with the inner cell membrane 67,68. The ATPase motor enables pilus retraction motion that moves

the bacteria on surfaces69. The flagellum is involved in bacterial movement in liquid

environment, which is termed swimming motility. The flagellum is a long helical filament made

of thousands of flagellins (FliC), and the swimming motility is achieved by reversibly rotating

the flagellum driven by membrane rotor proteins (MotA and MotB)70. The expressions of both

the T4P and the flagellum are regulated by the quorum sensing system in P. aeruginosa.

Twitching and swimming motility is important in P. aeruginosa virulence, because it allows the

bacterial cells to be motile, meaning that the bacteria can more easily gain access to nutrients and

host cells during infection. Bacterial pili and flagella are also involved in adhesion71, biofilm and

microcolony formation72, which protect the bacteria from external threats, such as anti-microbial

agents. In addition, since these surface structures are often targets for the host immune system

and phage attachment, through down-regulation of gene expression or alteration of their

structures, the bacteria can use pilus and flagellum as a method for immune escape and phage

resistance73. Indeed, studies have shown that there are significant differences in pilin alleles

between P. aeruginosa isolates from CF and non-CF patients74, suggesting that bacteria

possesses different pilin types that may be advantageous in different situations.

9

1.5.2.2 Biofilm

Biofilm is an extracellular polymeric substance (EPS) matrix produced by P. aeruginosa and

many other bacteria to facilitate growth in a protected environment. The EPS matrix is composed

of polysaccharides, extracellular DNA and proteins75,76. They function to hold the bacterial cells

together, forming a microcolony. The production of biofilm in P. aeruginosa is mainly regulated

by the quorum sensing system77, but there are also reports of the involvement of T4P and

flagellum in biofilm development and maturation78. As an important virulence factor, biofilm

protects the bacteria from a wide variety of environmental stressors, including antibiotics and the

host immune responses79,80. The biofilm also promotes genetic exchanges between bacterial

cells81 and aids in the acquisition of nutrients by concentrating cell densities. Due to the nature

and function of the biofilm, it plays a key role in the establishment of chronic infections in CF

patient lungs82.

1.5.2.3 Pyocyanin

Pyocyanin is a redox-active secondary metabolite produced by P. aeruginosa, and it is also a

blue color pigment that gives the bacterium its unique bluish-green color. Pyocyanin is important

for the bacteria as it is known to be involved in various biological processes, for example, it

participates in the reduction and release of iron from transferrin in bacterial iron metabolism, and

it acts as an electron shuttle during bacterial respiration83. The biosynthesis of pyocyanin is

mainly mediated by the phzABCDEFG operons and extra genes, and the expression of these

genes are regulated by the quorum sensing network involving multiple regulatory pathways84,85.

As a virulence factor and a bacterial toxin, pyocyanin contributes greatly during P. aeruginosa

infections; it has been recovered from CF patient sputum in large quantities and was shown to be

crucial for lung infection in mice84,86. Pyocyanin generates reactive oxygen intermediates,

creating oxidative stress inside target cells, which results in target cell death84. In the mammalian

system, pyocyanin is known to cause multiple cellular damages, including inhibition of cell

respiration, interference with ciliary function87 and disruption of calcium homeostasis88. In

addition to its role in bacterial infections, pyocyanin has anti-bacterial and anti-fungal

activities83,89,90, which provides advantages for P. aeruginosa when competing with other micro-

organisms.

10

1.6 P. aeruginosa Prophages and PrAGs

Although there are numerous examples of prophages involved in the modification of bacterial

phenotypes, there have been no systematic studies performed. Joe Bondy-Denomy, a former

student in the Davidson lab, conducted a systematic study of prophages in P. aeruginosa, where

he created lysogens, each with a different phage in the same bacterial strain background91. He

observed differences in various bacterial phenotypes related to the virulence and survivability of

P. aeruginosa, in the presence compared to the absence of prophages. For example, when Joe

inserted phage JBD26 into P. aeruginosa strain PA14, the bacteria gained increased resistance

against several phages, including JBD24, JBD23 and JBD591. JBD26 and JBD16C lysogens

were also observed to cause defects in twitching and swimming motility, respectively91. Even the

insertions of closely related phages in the host genome could result in different host phenotypes.

For example, phages JBD26 and JBD30 have very similar genomic composition, however, their

lysogens display significantly different phage resistance patterns91. Since related phages have

conserved gene orders, Joe was interested in pinpointing the cause of phenotypic modifications

through comparisons of the phage genomes. Because of the high sequence identity conserved

between these phages, Joe was able to identify a number of PrAGs that were not conserved

between strains, and thus were likely responsible for the host phenotypic changes. Through

studies of PrAGs and their effects on bacterial physiology, we hope to gain insight into P.

aeruginosa infections and the development of successful therapies.

1.7 Thesis Objectives

My thesis project aims to determine the specific genes responsible for the phenotypic changes

arising from prophage integration. I conducted a systematic study of PrAGs cloned from the

collection of JBD phages and characterized the effects of their expression on a variety of host

cell phenotypes, including: growth, resistance to phage infection and antibiotics, twitching and

swimming motility, biofilm production, pyocyanin production, and virulence in a Drosophila

malenogaster model of disease.

As the expression of phage genes and the movement of phages between bacterial strains are

common, they can cause variability in bacterial infections and make them difficult to treat.

Depending on the phages present in a given clinical isolate, different types of treatments may be

most effective. A catalog of PrAG phenotypes and other knowledge gained from my work will

11

help in understanding the contribution of phages to bacterial pathogenesis, and form the basis of

future research for personalized treatment against P. aeruginosa infections.

12

Chapter 2 Materials and Methods

2.1 Construction of Expression Plasmids and Protein Test Expression in E. coli

The 17 PrAGs were cloned into p15TV-L E. coli expression plasmids, expressed under the

control of the T7 promoter and tested for protein expression by Diane Bona. In brief, the gene

coding sequences were amplified from PA14 lysogens using pfx DNA polymerase (Invitrogen)

in polymerase chain reactions (PCR) and inserted into p15TV-L plasmids with In-Fusion HD

EcoDryTM Mix (Clontech) following the product protocol. The ligation mixtures were then

transformation into E. coli calcium chloride competent DH5α cells. The colonies obtained were

PCR screened for the correct size gene insert. Positive plasmids were purified using Qiagen

technologies and the correct gene insert was confirmed by DNA sequencing.

Protein test expression was performed to confirm the ability of the PrAGs to produce proteins in

E. coli strain BL21. Overnight cultures grown in LB media were sub-cultured 3 in 100 into fresh

LB media and grown at 37°C shaking until OD600 = 1. 0.8 mM IPTG was then added to the

culture to induce protein expression at 37°C for 3-4 hours. To assess solubility, 1 mL of induced

culture was spun down and resuspended in 200μL BugBuster Protein Extraction Reagent

(Novagen). The samples were incubated at room temperature for 30 minutes, and the cell debris

was collected by contrifugation. 50 μL of supernatant was mixed with 50 μL of 2x SDS-PAGE

loading buffer to prepare SDS-PAGE samples. For the whole cell fractions, 500μL of induced

culture was spun down and resuspended in 200μL 2x SDS-PAGE loading buffer. All SDS-

PAGE samples were boiled for 5 minutes at 100°C before being loaded on to the SDS-PAGE

gel. The expressions of the PrAG proteins were confirmed if protein bands of the expected size

were observed on the coomassie brilliant blue-stained SDS-PAGE gels.

2.2 Transformation of plasmids into P. aeruginosa

The 17 PrAG constructs were obtained from Joe Bondy-Denomy. The constructs have each of

the 17 PrAGs cloned into the pHerd30T expression plasmids individually. The expressions of the

PrAGs were under the control of a pBad promoter, which can be turned on by the addition of

arabinose. The plasmids were transformed into P. aeruginosa PA14 or PAO1 strains through

13

electroporation. To create electro-competent P. aeruginosa cells, overnight culture grown in LB

was spin down and washed twice with 300 mM sucrose, and then resuspended in 1/10th the initial

volume of 300 mM sucrose solution. Fresh made electro-competent cells were placed on ice for

5 minutes before the addition of 1 μL plasmids into 100 μL of cells. The mixture was incubated

on ice for 20 minutes, followed by electroporation at 2500 V. The electroporated cells were

immediately transferred into 900 μL LB and incubated at 37°C, with shaking for 1 hour. The

cells were spin down and plated on LB + gentamycin (50 μM/mL) plates. After overnight 37°C

incubation, any colonies grown on the plate could contain the desired plasmids, and were further

confirmed using PCR colony screens and sequencing.

2.3 PCR Reactions and Colony Screening

Bacterial colonies potentially containing the desired plasmids were screened using PCR before

sending for sequencing. PCR reactions were set-up with 1x Taq buffer with ammonium sulphate

(from 10x Thermo Scientific buffer stock), 3.5 mM magnesium chloride, 2.5 U Taq DNA

polymerase, 0.2 mM dNTPs, 1x PCR Enhancer solution and 2 μM of appropriate primers, and

inoculated with bacteria colonies. pBad forward and M13 reverse primers were used in all

colony PCR reactions with pHerd30T constructs, and the T7 promoter forward and reverse

primers were used in colony PCR reactions with p15TV-L constructs. The PCR program used

consisted of 94°C for 2 minutes, 30 cycles of: 30 seconds at 94°C, 30 seconds at 50°C, 1 minute

at 68°C, followed by 72°C for 10 minutes. The sizes of amplified PCR fragments were

determined by running on standard DNA gels.

2.4 Bacterial Growth

The growth rates of different strains of bacteria were determined by monitoring cell densities at

multiple time points through growth. Overnight cultures were diluted 1:100 into LB +

Gentamycin (50 μM/mL) + 0.1% (w/v) arabinose. 200 μL of diluted cultures were grown in 96-

well plates in Infinite F200 microplate reader (Tecan) with shaking at 37°C for 8 hours. The

growth curves were generated by measuring cell density at OD595 every 15 minutes during

growth.

14

2.5 Twitching Motility Assay

Twitching motility assay was used to assess the ability of the bacteria to move on solid media.

Thin plates containing LB broth + 1% (w/v) agar were prepared on the same day of the assay.

Gentamycin (50 μM/mL) and 0.1% (w/v) arabinose were added to the media prior to plate

pouring. Freshly poured plates were solidified in the biosafety cabinet with lids off for 20

minutes. Plates were inoculated by stabbing isolated colonies grown on overnight LB plates to

the bottom of the twitch plate, and the plates were incubated for 24 hours at 37°C. The agar was

carefully removed from the plates with a spatula, and the cells adhered to the plates were stained

with 1% crystal violet for 1 minute followed by 3 washes with water. The plates were dried

inverted overnight. The distance traveled by the bacteria was then determined by measuring the

diameter of the stain.

The twitching motility assay was performed with 3 technical replicates in each experiment, and

the experiments were repeated 3 separate times (biological replicates). The measurements from

all experiments were analyzed using a T-test, and the values were considered significantly

different from the empty vector control if P-value < 0.005.

2.6 Swimming Motility Assay

To study the movement of the bacteria in liquid media, swimming motility assays were

performed. The media used in this assay contained: 0.3% (w/v) agar, 1% (w/v) tryptone and

0.5% (w/v) NaCl. Thin plates were prepared on the same day of the assay. Gentamycin (50

μM/mL) and 0.1% (w/v) arabinose were added to the media prior to plate pouring. Freshly

poured plates were solidified in the biosafety cabinet with lids off for 20 minutes. Plates were

inoculated by stabbing isolated colonies grown on overnight LB plates to the bottom of the

swimming plate, and the plates were incubated for 24 hours at 37°C. The diameter of the area

traveled by the bacteria was then measured.

The swimming motility assay was performed with 3 technical replicates in each experiment, and

the experiments were repeated 3 separate times (biological replicates). The measurements from

all experiments were analyzed using a T-test, and the values were considered significantly

different from the empty vector control if P-value < 0.005.

15

2.7 Induction of P. aeruginosa Lysogens and Plaquing Assays

P. aeruginosa lysogens were grown overnight in LB media at 37°C. A few drops of chloroform

were added to the overnight culture, and cells were shaken for 30 minutes at 37°C. Cell debris

was collected by centrifugation and the supernatant containing phages was collected and stored

at 4°C.

The sensitivities of bacteria to various phages were assessed using the spotting assay. 150 μL of

overnight bacteria culture was mixed with 3 mL top agar (LB broth + 0.7% (w/v) agar) and

poured on top of thick 1.5% LB agar plates. 2 μL serial dilutions of freshly prepared P.

aeruginosa phage lysates were spotted on the surface of the plates. The plates were incubated

inverted at 37°C overnight. The following day, the phage titres were calculated by noting the

highest dilution where plaques were observed. The phage titres were compared between different

bacterial strains, and the fold changes in phage sensitivities were recorded.

2.8 Drug Resistance Assay

150 μL of overnight culture grown in LB media was mixed with top agar (LB broth + 0.7%

(w/v) agar) and poured on top of thick LB plates (LB broth + 1.5% (w/v) agar + 50 μM/mL

gentamycin + 0.1% (w/v) arabinose). Circular drug-containing discs (Sensi-Disc, BD) were

placed on top of the dried plates, and the plates were incubated at 37°C overnight. The drug

sensitivity of the bacteria was assessed by measuring the diameters of clear zones around the

drug discs.

2.9 Biofilm Production Assay

The assay protocol was adapted from O’Toole (2011)92. Overnight bacterial culture grown in LB

medium was diluted 1:100 into M63 minimum medium (3g monobasic potassium phosphate +

7g dibasic potassium phosphate + 2g ammonium sulphate dissolved in water into 1L solution)

supplemented with 1mM magnesium sulphate, 0.2% (w/v) glucose and 0.5% (w/v) casamino

acids. 100 μL of diluted culture was distributed into 96-well vinyl microtitre plates (Corning) and

incubated at 37°C for 24 hours without shaking. After incubation, the bacterial culture was

removed, the plates were washed 3 times with water and air dried. The wells were stained with

100 μL 0.1% crystal violet solution for 15 minutes at room temperature, followed by 3 washes

with water to get rid of excess stain. The plates were again air dried and the dye was dissolved in

16

125 μL 30% acetic acid for 15 minutes at room temperature. The dissolved dye was transferred

to Infinite F200 microplate reader (Tecan)-compatible 96-well plates, and the levels of biofilm

produced were determined through quatifying the amount of dye present in the wells by

measuring the OD at 595nm.

The biofilm production assay was performed with 3 technical replicates in each experiment, and

the experiments were repeated 3 times at different occasions (biological replicates). The

measurements from all experiments were analyzed using T-test, and the values were considered

significantly different from the empty vector control if P-value < 0.05.

2.10 Virulence Assay

The fly feeding virulence assay was used to determine the virulence of different P. aeruginosa

strains. This assay models the chronic P. aeruginosa infection in cystic fibrosis patient lungs.

The assay protocol was modified from Shen et al. (2012)57. In preparation for the fly feeding

experiment, bacteria were inoculated in LB and grown overnight at 37°C. 5 mL of molten

sucrose agar solution (5% (w/v) sucrose + 2.5% (w/v) agar) was poured into polystyrene

Drosophila vials (VWR) and let solidify overnight. The following day, bacterial cultures were

spun down, resuspended in 100μL 5% sucrose, and spotted onto sterile filter paper (Whatman

GF/A, 21 mm) on top of solidified sucrose agar in the fly vials. The volume of bacterial culture

used varied according to the cell density of different strains so that similar numbers of cells were

distributed in each vial. The vials were incubated at 37°C for 30 minutes, followed by 25°C for

another 30 minutes. Twenty pre-starved 3-5 day old male W1118 Drosophila melanogaster flies

were transferred into each vial using carbon dioxide pads. The vials were placed at 25°C

humidity-controlled dark environment, and the number of dead flies were recorded every 24

hours. Each experiment was performed with 100 flies (5 vials with 20 flies each), and the assay

includes at least 3 individual experiments.

17

Chapter 3 Results

3.1 Acknowledgement

A number of experiments in the study was performed with the help of Diane Bona and Smriti

Kala as outlined below:

(1) The cloning and protein test expression of PrAGs in E. coli was performed by Diane Bona.

(2) The drug resistance assay was conducted by Diane Bona and Smriti Kala.

(3) The virulence assay (fly feeding assay) was performed with the help of Diane Bona and

Smriti Kala.

3.2 P. aeruginosa PrAGs possess diverse genomic and bioinformatics properties

The PrAGs identified in P. aeruginosa phages were distributed throughout the entire phage

genomes (Figure 3). They were commonly found in the transposase, gam, terminase and tail

regions of the Mu-like phage genome (Figure 3A and B), and between the DNA helicase and

protein NinG in phage JBD44 (Figure 3C). Some of the PrAGs had homologs in several phages,

while some were unique and only present in one phage genome. Each of the 17 PrAGs I studied

are small in terms of gene sizes, and they produce proteins of less than 300 amino acids (Table

1). PrAGs are diverse and highly mobile genetic elements, they can frequently be transferred

between bacterial and phage genomes, even among different bacterial species. Some of the

PrAGs I studied have over 100 homologs found in a wide variety of Gram-negative bacterial

strains (Table 1). Within the P. aeruginosa phages I examined, the percent identity among PrAG

proteins range anywhere from 47% to 100% (Table 1). The genomic positions, sequence

identities and numbers of homologs suggests that PrAGs plays important roles in the life cycle of

P. aeruginosa and their presence is greatly influenced by evolution.

18

Figure 3. Genome comparison between related phage strains and positions of PrAGs.

The genome alignment of (A and B) Mu-like phages: JBD26, JBD30, JBD23, JBD5 and JBD24

and (C) phages JBD44 and JBD25. Genomes of close related P. aeruginosa phages were aligned

based on sequence similarities and gene functions. The light grey arrows represent genes of

known functions, and the dark grey arrows represent hypothetical genes. The putative PrAGs

were colored differently according to their homologies. The PrAGs studied in this project were

numbered.

19

Table 1. Summary of PrAGs included in this study.

PrAG

Number PrAG GenBank # Protein Length

(amino acids) Protein Size

(kDa)

% Identity range

for homologues

examined

Number of

homologs by

BLAST* 1 JBD30-4 AFQ21918.1 108 12.5 97-98% 4 2A JBD26-5 AGC24045.1 136 14.4 47-94% 16 2B JBD5-4 AFQ21801.1 69 7.6 47-94% 16 2C JBD24-4 AFQ21860.1 143 15.1 47-94% 29 3 JBD30-9 AFQ21923.1 211 23.6 95-100% 78 4 JBD23-13 N/A 125 13.6 97% 36 5 JBD26-14 AEY99459.1 93 10.0 77-100% 75 6 JBD30-14 AFQ21928.1 67 7.5 N/A 4 7 JBD26-15 AEY99428.1 102 11.2 100% 14 8 JBD5-15 AFQ21812.1 73 8.1 N/A 33 9 JBD24-17 AFQ21873.1 223 23.9 N/A >100 10 JBD26-30 AEY99449.1 83 9.0 100% 2 11 JBD26-31 AEY99485.1 83 9.3 100% 0 12 JBD30-30 AFQ21944.1 164 18.3 100% >100 13 JBD26-61 AEY99477.1 74 8.5 85-99% 32 14 JBD44-8 N/A 52 5.7 N/A 2 15 JBD44-9 N/A 77 8.6 N/A 1 *Proteins with BLAST E-value lower than 10-4 are considered homologs.

3.3 The PrAGs all express and make proteins in E. coli

Since the PrAGs that I am screening were mostly identified through genome comparison and

bioinformatics, it is important to determine if these open reading frames (ORFs) actually encode

proteins. All 17 PrAGs were cloned into Escherichia coli expression vectors and most of them

were shown to express and produce proteins (Figure 4). JBD30-9 may be expressed at a low

level, which could not be captured by the Coomassie blue dye.

20

Figure 4. Protein expression of PrAGs in E. coli.

The PrAGs were cloned into p15TV-L expression vectors and induced in E. coli BL21 strain.

Protein expression of individual PrAGs was examined in whole cell lysate (w) and soluble

fraction (s) on Coomassie blue-stained sodium dodecyl sulfate polyacrylamide gel

electrophoresis (SDS-PAGE) gels. All PrAG protein bands observed are close to the expected

sizes (Refer to Table 1 for PrAG protein sizes). Note that JBD26-61is known to proteolyze at a

fast rate, resulting in two protein bands observed in the soluble fraction.

21

3.4 Some PrAGs introduce growth defects in the bacterial host

To determine if the expression of any individual PrAG affects the growth of P. aeruginosa, I

examined the growth rates of each of the strains expressing a PrAG from a plasmid. P.

aeruginosa PA14 and PAO1 strains overexpressing individual PrAGs of interest were grown in

LB medium at 37°C for 8 hours, and cell density was monitored at OD595 in the TECAN

microplate reader during growth. While most PrAGs have little effects on host growth, four

PrAGs, JBD26-15 and JBD30-14 in PA14 and JBD44-8 and JBD5-4 in PAO1, slowed bacterial

cell growth (Figure 5). The four PrAGs have varying levels of effects on bacterial growth. At the

eighth hour, the cell densities of bacterial strains expressing JBD30-14 and JBD5-4 were 19%

and 17% lower than strains expressing no PrAGs (empty vector control), respectively, which is

mild compared to the effects of JBD44-8 and JBD26-15. When JBD44-8 was expressed in

PAO1, the bacterial cell density was more than 30% lower than PAO1 empty vector control at

the 8-hour mark (Figure 5D). The expression of JBD26-15 in PA14 strain causes the most

serious growth defect among all the PrAGs examined. After 8 hours of growth, the cell density

of PA14 strain expressing JBD26-15 was less than 45% of the empty vector control strain, and it

reached the bacterial growth log phase after 6 hours of growth, which was approximately 4 hours

later than the control strain (Figure 5B). These bacterial growth experiments demonstrated that

PrAGs are able to influence the growth rate of bacterial hosts. It is unclear if the growth defects

affected other phenotypic screens performed, but it is important to take them into account during

analysis and interpretation.

22

23

Figure 5. PrAG expression affect bacterial growth rate.

Different PrAGs were expressed in (A and B) PA14 or (C and D) PAO1 P. aeruginosa strains,

and the cell density of the growing liquid culture was monitored every 15 minutes for 8 hours.

Most of the PrAGs had little effects on bacterial growth rate (A and C), while some slowed the

growth of the host (B and D).

24

3.5 PrAGs increase bacterial survivability by protecting their hosts from phage infection

As prophages and their bacterial hosts exist in a symbiotic relationship, the survivability of the

bacteria is important to both. The longer the lysogen lives and replicates, the more copies of the

phage genome there will be. Since phage infection and killing is one of the most common factors

in limiting the sizes of bacteria populations, resistance against further phage infection can

therefore provide a great advantage in bacterial survivability. Thus, I examined the effects of

PrAG expression on bacterial resistance against a panel of P. aeruginosa phages. I spotted serial

dilutions of 31 different phages atop lawns of PA14 and 18 phages on lawns of PAO1 expressing

individual PrAGs. I found five PrAGs in PA14 (Table 2) and six PrAGs in PAO1 (Table 3) that

increased the bacterial resistance against several phages. JBD30-30, JBD26-5, JBD26-31,

JBD26-61 and JBD24-4 all caused increased phage resistance in both PA14 and PAO1, while

JBD44-8 only had an effect in PAO1. The levels and patterns of phage resistance caused by

PrAGs differ from PrAG to PrAG even within the same bacterial strain. For example, the

expression of JBD26-61 and JBD24-4 led to increased resistance against more phages than the

expression of JBD30-30, JBD26-5 and JBD26-31 in PA14 (Table 2). In addition, JBD44-8 only

caused a slight increase in phage resistance in PAO1, while JBD24-4 was able to protect its host

completely from infections by most phages in our collection (Table 3). Interestingly, the

expression of JBD26-61 and JBD24-4 resulted in identical phage resistance patterns, suggesting

that they may be modifying the host phenotypes via similar mechanisms. Since phage infection

largely depend on bacterial cell surface structures – T4P and O-antigen, as they serve as common

phage binding sites during initial adsorption, the data of phage spotting on PA14 knockouts of

T4P (pilA) and O-antigen (wbpL) adapted from study done by Bondy-Denomy91 is also included

for clarity (Table 2).

25

Table 2. Increase in phage resistance caused by PrAG expression in PA14 compared to no

PrAG expression.

PA14 Knockout* Phage

PrAG

wbpL pilA JBD30-30 JBD26-5 JBD26-31 JBD26-61 JBD24-4

JBD16C

JBD63

JBD5

JBD10**

JBD23

JBD26

JBD32

JBD33

JBD35C

JBD59a

JBD60a

DMS3

MP22

MP29

JBD95b

JBD86

JBD63c

JBD93a

JBD8

JBD13

JBD30

JBD88a

JBD88b

JBD16

JBD1

JBD24

JBD44a

JBD62b

JBD69a

JBD70a

JBD80

*Data adapted from study done by Bondy-Denomy91. Only complete resistance is

shown.

**JBD10 infects PA14 poorly.

Legend

Sensitive

Slightly resistant (>100-fold)

Intermediately resistant (>10^4-fold)

Highly resistant (>10^6-fold)

Complete resistant

26

Table 3. Fold increase in phage resistance caused by PrAG expression in PAO1 compared

to no PrAG expression.

Phage PrAG

JBD30-30 JBD44-8 JBD26-5 JBD26-31 JBD26-61 JBD24-4

JBD16C

JBD63

JBD5

JBD10

JBD23

JBD26

JBD32

JBD33

JBD35C

JBD59a

JBD60a

DMS3

MP22

MP29

JBD95b

JBD86

JBD63c

JBD93a

JBD8

JBD13

JBD30

JBD88a

JBD88b

JBD16

JBD1

JBD24

JBD44a

JBD62b

JBD69a

JBD70a

JBD80

Legend

Sensitive

Slightly resistant

(>100-fold)

Intermediately resistant

(>10^4-fold)

Highly resistant

(>10^6-fold)

Complete resistant

Phages that do not

infect wild-type PAO1

27

Some PrAGs were found to have different effects on phage resistance in different P. aeruginosa

strains. I compared phage resistance caused by PrAGs in PA14 and PAO1 and found that in most

cases where PrAGs caused increased phage resistance, it was observed in both strain

backgrounds tested. However, there were some examples of resistance only seen in PAO1 (Table

4). Among the phages that are able to infect both PAO1 and PA14 strains, the only case of a

PrAG affecting phage resistance only in PA14 is the case of JBD26-31, which only causes

resistance in PA14 against phage JBD60a (Table 4). I have also noted that in PAO1 three PrAGs

that caused complete resistance against all the phages I tested: JBD26-5, JBD26-61 and JBD24-

4. However, these PrAGs did not cause complete resistance in PA14 (Table 2 and 3). The results

from the phage resistance screens suggests that multiple PrAGs function to increase the

resistance of the bacteria against various phages. As the patterns of resistance are unique for each

PrAG, they are likely acting through different mechanisms, or interacting with different host

components in the bacterial cell.

28

Table 4. Difference in phage resistance caused by PrAG expression in PA14 and PAO1.

Phage PrAG

JBD30-30 JBD44-8 JBD26-5 JBD26-31 JBD26-61 JBD24-4

JBD16C

JBD63

JBD5

JBD10

JBD23

JBD26

JBD32

JBD33

JBD35C

JBD59a

JBD60a

DMS3

MP22

MP29

JBD95b

JBD86

JBD63c

JBD93a

JBD8

JBD13

JBD30

JBD88a

JBD88b

JBD16

JBD1

JBD24

JBD44a

JBD62b

JBD69a

JBD70a

JBD80

Another major factor determining the survivability of the bacteria is how well the bacteria can

survive and replicate in the presence of antibiotics. The ability to withstand antibiotics increases

the chance of successful infection and propagation of the bacteria inside mammalian systems.

We examined 9 antibiotics commonly used to treat bacterial infections (Table 5) using the disk

diffusion method and determined that none of the 17 PrAGs had effects on drug resistance.

Legend

Sensitive in both PA14

and PAO1

Resistance observed in

PA14 only

Resistance observed in

PAO1 only

Resistance observed in

both PA14 and PAO1

Phages that infect wild-

type PA14 but not PAO1

29

Table 5. List of drugs tested in drug resistance assay.

Drug Name Amount (μg) Azithromycin 15 Cefoperazone 75 Ceftazidime 30 Imipenem 10 Ticarcillin 75 Piperacillin 100 Colistin 10 Amikacin 30 Aztreonam 30

3.6 Bacterial motility is often modified by PrAGs

Bacterial motility is considered a virulence factor for P. aeruginosa as the motility of the

bacterial facilitates invasive infection of their mammalian hosts71,73. I performed two motility

screens to study the ability of the bacteria to travel in different environments when expressing

PrAGs of interest. Twitching motility describes the motility of the bacteria on solid media.

Through the twitching motility screen, I found that more than half of the PrAGs I screened

decrease bacterial twitching. JBD26-15, JBD23-13, JBD44-8, JBD26-5, JBD26-61, JBD5-4 and

JBD24-4 cause twitching defects in both PA14 and PAO1 strains, and JBD30-4 and JBD24-17

only affects PAO1 (Figure 6). Within the same P. aeruginosa strain, different PrAGs modified

twitching motility to different degrees. JBD26-5, JBD26-61, JBD5-4 and JBD24-4 completely

abolished twitching motility, comparable to the control where the pili was knocked out, while the

other PrAGs only cause mild to moderate defects (Figure 6). Given the differences in degree of

modification within and between bacterial strains, it is possible that the PrAGs are able to work

through several different mechanisms to alter bacterial motility.

30

Figure 6. Some PrAGs affect twitching motility of P. aeruginosa.

Twitching motility of P. aeruginosa (A) PA14 and (B) PAO1 strains over-expressing different

PrAGs were measured as diameter traveled by the bacteria over 24 hr on solid medium. P.

aeruginosa strains with no PrAG expressed (empty vector) and Type IV pilus knockout control

(pilA knockout) were also included. T-test was used to determine statistically significant results

compared to empty vector control and they are marked with asterisks (P<0.005), error bars

represent the standard deviation of sample groups.

31

In addition to twitching motility, I examined the effects of PrAG expression on bacterial

swimming motility, which describes the ability of bacteria to move in liquid media. I performed

the swimming motility assay using semi-solid medium to mimic liquid environments, which

allow the measurement of distance traveled by the bacteria. Three PrAGs caused swimming

defects in both PA14 and PAO1, including JBD30-4, JBD26-15 and JBD44-8, while JBD23-13

only has an effect in PAO1 (Figure 7). Similar to the twitching motility screen, the PrAGs altered

bacterial swimming motility to different degrees. However, the effects of PrAGs on swimming

motility were mostly mild to moderate, and no PrAGs were able to cause swimming defects as

serious as the flagellum knockout negative control (Figure 7).

Using the bacterial motility screens, I found a number of PrAGs can modify the ability of the

bacteria to travel on solid or in semi-solid medium, and some can affect both.

32

Figure 7. PrAGs could alter swimming motility of the bacteria.

Swimming motility of P. aeruginosa (A) PA14 and (B) PAO1 strains over-expressing different

PrAGs were measured as diameter traveled from central point by the bacteria over 24 hr in low

percentage medium. P. aeruginosa strains with no PrAG expressed (empty vector) and flagellum

knockout control (fliC knockout) were also included. T-test was used to determine statistically

significant results compared to empty vector control and they are marked with asterisks

(P<0.005), error bars represent the standard deviation of sample groups.

33

3.7 PrAGs are capable of altering the production of bacterial virulence factors

P. aeruginosa produces a number of virulence factors to aid in the invasion of mammalian

systems, which include biofilm and pyocyanin. Biofilm mediates bacterial attachment to target

mammalian cells and it can also help the bacteria in evading mammalian immune systems and

antibiotics79,80. To investigate if PrAG expression alters the level of biofilm produced by the

bacterial host, I measured the biofilm production of PA14 and PAO1 strains expressing different

PrAGs. I found seven PrAGs affected the amount of biofilm produced by P. aeruginosa. JBD23-

13, JBD26-61, JBD5-4 and JBD24-4 in PA14 and JBD30-4, JBD26-15, JBD26-5 and JBD26-61

in PAO1 decreased bacterial biofilm production (Figure 8). The only case where PrAG increased

level of biofilm production by the host is when expressing JBD23-13 in PAO1 (Figure 8B).

Interestingly, JBD23-13 has an opposite effect when expressed in PA14 where it decreased

biofilm production (Figure 8A). JBD26-61 is the only PrAG that behaved similarly in both PA14

and PAO1 strains, where it was able to lower the production of biofilm to the level comparable

with the pili and flagella knockout controls (pilA and fliC knockout, Figure 8). One thing to note

here is that PAO1 strains seem to grow better in the minimal media I used for this particular

assay, therefore they have overall higher biofilm production than PA14 strains. The results of the

experiments show that PrAGs are able to either increase or decrease the production of biofilm by

their hosts, and the effects can vary depending on the bacterial strain.

34

Figure 8. PrAGs cause changes in bacterial biofilm production.

The level of biofilm production for P. aeruginosa (A) PA14 and (B) PAO1 strains over-

expressing different PrAGs were measured. P. aeruginosa strains with no PrAG expressed

(empty vector) serves as wild-type control. As controls, Type IV pilus and flagellum knockouts

(pilA and fliC knockout) were included. T-test was used to determine statistically significant

results compared to empty vector control and they are marked with asterisks (P<0.05), error bars

represent the standard deviation of sample groups.

Pyocyanin produced by P. aeruginosa is a toxin as well as a colour pigment that is responsible

for the greenish blue color of the bacteria. The amount of pyocyanin produced by the bacteria is

35

dependent on many factors, including the growth conditions and quorum sensing systems.

However, because of the color of pyoaynin, a significant change in the production level can be

observed by eye. One really interesting PrAG – JBD5-4 changed the level of pyocyanin

production in both PA14 and PAO1 depending on the environment the bacteria is growing in.

When the bacterial host was grown on solid medium, the expression of JBD5-4 greatly increased

pyocyanin production (Figure 9A), while JBD5-4 decreased pyocyanin production when the

bacteria were grown in liquid medium (Figure 9B). This observation suggested that JBD5-4 may

be able to interact with host components that the bacteria uses to sense growing environment,

and alter the amount of pyocyanin produced accordingly.

Figure 9. JBD5-4 alters the levels of pyocyanin produced by P. aeruginosa.

P. aeruginosa PA14 strains expressing PrAG JBD5-4 or no PrAG were grown on (A) solid

medium and (B) liquid medium, and the amount of pigment produced was compared.

36

3.8 Two PrAGs modify bacterial virulence in Drosophila melanogaster

Phages are known to alter the degree of bacterial virulence through a variety of different

mechanisms, ranging from encoding genes for the production of toxins in their genomes to

helping their host evade mammalian immune system. To investigate if the expression of PrAGs

has an effect on the virulence level of P. aeruginosa, I infected Drosophila melanogaster (fruit

flies) with P. aeruginosa PA14 strain expressing different PrAGs to mimic the bacterial chronic

infection seen in the lungs of cystic fibrosis patients. Because of the striking similarities between

mammalian and Drosophila innate immunity responses, the flies provide an excellent model

organism to study pathogenesis of chronic P. aeruginosa infections58. Through the virulence

assay, we found two PrAGs that were able to modify the bacterial host virulence. I observed that

fruit flies infected with PA14 expressing JBD5-4 live longer than flies fed with PA14 empty

vector control. Approximately half of the flies in the control group died within the first eight

days post infection, when 91% of the flies were still alive in the JBD5-4 group (Figure 10A). By

contrast, JBD44-8 had the opposite effect on bacterial virulence. JBD44-8 increased the rate of

fly killing by the bacteria. By day 9, almost all the flies infected by JBD44-8-expressing PA14

strain were all dead, while 67% of the flies infected with the empty vector strain survived (Figure

10B). There exists some discrepancies between the fly survival profiles in the control group from

experiments done on different days, which were most likely due to the conditions of the bacteria

and flies on particular days. Since the experiments were repeated on at least three different

occasions, and we observed the same trend in all of them, the differences in results between

controls and samples we seen were likely due to PrAG expression. The virulence assay

demonstrated that the effects of PrAGs on their host are significant enough to modify bacterial

virulence in the chronic infection of fruit flies, which suggests that PrAGs could also play crucial

roles in bacterial infection of mammalian systems.

37

Figure 10. PrAGs have effects on bacterial virulence towards Drosophila malenogaster.

P. aeruginosa PA14 strains expressing different PrAGs or no PrAG (empty vector, EV) were fed

to D. malenogaster. The number of dead flies were recorded every 24 hours. (A) The survival

rate of flies fed with PA14 strain expressing JBD5-4 (○) was compared with PA14 strain

harbouring an empty expression vector (■). (B) The survival rate of flies fed with PA14 strain

expressing JBD44-8 (∆) was compared to PA14 strain harbouring an empty expression vector

(■).

38

Chapter 4 Discussion and Future Directions

Phages have been shown to play very important roles in bacterial ecosystems. While prophages

are known to express PrAGs that function to alter the physiology of their hosts, no systematic

studies had investigated the bacterial phenotypes caused by expressions of a variety of PrAGs.

The experiments described in this thesis aimed to assess the diversity of PrAGs in a closely

related group of P. aeruginosa phages and determine their effects on bacterial phenotypes related

to survivability and virulence in a systematic manner. Through a number of phenotypic screens, I

found that P. aeruginosa PrAGs are able to modify various host phenotypes, including

swimming and twitching motility, phage resistance, biofilm production, pyocyanin production,

and degree of virulence against Drosophila fruit flies. More than half (11/17) of the PrAGs I

studied were observed to affect host phenotypes in at least one screen (Table 6), and they

possibly alter other bacterial phenotypes that were not tested in this study, which suggests that

the role of PrAGs in bacterial biology may be more prevalent than previously appreciated. The

phenotypic effects of PrAGs vary based on the individual PrAGs and the bacterial strain that the

PrAG is expressed in. Some PrAGs, such as JBD30-30 and JBD26-31, were only able to modify

a single host phenotype while others, like JBD26-61 and JBD24-4, can alter multiple phenotypes

(Table 6), which likely depends on the mechanism by which each PrAG functions. It is also

noted that a number of phenotypic modification are bacterial strain-specific. For example, the

expression of JBD23-13 in strain PA14 resulted in decreased biofilm production but it increased

the production of biofilm in strain PAO1 (Table 6), suggesting that PrAGs interact with host

elements to cause changes in phenotypes.

Although various functional studies are still required to characterize the biological roles of the

PrAGs, it is possible to make predictions on how the PrAGs function to alter host phenotypes

based on the results of the phenotypic screens. One example includes PrAGs JBD24-4, JBD26-5

and JBD26-61, all of which caused decreased twitching motility and biofilm production and

increased bacterial resistance against phages (Table 6). The bacterial T4P mediates twitching

motility68 and it is involved in the biofilm synthesis72. In addition, as a cell surface structure

which serves as a binding site during phage infections93, the pilus is essential for certain phages

to infect. It is very likely that these three PrAGs function to down-regulate or affect the proper

functioning of the host T4P, which leads to the reduction in twitching motility and biofilm

39

production, and protects the host from phages that use pili to infect bacteria. A recent study has

shown that protein gp05 of P. aeruginosa phage D3112, which is identical to JBD26-5 in

sequence, interacts and interferes with the activity of the host T4P assembly/extension ATPase,

PilB94. This result is consistent with the phenotypes I observed through the screens. However,

the functional amino acids or regions in gp05 responsible for the interaction are still unclear, and

the PrAG may have other functions in bacterial cells that are not yet identified.

Another example is JBD5-4, which was observed to decrease bacterial twitching motility,

biofilm production and cause increased or decreased level of host pyocyanin production when

grown in liquid or on solid medium, respectively (Table 6). All three biological processes of the

bacteria are controlled by the quorum-sensing system66. Therefore JBD5-4 may be interacting

with host components regulating the quorum-sensing system. In their 2002 publication, Scott A.

Beatson and his colleagues found that a regulator of quorum-sensing in P. aeruginosa, Vfr, was

able to alter level of pyocyanin production based on different growth media95. It is possible that

there exist a quorum-sensing regulator that can sense the difference between liquid and solid

medium which JBD5-4 is interacting with. Indeed, a more recent study has shown that mutations

in the mexT or mexF gene in P. aeruginosa strain PAO1 could lead to defective MexEF-OprN

pump, and results in enhanced pyocyanin production in liquid medium and attenuated production

on solid medium96,97. Since the MexEG-OprN pump has been suggested to be involved in the

efflux of quorum sensing signalling molecules98, it is a likely candidate to interact with JBD5-4.

According to protein sequence alignments, JBD5-4, JBD26-5 and JBD24-4 are protein

homologs. They share a central domain with 40-60% identity, while JBD26-5 has an extra N-

terminal domain, and JBD24-4 has a C-terminal domain. Although the three PrAGs all caused

twitching defects, their effects on phage resistance, biofilm and pyocyanin production differ,

despite being protein homologs. The results of the screens suggested that the differences in

phenotypic modifications may be caused by the non-homologous sequences in the PrAGs, and

this also suggests the possibility of one PrAG interacting with multiple host components to affect

bacterial phenotypes.

Among all the PrAGs I studied, two sets of PrAGs have very similar phenotypic effects on P.

aeruginosa. First, JBD24-4 and JBD26-61 both abolished twitching motility, increased host

resistance against the same phages and decreased biofilm production in the PA14 strain (Table

40

6). The only difference was JBD26-61 also affected biofilm production in PAO1, but JBD24-4

did not. This is interesting because JBD26-61 and JBD24-4 have no sequence similarity, and

their genome positions are very different. The other set of PrAGs: JBD26-31 and JBD30-30 both

increased phage resistance of the host without causing defects in bacterial motility or biofilm

production (Table 6), which suggests that they might function at subsequent steps downstream of

cell binding during phage infections. JBD26-31 and JBD30-30 have no sequence similarity,

however they occupy the same genome position right between the small and large terminase

(Figure 3). It is fascinating in the evolutionary point of view how different PrAGs may evolve to

function through similar mechanisms to affect host phenotypes in similar ways, which indicates

that the biological roles of PrAGs may be more relevant than previously known.

One PrAG, JBD23-13, has opposite effects on level of biofilm production under PA14 and

PAO1 background. It decreased host biofilm production in PA14, however when expressed in

PAO1, it increased the production of biofilm. In addition, JBD23-13 causes a twitching defect in

strain PA14 and a swimming defect in strain PAO1 (Table 6). Since all the phenotypic effects

caused by JBD23-13 were strain-specific, it is possible that the PrAG interacts with strain-

specific host components or its activity is induced by strain-specific signals. More studies are

required to explain how JBD23-13 functions in the host cell. Besides JBD23-13, differences

between phenotypes observed in strains PA14 and PAO1 were also seen for other PrAGs. For

examples, JBD30-4 caused twitching defects in PAO1 only, and similarly, JBD5-4, decreased

biofilm production in PA14 but not in PAO1 (Table 6). Although the two commonly used lab

strains are very similar in genome sequences, approximately 4-8% of the genome is unique to

only one strain and may be responsible for the differences observed in this study99. It is known

that the PA14-specific genomic regions contain a few pathogenicity islands, which account for

the difference in pathogenicity between PA14 and PAO1100. PrAGs that were observed to cause

strain-specific effects may interact with components found in unique regions, including Type IV

pilin, Type IV fimbrial assembly protein, ABC transporter-like protein, DNA helicase-like

protein, O-antigen biosynthesis proteins, putative transcriptional regulators and other genes of

unknown functions100,101. For example, in the phage resistance screen, the expression of JBD24-4

in PAO1 resulted in complete host resistance against all phages tested. However, JBD24-4 only

increased the resistance of PA14 against some of the phages. Since the bacterial T4P is involved

41

in initial phage infections, JBD24-4 possibly interferes with strain-specific components of the

pilus system, such as the pilin (PilA) or fimbrial assembly protein (pilC), or related regulators.

Two other PrAGs, JBD26-15 and JBD44-8, have effects on twitching and swimming motility in

both PA14 and PAO1 strains (Table 6), suggesting that they may play a role in quorum-sensing,

which is essential for T4P and flagellum functions. Although JBD26-15 and JBD44-8 interfered

with bacterial pili and flagella, they did not have significant effect on phage resistance except

JBD44-8 slightly increased the phage resistance of PAO1 strain. The PrAGs possibly affect the

proper functioning of the pili and flagella rather than the synthesis, so phages can still bind to the

host cell during infections. While pilus and flagellum are considered to be important virulence

factors during bacterial infections, it is interesting how JBD44-8 causes motility defects but

increased virulence in fruit flies. As previously mentioned, JBD44-8 likely interacts with

components in the quorum-sensing system. Therefore one possible explanation is that while it

interferes with bacterial motility, JBD44-8 enhances the functions of other virulence factors

controlled by quorum-sensing (e.g. exotoxin, protease), which were not captured by the screens

performed in this study.

Interestingly, the motility and biofilm production screens generated results which were opposite

to what is expected. Since bacterial motility and the production of biofilm often translates into

bacterial virulence, I was expecting that the PrAGs increase bacterial motility and the level of

biofilm production to aid bacterial infection, so the phages can replicate with their hosts. It is

possible that the phages are sacrificing these two virulence factors for something that provides

more advantages to themselves. For example, as previously mentioned in the case of JBD24-4,

JBD26-5 and JBD26-61, the phages might be down-regulating bacterial T4P to protect the host

from further phage infections, but it also resulted in motility and biofilm production defects. One

thing to note is that most phages and their P. aeruginosa host live in natural environments, and

Pseudomonas does not need to infect to survive. Therefore, the ability to resist phage infections

and increase survivability is likely more important for both the phage and its host than virulence

against other organisms.

Through the experiments done in this study, several PrAGs with interesting phenotypic effects in

P. aeruginosa were identified. However, only a small number of PrAGs were screened for

specific phenotypes, there are a lot more PrAGs identified in P. aeruginosa that were not

42

included in this study, and PrAGs likely affect the hosts in other ways that were not captured by

the screens performed. Another limitation of the study is that the phenotypes observed in the

screens were due to the overexpression of PrAGs, and it is not clear if the same phenotype

modifications occur outside of lab environments, since we do not know the endogenous

expression levels of PrAGs in P. aeruginosa at this point. However, the study does show that the

PrAGs have the potential to cause bacterial phenotypic changes, and whether they happen or not

depends on multiple factors, including environmental conditions, external cues and interactions

between PrAGs and other prophage or host components. In the study, I have only investigated

the phenotypic effects of the expression of individual PrAGs. In natural environments, it is

common for more than one PrAG to be expressed inside the host cell. Based on the findings of

the study, I cannot draw any conclusion on how the expression of multiple PrAGs interact with

each other and what the resulting bacterial phenotypes will be.

In order to expand our knowledge of the biological roles of PrAGs in P. aeruginosa, follow-up

studies are required to understand the mechanisms employed by the PrAGs to modify bacterial

physiology. Based on the phenotypic screen results, it is likely that PrAGs function through

interacting with host components. Therefore a potential next step in the study will be to identify

the physical interacting partners of PrAGs through protein pull-down assay, or use mutagenesis

to pinpoint genetic interactions between PrAGs and host genes. One can also take advantage of

RNAseq technology to study the effects of PrAGs on host genetic expression profile, which may

provide clues for the functions of the PrAGs. In addition, structural studies may prove useful in

determining signature motifs and PrAG function predictions. Combining phenotypic screens

with various biochemical and genetic approaches, one can more efficiently characterize the

functions of the PrAGs. As previously mentioned, in addition to the PrAGs studied in my thesis,

there are a lot of putative PrAGs with unknown functions, and new PrAGs are continuously

being identified. In the future, the systematic study can be expanded to include more PrAGs and

screen for more host phenotypes, such as swarming motility and production of other virulence

factors like pyoverdine and elastase. It will also be interesting to investigate the interaction

between different PrAGs when expressed simultaneously and the phenotypic consequences.

PrAGs are known to play important roles in bacterial physiology by altering different biological

processes of the host life cycle, however no systematic studies have been done to address the

effects of PrAGs on their bacterial hosts. Multiple screens were employed in this study to

43

systematically examine the phenotypic effects of PrAG expression in bacterial cells. The results

have shown that many P. aeruginosa PrAGs have the ability to modify bacterial phenotypes

related to survivability and virulence (Figure 11), and these events may contribute significantly

to bacterial infection and disease progression. Therefore, when studying human diseases caused

by bacteria, it is important to take into account the bacteriophages and what they can do through

PrAG expression. I hope the facts learned from this study will offer inspiration for more research

on bacteriophages and their contribution in human diseases, which may become the foundation

for possible treatment in the future.

Table 6. Summary of host phenotypic changes caused by PrAGs in P. aeruginosa PA14 and

PAO1 strains.

44

Figure 11. PrAG expression affect various bacterial elements.

The phenotypic screens conducted in this study have shown that the expression of PrAGs

modifies multiple host elements related to survivability and virulence. The PrAGs found to affect

specific elements are listed.

45

References

1 Hendrix, R. W., Smith, M. C., Burns, R. N., Ford, M. E. & Hatfull, G. F. Evolutionary

relationships among diverse bacteriophages and prophages: all the world's a phage.

Proceedings of the National Academy of Sciences of the United States of America 96,

2192-2197 (1999).

2 Letarov, A. & Kulikov, E. The bacteriophages in human- and animal body-associated

microbial communities. Journal of applied microbiology 107, 1-13, doi:10.1111/j.1365-

2672.2009.04143.x (2009).

3 Breitbart, M. et al. Metagenomic analyses of an uncultured viral community from human

feces. Journal of bacteriology 185, 6220-6223 (2003).

4 King, A. M. Q., Adams, M.J., Carstens, E.B. and Lefkowitz, E.J. Virus taxonomy:

classification and nomenclature of viruses: Ninth Report of the International Committee

on Taxonomy of Viruses. 1st edn, (Elsevier Academic Press, 2012).

5 Calendar, R. The Bacteriophages. 2nd edn, (Oxford University Press, 2006).

6 Hyman, P. & Abedon, S. T. Bacteriophage host range and bacterial resistance. Advances

in applied microbiology 70, 217-248, doi:10.1016/S0065-2164(10)70007-1 (2010).

7 Suttle, C. A. Viruses in the sea. Nature 437, 356-361, doi:10.1038/nature04160 (2005).

8 Borek, E. & Ryan, A. Lysogenic induction. Progress in nucleic acid research and

molecular biology 13, 249-300 (1973).

9 Choi, J., Kotay, S. M. & Goel, R. Various physico-chemical stress factors cause prophage

induction in Nitrosospira multiformis 25196--an ammonia oxidizing bacteria. Water

research 44, 4550-4558, doi:10.1016/j.watres.2010.04.040 (2010).

10 Banks, D. J., Lei, B. & Musser, J. M. Prophage induction and expression of prophage-

encoded virulence factors in group A Streptococcus serotype M3 strain MGAS315.

Infection and immunity 71, 7079-7086 (2003).

11 Dalmasso, M., Hill, C. & Ross, R. P. Exploiting gut bacteriophages for human health.

Trends in microbiology 22, 399-405, doi:10.1016/j.tim.2014.02.010 (2014).

12 Cumby, N., Davidson, A. R. & Maxwell, K. L. The moron comes of age. Bacteriophage

2, 225-228, doi:10.4161/bact.23146 (2012).

13 Wagner, J. et al. Bacteriophages in gut samples from pediatric Crohn's disease patients:

metagenomic analysis using 454 pyrosequencing. Inflammatory bowel diseases 19, 1598-

1608, doi:10.1097/MIB.0b013e318292477c (2013).

14 Hedin, C. R. et al. Altered intestinal microbiota and blood T cell phenotype are shared by

patients with Crohn's disease and their unaffected siblings. Gut 63, 1578-1586,

doi:10.1136/gutjnl-2013-306226 (2014).

46

15 Xavier, R. J. & Podolsky, D. K. Unravelling the pathogenesis of inflammatory bowel

disease. Nature 448, 427-434, doi:10.1038/nature06005 (2007).

16 Lepage, P. et al. Dysbiosis in inflammatory bowel disease: a role for bacteriophages? Gut

57, 424-425, doi:10.1136/gut.2007.134668 (2008).

17 Wittebole, X., De Roock, S. & Opal, S. M. A historical overview of bacteriophage

therapy as an alternative to antibiotics for the treatment of bacterial pathogens. Virulence

5, 226-235, doi:10.4161/viru.25991 (2014).

18 Alemayehu, D. et al. Bacteriophages phiMR299-2 and phiNH-4 can eliminate

Pseudomonas aeruginosa in the murine lung and on cystic fibrosis lung airway cells.

mBio 3, e00029-00012, doi:10.1128/mBio.00029-12 (2012).

19 Chhibber, S., Kaur, T. & Sandeep, K. Co-therapy using lytic bacteriophage and linezolid:

effective treatment in eliminating methicillin resistant Staphylococcus aureus (MRSA)

from diabetic foot infections. PloS one 8, e56022, doi:10.1371/journal.pone.0056022

(2013).

20 Krylov, V., Shaburova, O., Krylov, S. & Pleteneva, E. A genetic approach to the

development of new therapeutic phages to fight pseudomonas aeruginosa in wound

infections. Viruses 5, 15-53, doi:10.3390/v5010015 (2013).

21 Mendes, J. J. et al. Wound healing potential of topical bacteriophage therapy on diabetic

cutaneous wounds. Wound repair and regeneration : official publication of the Wound

Healing Society [and] the European Tissue Repair Society 21, 595-603,

doi:10.1111/wrr.12056 (2013).

22 Ksendzovsky, A. et al. Convection-enhanced delivery of M13 bacteriophage to the brain.

Journal of neurosurgery 117, 197-203, doi:10.3171/2012.4.JNS111528 (2012).

23 Kia, A., Yata, T., Hajji, N. & Hajitou, A. Inhibition of histone deacetylation and DNA

methylation improves gene expression mediated by the adeno-associated virus/phage in

cancer cells. Viruses 5, 2561-2572, doi:10.3390/v5102561 (2013).

24 Shoae-Hassani, A. et al. lambda Phage nanobioparticle expressing apoptin efficiently

suppress human breast carcinoma tumor growth in vivo. PloS one 8, e79907,

doi:10.1371/journal.pone.0079907 (2013).

25 Johnson, R. P. et al. Bacteriophages for prophylaxis and therapy in cattle, poultry and

pigs. Animal health research reviews / Conference of Research Workers in Animal

Diseases 9, 201-215, doi:10.1017/S1466252308001576 (2008).

26 Chibeu, A. et al. Efficacy of bacteriophage LISTEXP100 combined with chemical

antimicrobials in reducing Listeria monocytogenes in cooked turkey and roast beef.

International journal of food microbiology 167, 208-214,

doi:10.1016/j.ijfoodmicro.2013.08.018 (2013).

47

27 Fujii, N., K. Oguma, N. Yokosawa, K. Kimura, and K. Tsuzuki. Characterization of

bacteriophage nucleic acids obtained from Clostridium botulinum types C and D. Appl.

Environ. Microbiol. 54, 69-73 (1988).

28 Holmes, R. K. & Barksdale, L. Genetic analysis of tox+ and tox- bacteriophages of

Corynebacterium diphtheriae. Journal of virology 3, 586-598 (1969).

29 Uchida, T., Gill, D. M. & Pappenheimer, A. M., Jr. Mutation in the structural gene for

diphtheria toxin carried by temperate phage. Nature: New biology 233, 8-11 (1971).

30 Waldor, M. K. & Mekalanos, J. J. Lysogenic conversion by a filamentous phage

encoding cholera toxin. Science 272, 1910-1914 (1996).

31 Huang, A. et al. Cloning and expression of the genes specifying Shiga-like toxin

production in Escherichia coli H19. Journal of bacteriology 166, 375-379 (1986).

32 Newland, J. W., Strockbine, N. A., Miller, S. F., O'Brien, A. D. & Holmes, R. K. Cloning

of Shiga-like toxin structural genes from a toxin converting phage of Escherichia coli.

Science 230, 179-181 (1985).

33 Willshaw, G. A., Smith, H. R., Scotland, S. M. & Rowe, B. Cloning of genes determining

the production of vero cytotoxin by Escherichia coli. Journal of general microbiology

131, 3047-3053 (1985).

34 Sandvig, K. Shiga toxins. Toxicon : official journal of the International Society on

Toxinology 39, 1629-1635 (2001).

35 Moulding, D. A., Walter, C., Hart, C. A. & Edwards, S. W. Effects of staphylococcal

enterotoxins on human neutrophil functions and apoptosis. Infection and immunity 67,

2312-2318 (1999).

36 Bensing, B. A., Rubens, C. E. & Sullam, P. M. Genetic loci of Streptococcus mitis that

mediate binding to human platelets. Infection and immunity 69, 1373-1380,

doi:10.1128/IAI.69.3.1373-1380.2001 (2001).

37 Bensing, B. A., Siboo, I. R. & Sullam, P. M. Proteins PblA and PblB of Streptococcus

mitis, which promote binding to human platelets, are encoded within a lysogenic

bacteriophage. Infection and immunity 69, 6186-6192, doi:10.1128/IAI.69.10.6186-

6192.2001 (2001).

38 Hynes, W. L. & Ferretti, J. J. Sequence analysis and expression in Escherichia coli of the

hyaluronidase gene of Streptococcus pyogenes bacteriophage H4489A. Infection and

immunity 57, 533-539 (1989).

39 Verma, N. K., Brandt, J. M., Verma, D. J. & Lindberg, A. A. Molecular characterization

of the O-acetyl transferase gene of converting bacteriophage SF6 that adds group antigen

6 to Shigella flexneri. Molecular microbiology 5, 71-75 (1991).

48

40 Clark, C. A., Beltrame, J. & Manning, P. A. The oac gene encoding a lipopolysaccharide

O-antigen acetylase maps adjacent to the integrase-encoding gene on the genome of

Shigella flexneri bacteriophage Sf6. Gene 107, 43-52 (1991).

41 van Wamel, W. J., Rooijakkers, S. H., Ruyken, M., van Kessel, K. P. & van Strijp, J. A.

The innate immune modulators staphylococcal complement inhibitor and chemotaxis

inhibitory protein of Staphylococcus aureus are located on beta-hemolysin-converting

bacteriophages. Journal of bacteriology 188, 1310-1315, doi:10.1128/JB.188.4.1310-

1315.2006 (2006).

42 Kaneko, J., Muramoto, K. & Kamio, Y. Gene of LukF-PV-like component of Panton-

Valentine leukocidin in Staphylococcus aureus P83 is linked with lukM. Bioscience,

biotechnology, and biochemistry 61, 541-544, doi:10.1271/bbb.61.541 (1997).

43 Figueroa-Bossi, N. & Bossi, L. Inducible prophages contribute to Salmonella virulence in

mice. Molecular microbiology 33, 167-176 (1999).

44 Cumby, N., Edwards, A. M., Davidson, A. R. & Maxwell, K. L. The bacteriophage

HK97 gp15 moron element encodes a novel superinfection exclusion protein. Journal of

bacteriology 194, 5012-5019, doi:10.1128/JB.00843-12 (2012).

45 Kenzaka, T., Tani, K. & Nasu, M. High-frequency phage-mediated gene transfer in

freshwater environments determined at single-cell level. The ISME journal 4, 648-659,

doi:10.1038/ismej.2009.145 (2010).

46 Muniesa, M., Colomer-Lluch, M. & Jofre, J. Could bacteriophages transfer antibiotic

resistance genes from environmental bacteria to human-body associated bacterial

populations? Mobile genetic elements 3, e25847, doi:10.4161/mge.25847 (2013).

47 Modi, S. R., Lee, H. H., Spina, C. S. & Collins, J. J. Antibiotic treatment expands the

resistance reservoir and ecological network of the phage metagenome. Nature 499, 219-

222, doi:10.1038/nature12212 (2013).

48 Casjens, S. Prophages and bacterial genomics: what have we learned so far? Molecular

microbiology 49, 277-300 (2003).

49 Perna, N. T. et al. Genome sequence of enterohaemorrhagic Escherichia coli O157:H7.

Nature 409, 529-533, doi:10.1038/35054089 (2001).

50 Juhala, R. J. et al. Genomic sequences of bacteriophages HK97 and HK022: pervasive

genetic mosaicism in the lambdoid bacteriophages. Journal of molecular biology 299, 27-

51, doi:10.1006/jmbi.2000.3729 (2000).

51 Burrows, L. L., Charter, D. F. & Lam, J. S. Molecular characterization of the

Pseudomonas aeruginosa serotype O5 (PAO1) B-band lipopolysaccharide gene cluster.

Molecular microbiology 22, 481-495 (1996).

49

52 Uc-Mass, A. et al. An orthologue of the cor gene is involved in the exclusion of

temperate lambdoid phages. Evidence that Cor inactivates FhuA receptor functions.

Virology 329, 425-433, doi:10.1016/j.virol.2004.09.005 (2004).

53 Cumby, N., Reimer, K., Mengin-Lecreulx, D., Davidson, A. R. & Maxwell, K. L. The

Phage Tail Tape Measure Protein, an Inner Membrane Protein, and a Periplasmic

Chaperone Play Connected Roles in the Genome Injection Process of E. coli Phage

HK97. Molecular microbiology, doi:10.1111/mmi.12918 (2014).

54 Vica Pacheco, S., Garcia Gonzalez, O. & Paniagua Contreras, G. L. The lom gene of

bacteriophage lambda is involved in Escherichia coli K12 adhesion to human buccal

epithelial cells. FEMS microbiology letters 156, 129-132 (1997).

55 Mirold, S. et al. Isolation of a temperate bacteriophage encoding the type III effector

protein SopE from an epidemic Salmonella typhimurium strain. Proceedings of the

National Academy of Sciences of the United States of America 96, 9845-9850 (1999).

56 Barondess, J. J. & Beckwith, J. bor gene of phage lambda, involved in serum resistance,

encodes a widely conserved outer membrane lipoprotein. Journal of bacteriology 177,

1247-1253 (1995).

57 Shen, L. et al. PA2800 plays an important role in both antibiotic susceptibility and

virulence in Pseudomonas aeruginosa. Current microbiology 65, 601-609,

doi:10.1007/s00284-012-0196-2 (2012).

58 Sibley, C. D. et al. Discerning the complexity of community interactions using a

Drosophila model of polymicrobial infections. PLoS pathogens 4, e1000184,

doi:10.1371/journal.ppat.1000184 (2008).

59 Kirienko, N. V., Cezairliyan, B. O., Ausubel, F. M. & Powell, J. R. Pseudomonas

aeruginosa PA14 pathogenesis in Caenorhabditis elegans. Methods Mol Biol 1149, 653-

669, doi:10.1007/978-1-4939-0473-0_50 (2014).

60 Tsui, L. C. & Dorfman, R. The cystic fibrosis gene: a molecular genetic perspective. Cold

Spring Harbor perspectives in medicine 3, a009472, doi:10.1101/cshperspect.a009472

(2013).

61 Margaret Hodson, D. G., Andrew Bush. Cystic Fibrosis. 3rd edn, (CRC Press, Taylor &

Francis Group, 2007).

62 Staudinger, B. J. et al. Conditions associated with the cystic fibrosis defect promote

chronic Pseudomonas aeruginosa infection. American journal of respiratory and critical

care medicine 189, 812-824, doi:10.1164/rccm.201312-2142OC (2014).

63 Lyczak, J. B., Cannon, C. L. & Pier, G. B. Establishment of Pseudomonas aeruginosa

infection: lessons from a versatile opportunist. Microbes and infection / Institut Pasteur

2, 1051-1060 (2000).

50

64 Langton Hewer, S. C. & Smyth, A. R. Antibiotic strategies for eradicating Pseudomonas

aeruginosa in people with cystic fibrosis. The Cochrane database of systematic reviews

11, CD004197, doi:10.1002/14651858.CD004197.pub4 (2014).

65 Johansen, H. K. & Gotzsche, P. C. Vaccines for preventing infection with Pseudomonas

aeruginosa in cystic fibrosis. The Cochrane database of systematic reviews 6, CD001399,

doi:10.1002/14651858.CD001399.pub3 (2013).

66 Lee, J. & Zhang, L. The hierarchy quorum sensing network in Pseudomonas aeruginosa.

Protein & cell 6, 26-41, doi:10.1007/s13238-014-0100-x (2015).

67 Craig, L., Pique, M. E. & Tainer, J. A. Type IV pilus structure and bacterial

pathogenicity. Nature reviews. Microbiology 2, 363-378, doi:10.1038/nrmicro885 (2004).

68 Chiang, P., Habash, M. & Burrows, L. L. Disparate subcellular localization patterns of

Pseudomonas aeruginosa Type IV pilus ATPases involved in twitching motility. Journal

of bacteriology 187, 829-839, doi:10.1128/JB.187.3.829-839.2005 (2005).

69 Skerker, J. M. & Berg, H. C. Direct observation of extension and retraction of type IV

pili. Proceedings of the National Academy of Sciences of the United States of America 98,

6901-6904, doi:10.1073/pnas.121171698 (2001).

70 Doyle, T. B., Hawkins, A. C. & McCarter, L. L. The complex flagellar torque generator

of Pseudomonas aeruginosa. Journal of bacteriology 186, 6341-6350,

doi:10.1128/JB.186.19.6341-6350.2004 (2004).

71 Doig, P. et al. Role of pili in adhesion of Pseudomonas aeruginosa to human respiratory

epithelial cells. Infection and immunity 56, 1641-1646 (1988).

72 Barken, K. B. et al. Roles of type IV pili, flagellum-mediated motility and extracellular

DNA in the formation of mature multicellular structures in Pseudomonas aeruginosa

biofilms. Environmental microbiology 10, 2331-2343, doi:10.1111/j.1462-

2920.2008.01658.x (2008).

73 Comer, J. E., Marshall, M. A., Blanch, V. J., Deal, C. D. & Castric, P. Identification of

the Pseudomonas aeruginosa 1244 pilin glycosylation site. Infection and immunity 70,

2837-2845 (2002).

74 Kus, J. V., Tullis, E., Cvitkovitch, D. G. & Burrows, L. L. Significant differences in type

IV pilin allele distribution among Pseudomonas aeruginosa isolates from cystic fibrosis

(CF) versus non-CF patients. Microbiology 150, 1315-1326 (2004).

75 Wei, Q. & Ma, L. Z. Biofilm matrix and its regulation in Pseudomonas aeruginosa.

International journal of molecular sciences 14, 20983-21005,

doi:10.3390/ijms141020983 (2013).

76 Ma, L. et al. Assembly and development of the Pseudomonas aeruginosa biofilm matrix.

PLoS pathogens 5, e1000354, doi:10.1371/journal.ppat.1000354 (2009).

51

77 Sakuragi, Y. & Kolter, R. Quorum-sensing regulation of the biofilm matrix genes (pel) of

Pseudomonas aeruginosa. Journal of bacteriology 189, 5383-5386,

doi:10.1128/JB.00137-07 (2007).

78 Klausen, M. et al. Biofilm formation by Pseudomonas aeruginosa wild type, flagella and

type IV pili mutants. Molecular microbiology 48, 1511-1524 (2003).

79 Pamp, S. J., Gjermansen, M., Johansen, H. K. & Tolker-Nielsen, T. Tolerance to the

antimicrobial peptide colistin in Pseudomonas aeruginosa biofilms is linked to

metabolically active cells, and depends on the pmr and mexAB-oprM genes. Molecular

microbiology 68, 223-240, doi:10.1111/j.1365-2958.2008.06152.x (2008).

80 Mulcahy, H., Charron-Mazenod, L. & Lewenza, S. Extracellular DNA chelates cations

and induces antibiotic resistance in Pseudomonas aeruginosa biofilms. PLoS pathogens 4,

e1000213, doi:10.1371/journal.ppat.1000213 (2008).

81 Molin, S. & Tolker-Nielsen, T. Gene transfer occurs with enhanced efficiency in biofilms

and induces enhanced stabilisation of the biofilm structure. Current opinion in

biotechnology 14, 255-261 (2003).

82 Costerton, J. W., Stewart, P. S. & Greenberg, E. P. Bacterial biofilms: a common cause

of persistent infections. Science 284, 1318-1322 (1999).

83 Jayaseelan, S., Ramaswamy, D. & Dharmaraj, S. Pyocyanin: production, applications,

challenges and new insights. World journal of microbiology & biotechnology 30, 1159-

1168, doi:10.1007/s11274-013-1552-5 (2014).

84 Lau, G. W., Hassett, D. J., Ran, H. & Kong, F. The role of pyocyanin in Pseudomonas

aeruginosa infection. Trends in molecular medicine 10, 599-606,

doi:10.1016/j.molmed.2004.10.002 (2004).

85 Mavrodi, D. V. et al. Functional analysis of genes for biosynthesis of pyocyanin and

phenazine-1-carboxamide from Pseudomonas aeruginosa PAO1. Journal of bacteriology

183, 6454-6465, doi:10.1128/JB.183.21.6454-6465.2001 (2001).

86 Lau, G. W., Ran, H., Kong, F., Hassett, D. J. & Mavrodi, D. Pseudomonas aeruginosa

pyocyanin is critical for lung infection in mice. Infection and immunity 72, 4275-4278,

doi:10.1128/IAI.72.7.4275-4278.2004 (2004).

87 Sorensen, R. U. & Klinger, J. D. Biological effects of Pseudomonas aeruginosa

phenazine pigments. Antibiotics and chemotherapy 39, 113-124 (1987).

88 Denning, G. M., Railsback, M. A., Rasmussen, G. T., Cox, C. D. & Britigan, B. E.

Pseudomonas pyocyanine alters calcium signaling in human airway epithelial cells. The

American journal of physiology 274, L893-900 (1998).

89 Baron, S. S. & Rowe, J. J. Antibiotic action of pyocyanin. Antimicrob Agents Chemother

20, 814-820 (1981).

52

90 Kerr, J. R. et al. Pseudomonas aeruginosa pyocyanin and 1-hydroxyphenazine inhibit

fungal growth. Journal of clinical pathology 52, 385-387 (1999).

91 Bondy-Denomy, J. The diverse impact of bacteriophages on the bacterial host Doctor of

Science thesis, University of Toronto, (2014).

92 O'Toole, G. A. Microtiter dish biofilm formation assay. Journal of visualized experiments

: JoVE, doi:10.3791/2437 (2011).

93 Chibeu, A. et al. The adsorption of Pseudomonas aeruginosa bacteriophage phiKMV is

dependent on expression regulation of type IV pili genes. FEMS microbiology letters

296, 210-218, doi:10.1111/j.1574-6968.2009.01640.x (2009).

94 Chung, I. Y., Jang, H. J., Bae, H. W. & Cho, Y. H. A phage protein that inhibits the

bacterial ATPase required for type IV pilus assembly. Proceedings of the National

Academy of Sciences of the United States of America 111, 11503-11508,

doi:10.1073/pnas.1403537111 (2014).

95 Beatson, S. A., Whitchurch, C. B., Sargent, J. L., Levesque, R. C. & Mattick, J. S.

Differential regulation of twitching motility and elastase production by Vfr in

Pseudomonas aeruginosa. Journal of bacteriology 184, 3605-3613 (2002).

96 Luong, P. M. et al. Emergence of the P2 phenotype in Pseudomonas aeruginosa PAO1

strains involves various mutations in mexT or mexF. Journal of bacteriology 196, 504-

513, doi:10.1128/JB.01050-13 (2014).

97 Olivas, A. D. et al. Intestinal tissues induce an SNP mutation in Pseudomonas aeruginosa

that enhances its virulence: possible role in anastomotic leak. PloS one 7, e44326,

doi:10.1371/journal.pone.0044326 (2012).

98 Lamarche, M. G. & Deziel, E. MexEF-OprN efflux pump exports the Pseudomonas

quinolone signal (PQS) precursor HHQ (4-hydroxy-2-heptylquinoline). PloS one 6,

e24310, doi:10.1371/journal.pone.0024310 (2011).

99 MGH-ParaBioSys:NHLBI Program for Genomic Applications, M. G. H. a. H. M. S.,

Boston, MA. P. aeruginosa PA14 Genomic Sequencing Project,

<http://pga.mgh.harvard.edu> (

100 He, J. et al. The broad host range pathogen Pseudomonas aeruginosa strain PA14 carries

two pathogenicity islands harboring plant and animal virulence genes. Proceedings of the

National Academy of Sciences of the United States of America 101, 2530-2535 (2004).

101 Choi, J. Y. et al. Identification of virulence genes in a pathogenic strain of Pseudomonas

aeruginosa by representational difference analysis. Journal of bacteriology 184, 952-961

(2002).

53

Appendix

Figure 12. Detail genome alignment for phages used in this study.

Genomes of closely related JBD phages were aligned and known gene functions were labeled.

Genes without function labeled represent hypothetical genes (ORFs) or genes with unknown

function. The numbers in black bordered boxes represent gene number, and blank boxes indicate

that the gene homolog does not exist in a specific phage. Phage JBD26 was used as the reference

genome, and the percentages shown are the differences in DNA sequence of phage genes

compared to homologs in phage JBD26. However, for genes with no homologs in JBD26, I

compared them to their homologs in JBD5. The genes labeled in yellow are the PrAGs examined

in this study.