Investigating Amrz-Mediated Activation of Pseudomonas Aeruginosa Twitching

Investigating AmrZ-mediated activation of Pseudomonas aeruginosa twitching

motility and alginate production

DISSERTATION

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University

Binjie Xu

Graduate Program in Microbiology

The Ohio State University

2015

Dissertation Committee:

Daniel J. Wozniak, Ph.D., Advisor

Irina Artsimovitch, Ph.D.

Robert S. Munson, Jr., Ph.D.

John S. Gunn, Ph.D.

Binjie Xu

2015

Abstract

The Gram negative bacterium Pseudomonas aeruginosa is ubiquitous in the natural

environment and responsible for various human infections. Successful P. aeruginosa

infections require many virulence factors such as flagella, type IV pilus and alginate, whose expression is under tight control by transcription factors such as AmrZ. AmrZ activates

type IV pilus-mediated twitching motility and alginate production while repressing the

flagella machinery. However, molecular mechanisms of AmrZ-mediated activation of

twitching motility and the alginate biosynthesis operon promoter (algD promoter) remain

elusive and are the emphases of this study.

Specifically, DNA microarray analyses were performed in order to identify AmrZ-

dependent gene(s) that is required for twitching motility. The AmrZ regulon contains 112

genes, many of which are involved in virulence. However, this regulon does not contain a

gene that is known to be necessary for twitching motility. In addition, we used multiple

biochemical methods to show that AmrZ tetramerizes via its C-terminal domain, the loss

of which results in diminished DNA binding and reduced efficiency during AmrZ- ii

mediated activation and repression. Demonstration of AmrZ tetramerization also assists in understanding the activation mechanism of the algD promoter. This complex promoter

contains a number of transcription factor binding sites with many in the far upstream

region. It is unclear how those transcription factors from far upstream contribute to

transcription activation. This study identified four AmrZ binding sites within this promoter

and showed that each is required for algD transcription activation. We also illustrated

phase-dependent activation of this promoter, suggesting interactions between upstream and

downstream factors. Furthermore, in vitro fluorescence resonance energy transfer

experiments indicate the formation of trans DNA-AmrZ complexes, suggesting that two

DNA molecules are held together by AmrZ, likely via AmrZ oligomerization. These results

thus provide new support for the DNA looping model of the algD promoter.

Taken together, the work described in this document has significantly enhanced our

knowledge of AmrZ-mediated regulation, full understanding of which may provide

insights into discovering new means of interfering this regulation and successful control of

P. aeruginosa infections.

iii

Dedication

This document is dedicated to my family and friends for their love, care and support

Acknowledgments

First and foremost, I would like to express my greatest gratitude to my Ph.D.

advisor Dr. Daniel J. Wozniak, who has been a phenomenal scientific mentor inside the

lab as well as a precious friend off work. Besides Dr. Wozniak, all current and previous

“Woz Lab” members have fostered my growth and maturity as a scientist and as a person.

They have offered substantial assistance to promote my professional development. I would

like to specially thank Mohini Bhattacharya, Sheri Dellos-Nolan, Dr. Dominique H. Limoli

and Preston J. Hill for helping with numerous presentation practices and paper editing. I

was also lucky to work with the undergraduate student Amyla Tan and the rotation student

Qian Shen. Both of them are smart, self-motivated and hard-working, and should take the credit of multiple experiments.

I would also like to appreciate the guidance of my dissertation committee throughout graduate school. Bob, Irina and John have offered numerous insightful advices, all of which are instrumental during the completion of my dissertation projects. I would like to thank Dr. Jesse Kwiek for his helpful career advices. All members in the Center for v

Microbial Interface Biology have created a unique work environment that is packed with expertise and fun.

Many experiments were conducted in collaboration with scientists from other disciplines. We thank Dr. Richard Fishel and Randal Soukup for gel filtration and FRET experiments, and appreciate the collaboration of Dr. Vicki H. Wysocki and Yue Ju for mass spectrometry experiments. Laboratories of Dr. Fishel and Dr. Wysocki are both located at

OSU. Whole genome sequencing was conducted and analyzed by Benjamin Kelly from the

Peter White laboratory at Center for Microbial Pathogenesis in Nationwide Children’s

Hospital, Columbus, OH.

We want to thank Genomics Shared Resource of OSU Comprehensive Cancer

Center for performing DNA microarray experiments. We would also like to thank Dr.

Thomas Hollis (Wake Forest School of Medicine, Winston-Salem, NC) for sharing recombinant AmrZ and its variants, and express our gratitude to Dr. Laura Jennings in the

Matthew Parsek laboratory (University of Washington, Seattle, WA) for measuring c-di-

GMP concentrations for us. We also thank Dr. Matthew Wolfgang (University of North

Carolina, Chapel Hill, NC), Drs. Reuben Ramphal/Jeevan Jyot (University of Florida,

Gainesville), and Dr. Joe J. Harrison (University of Calgary, Calgary, Alberta, Canada) for sharing strains and plasmids. We acknowledge Dr. Gayan Senavirathne for valuable scientific discussions, and Dr. Nrusingh Mohapatra for assistance with the β-galactosidase assay.

Vita

July 2005 ...... Leping No.1 High School, Jiangxi, China

2009...... B.S. Life Sciences, University of Science

and Technology of China, Hefei, China

2009 to present ...... Graduate Assistant, Department of

Microbiology, The Ohio State University,

Columbus, OH, USA

Publications

1. Jones, C. J., Newsom, D., Kelly, B., Irie, Y., Jennings, L. K., Xu, B., … Wozniak, D.

J. (2014). ChIP-Seq and RNA-Seq reveal an AmrZ-mediated mechanism for cyclic di-

GMP synthesis and biofilm development by Pseudomonas aeruginosa. PLoS

Pathogens, 10(3), e1003984. http://doi.org/10.1371/journal.ppat.1003984

vii

2. Pryor, E. E., Waligora, E. A., Xu, B., Dellos-Nolan, S., Wozniak, D. J., & Hollis, T.

(2012). The transcription factor AmrZ utilizes multiple DNA binding modes to

recognize activator and repressor sequences of Pseudomonas aeruginosa virulence

genes. PLoS Pathogens, 8(4), e1002648. http://doi.org/10.1371/journal.ppat.1002648

3. Xu, B., Ju, Y., Soukup, R. J., Ramsey, D. M., Fishel, R., Wysocki, V. H., & Wozniak,

D. J. (2015). The Pseudomonas aeruginosa AmrZ C-terminal domain mediates

tetramerization and is required for its activator and repressor functions. Environmental

Microbiology Reports. http://doi.org/10.1111/1758-2229.12354

4. Xu, B., & Wozniak, D. J. (2015). Development of a novel method for analyzing

Pseudomonas aeruginosa twitching motility and its application to define the AmrZ

regulon. PLoS One, 10(8). http://doi.org/10.1371/journal.pone.0136426

Fields of Study

Major Field: Microbiology

viii

Table of Contents

Abstract ...... ii

Dedication ...... iv

Acknowledgments...... v

Vita ...... vii

Table of Contents ...... ix

List of Tables ...... xiii

List of Figures ...... xiv

Chapter 1. Introduction ...... 1

Pseudomonads and Pseudomonas ...... 1

Pseudomonas aeruginosa and its clinical significance ...... 3

Pseudomonas aeruginosa pathogenesis in acute and chronic infections ...... 5

Virulence factors of Pseudomonas aeruginosa ...... 9

Alginate production and its regulation ...... 9

Flagella and FleQ...... 11

Type IV pilus and twitching motility ...... 13

Biofilm and its significance ...... 16

Control by transcription factors ...... 19

The versatile transcription factor AmrZ ...... 23

Key methods used in this study ...... 25

DNA microarray analysis ...... 26

Next-generation sequencing ...... 27

Fluorescence resonance energy transfer ...... 29

Overall goals in this dissertation ...... 30

Chapter 2. Development of a novel method for analyzing Pseudomonas aeruginosa twitching motility and its application to define the AmrZ regulon ...... 39

Abstract ...... 39

Introduction ...... 40

Materials & methods ...... 43

Results ...... 47

Discussion ...... 53

Acknowledgement ...... 59

Chapter 3. The Pseudomonas aeruginosa AmrZ C-terminal domain mediates tetramerization and is required for its activator and repressor functions ...... 74

Abstract ...... 74

Introduction ...... 75

Materials and Methods ...... 77

Results ...... 83

Discussion ...... 88

Acknowledgement ...... 92

Chapter 4. Pseudomonas aeruginosa AmrZ binds to four sites in the algD promoter and induces trans DNA-AmrZ complex formation ...... 111

Abstract ...... 111

Introduction ...... 112

Materials & Methods ...... 115

Results ...... 118

Discussion ...... 123

Acknowledgement ...... 128

Chapter 5. Overall discussion and synopsis ...... 141

References ...... 148

Appendix A. Spontaneous suppressor screening ...... 175

xii

List of Tables

Table 1.1. Rates of antibacterial resistance among P. aeruginosa isolates from hospitals

and ICUs ...... 38

Table 2.1. Strains, plasmids, and oligonucleotides used in Chapter 2 ...... 67

Table 2.2. Complete gene list from microarray analyses ...... 70

Table 3.1. Strains, plasmids, and oligonucleotides used in Chapter 3 ...... 108

Table 3.2. The AmrZ CTD is required for efficient activation of twitching motility ..... 110

Table 4.1. Strains, plasmids, and oligonucleotides used in Chapter 4 ...... 138

Table A1. Point mutations identified in three suppressor mutants ...... 190

Table A2. Strains, plasmids, and oligonucleotides used in Appendix A ...... 191

xiii

List of Figures

Figure 1.1. Virulence factors in acute and chronic infections...... 32

Figure 1.2. Assembly of the type IV pilus apparatus...... 33

Figure 1.3. The alginate biosynthesis operon and its promoter region...... 34

Figure 1.4. Simplified life cycle of a biofilm...... 35

Figure 1.5. Examples of proteins in the Arc superfamily...... 36

Figure 1.6. AmrZ primary structure and crystal structure when bound to DNA...... 37

Figure 2.1. Cellophane-air interface stimulates P. aeruginosa twitching motility...... 60

Figure 2.2. TM-proficient and TM-deficient strains retain their TM phenotypes on

cellophane...... 61

Figure 2.3. Identification of the AmrZ regulon from the TM-active “edge” sub-population.

...... 62 xiv

Figure 2.4. Verification of representative AmrZ-dependent genes...... 63

Figure 2.5. AmrZ specifically binds to the lecB promoter...... 64

Figure 2.6. The lecB gene is required for TM in P. aeruginosa strain PAK, but not in strain

PAO1...... 65

Figure 2.7. lecB is required for TM in PAK, but not in PAO1 using the cellophane-based method...... 66

Figure 3.1. The primary form of AmrZ in solution is tetrameric...... 94

Figure 3.2. The C-terminal domain of AmrZ mediates tetramerization...... 95

Figure 3.3. AmrZ binds to DNA as a tetramer...... 96

Figure 3.4. The C-terminal domain of AmrZ is required for tetramerization...... 97

Figure 3.5. AmrZR22A forms tetramers in solution...... 98

Figure 3.6. Similar amounts of AmrZR22A and WT AmrZ protein levels were produced after arabinose induction...... 99

Figure 3.7. Dominant negative effect of AmrZR22A...... 100

Figure 3.8. Truncation of the C-terminal domain results in reduced binding to PalgD. 101

Figure 3.9. AmrZ1-66 displays reduced binding to the gcbA promoter...... 102

Figure 3.10. Efficient activation of PalgD requires the AmrZ C-terminal domain...... 103

Figure 3.11. Western blot analyses of AmrZ1-66 after arabinose induction...... 104

Figure 3.12. The AmrZ C-terminal domain is required for efficient repression and activation of its targets...... 105

Figure 3.13. ClustalW2 alignment of AmrZ with other Arc proteins...... 106

Figure 3.14. Predicted AmrZ secondary structure and polarity of the AmrZ CTD...... 107

Figure 4.1. Identification of a second AmrZ binding site in PalgD...... 129

Figure 4.2. Scrambling of ZBS2 and ZBS3 results in reduced DNA binding and impaired

PalgD activation...... 130

Figure 4.3. Deletion of ZBS2 or replacement by an algB sequence does not eliminate AmrZ binding...... 131

Figure 4.4. Identification of two additional ZBSs in PalgD...... 132

Figure 4.5. Proposed positions and nucleotide sequences of four ZBSs...... 133

Figure 4.6. Scrambling of all ZBSs results in abolished PalgD activation...... 134

Figure 4.7. Correct DNA phasing between ZBSs is critical for PalgD activation...... 135

Figure 4.8. AmrZ binding results in formation of trans DNA-AmrZ complexes...... 136

xvi

Figure 4.9. The 105bp truncation between ZBS2 and ZBS3 disrupts PalgD activation. 137

Figure A1. Flowchart of the suppressor screening process...... 185

Figure A2. Three suppressors exhibit restored TM and no intragenic mutation within amrZ.

...... 186

Figure A3. PilAS73A fails to restore TM in the ΔpilA&ΔamrZ mutant...... 187

Figure A4. The suppressor M1 displays a reduced c-di-GMP level while gcbA overexpression in this strain fails to inhibit TM...... 188

Figure A5. TM of the ΔamrZ mutant after expressing the plasmid-borne phosphodiesterase

PA2133...... 189

xvii

Chapter 1

Chapter 1. Introduction

Pseudomonads and Pseudomonas

Pseudomonads are amongst the earliest groups of bacteria discovered due to their versatile lifestyles and diverse niches. In 1894, the genus Pseudomonas was first defined by Walter Migula with the broad definition: “Cells with polar organs of motility. Formation of spores occurs in some species, but it is rare” (Migula, 1894; Sokatch, Gunsalus, &

Ornston, 1986). Thereafter, significantly more bacterial species were discovered and classified as pseudomonads. In 1966, Stanier et al. (1966) conducted a comprehensive study characterizing phenotypic traits of pseudomonads, which by this point were defined as “unicellular rods, with the long axis straight or curved, but not helical. Motile by means of one or more polar flagella. Gram-negative. Do not form spores, stalks, or sheaths. The energy-yielding metabolism is respiratory, never fermentative or photosynthetic. All use molecular oxygen as a terminal oxidant; and some can use denitrification as an alternative, anaerobic respiratory mechanism. All are chemo-organotrophs; some are facultative chemo-lithotrophs that use H, as an energy source.” (Stanier et al., 1966). This more 1

detailed definition excluded many species from the pseudomonad group. Later more

bacterial species were removed from this genus due to the advancement of molecular

phylogenetics. For example, the pseudomonad Burkholderia cepacia (formerly known as

Pseudomonas cepacia) was transferred into the new genus Burkholderia in 1992

(Yabuuchi et al., 1992). Currently Pseudomonas is the largest Gram-negative genus with

144 species (Gomila, Peña, Mulet, Lalucat, & García-Valdés, 2015).

Pseudomonas species and many other pseudomonads are prevalent in natural

environments such as soil and water as well as in/on animal and plant hosts. Strikingly,

pseudomonads are capable of growing with very limited or unconventional nutrients,

including hydrocarbons (such as n-dodecane and n-hexadecane) rarely utilized by other

bacteria (Stanier et al., 1966). As a result, they have been widely used in bioremediation

applications.

Amongst all pseudomonads, the most studied species are P. aeruginosa, P.

fluorescens and P. putida. All three species belong to the fluorescent pseudomonad

subgroup that produces water-soluble, yellow-green fluorescent pigments. There are

multiple shared traits between P. fluorescens and P. putida. Both are easily found in soil, water and plants. Their colonization on plants promotes plant growth and suppresses infections by fungal pathogens (O’Sullivan & O’Gara, 1992). They both harbor

multitrichous polar flagella and grow optimally at 25-30 °C. These two species can be distinguished by the ability to liquefy gelatin, with only P. fluorescens able to do so

(Stanier et al., 1966). It is noteworthy that many P. putida sub-species have wide industrial applications such as bioremediation.

Pseudomonas aeruginosa and its clinical significance

P. aeruginosa is the type species of the genus Pseudomonas, but also is distinct from all other Pseudomonas species. Exclusively, P. aeruginosa has one polar flagellum and produces the blue-green pigment named pyocyanin. In addition, it can grow at higher temperatures such as 41°C and is the only Pseudomonas species that poses a significant threat to mammals (Stanier et al., 1966).

P. aeruginosa is an opportunistic pathogen that rarely causes disease in healthy human beings. This bacterium is occasionally implicated in outbreaks associated with hot

tubs, whirlpools and swimming pools. In one clinical case, a previously healthy woman

died of acute pneumonia caused by P. aeruginosa, with the hot tub being the most likely source of infection (Huhulescu et al., 2011). Colonization by this bacterium in compromised patients however, can result in a variety of severe acute and chronic infections. P. aeruginosa is a significant pathogen that causes community-acquired pneumonia, and can infect virtually any traumatic wound (Hauser & Rello, 2003). It also poses a significant threat to individuals with indwelling medical devices, transplants, diabetes, and patients in intensive care units (Hauser & Rello, 2003). This pathogen is able to cause infection in multiple tissue environments including but not limited to the 3

respiratory tract (especially lower respiratory tract), heart, eye, ear, urinary tract and skin

(Gellatly & Hancock, 2013; Hauser & Rello, 2003). In addition, in the lungs of cystic

fibrosis (CF) patients, P. aeruginosa is a primary pathogen associated with pulmonary

exacerbations and high mortality rates (Ahlgren et al., 2015; Folkesson et al., 2012).

Two major contributing factors to the clinical significance of this bacterium are its

prevalence and high antimicrobial resistance. P. aeruginosa is a common resident in all

types of non-desiccated niches. This bacterium can be found in feces of 1.2-2.3% of healthy

individuals and oral antibiotic intake often results in increased incidence of P. aeruginosa-

positive stools (Noble and White, 1969; Campa et al., 1993). More importantly, P.

aeruginosa is easily detectable in various locations inside and outside hospitals. Outside

hospitals, the most common source of P. aeruginosa is water reservoirs polluted by human

or animal sources. P. aeruginosa isolates are frequently found in swimming pools and whirlpools. In private homes, this bacterium is present in places such as sink drains, toilets, showers, air humidifiers and many other devices requiring water to function. Inside hospitals, medical devices that work with water are frequently contaminated with P. aeruginosa (Campa et al., 1993). These can render immunocompromised patients highly susceptible to P. aeruginosa infections. Another trait that contributes to P. aeruginosa pathogenesis is its high resistance and recalcitrance to antimicrobial reagents. Disinfectants such as Cetrimide as well as numerous antibiotics have little effect on P. aeruginosa growth

(Noble & White, 1969). During the last decade P. aeruginosa clinical isolates displayed

increased resistance to antibiotics such as β-lactams, fluoroquinolones and

aminoglycosides (Table 1.1), making P. aeruginosa infections difficult to treat. It should be noted that although certain antibiotics remain active against free-living (planktonic) P. aeruginosa, this bacterium usually displays significantly higher recalcitrance to these agents when encased within biofilms (see details in the biofilm sub-section in Chapter 1).

Pseudomonas aeruginosa pathogenesis in acute and chronic infections

P. aeruginosa causes many infections with varying clinical manifestations and outcomes. Examples of acute infections include acute pneumonia, urinary tract infections, bacteremia and keratitis, while chronic diseases include wound infections, lung infections of CF patients, and infections on/in indwelling medical devices, etc. This section discusses how numerous P. aeruginosa virulence factors contribute to different forms of infections.

During acute infections, P. aeruginosa expresses various virulence factors that combat against the host immune defenses and aid in survival within the host. These factors include but are not limited to alginate, flagella, type IV pili, quorum sensing, phenazines, the type III secretion system (T3SS), siderophores, proteases and exotoxins. Since there are detailed descriptions in the next section, alginate, flagella and type IV pili will be not discussed here.

The quorum sensing (QS) signaling cascade is activated when concentrations of QS signaling molecules (homoserine lactones and quinolones in Gram-negatives; small

peptides in Gram-positives) reach the quorum. QS regulates a number of targets in P.

aeruginosa and has significant impacts on its virulence (Bjarnsholt et al., 2005; Rutherford

& Bassler, 2012). P. aeruginosa possesses three QS systems: LasR/I, RhlR/I and PQS

(Pseudomonas quinolone signal). Examples of QS targets include proteases, pyocyanin and toxins (Rutherford & Bassler, 2012), and QS signaling also controls biofilm formation and antibiotic sensitivity (Bjarnsholt et al., 2005; Shrout et al., 2006; Singh et al., 2000).

Phenazines such as pyocyanin are another class of virulence factors with broad

antimicrobial activities and cytotoxicity to host cells including neutrophils and

lymphocytes (Lau, Hassett, Ran, & Kong, 2004; Sorensen & Klinger, 1987). T3SS punches

through the host cell membrane via a needle-like machinery and delivers cytotoxic effectors (Hauser, 2009a). Two types of siderophores (pyoverdine and pyochelin) are required for scavenging iron from host sequestration (Cornelis & Dingemans, 2013). A fine control of these virulence factors is critical for successful infections in the host. For instance, P. aeruginosa is able to switch between different iron uptake systems depending on the type of infections (Cornelis & Dingemans, 2013; Marvig et al., 2014). Similarly, an increase in temperature from 28°C to 37°C leads to elevated expression of genes involved in T3SS and phenazines (Wurtzel et al., 2012). Distinct sets of virulence factors play important roles during acute and chronic infections (Figure 1.1). For example, T3SS and exotoxins are important for host cell invasion and toxicity (Hauser, 2009b), while exopolysaccharide production results in enhanced immune evasion (Leid et al., 2005).

During chronic pulmonary infections in CF patients, P. aeruginosa is the dominant

pathogen, colonizing 60-80% of CF patients (Cystic Fibrosis Foundation, 2013). CF is the

most common lethal genetic disorder in Caucasians. About 1 in 25 Caucasians are

heterozygous carriers, and 1 in 2,500 newborns carry this disease (Döring et al., 2000). CF

patients have defects in the chloride ion channel named CFTR (CF transmembrane

conductance regulator) and diseases manifest in multiple organs including skin, pancreas

and lungs. CFTR mutations result in defects of ions and water transport across cell

membranes in multiple locations. Specifically in the lung, mucus accumulation and

impaired mucociliary clearance render CF airways highly susceptible to recurrent and

chronic microbial infections, which are de facto the most significant cause of mortality in

CF patients (Hoiby, 1982). CF lungs are infected by multiple bacterial species with the

most common ones being P. aeruginosa, Staphylococcus aureus and Haemophilus

influenzae (Coburn et al., 2015; Govan & Deretic, 1996). Identification of P. aeruginosa

in CF patients is often a poor prognostic indicator (Govan & Deretic, 1996).

Since initial colonization, P. aeruginosa in CF patients suffers hostile attacks throughout infections including immune responses, oxidative stress, and antibiotic

treatment (Snitkin & Segre, 2014). Long-term adaptation leads to distinct traits in

descendants compared to initial colonizers, including hyper-mutability, mucoid conversion, emergence of small colony variants, loss of motility and alteration of quorum sensing (Singh et al., 2000; Cullen and McClean, 2015). Although initial infections in CF

patients can be attributed to a single P. aeruginosa lineage, there is significant

heterogeneity and diversification within one patient at later stages of infection (Markussen et al., 2014). Higher mutation rates enable bacteria to diversify and be prepared, as a community, against selective pressures from the host immunity and antimicrobial treatments. Typically, P. aeruginosa isolates found in chronic infections are less invasive but are significantly more immune evasive and persistent compared to those isolated from acute infections (Byrd et al., 2011; Wagner & Iglewski, 2008).

The transition between acute and chronic infections is a multi-factorial practice. P. aeruginosa isolates from acute infections are considered similar to environmental isolates and exhibit a more invasive and motile lifestyle, while those involved in chronic infections resemble bacteria in biofilms (Singh et al., 2000; Coggan and Wolfgang, 2012). Bis-

(3’,5’)-cyclic diguanylate (c-di-GMP) for example is a major determining factor between these lifestyles in most bacteria including pathogens such as Salmonella species, Vibrio cholerae and P. aeruginosa (Hengge, 2009). A high level of c-di-GMP represses motility and expression of acute virulence factors while promoting biofilm formation and the sessile lifestyle. The Gac/Rsm pathway serves as another mechanism to control the switch between acute and chronic infections. This pathway relies on two sRNA molecules, RsmY and RsmZ, for modulating target mRNA stability and translation efficiency. In this regulatory pathway, acute virulence factors (T3SS) and chronic virulence factors (type VI secretion and exopolysaccharides) are inversely regulated (Coggan & Wolfgang, 2012).

QS signaling is altered during chronic infections with many CF P. aeruginosa isolates displaying loss-of-function lasR mutations (Singh et al., 2000; Sousa & Pereira, 2014).

Meanwhile, other factors, such as the periplasmic protease CtpA and the regulator of β- lactam resistance named AmpR, are implicated during this transition (Balasubramanian,

Kumari, & Mathee, 2014; Seo & Darwin, 2013). Of note, these signaling pathways work synergistically to provide a fitness advantage for P. aeruginosa during infections.

Virulence factors of Pseudomonas aeruginosa

The ability of this bacterium to withstand the host immune defense and antimicrobial treatments while successfully colonizing and establishing an infection is attributed to its large pool of virulence factors. Due to the length limit of this document, only those relevant in later chapters will be discussed here.

Alginate production and its regulation

During initial colonization in CF lungs, P. aeruginosa exhibits a non-mucoid colony phenotype. However, mucoid variants are increasingly isolated as the disease progresses (Hoiby, 1982). Strikingly, the percentage of mucoid isolates may account up to

90% of total P. aeruginosa isolates from CF patients (Govan & Deretic, 1996). Emergence of the mucoid phenotype is typically associated with prolonged antibody response and deterioration of lung functions (Hoiby, 1982).

The mucoid phenotype is the outcome of overproduction of the exopolysaccharide alginate. Polymerized from the uronic acid β-D-mannuronate and C-5 epimer α-L-

guluronate, alginate is polyanionic, highly hydrated and viscous (Gacesa, 1998). Alginate

is known to possess anti-phagocytic properties and the polyanionic nature of alginate

renders itself a barrier to antimicrobial agents (Hoiby, 1982; Govan and Deretic, 1996;

Leid et al., 2005). Although often found in CF patients, anti-alginate antibodies are unable to clear mucoid P. aeruginosa infections. As a result of these protective functions conferred

by alginate, mucoid P. aeruginosa is even more difficult, if not impossible, to eradicate,

with infections lasting throughout the lifetime of a colonized CF patient. Alginate

production is however, not exclusively found in CF patients, and mucoid P. aeruginosa

variants have been isolated in other infected sites such as gallbladders (Govan & Deretic,

1996). Considering that the human host is not its natural niche, alginate likely plays other

roles when P. aeruginosa grows in fresh water or soil. It was previously shown in P.

aeruginosa and other Pseudomonas species that alginate offers protection and increases survival in the soil and against desiccation (Chang et al., 2007; Hall, McLoughlin, Leung,

Trevors, & Lee, 1998). It would be instrumental to determine other roles of alginate during

P. aeruginosa growth in its natural niches. Interestingly, alginate is also produced by other

bacteria such as the nitrogen fixing bacteria Azotobacter. In Azotobacter, alginate is

produced during vegetative phases of these bacteria and the alginate casing preserves

dormant cysts from mechanical stress and desiccation (Clementi, 1997).

Alginate production is an energy-exhaustive process, requiring a number of enzymes and precursor substrates. Bacteria therefore implement a tight control on its

production. All enzyme-encoding genes except algC are within the alginate biosynthesis

operon, which is thereby a focal point of regulation (Figure 1.2) (Gacesa, 1998; Ramsey &

Wozniak, 2005). Expression of this operon is regulated by a number of transcription factors

(TFs), including AlgR, Vfr, AlgB, AlgP (Hp-1), IHF, CysB and AmrZ (Figure 1.2)

(Wozniak, 1994; Wozniak and Ohman, 1994; Baynham et al., 1999; Whitchurchet al.,

2005; Leech et al., 2008; Schurr, 2013). Previous studies have shown that the sigma factor

AlgT (also referred to as AlgU/σ22) directs the transcription of the alginate biosynthesis

operon (Boucher, Schurr, & Deretic, 2000; M J Schurr et al., 1993; D J Wozniak & Ohman,

1994). Freed from the sequestration by the anti-sigma factor MucA, AlgT initiates transcription of genes such as amrZ, algB and algR (P. J. Baynham & Wozniak, 1996; A.

K. Jones et al., 2010). These three and four additional TFs bind to the promoter region of

the alginate biosynthesis operon (PalgD) and initiate transcription (Deretic & Konyecsni,

1990; Kato, Misra, & Chakrabarty, 1990; Konyecsni & Deretic, 1990; Mohr et al., 1991).

Interestingly, in the region over 100 base pairs (bp) upstream of the transcription start site,

there are at least seven TF binding sites, exact functions of which are yet characterized.

This promoter arrangement is rare in bacteria, and one of the aims in this study is to unveil

the molecular mechanism of this activation (Chapter 4).

Flagella and FleQ

Mucoid P. aeruginosa strains are typically aflagellated, with the absence of

functional flagella as an immune evasion mechanism (Cobb et al., 2004; Tart.et al., 2006;

Amiel.et al., 2010). However, in non-mucoid P. aeruginosa as well as other bacterial

pathogens this cell surface appendage plays essential roles during infections. The single

polar flagellum of P. aeruginosa is required for dissemination, chemotaxis, initial surface

attachment and robust biofilm formation (Klausen, Heydorn, et al., 2003; O’Toole &

Kolter, 1998; Q. Wang, Suzuki, Mariconda, Porwollik, & Harshey, 2005). In an acute

mouse burn wound model, flagella mutants displayed significantly reduced invasiveness

in comparison to their flagellated counterparts, thereby underscoring its essentiality during

acute infections (D. Drake & Montie, 1988).

The flagellum apparatus contains three main sub-structures: the motor-containing

basal body that anchor the whole structure to the cell wall, the flagellin filament as the

propeller and the hook connecting above two sub-structures (Jarrell & McBride, 2008).

The flagellum apparatus is assembled starting with the basal body, the hook and lastly the

filament. In P. aeruginosa, flagella assembly is regulated in four tiers, with FleQ being the

master regulator in the first tier. FleQ directly or indirectly regulates most flagella

promoters, and the absence of functional FleQ results in reduced expression of most

flagella genes and loss of flagella-mediated swimming motility (Dasgupta et al., 2003). On the other hand, fleQ expression is subject to external regulation. It is well documented that both Vfr, a P. aeruginosa homolog of the Escherichia coli cAMP receptor CRP, and AmrZ

(Alginate and Motility Regulator Z), repress fleQ transcription (Dasgupta et al., 2002; Tart et al., 2006). Additionally, FleQ receives signals from c-di-GMP and modulates expression of the pel operon in a c-di-GMP dependent manner (Baraquet, Murakami, Parsek, &

Harwood, 2012; J. Hickman & Harwood, 2008). These findings suggest an active crosstalk between flagella and other systems in the cell.

Type IV pilus and twitching motility

In addition to flagella-mediated swimming motility, P. aeruginosa also conducts twitching motility (TM) via its type IV pilus (TFP). This polar apparatus mobilizes the cell by means of repeating cycles of extension, tethering and retraction. Besides TM, TFP mediates surface adherence and biofilm formation (O’Toole and Kolter, 1998; Klausen,

Heydorn, et al., 2003). It also plays important roles in P. aeruginosa virulence (Comolli et al., 1999; Zolfaghar, Evans, & Fleiszig, 2003) and is involved in swarming motility, DNA binding and chemotaxis (Anyan et al., 2014; Kearns, Robinson, Lawrence, Kearns, &

Robinson, 2001; Schaik et al., 2005). Recent studies demonstrated that TFP plays important roles in surface sensing (Luo et al., 2015; Persat, Inclan, Engel, Stone, & Gitai,

2015; Siryaporn, Kuchma, O’Toole, & Gitai, 2014). TFP is present in many other Gram- negative bacteria besides P. aeruginosa such as Neisseria gonnorrhoeae and Myxococcus xanthus (Friedrich, Bulyha, & Søgaard-Andersen, 2013; Giltner, Nguyen, & Burrows,

2012; Hoyne, Haas, Meyer, Davies, & Elleman, 1992). Recent studies have also identified this organelle in a few Gram-positive species, indicating its prevalence in bacteria with diverse evolutionary origins (Melville & Craig, 2013).

In P. aeruginosa nearly forty genes are required for TFP functions, suggesting its complex nature (Alm & Mattick, 1997). The current model for TFP assembly is shown in

Figure 1.3. The main pilus filament is comprised of the major pilin PilA and five minor pilins (FimU, PilV, PilW, PilX and PilE). Exact roles of these minor pilins are unclear, but they are required for pili assembly and twitching motility (Kuchma et al., 2012). PilY1, a homolog to the gonococcal fimbrial tip adhesin PilC, is considered the tip adhesion in P. aeruginosa and shows affinity to integrin on host cells (Aim, Hallinan, Watson, & Mattick,

1996; Johnson et al., 2011; Orans et al., 2010). Among these minor pilins, the PilVWXY1 complex is proposed as the primer for pilus assembly (Nguyen et al., 2015). Prepilins, including the major pilin PilA and these minor pilins, are processed by the peptidase PilD, assembled by the ATPase PilB, and transported across the cytoplasmic membrane through

PilC. Thereafter, assembled pilin subunits are extended beyond the outer membrane via

PilQ. PilQ and the inner membrane complex are aligned by the PilMNOP alignment complex (Burrows, 2012). Distinct from flagella, assembly of the TFP machinery begins with the outer membrane complex PilQ, followed by the alignment complex and finally the inner membrane complex (Friedrich et al., 2013). Known tethering targets of TFP include inert surfaces as well as host proteins such as asialo GM1 receptors on epithelial

cells and integrin proteins (de Bentzmann et al., 1996; Jenkins, Buckling, McGhee, &

Ffrench-Constant, 2005; Johnson et al., 2011; Sheth et al., 1994). After adhering to specific

targets, the filament is retracted back into cells by the ATPases PilT and PilU (Whitchurch

& Mattick, 1994).

Transcription of the major pilin gene pilA requires the sigma factor RpoN and the

two-component system PilR/S (Hobbs, Collie, Free, Livingston, & Mattick, 1993). TFP- mediated TM also requires the Chp chemotaxis system, which is composed of a histidine kinase ChpA and two CheY-like response regulators PilG and PilH (Bertrand, West, &

Engel, 2010).

The TFP machinery is also subject to control by a number of external regulators, such as the AlgR/FimS two-component system, c-di-GMP, and AmrZ (Alm, Bodero, Free,

& Mattick, 1996; P. J. Baynham, Ramsey, Gvozdyev, Cordonnier, & Wozniak, 2006;

Belete, Lu, & Wozniak, 2008). In the AlgR/FimS two-component system, AlgR phosphorylation is necessary for AlgR-mediated activation of TM (Whitchurch, Erova, et al., 2002). Since this phosphorylation is dispensable for AlgR-mediated activation of alginate production, it is intriguing what TM-pertinent signals FimS senses (S. Ma et al.,

1998). A recent study identified the interaction between FimS and the methyl-accepting protein PilJ, suggesting FimS might receive signals from PilJ (Luo et al., 2015). Although it remains undetermined how c-di-GMP impacts TFP, two TFP proteins FimX and PilZ bind to c-di-GMP while c-di-GMP production is activated by PilY1 signaling, suggesting communications between these two systems (Amikam & Galperin, 2006; Huang,

Whitchurch, & Mattick, 2003; Jain, Behrens, Kaever, & Kazmierczak, 2012; Kazmierczak,

Lebron, & Murray, 2006; Luo et al., 2015). The mechanism of AmrZ-mediated activation of TM is unknown and is one focus of this study (see Chapter 2 and Appendix A).

Biofilm and its significance

Since the last century, there have been tremendous advances in our knowledge of

microorganisms, but most research has focused on free-living (planktonic) cells. In fact,

microorganisms rarely exist as single-cell suspensions; instead, they are mainly found

within complex communities called biofilms (Donlan & Costerton, 2002; Hall-Stoodley,

Costerton, & Stoodley, 2004). Biofilms were defined by IUPAC in 2002 as “aggregates of

microorganisms in which cells that are frequently embedded within a self-produced matrix

of extracellular polymeric substance (EPS) adhere to each other and/or to a surface” (Vert

et al., 2012). This lifestyle however, is not simply passive aggregation of cells, but an active

and complex process requiring cooperation across the community. These communities can

be formed by a single or multiple species, the latter of which is likely to be the norm in

natural environment and during infections (Hall-Stoodley, Costerton, & Stoodley, 2004).

In bacterial biofilms, EPS is composed of a variety of biopolymers including proteins, polysaccharide and extracellular DNA (eDNA) (Flemming et al., 2007). In the

model organism P. aeruginosa, three exopolysaccharides (alginate, Psl and Pel) are

involved in biofilm formation (Chew et al., 2014; Mann & Wozniak, 2012). For instance,

disruption of Psl biosynthesis abolishes biofilm architecture (L. Ma, Jackson, Landry,

Parsek, & Wozniak, 2006). Studies have also revealed important roles of proteins such as

IHF and CdrA in biofilms (Borlee et al., 2010; Brockson et al., 2014; Devaraj, Justice,

Bakaletz, & Goodman, 2015). Antibodies targeting the DNA binding protein IHF disrupt

biofilms and offer potential as a future therapeutic option (Brockson et al., 2014). In

addition, eDNA is abundant in biofilms with DNase I treatments preventing biofilm

formation and disrupting pre-formed biofilms (Gloag et al., 2013; Whitchurch, Tolker-

Nielsen, Ragas, & Mattick, 2002).

In general biofilm formation is divided into three stages: initial attachment to

surfaces, growth and maturation, and dispersal (Figure 1.4) (Sauer et al., 2002; Stoodley et al., 2002). Initial attachment may occur on a biotic or abiotic surface and requires flagella, type IV pili and Psl (L. Ma et al., 2006; O’Toole & Kolter, 1998). Attached bacteria undergo subsequent proliferation and develop three-dimensional biofilm architecture.

Matured P. aeruginosa biofilms can be flat or “mushroom”-shaped depending on nutrition sources (Shrout et al., 2006). The biofilm structure is also modulated by QS, alginate, Psl, flagella, TFP, and the catabolite repression control (Crc) protein (D G Davies et al., 1998;

Klausen, Aaes-Jørgensen, Molin, & Tolker-Nielsen, 2003; Klausen, Heydorn, et al., 2003;

O’Toole, Gibbs, Hager, Phibbs, & Kolter, 2000; Stoodley et al., 2002). After biofilm maturation, small quantities of cells may detach from biofilms and become free-flowing again. These cells may find a new niche and re-initiate the biofilm cycle. Successful detachment is dependent on factors such as flagella, TFP and rhamnolipid (Boles,

Thoendel, & Singh, 2005; Hall-Stoodley, Costerton, & Stoodley, 2004). Biofilm dispersal is also affected by carbon sources (K Sauer et al., 2004), nitric oxide (Barraud et al., 2006), metal chelators (Banin, Brady, & Greenberg, 2006), the fatty acid cis-2-decenoic acid (D.

G. Davies & Marques, 2009), and the Pf4 prophage (Rice et al., 2009). Importantly, Chua et al. showed that dispersed cells exhibited higher virulence against macrophages and

Caenorhabditis elegans compared to planktonic cells, indicating that cells dispersed from

biofilms are distinct from planktonic ones (Chua et al., 2014).

Compared to the planktonic counterpart, bacteria in biofilms display distinct

phenotypes and significantly higher survival rates in adverse environments. In 2001,

Whiteley et al. compared transcriptional profiles between biofilm-grown and planktonic

bacteria and 73 genes exhibited differential expression (Whiteley et al., 2001). This suggests the lifestyle of bacteria in biofilms is distinct from planktonic cells. Bacteria in

biofilms are more tolerant to various detrimental forces such as antibiotics, phagocytosis

and reactive oxygen species. As an example, planktonic P. aeruginosa is susceptible to

imipenem, but the same strain becomes 1,000-fold more tolerant when grown in biofilms

(Donlan & Costerton, 2002). Increased resistance is a result of EPS protection and natural

persistence in this growth mode (Hall-Stoodley et al., 2004; Ryder et al., 2007). A thick

layer of EPS may act as a physical barrier preventing/delaying the penetration of

antimicrobial agents. In addition, bacteria in mature biofilms grow significantly more

slowly than planktonic cells and are thereby naturally more recalcitrant to antimicrobials,

especially β-lactams (Donlan & Costerton, 2002; Hall-Stoodley et al., 2004). Another

contributing factor is an increased proportion of persister cells in biofilms. Albeit being a

tiny fraction of cells in exponential phases during planktonic growth, persister cells can

account up to 1% in biofilms and stationary phases. These persister cells are not

metabolically active and thus insensitive to antibiotic treatment. However, they can revert

to metabolically active cells and repopulate the community after antibiotic treatment

(Wood, Knabel, & Kwan, 2013).

Control by transcription factors

The Central Dogma of molecular biology dictates that transcription is an essential

step to decode the information carried by DNA and products of this process (mRNAs)

direct protein synthesis (Crick, 1970). During transcription, the RNA polymerase (RNAP)

recognizes specific transcription initiation DNA sequences (i.e., promoters), unwinds

double stranded DNA, and uses it as the template to synthesize RNA. In bacteria, a RNAP

holoenzyme is composed of a sigma factor and the core enzyme. Most prokaryotic

organisms encode only one type of the core enzyme with five subunits (2 α, β, β’ and ω).

The core enzyme is catalytically competent, but requires the sigma factor for promoter

recognition and transcription initiation. The active site of the polymerase that mediates

binding of the template DNA and the RNA product, is formed by β and β’ subunits. Two

α subunits dimerize via their N-terminal domains, which are also responsible for the assembly of the β and β’ subunits. C-terminal domains of α subunits (αCTD) bind to upstream DNA and are important sites for TFs to interact with and then execute regulation

(Browning & Busby, 2004). Specific roles of the ω subunit are not known, but the ω subunit binds ppGpp (Ross et al., 2013) and increases transcription efficiency (Gunnelius et al.,

2014). Some Gram-positive bacteria express additional accessary RNAP components

including δ and ε. These subunits are not essential for RNAP functions but contribute to

proper functions of RNAPs (Weiss & Shaw, 2015). There are multiple types of sigma

factors in the cell, with σ70 being the predominant one and the focus in this section. During

transcription initiation, the sigma factor and the RNAP core form the holoenzyme and

recognize specific promoter sequences. The holoenzyme unwinds the promoter DNA and forms the open complex. Transcription is initiated by the synthesis of a nascent RNA chain from the DNA template strand. When transcription proceeds into elongation, the sigma factor is released and the RNAP core completes synthesis of remaining RNA (Borukhov

& Nudler, 2008). Transcription can be terminated by protein-dependent and protein-

independent mechanisms (Cooper GM., 2000).

In all forms of life, transcription regulation serves as a major and efficient form of

gene regulation. Both prokaryotic and eukaryotic organisms have complex mechanisms to

fine-tune transcription. Most transcription control events occur during initiation in which

TFs bind to target DNAs and activate/repress transcription. TFs are DNA-binding proteins that specifically interact with DNA and modulate gene expression. All organisms possess a significant number of TFs regulating almost all transcription events. For instance, in the model organism Escherichia coli, ~6.5% (300/4605) of open reading frames encode TFs

(Martínez-Antonio, 2011). Organismal complexity positively correlates with the ratio and number of TFs per genome (Levine & Tjian, 2003).

TFs have evolved to use various mechanisms to mediate regulation and different

classes of these proteins include helix-turn-helix (HTH) proteins, zinc-finger and steroid- binding proteins, leucine zipper proteins, homeodomains and β-sheet DNA binding proteins (Pabo & Sauer, 1992). HTH proteins are the dominant family of TFs, and their

HTH motifs are composed of two α-helices separated by a turn. Although the second helix directly interacts with the major groove of target DNA, other contacts by the first helix and other portions of TFs are important for specific recognition (Pabo & Sauer, 1992). Most

TFs, including all types of proteins except β-sheet DNA binding proteins, contact with major grooves of DNA via their α-helices, indicating the predominance of α-helices in protein-DNA interactions. The α-helix was thought as the only motif to interact with DNA until the discovery of the ribbon-helix-helix (RHH) protein superfamily, in which each protein contains a β-sheet followed by two α-helices. In this protein superfamily, β-sheets interact with DNA major grooves (Schreiter & Drennan, 2007). The RHH protein superfamily contains multiple proteins such as Arc and Mnt from the phage P22, MetJ from

E. coli, NikR from Helicobacter pylori and AmrZ from P. aeruginosa (Figure 1.5)

(Vershons et al., 1985; Rafferty et al., 1989; Chivers and Sauer, 1999; Waligora et al.,

2010).

Transcription can be positively or negatively regulated by TFs. Transcriptional repression is often mediated by physically blocking the access of the RNA polymerase to promoters. This hindrance can be achieved by either binding of repressors to the sigma factor binding sequence (for σ70, -35 bp to +1 bp relative to the transcription start site, same

denotations below) or altering DNA conformation, to prevent promoters from being

recognized by the RNAP. Most repressors fall into one of these two categories, and in

microorganisms such as E. coli, the region downstream of -30 is an “exclusive zone of repression” where few activators bind (Neidhardt & Curtiss, 1996). Some other repressors can, however, interact with transcriptional activators and prevent them from activating transcription (Browning & Busby, 2004). Transcriptional activators function via more diverse mechanisms. Transcription activation is typically accomplished by interactions between its activators and αCTDs or sigma factors (Neidhardt & Curtiss, 1996). Therefore, binding sites of transcription activators are typically found between -31 to -80 bp. Remote

elements however, exist in some promoters, but they usually pair with proximal factors.

TFs that bind to remote elements may make contact with proximal proteins by bending the intervening DNA or inducing DNA looping (Neidhardt & Curtiss, 1996). In rare examples certain TFs regulate transcription solely by interacting with RNAP without making contact with DNA (Beck et al., 2007).

The majority of these TFs share a similar modular arrangement containing DNA- binding domains and modules with other functions, the latter of which may mediate protein-protein interactions, small molecule binding, or signal transduction (Balleza et al.,

2009). Many TFs bind to DNA as homo- or hetero- oligomers via their oligomerization domains. On the other hand, DNA binding affinity of some TFs is modulated by binding of small molecules. The lac repressor loses DNA binding affinity when bound by allolactose, relieving repression of the lac operon (Lewis et al., 1996). In contrast, nickel

binding to H. pylori NikR stimulates DNA binding (Schreiter et al., 2003). On the other hand, some TFs such as DNA-binding response regulators use their signal reception domains as an on/off switch for DNA binding in the presence/absence of certain signals.

These response regulators, when phosphorylated by their cognate kinases, become activated and execute transcriptional control (Okkotsu, Tieku, Fitzsimmons, Churchill, &

Schurr, 2013; Whitchurch, Erova, et al., 2002). Modular arrangement in TFs has become a useful tool in biotechnology and various artificial TFs have been developed for application purposes (Ansari & Mapp, 2002; Sera, 2009).

The versatile transcription factor AmrZ

The global transcription factor AmrZ plays significant roles during P. aeruginosa pathogenesis. AmrZ contains 108 residues with the molecular weight of 12.34 kDa. It belongs to the Arc protein superfamily, in which proteins bind to DNA via their RHH domains and the basic functional unit is dimeric. This protein superfamily is composed of members from various origins, including several bacterial species and the bacteriophage

P22 (Schreiter & Drennan, 2007). Representative proteins in this superfamily are shown in

Figure 1.5. Albeit their RHH domains exhibit similar structures, these proteins have limited sequence similarity even within RHH domains (Figure 1.5). Their C-terminal domains

(CTDs) carry diverse functions (Figure 1.5). Proteins such as Arc and CopG are short (~50 residues) and contain solely RHH domains. These proteins bind to DNA as tetramers but

are dimeric in solution (Vershons et al., 1985; Brigitte E. Raumann Carl O. Pabo & Robert

T. Sauer, 1994). Mnt tetramerization via its CTD enhances DNA binding (Waldburger &

Sauer, 1995). Both Arc and Mnt are encoded in the genome of the bacteriophage P22 and are important to repress anti-repressor genes during lytic and lysogenic forms of the phage, respectively (Waldburger & Sauer, 1995). The repressor MetJ inhibits transcription of genes involved in methionine and S-adenosylmethionine (SAM) synthesis, and MetJ DNA binding activity is significantly enhanced when bound by SAM (Rafferty et al., 1989).

Similarly, nickel-bound NikR exhibits increased DNA binding and repressor activities, which are important during the control of nickel transport (P T Chivers & Sauer, 1999;

Muller et al., 2011; Schreiter et al., 2003).

AmrZ is composed of three segments: the N-terminal motif (residues 1-11), the ribbon-helix-helix domain (residues 12-66), and the CTD (residues 67-108) (Figure 1.6A).

The N-terminal motif is highly flexible and does not seem to be essential during AmrZ- mediated regulation (Waligora et al., 2010). The ribbon-helix-helix domain mediates DNA binding and AmrZ dimerization (Pryor et al., 2012). Specifically, a short β-sheet (residues

15-23) within this domain is critical for DNA binding, and single amino acid mutations at certain residues within this motif lead to abolished DNA binding (Waligora et al., 2010).

However, the role of the AmrZ CTD remains unknown and is one of the goals in this study

(Chapter 3). Based on the recently solved crystal structure, the CTD-truncated variant

AmrZ1-66 binds to its target DNA as a dimer of dimers (Figure 1.6B). In addition, residues

such as Lys18, Val20, Arg22 and Arg28 within the β sheet of each monomer make direct

contacts with the major groove of target DNA (Pryor et al., 2012).

While most Arc superfamily proteins act as repressors, AmrZ functions as both an

activator and repressor. AmrZ activates transcription of the alginate biosynthesis operon

and TM, yet represses flagella-mediated motility, Psl production and c-di-GMP synthesis

(Baynham and Wozniak, 1996; Ramsey et al., 2005; Baynham et al., 2006; Tart et al.,

2006; Jones et al., 2013, 2014). Specifically, increased levels of AmrZ in mucoid strains

activate alginate production and repress flagella-mediated motility and Psl production (C.

J. Jones et al., 2013; Tart et al., 2006). However, the intracellular AmrZ concentration is much lower in non-mucoid strains, allowing flagella assembly and Psl-mediated biofilm formation. In non-mucoid strains, AmrZ is also required for TM and repression of gcbA

(also referred to as adcA, PA4843), which encodes a diguanylate cyclase (C. J. Jones et al.,

2014; Petrova, Cherny, & Sauer, 2014). Deletion of amrZ in non-mucoid strains therefore

results in an increased c-di-GMP level and enhanced biofilm formation (C. J. Jones et al.,

2014).

Key methods used in this study

A number of laboratory techniques were used during this work, but only three

methods will be described here while the rest will be covered in following chapters.

DNA microarray analysis

Traditional approaches to study transcriptional regulation include genetic methods

such as reporter assays, and biochemical methods including northern blot, and quantitative

real-time PCR (qRT-PCR). However, all these methods are performed on the scale of

single genes. Developed in 1995, DNA microarrays revolutionized almost every biological

science field by enabling the examination of thousands of genes in parallel (Schena et al.,

1995). There are two types of microarrays: oligo-based DNA arrays and cDNA arrays. In cDNA arrays, mRNAs are reverse-transcribed into cDNAs, which are subsequently converted to double strands and amplified. These dsDNA molecules are then used as probes in microarray analyses (Trevino, Falciani, & Barrera-Saldana, 2007). In this document only oligo-based DNA arrays are used and discussed. Millions of oligonucleotides, i.e., probes, are covalently attached to a solid surface (glass or a silicon chip) in designated wells. These probes are ~25 nucleotides long and designed from protein open reading frames (ORFs) in a certain genome. Expression of each gene is monitored by multiple probes. During sample preparation, mRNAs are purified and reverse-transcribed to single stranded cDNA. This cDNA library is then labeled with fluorescent dyes and hybridized to probes in the microarray chip. Perfect matches with specific probes result in the formation of strong hydrogen bonds and are therefore resistant to subsequent washes.

Relative amounts of mRNAs are quantified by fluorescence intensity measurements of respective wells.

After twenty years of development, DNA microarrays have become a mature technology that is relatively easy to perform and analyze. Data analyses have become straightforward and efficient with sharing of open-source analysis tools across the research community. As a great example, the Bioconductor project has provided the platform for sharing extensible software packages in computational biology and bioinformatics

(Gentleman et al., 2004). Bioconductor has allowed researchers to collaborate and share

software packages rather than to repetitively design similar scripts. Currently over a

thousand open-source software packages are publically available for various data analyses.

However, there are a few limitations of DNA microarrays. First of all, data

interpretation can vary significantly using different analytical methods, and proper controls

as well as statistical analyses are required for drawing appropriate conclusions. Results

from DNA microarrays therefore often require validation by other methods. Moreover,

since DNA microarrays are based on sample hybridization to probes, issues with

hybridization efficiency and specificity have been great concerns. DNA microarrays rely

on existing knowledge of target genomes, so they are only capable of measuring expression

of known genes (Wang et al., 2009).

Next-generation sequencing

Next-generation sequencing (also referred to as second-generation sequencing)

(NGS) first appeared in the market in 2005 and is another revolution that has dramatically

altered the course of biological research (Margulies et al., 2005). NGS is referred to as non-

Sanger-based high-throughput DNA sequencing. The basic principle of Sanger-based

DNA sequencing is tricking DNA polymerase with dideoxynucleotides (ddNTPs), so new syntheses stop whenever ddNTPs are incorporated. A collection of terminated DNA fragments are generated after DNA amplification. These fragments were originally analyzed by slab gels, and later by capillaries (Mardis, 2013). Although it is a significant improvement to this technique, the Sanger-based sequencing method has the weakness of limited parallelism and high per-sample cost. The need for high throughput and low cost- yield ratios led to the development of NGS. Although there are multiple NGS platforms, the fundamental principle is similar. Sample DNA is randomly fragmented and ligated to adaptor sequences in arrays, generating a library of short labeled DNA fragments.

Sequencing of this library is performed by alternating cycles of amplification and data acquisition (Shendure & Ji, 2008). When compared to Sanger-based DNA sequencing,

NGS achieves high throughput via in vitro construction of the sequencing library and the use of arrays. Strikingly, the transition from Sanger-based DNA sequencing to NGS has reduced the cost per genome by over 1,000 fold (Wetterstrand, 2015).

The development of NGS has not only revolutionized the sequencing of whole genomes, but also fostered subsequent applications in ChIP-Seq (CHromatin

ImmunoPrecipitation-sequencing), RNA-Seq and metagenomic sequencing (Shendure &

Ji, 2008). ChIP-Seq is widely used to determine binding sites of a DNA binding protein throughout the genome. RNA-Seq, also known as transcriptome sequencing, is used to

quantify gene expression and to identify unknown transcripts, etc. RNA-Seq is considered a superior transcriptomics method than DNA microarrays owing to its minimized artifacts and applications in species with unknown genome sequences (Wang et al., 2009).

Metagenomic sequencing allows the study of multiple species in a microbial community,

and therefore has been applied extensively during microbiome research (Eloe-fadrosh et

al., 2015; Van Dijk, Auger, Jaszczyszyn, & Thermes, 2014).

Fluorescence resonance energy transfer

Fluorescence is a type of luminescence conferred by a certain molecule

(fluorophore) that when excited by light at a specific wavelength, absorbs this light and

then re-emits light (photons) at a longer wavelength (Ishikawa-Ankerhold et al., 2012).

Fluorescence/Föster resonance energy transfer (FRET) involves two fluorophores and occurs when the emission wavelength of one fluorophore (donor) matches the excitation wavelength of the other (acceptor). Emitted photons by the donor fluorophore are absorbed by and excite the acceptor fluorophore when two fluorophores are very close, leading to the loss of emission by the donor fluorophore and emission from the acceptor, i.e., energy transfer. Detectible FRET requires extreme proximity between two fluorophores (2-10 nm)

(Blair, Goodrich, & Kugel, 2013).

FRET has been widely used to determine proximity between molecules and is able to detect intra/inter-molecule interactions well below the resolution of normal optical

microscopy (Zadran et al., 2012). Its applications include analyzing protein-protein and protein-DNA interactions as well as monitoring protein conformational changes.

Importantly, these interactions can be assessed both in vitro and in vivo, offering more versatility for diverse applications.

Albeit a powerful tool, FRET has its limitations. First of all, there are a limited number of donor/acceptor pairs for FRET experiments, which can restrain its application from the studies of certain sensitive cellular mechanisms (Ishikawa-Ankerhold et al., 2012;

Zadran et al., 2012). In addition, other artifacts such as auto-fluorescence and protein- induced fluorescence enhancement may complicate FRET data interpretation (Hwang &

Myong, 2014; Zadran et al., 2012). FRET experiments require careful experimental design and proper controls, and concerns with fluorescence fading, fluorophore and analyte concentrations should be considered (Zadran et al., 2012).

Overall goals in this dissertation

The overall goal of this work is to further understand mechanisms of AmrZ- mediated regulation in Pseudomonas aeruginosa. Specifically, we have performed experiments with the aim of identifying the target(s) of AmrZ during its activation of twitching motility (Chapter 2 and Appendix A). In addition, we are also interested in determining the oligomeric state of AmrZ and identifying the domain(s) that mediates this oligomerization, which will be discussed in Chapter 3. AmrZ and many other transcription 30

factors are known to activate the transcription of the alginate biosynthesis operon; however, the mechanism remains elusive how remote factors activate transcription. Our new discovery in Chapter 4 has shed new insights into this activation mechanism.

Figure 1.1. Virulence factors in acute and chronic infections.

P. aeruginosa causes both acute and chronic infections with distinct sets of virulence factors. Expression of these factors is regulated by at least three signaling systems: c-di-

GMP, the Gac/Rsm system and quorum sensing. T3SS/T6SS: type III/VI secretion system.

Figure 1.2. Assembly of the type IV pilus apparatus.

This apparatus is composed of four sub-complexes shown in four shades. The assembled pilus (in white) contains the major pilin PilA, a few minor pilins (mp) and the tip adhesin

PilY1. During extension, prepilins are processed by the peptidase PilD and then assembled into the pilus by the ATPase PilB across the cytoplasm via PilC. During retraction, the

ATPases PilT and PilU mediate the disassembly. These five proteins (PilBCD, PilT and

PilU) are the inner membrane motor sub-complex (in light grey). The assembled pilus traverses the outer membrane via the outer membrane sub-complex PilQ and PilF (in black). In addition, the outer membrane sub-complex is aligned with the inner membrane motor sub-complex by the alignment sub-complex composed of PilMNOP and FimV (in dark grey). This image is adapted from (Burrows, 2012).

Figure 1.3. The alginate biosynthesis operon and its promoter region.

The string of colored boxes represents 12 genes within the alginate biosynthesis operon,

and boxes in the lower panel are binding sites of different transcription factors and the

sigma factor AlgT. Colors in the upper panel represent their subcellular localizations.

Yellow: periplasmic; orange: cytoplasmic membrane; green: outer membrane; red:

cytoplasmic. The picture of the operon is adapted from the pseudomonas.com website

(Winsor et al., 2011). Numbers in the lower panel are the distance in base pairs relative to

the transcription start site, which is marked as the black arrow. Two Hp-1 (AlgP) binding sites are not shown here due to their imprecisely mapped positions (Michael J Schurr,

2013).

Figure 1.4. Simplified life cycle of a biofilm.

Microorganisms attach to surfaces (Stage 1), and continue to grow and mature with distinct biofilm three-dimensional structures (Stage 2). A proportion of cells within the biofilm may detach and undergo the dispersal process (Stage 3), returning to the free-flowing phase and potentially reinitiating the biofilm life cycle. Adapted from (Stoodley et al., 2002).

Figure 1.5. Examples of proteins in the Arc superfamily.

Apart from the ribbon-helix-helix (RHH) domain, these proteins share limited sequence or function similarities. CTD: C-terminal domain. Adapted from (Schreiter & Drennan,

2007).

Figure 1.6. AmrZ primary structure and crystal structure when bound to DNA.

(A.) Full-length form of AmrZ and the truncated variant lacking its C-terminal domain. N:

N-terminus; B: DNA-binding domain; Z: oligomerization domain. (B.) The crystal structure of AmrZ1-66 when binding to its target DNA amrZ1. Each monomer within the dimer of dimers of AmrZ (shown in four colors) interacts with the major groove of DNA via its β-sheet. Panel (B.) is adapted from (Pryor et al., 2012).

Table 1.1. Rates of antibacterial resistance among P. aeruginosa isolates from hospitals and ICUs

(Adapted from (Lister et al., 2009))

Antibiotic % of P. aeruginosa strains exhibiting resistance

Hospital study, Hospital Hospital ICU study, ICU study, Hospital ICU study, Hospital 2006 study, 2005 study, 2002 2002 2000-2002 study, 2001 2001 (n ≥ study, 2000 (n = 606) (n = 589) (n = 9,896) (n = 951) (n ≥ 7,500) (n ≥ 2,157) 543) (n = 882) β-Lactams Cefepime 6 5 9 25 12 8 10 9 Ceftazidime 13 10 13 19 17 9 9 13 38 Piperacillin- 11 9 11 10 14 8 8 13 tazobactam Aztreonam 12 32 Imipenem 11 7 16 23 22 12 16 16 Meropenem 6 7 18 11 16 10 Fluoroquinolones Ciprofloxacin 21 22 35 32 33 26 25 25 Levofloxacin 22 22 34 32 27 25 27 Aminoglycosides Amikacin 5 10 4 3 Tobramycin 8 10 12 16 Gentamicin 12 12 16 22 15 15 14

38[G

Chapter 2

Chapter 2. Development of a novel method for analyzing Pseudomonas aeruginosa

twitching motility and its application to define the AmrZ regulon

Abstract

Twitching motility is an important migration mechanism for the Gram-negative

bacterium Pseudomonas aeruginosa. In the commonly used subsurface twitching assay, the sub-population of P. aeruginosa with active twitching motility is difficult to harvest for high-throughput studies. Here we describe the development of a novel method that allows efficient isolation of bacterial sub-populations conducting highly active twitching motility.

The transcription factor AmrZ regulates multiple P. aeruginosa virulence factors including twitching motility, yet the mechanism of this activation remains unclear. We therefore set out to understand this mechanism by defining the AmrZ regulon using DNA microarrays in combination with the newly developed twitching motility method. We discovered 112 genes in the AmrZ regulon and many encode virulence factors. One gene of interest and the subsequent focus was lecB, which encodes a fucose-binding lectin. DNA binding assays revealed that AmrZ activates lecB transcription by directly binding to its promoter. 39

The lecB gene was previously shown to be required for twitching motility in P. aeruginosa strain PAK; however, our lecB deletion had no effect on twitching motility in strain PAO1.

Collectively, in this study a novel condition was developed for quantitative studies of twitching motility, under which the AmrZ regulon was defined.

Introduction

Microorganisms are ubiquitous in all types of habitats, ranging from hot springs in deep seas to underneath thick ice sheets in the Antarctic continent. Occasionally, environmental bacteria such as Pseudomonas aeruginosa find a human host and become pathogenic. Efficient colonization and dissemination of P. aeruginosa is dependent on its multiple motility mechanisms –swimming, twitching, and swarming (Kohler et al., 2000;

Rashid and Kornberg, 2000; Jarrell and McBride, 2008; Murray and Kazmierczak, 2008).

Under laboratory settings, swimming motility occurs in liquid culture or on semi-solid surfaces (0.3% agar) yet this is suppressed on solid surfaces (Jarrell & McBride, 2008;

Murray & Kazmierczak, 2006). In contrast, swarming motility is observed on softer agar surfaces (0.5-0.7% agar) compared to twitching motility (TM) (1% agar) (Kohler et al.,

2000; Rashid & Kornberg, 2000). Therefore, growth conditions significantly impact activities of different motility organelles. TM is conducted by type IV pili (TFP), which also contribute to adherence onto epithelial cells, biofilm formation, DNA uptake, and virulence (Burrows, 2012; Doig et al., 1988; Huang et al., 2003; Klausen, Aaes-Jørgensen,

et al., 2003; Klausen, Heydorn, et al., 2003; R. M. Miller et al., 2008; Zolfaghar et al.,

2003).

TFP are present in a variety of bacterial genera including Proteobacteria,

Cyanobacteria and Firmicutes (Pelicic, 2008). The TFP machinery mediates cell motion

by repeating cycles of pili extension, tethering to surfaces and retraction. Almost 40 P.

aeruginosa genes have been shown to be involved in TM (J. Mattick, 2002), and assembly

of the TFP apparatus is efficiently coordinated by these gene products. Briefly, in P.

aeruginosa, pre-major pilin PilA and other minor pilins are processed by the peptidase

PilD, assembled into the pilus fiber by the ATPase PilB, and translocated across the

cytoplasmic membrane via the transmembrane protein PilC. With the guidance by the

alignment complex composed of PilMNOP and potentially FimV, polymerized pilus

filaments pass through the outer membrane complex (PilQ and PilF) (Burrows, 2012).

During retraction, pilus filaments are depolymerized by the ATPase PilT with probable

assistance from PilU (11,12).

TM has been studied by multiple approaches such as macroscopic observations of

colony edges, macro-/micro-scopic subsurface twitching assays and video-microscopy in

flow cells (Henrichsen, 1983; Semmler et al., 1999; Gibianskyet al., 2010; Turnbull and

Whitchurch, 2014). Macroscopic observations of colony edges provided the platform for

two independent transposon mutant screening studies (Hobbs et al., 1993; Jacobs et al.,

2003). Currently the most popular method is the macroscopic subsurface twitching assay,

in which bacteria are stabbed underneath the agar and grow at the agar/Petri dish interface

(J. Mattick, 2002; Semmler et al., 1999; Turnbull & Whitchurch, 2014). At interstitial

surfaces, TM-proficient P. aeruginosa produces both TM-active and TM-inactive sub-

populations (Semmler et al., 1999). Despite its convenience for imaging studies, this

method is limited by the difficulty to recover sufficient quantities of TM-active bacteria

from underneath the agar. This might explain why limited high-throughput (a.k.a. omics)

studies have been published focusing specifically on TM-active sub-populations. We therefore designed a novel approach to achieve this goal. In this method, cellophane sheets placed above the agar surface allow diffusion of nutrients from the underlying media, supporting bacterial growth; more importantly, cellophane sheets provide moist and smooth surfaces required for TM activation. We showed that TM-proficient P. aeruginosa strains exhibited active TM under this condition and were easily distinguishable from TM- deficient ones.

We next sought to take advantage of this approach to understand AmrZ-mediated

regulation in P. aeruginosa. AmrZ (alginate and motility regulator Z) is a global regulator

of multiple virulence factors, including the metabolism of bis-(3’,5’)-cyclic diguanylate (c- di-GMP), extracellular polysaccharide production, and flagella (Baynham and Wozniak,

1996; Tart et al., 2006; Joneset al., 2013, 2014). Although AmrZ was also shown to be necessary for TM, its mechanism requires further investigation (P. J. Baynham et al.,

2006). Therefore, we harvested TM-active bacterial sub-populations with the cellophane- based approach, and used DNA microarrays to compare transcriptional profiles between

WT P. aeruginosa and two isogenic amrZ mutants. The AmrZ regulon under this condition

contained 112 genes, and the lectin-encoding gene lecB was the only one known necessary

for TM (Sonawane, Jyot, & Ramphal, 2006) in this gene list. The fucose-specific LecB

(formerly known as PA-IIL) is abundantly present intracellularly and on the outer

membrane of P. aeruginosa, and it plays important roles in host cell adherence and

reduction of the ciliary beat frequency of the airway epithelium (Chemani et al., 2009;

Loris, Tielker, Jaeger, & Wyns, 2003; Tielker et al., 2005). We provide evidence that AmrZ

activates lecB expression by specifically binding to its promoter. However, lecB

overexpression was not sufficient to restore TM in the ΔamrZ mutant, indicating lecB is

either not required for TM in strain PAO1 or AmrZ-dependent genes in addition to lecB

are necessary for TM. In support of the former, we were unable to recapitulate the TM- deficient phenotype seen previously with the PAK ΔlecB mutant (Sonawane et al., 2006)

in our PAO1 strain background.

In conclusion, in this study we developed a novel approach to harvest actively

twitching P. aeruginosa cells, and applied this new method to define the AmrZ regulon.

Materials & methods

Bacterial strains, plasmids, oligonucleotides and growth conditions

Information of bacterial strains, plasmids, and oligonucleotides employed in this

study is summarized in Table 2.1. Bacteria (Escherichia coli and Pseudomonas 43

aeruginosa) were grown as previously (Waligora et al., 2010). Briefly, E. coli strains were

grown in LB (each liter LB contains 10 g tryptone, 5 g yeast extract, 5 g sodium chloride)

or on LA (LB with 1.5% (w/v) agar). 100 µg/ml ampicillin was used to maintain plasmids

in E. coli. All P. aeruginosa strains were grown in LBNS or on LANS (LB or LA without

sodium chloride, respectively). In P. aeruginosa, 300 µg/ml carbenicillin was used to

maintain plasmids. All strains were grown at 37°C unless stated otherwise. All

oligonucleotides were ordered desalted from Sigma Aldrich.

In-frame deletion mutants were generated through overlap extension PCR using the

gene replacement vector pEX18Ap (Hoang et al., 1998; Heckman and Pease, 2007). Multi- copy pHERD20T-based plasmids were transferred into P. aeruginosa strains through bi- parental mating with the E. coli helper strain S17-1 (Qiu, Damron, Mima, Schweizer, &

Yu, 2008; Simon, Priefer, & Pühler, 1983).

Subsurface twitching assays

Subsurface twitching assays were modified from a previously described study (J.

Mattick, 2002). Briefly, a single colony of a strain of interest was resuspended in 400 µl

LBNS, and P10 tips were immersed in the above suspension and used to stab through the agar of 1-day-old 1% LANS plates. Plates were incubated upright in a humid chamber at room temperature for 5 days, and imaged through white epiluminescence via a ChemicDoc imaging system (Bio-Rad).

Twitching motility assays on cellophane sheets.

Square sterile cellophane sheets (Bioexpress, catalog #: E-3077-14) were placed on top of 1-day-old 1% LANS plates. A single colony of each strain was resuspended in 400

µl LBNS, and 2 µl spotted in the middle of the sheet. After air-drying, plates were incubated upright in a humid chamber at room temperature for 48 h. Microscopic images were taken by the Olympus DP71 camera mounted onto the inverted light microscope

Olympus CKX41.

Quantitative real-time PCR (qRT-PCR)

To isolate total RNA, bacteria were harvested from different growth conditions.

Cells grown on LANS plates were harvested by flooding with LBNS, while those on cellophane sheets were scraped off using P200 tips and then resuspended in LBNS. 1 ml bacteria (OD600=1) were lysed by the addition of 100 µl TE containing 1 mg/ml RNase- free lysozyme (Sigma Aldrich), and then underwent RNA isolation (RNeasy RNA isolation kit, Qiagen). To obtain cDNA, ~1 µg RNA was used for reverse transcription (Superscript

III cDNA preparation kit, Invitrogen) following manufacturer protocols. Amounts of cDNA were determined by SYBR Green Mix (Bio-Rad) and CFX1000 thermal cyclers

(Bio-Rad). The housekeeping gene rpoD was used as reference, and the ΔΔCt method was used to calculate relative gene expression (Livak and Schmittgen, 2001; Jones et al., 2014).

DNA microarray analysis

Bacteria from the “edge” sub-population on cellophane sheets were scraped off by

P200 tips, and resuspended in fresh LBNS. Total RNA was isolated as for qRT-PCR.

Thereafter, 10 µg RNA for each strain was used to generate cDNA libraries and then

analyzed using GeneChip P. aeruginosa Genome Array (Affymetrix) following

manufacture instructions. Microarray “.cel” raw files were analyzed with RMA

normalization, BH adjust, cutoff of p.value=0.05, lfc=1, using “affy” and “limma”

packages in Bioconductor, in the environment of R 3.0.2 (Ihaka and Gentleman, 1996;

Gautier et al., 2004; Gentlemanet al., 2004; Smyth, 2005). Microarray raw and result files

have been deposited in NCBI Gene Expression Omnibus (Accession number: GSE68066).

DNA binding studies

Electrophoretic mobility shift assays (EMSA) were used to evaluate AmrZ-DNA

binding as previously described (Jones et al., 2014). Each EMSA reaction contained 5 nM

[FAM]-labeled DNA. After electrophoresis, EMSA gels were imaged via a Typhoon

scanner (GE Lifescience) with the following settings: fluorescence, PMT800, 520BP 40

CY2 Blue FAM, 100 µm pixel size.

Results

Development of a novel TM-promoting condition for easy cell harvest

Traditionally, TM activity is measured using subsurface twitching assays, in which bacteria grow at the agar-Petri dish interface (J. Mattick, 2002; Turnbull & Whitchurch,

2014). One caveat of this method is the difficulty of harvesting sufficient actively twitching cells for downstream high-throughput analyses, such as transcriptomics and proteomics.

Therefore, a novel growth condition using cellophane sheets was developed taking into consideration previous studies describing the significance of inert smooth surfaces (most important) and a humid environment (Semmler et al., 1999; Mattick, 2002). It is also more

convenient to harvest cells from the surface of cellophane sheets as opposed to the

interstitial space between agar and Petri dishes. The design of this approach is shown in

Figure 2.1A. Following 48-hour growth on cellophane, P. aeruginosa formed a large

smear-like colony and we observed two distinct sub-populations of bacteria (“edge” and

“center”; Figure 2.1A).

In order to evaluate if the cellophane-air interface promotes TM gene expression,

we harvested WT P. aeruginosa (strain PAO1) from both sub-populations, and measured

transcript levels of the type IV major pilin gene pilA. We used qRT-PCR to compare pilA

mRNA levels among four different conditions: overnight broth culture (stationary phase),

agar surfaces (heterogeneous populations), cellophane “edge”, and cellophane “center”.

On agar surfaces, pilA expression was significantly higher compared to broth culture

(Figure 2.1B), consistent with a previous study (Kuchma et al., 2012). Strikingly, bacteria

from the “edge” sub-population on cellophane sheets exhibited 14 or 8-fold elevation of

the pilA mRNA level compared to cells harvested directly from agar surfaces or the

“center” sub-population, respectively (Figure 2.1B). This observation is consistent with a

previous finding using subsurface TM assays that the edge subpopulation produced

significantly more PilA than the center (Semmler et al., 1999). Therefore, we conclude that

under this newly developed condition, the “edge” sub-population on cellophane surfaces

conducts highly active TM and is an ideal subject for downstream TM studies.

TM phenotypes of P. aeruginosa strains on cellophane surfaces

On surfaces where flagella-mediated swimming motility is suppressed, TFP

provide opportunities for bacteria to explore new territories via TM (9, 34). The subsurface

twitching assay is a valuable tool to distinguish TM-proficient from TM-deficient P.

aeruginosa. We therefore sought to determine whether there is a TFP-dependent phenotype

using the new cellophane-based method. Four P. aeruginosa strains were tested, including

two TM-proficient strains (PAO1 and the flagella mutant ΔfliC) and two TM-deficient

strains (ΔpilA and ΔamrZ). When the subsurface twitching method was used, TM-

proficient strains exclusively formed twitching zones outside of the central colony as

expected (Figure 2.2A). In cellophane-based assays, to discern differences between TM-

proficient and TM-deficient strains, it was necessary to use phase contrast microscopy.

These differences were not readily apparent by the naked eye. Microscopic examinations

revealed any strain capable of TM (PAO1 and ΔfliC) displayed tendril-like structures at

colony edges on cellophane sheets, which were absent in TM defective strains (ΔpilA and

ΔamrZ) (Figure 2.2B). Notably, formation of these tendril-like structures is flagella- independent, as the flagella mutant ΔfliC still formed tendrils similar to its parental strain

PAO1. In conclusion, this cellophane approach can serve as a new method to identify TM-

proficient from deficient strains.

Definition of the AmrZ regulon under this TM-promoting condition

It is unknown how the transcription factor AmrZ activates P. aeruginosa TM

(Baynham et al., 2006). We hypothesized that AmrZ activates TM by modulating the transcription of target TM gene(s). Therefore, we sought to apply this newly-developed method to understand the mechanism of AmrZ-mediated TM activation. DNA microarray

analyses were performed to identify AmrZ-dependent targets using cells harvested from

“edge” sub-populations of the parental strain PAO1, two isogenic amrZ mutants (ΔamrZ

and AmrZV20A), and a complemented ΔamrZ strain. AmrZV20A harbors a V20A point

mutation that renders the protein deficient in DNA binding, a property required for its role

in controlling TM (P. J. Baynham et al., 2006; Waligora et al., 2010). The complemented

ΔamrZ strain was constructed by placing the amrZ gene with its native promoter back into

the ΔamrZ mutant at the native amrZ locus (P. J. Baynham et al., 2006). Our rationale was

that any gene transcriptionally regulated by AmrZ would exhibit differential expression in

amrZ mutants including ΔamrZ and AmrZV20A (AmrZ-) compared to strains expressing

WT amrZ (PAO1 and the complemented strain; AmrZ+). It should be noted that during

bacterial cell harvest, although amrZ mutants do not twitch and their colony edges appeared

distinct from TM-proficient strains, we still harvested cells from their colony edge portions.

As shown in Figure 2.3A, the AmrZ regulon contains 112 genes.

Multiple virulence pathways were affected by amrZ mutations (Figure 2.3B). A complete list of AmrZ-dependent genes is summarized in Table 2.2. We confirmed mRNA levels of selected targets by qRT-PCR (Figure 2.4). Key virulence pathways regulated by

AmrZ are categorized and discussed below.

i) Diguanylate cyclase. AmrZ inhibited the expression of the diguanylate cyclase

(DGC)-encoding gene gcbA (PA4843, also referred to as adcA), confirming previous

findings (Jones et al., 2014; Petrovaet al., 2014). GcbA is required for surface attachment and regulates flagella-mediated motility in P. fluorescens, and gcbA deletion led to reduced biofilms in P. aeruginosa (C. J. Jones et al., 2014; Petrova et al., 2014). Deletion of this

DGC gene results in reduced c-di-GMP levels, which serves as a crucial second messenger regulating numerous processes involved in transitions between sessile and motile lifestyles

(Hengge, 2009; C. J. Jones et al., 2014; Luo et al., 2015).

ii) Lectins. Both lecA and lecB exhibited reduced expression in amrZ mutants.

Galactose-specific lectin LecA and fucose-specific lectin LecB mediate P. aeruginosa

adhesion and cytotoxicity towards epithelial cells, as well as contribute to lung damage in

an acute mouse infection model (Tielker et al., 2005; Chemaniet al., 2009). In addition, lecB was shown to be necessary for robust TM and caseinolysis (Sonawane et al., 2006).

Since lecB was the only AmrZ-dependent gene found in our profiling studies known to be

involved in TM, we focused on this in more detail (see below).

iii) Pyocin. AmrZ activated pyocin genes such as pys2 and prtN. The pys2 gene

encodes the large component of the bacteriocin pyocin 2, and exhibits DNase activity (Sano

et al., 1993). PrtN functions as the activator for all types of pyocins (Matsui, Sano, Ishihara,

& Shinomiya, 1993).

iv) Rhamnolipids. AmrZ also activated rhlA expression. The rhlA gene encodes

chain A of the rhamnosyltransferase, and is required for the synthesis of 3-(3-hy-

droxyalkanoyloxy) alkanoic acids (HAAs) and the fatty acid moiety of rhamnolipids

(Déziel et al., 2003; Soberón-Chávezet al., 2005). The biosurfactant rhamnolipids are

involved in multiple processes such as swarming and sliding motilities, biofilm

detachment, and have antimicrobial activities against other bacteria (Boles et al., 2005;

Kohler et al., 2000; Murray & Kazmierczak, 2008).

v) Iron. Under this condition, AmrZ modulated iron metabolism via activating the

expression of hasAp and repressing hemH. HasAp is necessary for haemoglobin uptake in

P. aeruginosa, while the ferrochelatase HemH is the final enzyme involved in the heme

biosynthesis pathway (Létoffé et al., 1998; Schobert and Jahn, 2002).

vi) Denitrification. AmrZ regulated transcription of multiple operons involved in denitrification. The majority of genes within nirS and nosR operons showed significantly reduced expression in amrZ mutants. These genes are required for anaerobic respiration using nitrate/nitrite as electron receptors, indicating the impact of AmrZ on denitrification.

AmrZ activates lecB transcription by directly binding to its promoter

One goal in this study was to use this novel approach to identify AmrZ-dependent genes necessary for TM. As discussed above, only lecB within the defined AmrZ regulon was reported to be necessary for TM (Sonawane et al., 2006). Therefore, we determined if

AmrZ activates lecB transcription directly by binding to its promoter. A 5’-[6FAM]- labeled DNA fragment containing the lecB promoter region was amplified (Figure 2.5A), and its binding by AmrZ was determined through the electrophoretic mobility shift assay

(EMSA). Addition of AmrZ caused mobility retardation of this DNA fragment, suggesting

AmrZ-DNA binding. Moreover, this retardation required a DNA binding-proficient AmrZ, since the DNA binding deficient protein AmrZR22A was unable to cause a shift. In this

EMSA assay, promoter fragments from algD and algB were used as positive and negative controls respectively. From this, we conclude that AmrZ specifically binds to the lecB promoter.

The lecB gene is not necessary for TM in P. aeruginosa PAO1

We hypothesized that since lecB was the only AmrZ-dependent gene known to be

required for TM, lecB ectopic expression in the ΔamrZ mutant would restore TM.

Therefore, we placed the lecB gene into the backbone of the arabinose-inducible vector

pHERD20T. However, pHERD20T-lecB failed to restore TM in the ΔamrZ mutant after

arabinose induction (Figure 2.6B).

In P. aeruginosa strain PAK, lecB deletion led to the absence of surface pili and

defective TM (Figure 2.6A) (Sonawane et al., 2006). Therefore, to examine the role of lecB

in the PAO1 strain, we constructed an in-frame ΔlecB mutant, and measured its TM via the

subsurface twitching assay. Surprisingly, the ΔlecB mutant displayed normal TM

compared to its parental strain PAO1 (Figure 2.6B). This result was confirmed by

microscopic observations using the cellophane-based method (Figure 2.7).

Discussion

In this study, we developed a novel cellophane-based approach for harvesting TM- active bacterial cells. This approach provides the convenience of recovering enough cells for downstream studies, the validity of which was confirmed by multiple assays. We then applied this method to define the AmrZ regulon through DNA microarray analyses and discover AmrZ-dependent genes. Absence of functional AmrZ led to global changes in

gene expression, including the lectin-encoding gene lecB. We also revealed that lecB was not required for TM in P. aeruginosa strain PAO1.

P. aeruginosa is capable of thriving under diverse environments. Adaptation into new environments usually requires detection of external stimuli, intracellular signal transduction, and corresponding responses. In this study, we showed that pilA transcript levels were 69 times higher when grown on cellophane sheets than in broth culture. This suggests that P. aeruginosa has a remarkable capability of maintaining low pilA expression in liquid culture, but activating it when grown on surfaces, especially smooth ones such as cellophane sheets. Presumably minimal pilA expression in broth culture may conserve resources for bacteria when TFP are not necessary. In contrast, on surfaces where TFP are critical for migration, pilA expression is enhanced. On cellophane sheets, cells appeared to differentiate into “edge” and “center” sub-populations, and the “edge” sub-population displayed significantly higher TM activities than “center”, similar to previous findings using the subsurface twitching assay (Semmler et al., 1999). Activation of P. aeruginosa

TFP responses on surfaces involves effective surface recognition, which requires multiple systems, including but likely not limited to the Wsp signal transduction system, the Chp chemotaxis system, and PilY1 in the TFP system (Bertrand et al., 2010; Fulcher, Holliday,

Klem, Cann, & Wolfgang, 2010; Güvener & Harwood, 2007; Luo et al., 2015; O’Connor,

Kuwada, Huangyutitham, Wiggins, & Harwood, 2012). Surface attachment resulted in phosphorylation and clustering of the response regulator WspR and an increased level of the second messenger cAMP, leading to a rise of the c-di-GMP concentration and the

activation of the cAMP-binding regulator Vfr, respectively (Güvener & Harwood, 2007;

Luo et al., 2015). Downstream effects involve elevated PilY1 expression and its secretion by the TFP system, serving as one plausible mechanism to explain our observation of increased pilA expression on surfaces versus liquid.

Twitching motility is a well-coordinated group behavior, resulting in the formation of “bacterial rafts” on interstitial spaces (Semmler et al., 1999; Gloaget al., 2013). Our microscopic observations on cellophane sheets were consistent with those findings.

Tendril-like structures were observed at leading edges of exclusively TM+ P. aeruginosa

similar to those rafts described previously. It is intriguing how bacteria aggregate and

properly arrange themselves in these rafts. The Psl exopolysaccharide fibers and

extracellular DNA (eDNA) are two factors involved in this phenomenon. Previous studies

observed that Psl fibers held bacteria together and linked aggregates, which were absent in

TM-deficient strains (L. Ma et al., 2009; S. Wang, Parsek, Wozniak, & Ma, 2013). In

another elegant study, Gloag et al. showed that DNase I treatment inhibited the formation

of lattice-like networks formed by TFP (Gloag et al., 2013). Potential roles of Psl and DNA

on TM under our cellophane condition require further investigation.

With this new TM-promoting approach, we defined the AmrZ regulon through

DNA microarray analyses. Absence of functional AmrZ led to global gene expression

changes, and multiple pathways associated with pathogenesis were impacted. Previously

we and others defined the AmrZ regulon via RNA-Seq of mid-log phase bacteria grown in

liquid culture, one in P. aeruginosa and the other one in P. fluorescens (Jones et al., 2014;

Martínez-Graneroet al., 2014). A comparison was made between the current DNA microarray analysis and the previous P. aeruginosa RNA-seq study (C. J. Jones et al.,

2014). The AmrZ regulons from both studies shared multiple genes, such as the DGC-

encoding gene gcbA, and hypothetical genes PA0102, PA3235, etc. (Table 2.2). However,

there were significant differences between these two studies. First, growth conditions were

dissimilar in these two studies. We and others have shown that pilA expression varies depending on growth conditions (activated on surfaces while repressed in liquid) (Figure

2.1B; (Kuchma et al., 2012)), and it is likely that other genes would behave differently in these conditions. Second, in the RNA-Seq study, the comparison was made between a

ΔamrZ mutant and an amrZ-overexpressing strain, which mimics alginate-overproducing mucoid P. aeruginosa (C. J. Jones et al., 2014). However, in the current study all strains are in the non-mucoid PAO1 background and have significantly lower AmrZ levels, which may lead to distinct regulatory outcomes.

Our DNA microarray results compared transcriptional profiles between AmrZ- and

AmrZ+ strains. Within the four strains used, one AmrZ- strain encodes a DNA binding-

deficient variant AmrZV20A and is TM deficient, while the complemented strain re- obtained WT AmrZ and has restored TM (Baynham et al., 2006). Thus, the comparison between AmrZ- and AmrZ+ strains should only reveal targets that are modulated by DNA

binding-proficient AmrZ. It was therefore surprising that 62 genes had differential

expression between the complemented strain and PAO1. However, the majority of these

genes had differential expression only slightly above the 2-fold threshold. Only 10 genes

showed more than 3-fold difference, and the largest difference was 6.4-fold. Nevertheless,

the possibility of unknown artifacts introduced during strain construction cannot be

excluded. Therefore, these 62 genes have been excluded from the AmrZ regulon.

This study discovered that both lectin-encoding genes lecA and lecB exhibited

differential expression between AmrZ- and AmrZ+ strains. In addition, AmrZ specifically

bound to the lecB promoter, indicating direct AmrZ activation. Interestingly, in a previous

study when ChIP-Seq was used to determine AmrZ binding sites throughout the P.

aeruginosa genome, AmrZ was not observed to bind to the lecB promoter (Jones et al.,

2014). This could also be attributed to different growth conditions. It is plausible that in broth culture and possibly other conditions, certain signals suppress AmrZ binding to the lecB promoter thereby preventing its transcriptional activation.

Our original hypothesis was that AmrZ activates TM through lecB, which was shown to be required for TM in P. aeruginosa strain PAK. Surprisingly, our ΔlecB mutant in PAO1 did not show a TM defect, in contrast to the study in the PAK strain background

(Sonawane et al., 2006). To determine if the lecB gene differs between these two strains, we sequenced the lecB coding and flanking sequences. There was a single nucleotide variation at the 48th nucleotide in the lecB coding sequence: a deoxythymidine in PAO1

and a deoxycytidine in PAK (data not shown). However, this polymorphism does not lead

to a change in the amino acid sequence, since both codons (ACU and ACC) encode

threonine. Nevertheless, it is possible that with a different genetic background the lecB

gene in PAK might play a different role compared to our PAO1 strain. A comprehensive

transposon mutant library was constructed in another PAO1 strain MPAO1 and screened

to identify TM-necessary genes (Jacobs et al., 2003). This screening did not identify any

lecB mutant that lost TM. Meanwhile, there was no overlap between tfp genes identified in

the transposon mutant screening (Jacobs et al., 2003) and AmrZ-dependent genes in the current study. Therefore, the AmrZ regulon described in this study does not contain any known gene(s) necessary for TM. Therefore, despite our efforts, the mechanism for AmrZ- mediated control of TM therefore remains unknown. It is a formal possibility that AmrZ may regulate TM at a post-transcriptional level, e.g. via small RNAs, during translation or through protein-protein interactions. This is less likely because of the stringent requirement of DNA binding-proficient AmrZ for TM. The most likely explanation therefore is that TM genes controlled by AmrZ have not been identified in previous genetic strategies to identify

TM-defective strains. Our current aim is to use alternative approaches to identify these targets.

Overall, this study described a novel TM-promoting approach that facilitates the harvest of TM-active cells for downstream transcriptomics studies. We defined the AmrZ regulon under this growth condition, and uncovered the role of AmrZ as a global regulator in P. aeruginosa. This novel approach may be applied to other bacterial species capable of

TM and facilitate more high-throughput research aiming at understanding regulatory mechanisms of this unique form of motility.

Acknowledgement

DNA microarray experiments were performed in Genomics Shared Resource of

OSU Comprehensive Cancer Center. We appreciate the critical readings by Mohini

Bhattacharya, Sheri Dellos-Nolan, Dr. Valerie A. Ray and Preston J. Hill. We also thank

Drs. Reuben Ramphal/Jeevan Jyot for sending PAK, and PAK ΔlecB strains and respective plasmids, and Dr. Joe. J. Harrison for the ΔfliC strain.

Figure 2.1. Cellophane-air interface stimulates P. aeruginosa twitching motility.

(A.) Illustration of the cellophane-based TM-promoting condition. (B.) Quantification of

pilA transcript levels from indicated growth conditions via qRT-PCR. “Broth” refers to overnight broth culture (stationary phase). Transcript levels of pilA were normalized to those isolated from single colonies on agar surfaces (“agar”) with rpoD as the reference gene. Unpaired two-tailed student t-test was used for statistical analysis of three independent experiments. *: P<0.05; **: P<0.01; ***: P<0.001.

Figure 2.2. TM-proficient and TM-deficient strains retain their TM phenotypes on cellophane.

(A.) TM measurement of four P. aeruginosa strains by subsurface twitching assays. Scale bar: 5 mm. (B.) Phase contrast microscopy of edge sub-populations of the same strains above cellophane sheets. Dashed lines illustrated tendril-like structures. Scale bar: 100 µm.

Figure 2.3. Identification of the AmrZ regulon from the TM-active “edge” sub- population.

(A.) Venn diagram of the AmrZ regulon under the TM-promoting condition. 112 genes in the red box were included in the AmrZ regulon as they displayed differential expression between both amrZ mutants (ΔamrZ and AmrZV20A) and strains encoding WT AmrZ

(PAO1 and the complemented strain ΔamrZ/amrZ). The other three numbers indicate the total number of genes that exhibited distinct expression from PAO1, respectively. (B.)

Representative virulence pathways regulated by AmrZ. Red arrows indicate activation, while green bars represent repression.

Figure 2.4. Verification of representative AmrZ-dependent genes.

Bacteria growth conditions and RNA isolations were the same as for microarray analyses.

Gene expression in the ΔamrZ mutant was compared to the parental strain PAO1 by qRT-

PCR. The horizontal dashed line represented normalized mRNA levels as seen in the parental strain PAO1. Unpaired two-tailed student t-test was used for statistical analysis of three independent experiments. *: P<0.05; **: P<0.01; ***: P<0.001.

Figure 2.5. AmrZ specifically binds to the lecB promoter.

(A.) Design of the lecB DNA fragment used in (B.). The star represents [6FAM] used to label the 5’ end of the lecB fragment. The arrow indicates the lecB transcription start site.

(B.) EMSA was used to determine AmrZ binding to the lecB promoter, the positive control

“algD” DNA, and the negative control “algB” DNA. Black arrows point at delayed

mobility of DNA. Two AmrZ variants (AmrZ and the DNA binding-deficient variant

AmrZR22A) were used in the experiment. “-“: No protein added; “+”: 50 nM protein;

“++”: 100nM protein.

Figure 2.6. The lecB gene is required for TM in P. aeruginosa strain PAK, but not in

strain PAO1.

(A.) TM of the parental strain PAK and its derivative lecB::GmR via subsurface twitching

assays (Sonawane et al., 2006). (B.) Subsurface twitching assays of strains in PAO1

background. In the ΔamrZ strain with the plecB plasmid, 1% arabinose was added as the inducer. Scale bar: 5 mm.

Figure 2.7. lecB is required for TM in PAK, but not in PAO1 using the cellophane- based method.

Phase contrast microscopy was used to examine edge subpopulations of PAO1, PAK, and their respective lecB mutants after growth using the cellophane-based method. Dashed lines and arrows illustrated tendril-like structures.

Table 2.1. Strains, plasmids, and oligonucleotides used in Chapter 2

Strains /Plasmids Relevant genotype/description/sequence Sources

Strains Escherichia coli NEB5α fhuA2 Δ(argF-lacZ)U169 phoA glnV44 Φ80 New England Δ(lacZ)M15 gyrA96 recA1 relA1 endA1 thi-1 Biolabs hsdR17 S17λpir pro thi hsdR+ TpR SmR; chromosome::RP4-2 (Simon et al., Tc::Mu-Kan::Tn7/λpir 1983) P. aeruginosa PAO1 strain background PAO1 PAO1 wild type (P. J. Baynham et al., 2006) WFPA205 ΔamrZ (P. J. Baynham et al., 2006) WFPA513 AmrZV20A (P. J. Baynham et al., 2006) WFPA510 amrZ complemented in WFPA205 (P. J. Baynham et al., 2006) JJH283 ΔfliC J.J. Harrison PAO-JP3 ΔlasR&ΔrhlR (Mittal, Sharma, Chhibber, & Harjai, 2006) OSUPA239 ΔlecB This study OSUPA268 ΔpilA This study

PAK strain background PAK PAK wild type (Sonawane et al., 2006) PAK ΔlecB PAK lecB::GmR (Sonawane et al., 2006) Plasmids pEX18Ap Deletion construct backbone. Cb300 (Hoang et al., 1998) pBX27 PAO1 lecB deletion construct in pEX18Ap. This This study plasmid was constructed by lecB_ABCD primers pBX38 PAO1 pilA deletion construct in pEX18Ap. This This study plasmid was constructed by pilA_ABCD primers

Continued 67

Table 2.1. Continued pBX39 PAO1 amrZ deletion construct in pEX18Ap. This This study plasmid was constructed by primers amrZ122-125 pHERD20T Arabinose-inducible vector (Qiu et al., 2008) plecB (pBX14) pHERD20T-lecB. Primers lecB_CDS_F and This study lecB_CDS_R were used to amplify the lecB coding sequence pJN105 Empty vector. Cb300 (J. W. Hickman, Tifrea, & Harwood, 2005) pJN2133 pJN105-PA2133. Cb300 (J. W. Hickman et al., 2005) Oligonucleotides lecB_CDS_F cgctctagaAGGCAGGCCAGGTATTCAGTG This study lecB_CDS_R cgcgcatgcCCATCCCGTCCCTTCCGAAC This study pilA_RT_F CGACGATCAACCCGCTGAAGAC This study pilA_RT_R ACTTGTTGGCATCCGGCTCGA This study lecA_RT_F CGTTTTGTGGTGCGCTGGTC This study lecA_RT_R TAGGTTCCGGGCACGTCGTT This study lecB_RT_F AGTGTTCACCCTTCCCGCCA This study lecB_RT_R GTGCTTTGCCCGCTGAAGGT This study pys2_RT_F GGAAGCAGAAGCCCAGCGAG This study pys2_RT_R TGGCGTTGGATTGGTCAGTTGAG This study prtN_RT_F GAACGCTGGTTTCGCAACCTG This study prtN_RT_R GGCGCTTCCGGCTTGGTTC This study rhlA_RT_F CGCTCAACGATCGGGGCTAC This study rhlA_RT_R CTCCACCCGCGAGAAACTGC This study nirS_RT_F GCGAATCGTGGAAGGTGCTG This study nirS_RT_R TGACCTTGACGATCTTCTTGCTG This study norC_RT_F CCTGACCTACCACACCGAGAAG This study norC_RT_R GCAGCCGACGCAGTTGTTC This study hemH_RT_F CGAGAACAGCCGCAACGTCA This study hemH_RT_R GCGACTGGAACGACACCGAC This study hasAp_RT_F GACAGCCAGGAGGTGAGCTTC This study hasAp_RT_R CGTCGATCTGCCCCTGCAG This study rpoD_RT_F GCCGAGCTGTTCATGCCGAT (C. J. Jones et al., 2014) rpoD_RT_R GAACAGGCGCAGGAAGTCGG (C. J. Jones et al., 2014) lecB_A cccctctagaTTGATCGGGAGTCATAGGGTAC This study

Continued 68

Table 2.1. Continued lecB_B GTCCCTTCCGAACTCCTAGCGTTGCCATGG This study TGTATCTCCACT lecB_C AGTGGAGATACACCATGGCAACGCTAGGA This study GTTCGGAAGGGAC lecB_D ccgaagcttGGAGGTGGCGGTGATCAAC This study pilA_A tgctctagaGCTTTTCGCTGATGGCGTC This study pilA_B AACAAGCCACCTTCGATCACCGAATCTCTC This study CGTTGATTATGTA pilA_C TACATAATCAACGGAGAGATTCGGTGATCG This study AAGGTGGCTTGTT pilA_D aactgcagCGCATAGCACCCGGCAAG This study amrZ122 tgctctagaGGAGCGTGGATTTGCCGGAG This study amrZ123 TACGCGTGGGCTTCGGCGCACATTGAACCT This study GTAGAGTCAGG amrZ124 CCTGACTCTACAGGTTCAATGTGCGCCGAA This study GCCCACGCGTA amrZ125 cccaagcttCCACAGATACAGGTAGGCGATC This study lecB_EMSA_F [6FAM]CGGCATCGGCAATAGATCGTTAC This study lecB_EMSA_R GAACCGGGTGTTGGCGGGAAG This study algB68 [6FAM]CGCATGCCAGCCTTTCTGA This study algB69 TTCACTGCCATTCGGCTTCTG This study algD71 [6FAM]ACCGTTCGTCTGCAAGTCATG This study algD73 ACCAACTTGATGGCCTTTCCG This study

Table 2.2. Complete gene list from microarray analyses

ID Fold change Adj.P.Val Notes (mutant/WT) PA0045_at 2.285 0.011 Hypothetical lipoprotein. Homolog of CsgG. PA0067_prlC_at 2.002 0.011 75% similarity to oligopeptidase A. PA0102_at 0.457 0.015 Probable carbonic anhydrase PA0129_gabP_at 2.603 0.001 bauD. Amino acid permease PA0130_at 2.098 0.008 bauC. 3-Oxopropanoate dehydrogenase PA0167_at 2.379 0.003 Bacterial regulatory proteins, tetR family signature PA0296_s_at 2.240 0.003 spuI/pauA1. Glutamyl polyamine synthetase PA0298_at 2.356 0.004 spuB/pauA2. Glutamyl polyamine synthetase PA0509_nirN_at 0.290 0.007 Gene involved in denitrification processes PA0510_at 0.237 0.002 Gene involved in denitrification processes PA0511_nirJ_at 0.171 0.006 Gene involved in denitrification processes PA0512_at 0.392 0.002 Gene involved in denitrification processes PA0513_at 0.218 0.003 Gene involved in denitrification processes PA0514_nirL_at 0.170 0.007 Gene involved in denitrification processes PA0515_at 0.112 0.010 Gene involved in denitrification processes PA0516_nirF_at 0.177 0.003 Gene involved in denitrification processes PA0517_nirC_at 0.113 0.013 Gene involved in denitrification processes PA0518_nirM_at 0.083 0.011 Gene involved in denitrification processes PA0519_nirS_at 0.075 0.012 Gene involved in denitrification processes PA0534_at 2.026 0.003 pauB1. FAD-dependent oxidoreductase PA0610_prtN_at 0.450 0.003 Transcriptional regulator PrtN. Pyocin activator PA0612_i_at 0.432 0.008 prtB. DnaK suppressor protein PA0614_at 0.282 0.002 Hypothetical protein. Transmembrane protein PA0617_at 0.362 0.002 Phage protein PA0618_at 0.392 0.002 Phage protein PA0620_at 0.373 0.001 Phage protein PA0622_at 0.368 0.001 Phage protein PA0639_at 0.378 0.002 Phage protein PA0643_at 0.384 0.001 Phage protein Continued

Table 2.2. Continued PA0743_at 0.427 0.016 Probable 3-hydroxyisobutyrate dehydrogenase PA0866_aroP2_at 3.862 0.002 Aromatic amino acid transport protein AroP2 PA0887_acsA_at 2.997 0.004 acsA. Acetyl-coenzyme A synthetase PA0911_at 0.374 0.002 Hypothetical protein. PA1070_braG_at 2.022 0.007 Branched-chain amino acid transport protein BraG PA1072_braE_at 2.285 0.007 Branched-chain amino acid transport protein BraE PA1150_pys2_at 0.415 0.001 pys2. Pyocin S2 PA1338_ggt_at 2.539 0.012 ggt. Gamma-glutamyl transpeptidase precursor. Amino acid biosynthesis and metabolism PA1494_at 0.346 0.001 Conserved hypothetical protein. Contains type I secretion signal. PA1545_at 0.423 0.002 Hypothetical protein PA1732_at 0.314 0.004 Conserved hypothetical protein. May be involved in amino acid transport and metabolism PA1736_at 0.480 0.009 Probable acyl-CoA thiolase PA1737_at 0.494 0.011 Probable 3-hydroxyacyl-CoA dehydrogenase PA1757_thrH_at 3.010 0.001 Homoserine kinase PA1818_at 2.749 0.002 ldcA. Lysine-specific pyridoxal 5'- phosphate-dependent carboxylase, LdcA PA1978_at 2.245 0.004 Response regulator ErbR. Involved in ethanol oxidation pathway. PA1983_exaB_at 2.336 0.006 Cytochrome c550 PA2015_at 2.186 0.005 liuA. Putative isovaleryl-CoA dehydrogenase PA2080_at 2.025 0.003 kynU. Kynureninase KynU. PA2081_at 2.326 0.007 Kynurenine formamidase, KynB PA2134_at 0.150 0.002 Hypothetical protein. Contains type I secretion signal. PA2146_i_at 0.100 0.001 Conserved hypothetical protein PA2169_at 0.341 0.016 Hypothetical protein PA2174_at 0.266 0.002 Hypothetical protein PA2190_at 0.171 0.002 Conserved hypothetical protein

Continued

Table 2.2. Continued PA2204_at 2.534 0.004 Probable binding protein component of ABC transporter. Periplasmic protein PA2329_at 2.660 0.001 Probable ATP-binding component of ABC transporter. Cytoplasmic membrane PA2445_gcvP2_at 2.094 0.002 gcvP2. Glycine cleavage system protein P2 PA2446_gcvH2_a 3.290 0.001 Glycine cleavage system protein H2 t PA2531_at 0.312 0.001 Probable aminotransferase PA2570_pa1L_at 0.122 0.008 lecA PA2711_at 2.053 0.008 Probable periplasmic spermidine/putrescine-binding protein PA2788_at 0.408 0.013 Probable chemotaxis transducer PA2843_at 2.781 0.009 Probable aldolase PA2862_lipA_at 2.619 0.040 Lactonizing lipase precursor PA2937_at 0.286 0.002 Hypothetical protein PA2945_at 2.557 0.001 Conserved hypothetical protein PA2946_at 2.542 0.002 Hypothetical protein PA3032_at 0.394 0.004 Cytochrome c Snr1 PA3234_at 2.209 0.021 Probable sodium: solute symporter PA3235_at 2.304 0.023 Conserved hypothetical protein PA3284_at 0.483 0.004 Hypothetical protein PA3356_at 2.086 0.003 Glutamyl polyamine synthetase PauA5 PA3361_at 0.198 0.003 lecB. Fucose-binding lectin PA-IIL. (lecB is required for TFP biogenesis) PA3365_at 2.176 0.013 Probable chaperone PA3393_nosD_at 0.376 0.004 Denitrification gene PA3394_nosF_at 0.281 0.006 Denitrification gene PA3396_nosL_at 0.450 0.007 Denitrification gene PA3407_hasAp_at 0.430 0.010 Heme acquisition protein HasAp PA3415_at 0.327 0.009 Probable dihydrolipoamide acetyltransferase PA3416_at 0.302 0.011 Probable pyruvate dehydrogenase E1 component, beta chain PA3417_at 0.314 0.009 Probable pyruvate dehydrogenase E1 component, alpha subunit PA3479_rhlA_at 0.221 0.009 rhlA PA3520_at 0.165 0.002 Hypothetical protein Continued

Table 2.2. Continued PA3539_at 2.603 0.023 Conserved hypothetical protein PA3836_at 2.122 0.002 hypothetical protein PA3837_at 2.767 0.001 Probable permease of ABC transporter PA3872_narI_at 0.270 0.048 Respiratory nitrate reductase gamma chain. narI PA3920_at 0.396 0.019 Probable metal transporting P-type ATPase PA3923_at 2.559 0.021 Hypothetical protein PA4033_at 4.045 0.002 MucE PA4129_at 2.275 0.021 Hypothetical protein PA4130_at 2.359 0.003 Probable sulfite or nitrite reductase PA4182_at 2.427 0.001 Hypothetical protein PA4211_g_at 0.386 0.036 phzB1. Probable phenazine biosynthesis protein PA4294_at 0.479 0.004 Hypothetical protein PA4442_cysN_at 2.277 0.006 ATP sulfurylase GTP-binding subunit/APS kinase PA4443_cysD_at 2.190 0.016 ATP sulfurylase small subunit PA4448_hisD_at 2.047 0.016 Histidinol dehydrogenase PA4655_hemH_at 2.467 0.007 hemH. Ferrochelatase. PA4675_at 2.223 0.002 Probable TonB-dependent receptor PA4703_at 0.293 0.011 Hypothetical protein PA4843_at 3.993 0.004 Probable two-component response regulator PA5098_hutH_at 2.168 0.019 Histidine ammonia-lyase PA5100_hutU_at 2.583 0.002 Urocanase PA5131_pgm_at 2.033 0.010 Phosphoglycerate mutase PA5154_at 2.252 0.002 Probable permease of ABC transporter PA5168_at 2.367 0.005 DctQ PA5169_at 2.990 0.002 DctM PA5312_at 2.099 0.004 PauC. Aldehyde dehydrogenase PA5313_at 2.358 0.007 gabT2. Transaminase PA5429_aspA_at 2.256 0.010 Aspartate ammonia-lyase PA5519_at 2.387 0.020 Conserved hypothetical protein

Chapter 3

Chapter 3. The Pseudomonas aeruginosa AmrZ C-terminal domain mediates

tetramerization and is required for its activator and repressor functions

Abstract

Pseudomonas aeruginosa is an important bacterial opportunistic pathogen,

presenting a significant threat towards individuals with underlying diseases such as cystic

fibrosis. The transcription factor AmrZ regulates expression of multiple P. aeruginosa

virulence factors. AmrZ belongs to the ribbon-helix-helix protein superfamily, in which

many members function as dimers, yet others form higher-order oligomers. In this study,

four independent approaches were undertaken and demonstrated that the primary AmrZ

form in solution is tetrameric. Deletion of the AmrZ C-terminal domain leads to loss of

tetramerization and reduced DNA binding affinity to both activated and repressed target

promoters. Additionally, transcriptional fusion experiments showed that the C-terminal

domain is essential for AmrZ-mediated activation of the alginate biosynthesis operon.

Repression of gcbA by AmrZ is also compromised upon removal of the C-terminal domain.

To summarize, this study identified that the AmrZ C-terminal domain mediates 74

tetramerization, and AmrZ tetramers comprised of DNA binding-proficient monomers are necessary for efficient activation and repression of AmrZ-dependent genes.

Introduction

The Gram-negative bacterium Pseudomonas aeruginosa is ubiquitous. It can be widely found in fresh water and soil, and as an opportunistic pathogen, also imposes a threat to immunocompromised human populations. As an example, emergence of this bacterium is considered poor prognosis in lung infections for cystic fibrosis patients

(Govan & Deretic, 1996). P. aeruginosa versatility is achieved by rapidly adapting to new environments, in which multiple levels of regulation are involved. A large pool of transcription factors participate in this regulation, and one such factor AmrZ (Alginate and

Motility Regulator Z) functions as a global activator and repressor. Well-studied AmrZ targets include those involved in exopolysaccharide production (alginate, Psl and Pel), flagella, twitching motility, and the metabolism of the second messenger bis-(3’, 5’) cyclic di-guanylate (c-di-GMP) (P. J. Baynham et al., 2006; P. J. Baynham & Wozniak, 1996; C.

J. Jones et al., 2014, 2013; Martínez-Granero et al., 2014; Ramsey et al., 2005; Tart et al.,

2006). For instance, AmrZ is necessary for transcription of the alginate biosynthesis operon, the promoter of which is designated as PalgD since algD is the first gene of this operon. Alginate is responsible for the mucoid phenotype and contributes to increased resistance to antimicrobials and host immunity (P. Baynham et al., 1999; Gacesa, 1998;

Ramsey & Wozniak, 2005). AmrZ also represses transcription of gcbA (also referred to as

adcA), which encodes a diguanylate cyclase enzyme that synthesizes c-di-GMP (Jones et

al., 2014; Petrova et al., 2014). Deletion of gcbA results in increased motility and reduced

biofilm production (C. J. Jones et al., 2014; Petrova et al., 2014). The ΔamrZ mutant displays increased levels of Psl and c-di-GMP, and therefore forms enhanced biofilms compared to the wild type strain (C. J. Jones et al., 2014, 2013).

AmrZ is composed of three segments: a flexible N terminus (residues 1-11), a ribbon-helix-helix domain (residues 12-66), and a C-terminal domain (CTD, residues 67-

108). The ribbon-helix-helix domain mediates DNA binding and its structure is highly similar to other members, such as Arc, Mnt and MetJ, in the Arc superfamily (Knight et al., 1989; Schreiter & Drennan, 2007; Vershons et al., 1985; Waligora et al., 2010). All proteins within this superfamily form dimers or higher order oligomers, which are often necessary for their functions (Schreiter & Drennan, 2007; Waldburger & Sauer, 1995). In our previous studies, glutaraldehyde cross-linking experiments suggested that cross-linked

AmrZ forms oligomers independent of its N terminus (Waligora et al., 2010). However, it remains unknown which AmrZ domain(s) mediates oligomerization, or what roles this may play during AmrZ-mediated activation or repression.

In this study, size exclusion chromatography (SEC) revealed that AmrZ exists primarily as tetramers in solution under the physiological ionic strength (150 mM NaCl).

This result was confirmed by nano electrospray ionization mass spectrometry (nano-ESI

MS). Additionally, as expected with proteins that function as oligomers, a DNA binding-

deficient variant AmrZR22A, when overproduced, exhibits a dominant-negative effect on expression of AmrZ-dependent genes. We showed that the AmrZ CTD is responsible for oligomerization, and loss of the domain leads to reduced DNA binding affinity as well as diminished ability of AmrZ to mediate activation and repression.

Taken together, our findings have unveiled that the primary form of AmrZ is tetrameric. Moreover, we highlight the significance of the AmrZ CTD during AmrZ- mediated regulation of multiple target genes.

Materials and Methods

Bacterial strains, plasmids, oligonucleotides, and culture conditions

All bacterial strains, plasmids, and DNA oligonucleotides are listed in Table 3.1.

Strains of Escherichia coli and Pseudomonas aeruginosa were incubated as previously described (Jones et al., 2013). All E. coli strains were cultured in LB (1 liter LB contains

10 g tryptone, 5 g yeast extract, 5 g sodium chloride) or on LA (LB with 1.5% (w/v) agar).

If necessary, 100 µg*ml-1 ampicillin or 15 µg*ml-1 tetracycline was used to maintain

plasmids in E. coli. All P. aeruginosa strains were cultured in LBNS or on LANS (LB or

LA without sodium chloride). For P. aeruginosa cultures, 300 µg*ml-1 carbenecillin or 100

µg*ml-1 tetracycline was used when needed. All strains were grown at 37°C unless stated

otherwise. All oligonucleotides were ordered desalted from Sigma Aldrich. 77

Size exclusion chromatography

A 29.4 nmol aliquot of purified AmrZ was loaded onto a Superose 12 10/300 GL gel filtration column equilibrated and eluted in the same protein buffer (100 mM Bis-Tris

pH 6.5, 150 mM NaCl, 5% glycerol) (Pryor et al., 2012). Gel filtration standards (Bio-Rad,

#151-1901) were used to generate a standard curve describing the relationship between

retention time and protein molecular weights for this column. The protein extinction

coefficient of AmrZ WT was calculated as 1490 M-1*cm-1 by the ExPASy ProtParam tool

(Artimo et al., 2012).

Mass Spectrometry

Purified proteins (native AmrZ or AmrZ1-66, monomeric concentrations at 147 µM and 150 µM respectively) were buffer exchanged into 500 mM ammonium acetate by a

size exclusion spin column (Micro Bio-Spin 6, Bio-Rad, Hercules, CA). Proteins were

charge reduced by mixing 100 mM triethylammonium acetate (Sigma, Milwaukee, WI)

with protein solutions (in 500 mM ammonium acetate) to 20% by mole. Nano-electrospray

ionization mass spectrometry was used to analyze protein solutions on a Synapt G2S

instrument (Waters MS Technologies, Manchester, U.K.). The quadrupole profile was set

to 2500 for AmrZ to minimize low m/z of impurities and salts after charge reduction. The

glass capillary was pulled in-house using a Sutter Instrument (Novato, CA) P-97

micropipette puller and a platinum wire was inserted into the capillary to apply a voltage

of 1.0~1.2 kV. The cone voltage was kept at 20 V and the source temperature at 25°C to

minimize in source activation and denaturation of proteins.

DNA binding studies

We used EMSAs (electrophoretic mobility shift assays) modified from a previously

described protocol (Jones et al., 2014) to study DNA-protein interactions. 5’-[6FAM]

labeled DNA used for EMSA was amplified using Quick-load® Taq 2X Mastermix (New

England Biolabs), [6FAM]-labeled forward primers and non-labeled reverse primers, and

PAO1 genomic DNA as the template. Each EMSA reaction contained 20 mM Bis-Tris pH

-1 6.5, 4 mM MgCl2, 200 µg*ml acetylated BSA, 130 mM NaCl, 1 mM DTT, 5% glycerol,

150 ng*µl-1 Poly d[(I-C)] (Roche), 10 nM [6FAM] labeled DNA, and a specified

concentration of purified AmrZ or AmrZ1-66. Protein-DNA binding was equilibrated at

room temperature (25°C) for 20 min after adding all reagents to each reaction. 10 µl of

each reaction was loaded onto a cold pre-equilibrated 4% non-denaturing acrylamide gel.

Electrophoresis was conducted for 25 min at 200 V in 0.5% TBE on ice. A Typhoon scanner (GE Lifescience) (Settings: fluorescence, PMT800, 520BP 40 CY2 Blue FAM,

100 µm pixel size) was used to detect FAM fluorescence of gels. DNA fragments used were amplified using following primers – algD: algD71 + algD73; non-specific control:

algD111 + algD112; gcbA: adcA_FAM_F_EMSA + adcA_R_EMSA (C. J. Jones et al.,

2014).

Transcriptional reporter assays

All transcriptional reporters were constructed in the mini-CTX-lacZ plasmid

(Becher & Schweizer, 2000). After digestion by HindIII and BamHI (New England

Biolabs), the algD promoter (PalgD) (-633 - +368 relative to its transcriptional start site) was cloned into mini-CTX-lacZ using primers algD65 and algD66. Transcriptional reporters were then transferred and integrated into the chromosomes of the mucoid P. aeruginosa strain FRD1 and its isogenic ΔamrZ mutant FRD1200 as previously described

(Wyckoff & Wozniak, 2001). A 96-well plate reader (Molecular Devices SpectraMax M2) was used to measure absorbance at 420 nm and 550 nm, and β-galactosidase activities were determined as previously described (J. H. Miller, 1992).

Quantitative real-time PCR

Transcript levels of genes of interest were measured through quantitative real-time

PCR (qRT-PCR) assays. To isolate total RNA, overnight LANS-grown bacteria were harvested after flooding with LBNS. 1 ml OD600=1.0 bacteria were lysed by addition of

100 µl TE containing 1 mg*ml-1 RNase-free lysozyme (Sigma Aldrich), and then

underwent RNA isolation (RNeasy RNA isolation kit, Qiagen). To obtain cDNA, ~1 µg 80

RNA was used for reverse transcription (Superscript III cDNA preparation kit, Invitrogen)

following manufacturer protocols. Amounts of cDNA were determined by SYBR Green

Mix (Bio-Rad) and CFX1000 thermal cyclers (Bio-Rad). Gene expression was normalized

to that of the housekeeping gene rpoD, and the ΔΔCt method was used to calculate relative

gene expression (Livak and Schmittgen, 2001; Jones et al., 2014).

Protein cross-linking experiments

Cross-linking by glutaraldehyde was performed as previously described (Waligora

et al., 2010). Briefly, purified AmrZ was incubated with 0.1 M glutaraldehyde for 5 min.

Reactions were reduced by an equimolar concentration of sodium borohydride, chilled, and

neutralized by 167 mM Tris-HCl (pH 8.0). Samples were resolved by 12% SDS-PAGE and stained with Coomassie Brilliant Blue.

Western blot analyses

AmrZ and His-tagged protein levels were measured by Western blot. After growing to exponential phases in the presence/absence of arabinose/Carbenecillin, bacteria equivalent to 150 µl OD600=1.0 were resuspended by 75 µl SDS-PAGE sample buffer

(100 mM Tris-HCl pH=6.8, 0.04% sodium dodecyl sulfate, 3.6% glycerol, 2% 2- mercaptoethanol, 0.008% Bromophenol Blue), and lysed at 100°C for 5 min. After cooling down to room temperature, cell debris was removed by centrifugation at top speed for 1 81

min. 30 µl cell lysates were loaded into 15% Criterion™ (Bio-Rad) Tris-glycine SDS-

PAGE gels. Proteins were transferred onto nitrocellulose membranes via a semi-dry transfer system (Bio-Rad), and detected by the rabbit anti-AmrZ polyclonal antibody

(1:5,000) or the mouse anti-His tag monoclonal antibody (1:2,000, Thermo Scientific,

Product #: MA1-21315). After secondary antibody was applied, membranes were imaged by the Clarity™ Western ECL Blotting Substrate (Bio-Rad) and ChemicDoc (Bio-Rad).

Subsurface twitching motility assays

Subsurface twitching assays were conducted as previously described (Baynham et al., 2006). In brief, a single bacterial colony was resuspended in 400 µl LBNS, and a P10 tip dipped into the resuspension was stabbed into 1% LANS with antibiotics and/or arabinose if necessary. Plates (at minimum triplicates for each strain) were then incubated at room temperature in a humid chamber for five days. Twitching zones were imaged through white epiluminescence by ChemicDoc (Bio-Rad). The twitching motility activity for each strain was represented by the ratio of the twitching zone diameter divided by that of the central colony.

Results

AmrZ forms tetramers in solution via its C-terminal domain

In the Arc protein superfamily, the basic functional unit is dimeric; however, proteins such as Mnt and TrwA form tetramers in solution and even higher-order oligomers when bound to DNA (Schreiter & Drennan, 2007). To determine the AmrZ oligomeric state in solution, we performed SEC of purified AmrZ at the physiological ionic strength

(150 mM NaCl). SEC is widely used for protein purification and molecular weight measurements of proteins/protein complexes without the need of changing protein buffers

(Berek, 2010). AmrZ eluted as one major species with a molecular weight of approximately

51.76 kDa, which is consistent with a slightly non-globular tetramer (49.36 kDa) (Figure

3.1A&B). Additionally, nano-ESI MS was used to determine the AmrZ oligomeric state.

This MS method is powerful due to its small sample requirement and measurement when proteins/protein complexes remain folded (Karas, Bahr, & Dülcks, 2000; X. Ma et al.,

2014). Nano-ESI MS showed that the majority of AmrZ exists as native tetramers, yet some dimers were also detected (Figure 3.2B). Collectively, results from both assays suggest that the primary form of AmrZ in solution is tetrameric. We also used nano-ESI MS to determine the stoichiometry when AmrZ binds to its DNA target amrZ1 from the amrZ

promoter (Pryor et al., 2012). We observed a complex with the size of ~64000 Da (Figure

3.3), corresponding to a DNA molecule bound by four AmrZ monomers. This is consistent with previous studies (Pryor et al., 2012).

We next determined which part of AmrZ mediates oligomerization. AmrZ is

comprised of three segments. Previous work showed that removal of N-terminal residues

1-11 does not prevent AmrZ oligomerization, and the ribbon-helix-helix domain (residues

12-66) is responsible for DNA binding and dimerization (Figure 3.2A) (Pryor et al., 2012;

Waligora et al., 2010). Specifically, AmrZ1-66 was used in determining the crystal

structure of DNA-bound AmrZ, and this truncation variant interacts with the same DNA

sequence as a dimer of dimers (Pryor et al., 2012). Therefore, the AmrZ CTD does not

appear to affect DNA recognition and we hypothesized that the AmrZ CTD mediates

tetramerization. To test this, we determined the oligomeric state of purified AmrZ in the

absence of the CTD (AmrZ1-66). Nano-ESI MS of this truncated protein revealed that

AmrZ1-66 exists as dimers and monomers, but no tetramer was observed (Figure 3.2C).

This is consistent with data obtained from protein cross-linking experiments with full length and CTD-truncated AmrZ (Figure 3.4). Based on the results above, we conclude that the AmrZ CTD mediates tetramerization in solution.

A genetic method was also used to indirectly probe the AmrZ oligomeric state. In most biological systems that require protein oligomers for function, expression of dysfunctional subunit(s) in complexes with wild type (WT) ones can exert a negative effect on the function of the entire protein complex (i.e. dominant negative). We therefore determined the effects of overproducing a DNA binding-deficient variant (AmrZR22A) in wild type P. aeruginosa. AmrZR22A harbors a point mutation at the 22nd residue (arginine to alanine), leading to loss of DNA binding affinity and subsequent effects on AmrZ-

dependent gene expression (Waligora et al., 2010). However, AmrZR22A still forms tetramers as revealed by nano ESI-MS (Figure 3.5). Plasmids containing amrZ genes encoding different AmrZ variants (AmrZ WT or AmrZR22A) were transferred into the

FRD1 strain carrying a chromosomal algD-lacZ transcriptional fusion (note, this strain harbors a chromosomal WT amrZ allele), and PalgD activities were measured after induction by various arabinose concentrations. The algD gene is the first gene in the alginate biosynthesis operon, and previously we showed transcriptional activation of this operon requires AmrZ (P. J. Baynham & Wozniak, 1996). Western blot analyses suggested that AmrZ and AmrZR22A were induced by arabinose at comparative levels (Figure 3.6).

The empty vector pHERD20T did not affect algD transcription (Figure 3.7B), while overproduction of plasmid-borne AmrZ elevated PalgD activation (Figure 3.7B), likely due to gene dosage. However, the PalgD activity was reduced significantly when

AmrZR22A was overproduced (Figure 3.7B), suggesting a dominant negative effect of

AmrZR22A.

Impact of the AmrZ C-terminal domain on DNA binding, activation and repression

We next sought to determine effects of removing the AmrZ CTD on DNA binding affinity. Purified AmrZ and AmrZ1-66 proteins were tested for DNA binding affinity to promoters of AmrZ targets. A [6FAM]-labeled DNA fragment containing the AmrZ binding site (ZBS) was amplified from its activated target PalgD (P. J. Baynham &

Wozniak, 1996), and the AmrZ binding affinity determined by EMSA (Figure 3.8A).

Results showed that AmrZ1-66 has significantly reduced affinity to PalgD compared to

full length AmrZ (Figure 3.8B&C). However, the CTD is not absolutely necessary for

DNA binding, since DNA binding activities were seen when higher AmrZ1-66

concentrations were used (Figure 3.8B&C). Both AmrZ and AmrZ1-66 recognize the DNA

target specifically, as no binding was seen when a non-specific DNA was tested in this

assay (Figure 3.8D). Similar results were observed when the repressed target PgcbA (Jones

et al., 2014) was used (Figure 3.9), indicating this reduced DNA binding affinity of

AmrZ1-66 is likely to be conserved for different AmrZ targets.

To test roles of the AmrZ CTD in vivo, arabinose-inducible plasmids encoding

either AmrZ or AmrZ1-66 were transferred into a P. aeruginosa FRD1 ΔamrZ strain that

contained a chromosome-borne algD-lacZ transcriptional fusion. PalgD activities were

measured after AmrZ or AmrZ1-66 production was induced by addition of arabinose.

PalgD activation required a significantly higher amount of AmrZ1-66 compared to full length AmrZ (Figure 3.10). For instance, substantially more arabinose was needed to restore 15% PalgD activity for pHERD20T-AmrZ1-66 (pAmrZ1-66) (0.5% arabinose) compared to pAmrZWT (0.1% arabinose). We also measured AmrZ amounts after 0.1% arabinose induction, which showed that the intracellular level of AmrZ1-66 was 53% of that of AmrZ after taking into account reduced antibody recognition in the absence of the

CTD (Figure 3.11). We therefore conclude that the observed reduction in PalgD activation by AmrZ1-66 is likely to be contributed by both decreased protein levels and reduced DNA binding affinity. In this transcriptional fusion assay, an empty vector (pHERD20T) was

introduced into the FRD1 ΔamrZ strain as a negative control, and no PalgD activity was

seen at any arabinose concentration (Figure 3.10).

Finally, we sought to investigate the significance of the AmrZ CTD in the type

strain PAO1. In a previous study, we showed that AmrZ activates PA2146 transcription

and represses gcbA transcription (Jones et al., 2014; Petrova et al., 2014). The hypothetical gene PA2146 is 92% similar to Escherichia. coli yciG, which is associated with bacterial responses to glucose pulses (Sunya, Gorret, Delvigne, Uribelarrea, & Molina-Jouve, 2012).

Moreover, PA2146 expression is repressed when P. aeruginosa is treated with inhibitory

compounds such as protoanemonin and azithromycin (25, 26). We compared the ability of

AmrZ1-66 and full length AmrZ to regulate these two targets. In the PAO1 ΔamrZ strain

with plasmids containing AmrZ or AmrZ1-66, mRNA levels of PA2146 and gcbA were

quantified at various arabinose concentrations. Consistent with findings in RNA-Seq (C. J.

Jones et al., 2014), gcbA transcription remained de-repressed in the PAO1 ΔamrZ strain

when the empty vector pHERD20T was introduced. However, even at 0% arabinose,

pHERD20T-WT AmrZ (pAmrZWT) was sufficient to restore AmrZ-mediated repression

of gcbA, while regulation by AmrZ1-66 was not observed until 0.1% arabinose was used

(Figure 3.12A). Similar results were observed with the activated target PA2146, as it

required a much higher amount of AmrZ1-66 for transcription compared to full length

AmrZ (Figure 3.12B).

Our prior studies in the type strain PAO1 also revealed a role for AmrZ in activation

of type IV-fimbriae dependent twitching motility (Baynham et al., 2006). Although the

mechanism of AmrZ-mediated activation of twitching motility (TM) has yet been

identified, the AmrZ CTD remains important during activation of this behavior (Table 3.2).

With all data above, we conclude that AmrZ tetramerization through the CTD is

essential for efficient activation and repression of AmrZ-dependent targets.

Discussion

In this study, we found that AmrZ exists primarily as tetramers in solution. In addition, the AmrZ CTD is responsible for tetramerization and necessary for efficient transcriptional regulation of activated and repressed targets of AmrZ.

Protein-protein interactions are prevalent throughout biological systems and are

critical for flexible and rapid regulation of biological functions (Matthews et al., 2012). A

common example of protein-protein interactions is homo- or hetero-

dimerization/oligomerization. In fact, monomers are likely to be the minority form of

proteins. For example, in the Brenda enzyme database (http://www.brenda-enzymes.org,

as of April 2015), when searched through the “subunits” field, enzymes known as

“monomer” only comprise ~15% of the total (1769/11964). Oligomerization not only

provides an economic way to form large complexes, but also enables proteins, such as

transcription factors, to flexibly modulate their DNA binding specificity and affinity.

Differential sequence specificity can be achieved through oligomerization of different

combinations of transcription factors (Matthews et al., 2012). In some circumstances

oligomerization significantly enhances DNA binding affinity. For example, the λ repressor

CTD is responsible for high affinity binding to its operator sites in the phage genome (Bell,

Frescura, Hochschild, & Lewis, 2000). In the present study, we provide evidence that after removing the CTD, AmrZ loses the ability to form tetramers and exhibits reduced DNA binding affinity to both activated and repressed targets, suggesting the significance of the

CTD during AmrZ-mediated regulation. Although it is clear that this domain mediates tetramerization, further investigations are required to determine if it has other functions.

Members in the Arc protein family represent diverse DNA binding proteins in various phage and bacterial species, and some also contain CTDs. CTDs in proteins such as Mnt and TrwA mediate tetramerization (Madl et al., 2006; Moncalián & De La Cruz, 2004;

Waldburger & Sauer, 1995), while at least two members (NikR and MetJ) exhibit significantly higher DNA binding activities when their CTDs are bound by ligands (nickel and S-adenosylmethionine, respectively) (Peter T. Chivers & Sauer, 2002; Rafferty et al.,

1989). To determine if the sequence of the AmrZ CTD is similar to its homologs within the Arc protein superfamily, ClustalW2 alignment was performed between AmrZ with other Arc family members including Mnt, TrwA, NikR and MetJ (W. Li et al., 2015).

However, no significant similarity was observed (Figure 3.13) and AmrZ does not appear to harbor motifs required for ligand (nickel and S-adenosylmethionine) binding. While our current data support the requirement of the AmrZ CTD for tetramerization, with effects on

DNA binding and subsequent gene expression, we cannot exclude a role of the CTD in other functions, such as ligand binding.

A number of biophysical, biochemical and genetic methods have been developed

to study protein oligomerization. In this study we used four independent approaches

(glutaraldehyde cross-linking, SEC, nano-ESI MS, and genetics) to demonstrate the

oligomeric state of native AmrZ as tetrameric, and identified that the CTD mediates

tetramerization. Chemical cross-linking can be achieved using reagents such as

glutaraldehyde to link different subunits of the same protein or different proteins. This

approach is convenient and cost-effective, but cross-linked products may not reflect native states of proteins/protein complexes, thus the results need wary interpretation (Kluger &

Alagic, 2004). In SEC, proteins are loaded into and eluted off columns in their native buffers, so proteins retain native folding and remain oligomerized. Molecular weights of proteins/complexes are calculated through the standard curve generated by globular standard proteins. Since protein shapes affect elution rates, calculated weights of proteins/complexes are typically estimated. In contrast, nano-ESI MS has the advantages of small sample volume requirement and precise detection of protein/complex masses. This method becomes even more powerful in structural elucidation of proteins/complexes when combined with tandem MS such as surface-induced dissociation (X. Ma et al., 2014).

However, MS requires protein buffers changed to volatile reagents such as ammonium acetate, which might affect native states of proteins/complexes. In the current study results of above three approaches are consistent, providing clear evidence of AmrZ tetramerization via its CTD. We also used a genetic approach as additional evidence. DNA binding- deficient AmrZR22A was overproduced in a wild type P. aeruginosa strain, and a

dominant negative effect on PalgD activation was observed. The most likely explanation

is that non-functional monomers are incorporated into hetero-tetramers that then are non-

functional. This dominant negative phenomenon has been widely observed in other

oligomer-forming regulators, such as E. coli LacI and the Shigella flexneri AraC homolog

VirF (Betz & Fall, 1988; Porter & Dorman, 2002).

To unveil how the AmrZ CTD contributes to tetramerization, we used multiple

tools to predict the secondary structure and properties of this domain. The NCBI conserved

domain database failed to identify any conserved domain homologous to this CTD

(Marchler-Bauer et al., 2014). The online prediction tools PSIPRED (Figure 3.14A) and

Jpred (B. Xu, unpublished) predicted that most residues in the AmrZ CTD form two α-

helices (Drozdetskiy, Cole, Procter, & Barton, 2015; D. T. Jones, 1999). Alpha-helices,

which mediate DNA binding and are essential for transmembrane domain formation, are

the most common secondary structure of proteins. Many proteins, such as G-protein coupled receptors, form dimers or oligomers via helix-helix packing (Breitwieser, 2004).

Therefore, these two α-helices are likely to be responsible for AmrZ oligomerization.

Typically a small number of residues, called hot spot residues, are important for successful helix-helix interactions (Bullock, Jochim, & Arora, 2011). Although hydrophobic and

aromatic amino acids constitute the majority of hot spot residues, polar and charged

residues such as arginine remain significant players at monomer-monomer interfaces

(Bogan & Thorn, 1998; Bullock et al., 2011; Matthews et al., 2012). The ExPASy

ProtScale tool (Gasteiger et al., 2005) predicted most residues within the AmrZ CTD

domain are polar/hydrophilic (Figure 3.14B). In addition, the overall hydropathicity of this

domain is hydrophilic, with the grand average of hydropathicity at -0.512 (minus values

suggest hydrophilic) (ProtParam, (Gasteiger et al., 2005; Kyte & Doolittle, 1982)). The

AmrZ CTD contains five arginine residues, which, together with other polar residues, may

contribute to oligomerization through hydrogen bonding and salt bridges (Ali & Imperiali,

2005; Matthews et al., 2012).

Overall, we have investigated the significance of the AmrZ CTD for AmrZ

oligomerization as well as AmrZ-mediated activation and repression. Tetramers composed of four DNA binding-proficient monomers are critical for AmrZ-mediated regulation.

Further work is necessary to understand which residues are at the interface between monomers and mediate inter-monomer interactions. Since this CTD does not appear to be present in other proteins, more understanding of this domain may provide insights of a novel oligomerization mechanism and potentially other unknown functions of this domain.

Acknowledgement

This work was completed in collaboration with laboratories of Richard Fishel and

Vicki H. Wysocki, and specifically Randal J. Soukup and Yue Ju have provided tremendous help during collaborations. Public Health Service awards AI097511 and

NR013898 (DJW), an NSF instrument development grant DBI0923551 (VHW) and funds provided from the Public Health Preparedness for Infectious Diseases (PHPID) program 92

(phpid.osu.edu) supported this work. BX was partially supported by the student fellowship

grant from the cystic fibrosis foundation (XU13H0).

We thank Dr. Thomas Hollis for kindly providing AmrZ and AmrZ1-66, Dr. Joe J.

Harrison for providing the E. coli S17λpir strain, and Dr. Christopher J. Jones for the pCJ4 plasmid. We are grateful to Sheri Dellos-Nolan and Dr. Meenu Mishra for critical reading of this manuscript. We also acknowledge Dr. Nrusingh Mohapatra for assistance with β- galactosidase assays.

Figure 3.1. The primary form of AmrZ in solution is tetrameric.

Size exclusion chromatography was used to determine the oligomerization state of AmrZ under physiological ionic strength. (A.) AmrZ was loaded into the gel filtration column, and eluted proteins were monitored by absorbance at 280 nm. The vertical dashed line indicates a single peak was observed with the AmrZ sample. The small peaks at ~20 mL were from the buffer wash. (B.) A standard curve depicting the relationship between molecular weight (Y-axis) and elution volume (X-axis) was generated using protein standards. The molecular weight of the peak at 12.76 mL corresponded to 51.76 kDa.

Figure 3.2. The C-terminal domain of AmrZ mediates tetramerization.

(A.) Proposed domains of full length AmrZ and AmrZ1-66. N: N terminus; B: DNA binding domain; Z: oligomerization domain. Numbers above indicate residues flanking each domain. (B.) & (C.) Nano-ESI MS measurements of purified AmrZ (B.) and AmrZ1-

66 (C.). The X-axis represents mass/charge (m/z) ratios, while the Y-axis was calculated by normalizing to the highest signal intensity.

Figure 3.3. AmrZ binds to DNA as a tetramer.

Nano-ESI MS was used to determine the molecular weight of the AmrZ-DNA complex.

10 µM 18-bp amrZ1 dsDNA was incubated with 40 µM recombinant AmrZ. Black arrows point to a complex with the size ~64246 Da, which corresponds to the combined size of one DNA molecule and four AmrZ monomers. The Y-axis was calculated by normalizing to the highest signal intensity.

Figure 3.4. The C-terminal domain of AmrZ is required for tetramerization.

(A.) Domain structure of AmrZ. N: N-terminal tail. B: DNA binding domain. Z: oligomerization domain. (B.) & (C.) His-tagged AmrZ was cross-linked by glutaraldehyde.

Samples were resolved in 12% SDS-PAGE gels and stained with Coomassie Brilliant Blue.

The protein amount in each reaction was as below: (-) glutaraldehyde 500 ng; (+) glutaraldehyde 250 ng, 500 ng, 750 ng, 1 µg, 1.5 µg, 2 µg, 2.5 µg, and 5 µg. M: monomer;

D: dimer; T: tetramer.

Figure 3.5. AmrZR22A forms tetramers in solution.

Nano-ESI MS of purified AmrZR22A after buffer exchange into 500 mM ammonium

acetate. The X-axis represents mass/charge (m/z) ratios, while the Y-axis was calculated by normalizing to the highest signal intensity.

Figure 3.6. Similar amounts of AmrZR22A and WT AmrZ protein levels were

produced after arabinose induction.

(A.) P. aeruginosa strains were grown with/without arabinose to mid-log phases. Identical

amounts of bacterial cells were lysed, loaded on 15% SDS-PAGE, followed by western blots with the α-AmrZ antibody. 0%, 0.1%, and 1% represent arabinose concentrations. A non-specific cross-reacting band (~25 kDa) was used as the loading control (LC) for each western blot assay. (B.) AmrZ and AmrZR22A band intensities after 1% arabinose induction from panel A were quantified by densitometry via ImageJ (v. 1.46r) and normalized to the loading control.

Figure 3.7. Dominant negative effect of AmrZR22A.

(A.) Depiction of the point mutation (arginine to alanine) at the 22nd residue of AmrZ. (B.)

Three arabinose-inducible plasmids (pHERD20T, pAmrZWT and pHis-AmrZR22A) were

transferred into the FRD1 strain with integrated algD-lacZ transcriptional fusion. Various

arabinose concentrations were used to induce plasmid-borne protein production in broth culture, and algD promoter activities were measured and normalized to that in the wild type FRD1 strain with the empty vector without arabinose. Error bars (only above bars were shown for a cleaner graph) represent standard errors of the mean from three independent experiments. Unpaired two-tailed t-tests of three independent experiments were performed to determine statistically significant differences. **: P<0.01; ***:

P<0.001.

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Figure 3.8. Truncation of the C-terminal domain results in reduced binding to PalgD.

[6FAM]-labeled DNA fragments containing the ZBS in PalgD (B) and a control DNA (D) were incubated with various concentrations of AmrZ or AmrZ1-66 and analyzed by EMSA

(B. & D.). Relative amounts of free DNA in (B.) were quantified via densitometry of non- shift bands using ImageJ (v1.46r) and plotted at various AmrZ concentrations (C.). AmrZ monomeric protein concentrations were as below: Lane 1, 10: 0 nM; 2, 6, 11, 15: 10 nM;

3, 7, 12, 16: 20 nM; 4, 8, 13, 17: 50 nM; 5, 9, 14, 18: 100 nM. The arrow in (A.) represents the transcription start site of PalgD. Arrows in (B.) indicate DNA mobility shifts induced by AmrZ or AmrZ1-66 binding. Triangles in (B. & D.) indicate non-specific bands, which are also present in no protein control lanes.

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Figure 3.9. AmrZ1-66 displays reduced binding to the gcbA promoter.

A [6FAM] labeled DNA fragment containing the AmrZ binding site (ZBS) was amplified, purified and tested for AmrZ binding. AmrZ protein concentrations were as below: Lane

1, 5: 0 nM; 2, 6: 10 nM; 3, 7: 20 nM; 4, 8: 30 nM. The arrow in (A.) represents the transcription start site of the gcbA promoter. Arrows in (B.) indicate DNA mobility shifts induced by AmrZ or AmrZ1-66 binding.

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Figure 3.10. Efficient activation of PalgD requires the AmrZ C-terminal domain.

Arabinose-inducible plasmids encoding AmrZ or AmrZ1-66 were transferred into a FRD1

∆amrZ mutant (FRD1200) harboring an algD-lacZ transcriptional fusion. Levels of AmrZ or AmrZ1-66 were modulated using indicated arabinose concentrations (0%-1%). PalgD activities were measured by β-galactosidase assays and normalized to that in the ∆amrZ mutant with the empty vector without arabinose. Unpaired two-tailed student t-tests were used for statistical analyses of three independent experiments. ***: P<0.001.

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Figure 3.11. Western blot analyses of AmrZ1-66 after arabinose induction.

(A.) Western blots of purified AmrZ and AmrZ1-66 using the polyclonal α-AmrZ antibody.

Protein concentrations used for both AmrZ and AmrZ1-66 from left to right are as below:

2 µg, 1 µg, 500 ng and 200 ng. (B.) Western blots of OSUPA211 with different plasmids with/without induction by 0.1% arabinose. Same amounts of cells were used. (C.)

Densitometry of band intensities in (A.) and (B.) by ImageJ (v. 1.46r). The first (antibody recognition ratio) and second columns (apparent recognition ratio) were calculated and averaged from (A.) and (B.) (0.1% arabinose), respectively. The third column (actual ratio of protein amount) was calculated by dividing the apparent recognition ratio by the antibody recognition ratio in order to take into account the effect of reduced antibody recognition of AmrZ1-66.

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Figure 3.12. The AmrZ C-terminal domain is required for efficient repression and activation of its targets.

Arabinose-inducible plasmids encoding AmrZ or AmrZ1-66 were transferred into a PAO1

∆amrZ mutant. After induction by various arabinose concentrations, cells were harvested, and mRNA levels of gcbA (A.) and PA2146 (B.) quantified by qRT-PCR. Relative gene expression was compared to the PAO1 ∆amrZ mutant with the empty vector in the absence of arabinose. Unpaired two-tailed student t-tests were used for statistical analyses of three independent experiments. ***: P<0.001; ns: not significant.

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Figure 3.13. ClustalW2 alignment of AmrZ with other Arc proteins.

ClustalW2 alignment v2.1 was used to align above protein sequences using the Gonnet protein weight index (W. Li et al., 2015). The colon symbol “:” and the period symbol “.” indicate conservation of strongly and weakly similar properties –scoring >0.5 and =<0.5 in the Gonnet PAM 250 matrix, respectively.

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Figure 3.14. Predicted AmrZ secondary structure and polarity of the AmrZ CTD.

(A.) The AmrZ secondary structure was predicted by Psi Pred (D. T. Jones, 1999). (B.)

Polarity scores of the AmrZ CTD (residues 67-108) by ProScale (Gasteiger et al., 2005).

Higher scores (Y-axis) represent more polar amino acid residues. The X-axis represents the position of a certain residue within the domain.

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Table 3.1. Strains, plasmids, and oligonucleotides used in Chapter 3

Strains Relevant genotype/description/sequence Sources /Plasmids Strains Escherichia coli NEB5α fhuA2 Δ(argF-lacZ)U169 phoA glnV44 Φ80 New England Δ(lacZ)M15 gyrA96 recA1 relA1 endA1 thi-1 hsdR17 Biolabs S17λpir pro thi hsdR+ TpR SmR; chromosome::RP4-2 Tc::Mu- (Simon et al., Kan::Tn7/λpir 1983) P. aeruginosa FRD1 strain background FRD1 mucA22 (Ohman & Chakrabarty, 1981) FRD1200 amrZ::xylE-aacC1 in FRD1 (Wozniak et al., 2003) OSUPA211 algD-lacZ transcriptional fusion at the attB site in This study (FRD1/PalgD-lacZ) FRD1 OSUPA213 Promoterless lacZ at the attB site in FRD1 This study OSUPA226 amrZ deletion; WT algD-lacZ transcriptional fusion at This study the attB site in FRD1 PAO1 strain background

PAO1 PAO1 wild type D. J. Wozniak WFPA205 amrZ::tet (P. J. Baynham et al., 2006) Plasmids mini-CTX-lacZ Vector for transcriptional fusion analyses. TetR (Hoang TT Becher A, Schweizer HP., 2000) pBX1 mini-CTX-PalgD-lacZ. Two primers (algD65, This study algD66) were used to amplify the PalgD pHERD20T Arabinose-inducible vector (Qiu et al., 2008) pBX29 (pAmrZWT) pHERD20T-WT AmrZ. Primers amrZ118, amrZ119 This study were used to amplify the amrZ coding sequence. pBX30 (pAmrZ1-66) pHERD20T-AmrZ1-66. Primers amrZ118-121 were This study used to construct an amrZ mutant encoding a stop codon TGA at 67th residue Continued 108

Table 3.1. Continued pCJ3 (pHis- pHERD20T-His6-AmrZ WT (C. J. Jones AmrZWT) et al., 2014) pCJ4 (pHis- pHERD20T-His6-AmrZR22A C. J. Jones AmrZR22A) Oligonucleotides algD65 CCCCAAGCTTCTCTTTCGGCACGCCGAC This study algD66 CCGGGATCCCCGACATAGCCCAAACCAAAG This study algD71 [6FAM] ACCGTTCGTCTGCAAGTCATG This study algD73 ACCAACTTGATGGCCTTTCCG This study algD111 [6FAM] CAAACGGCCGGAACTTCCCT (C. J. Jones et al., 2014) algD112 TAGTTCGGTCCATAGAATTCAAG (C. J. Jones et al., 2014) amrZ118 TGCTCTAGACGAGAACAATGAACGCTTCCTC This study amrZ119 CCCAAGCTTCTTCGGCGCTCAGGCCTG This study amrZ120 CTGTCCAGTCAAACACCGAG This study amrZ121 CTCGGTGTTTGACTGGACAG This study CTX1 CCTCGTTCCCAGTTTGTTCC (Tart et al., 2006) attB2 GTCGCCGCCGGCGATGC (Tart et al., 2006) attB4 CGCCCTATAGTGAGTCG (Tart et al., 2006) attB5 CGCCCCAACCTCGCTGG (Tart et al., 2006) rpoD_F2 GCCGAGCTGTTCATGCCGAT (C. J. Jones et al., 2013) rpoD_R2 GAACAGGCGCAGGAAGTCGG (C. J. Jones et al., 2013) gcbA_RT_F GGTTCTGGAGGTCTGGCAAC This study gcbA_RT_R CTGGGCATGCTCGGCTTG This study adcA_FAM_F_EMS [6FAM] CGTAGTCCGTCGCACAAAG (C. J. Jones A et al., 2014) adcA_R_EMSA GCGCTTCTTTCGTGGTCTTC (C. J. Jones et al., 2014) PA2146_RT_F GCACAGCATCAAGGTGGTAAAG This study PA2146_RT_R CTTTCTTGCCAGCCTCGGAG This study

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Table 3.2. The AmrZ CTD is required for efficient activation of twitching motility

0%a 0.01% 0.1%

∆amrZ/vector 0.0b ± 0.0c 0.0 ± 0.0 0.0 ± 0.0 ∆amrZ/pAmrZWT 65.7 ± 8.7 65.7 ± 13.9 97.3 ± 4.4

∆amrZ/pAmrZ1-66 0.0 ± 0.0 1.0 ± 2.3 93.9 ± 16.4

a Arabinose concentration. b TM was measured and normalized to percentages of PAO1. c Standard errors of the mean were calculated from three independent experiments, each with three replicates. 110

Chapter 4

Chapter 4. Pseudomonas aeruginosa AmrZ binds to four sites in the algD promoter and

induces trans DNA-AmrZ complex formation

Abstract

During late stages of cystic fibrosis pulmonary infections, Pseudomonas aeruginosa often overproduces the exopolysaccharide alginate, protecting the bacterial community from host immunity and antimicrobials. The major target of regulation is the alginate biosynthesis operon, whose transcription is under tight control by a number of transcription factors including AmrZ. Interestingly, multiple transcription factors interact with the far upstream region of this promoter (PalgD). Transcription is proposed to be activated via DNA looping, yet this model has little supporting evidence. Prior, ChIP-Seq analyses revealed what appeared to be two AmrZ binding regions in PalgD. Further examinations here allowed the identification of four AmrZ binding sites in this promoter, disruption of which eliminated AmrZ binding and promoter activation. We proposed that

AmrZ induces DNA looping of PalgD. In support of this, PalgD transcription was abolished when the relative phase was reversed (5 bp), while a full-DNA-turn insertion (10 bp) restored promoter activity. Since AmrZ forms oligomers in solution and when bound

111

to DNA, we sought to determine if AmrZ brings its binding sites together and induces

DNA-AmrZ complex formation. Our in vitro fluorescence resonance energy transfer

experiments suggested that AmrZ holds two binding sites together and induces trans DNA-

AmrZ complex formation. Taken together, our studies have unveiled three previously

unknown AmrZ binding sites in the PalgD promoter. We also highlight the significance of

correct phasing and demonstrate the formation of trans DNA-AmrZ complexes, suggesting looping of the algD promoter in vivo.

Introduction

In cystic fibrosis (CF) patients, the Gram negative bacterium Pseudomonas aeruginosa is a major pathogen. Infection with P. aeruginosa, especially the mucoid strain that overproduces the exopolysaccharide alginate, often results in a poor prognosis for those patients (Govan & Deretic, 1996). During CF lung infections, P. aeruginosa first colonizes as a non-mucoid strain with minimal alginate production. However, at late stages, up to 70% of P. aeruginosa isolates are mucoid (Cystic Fibrosis Foundation, 2013;

Doggett, 1969; Leid et al., 2005; Ramsey et al., 2005; Stapper et al., 2004). Alginate- dominant biofilms form a thick protective layer against host immune attacks and antimicrobials (Leid et al., 2005; Lovewell, Patankar, & Berwin, 2014). However, overproduction of this exopolysaccharide is an energy-exhaustive process, requiring a number of enzymes and precursor substrates. Therefore, alginate production is under tight regulation involving multiple regulatory factors (Ramsey & Wozniak, 2005). Previous

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studies have shown that the sigma factor AlgT (also referred to as AlgU/σ22) is the key component necessary for expression of the alginate biosynthesis operon (Boucher et al.,

2000; M J Schurr et al., 1993; D J Wozniak & Ohman, 1994). Freed from sequestration by

the anti-sigma factor MucA, AlgT initiates transcription of genes encoding transcription

factors (TFs) AmrZ, AlgB and AlgR (P. J. Baynham & Wozniak, 1996; A. K. Jones et al.,

2010). These three and at least four other TFs bind to the promoter of the alginate operon

(PalgD) and initiate transcription (Deretic & Konyecsni, 1990; Kato et al., 1990;

Konyecsni & Deretic, 1990; Mohr et al., 1991). Interestingly, the region over 100 base pairs (bp) upstream of the transcription start site contains at least seven TF binding sites, and it is yet known how these TFs mediate activation from such a long distance. Although a DNA loop model has been proposed, there is little supporting evidence (Michael J Schurr,

2013).

Remote regulatory elements are commonly seen in bacterial promoters, but these elements are almost obligated to couple with proximal elements (Neidhardt & Curtiss,

1996). Generally speaking proteins between remote and proximal elements make contacts via looping out of the intervening DNA. This DNA looping phenomenon has been widely observed in different transcription regulatory systems such as the regulation by the lac repressor, the deo repressor and AraC (Amouyal, Mortensen, Buc, & Hammer, 1989;

Krämer et al., 1987; Lobell & Schleif, 1990). DNA looping plays important roles during transcription regulation, such as allowing remote proteins to execute regulation and blocking the access of RNA polymerase to the promoter.

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As a global regulator, AmrZ functions as both an activator and a repressor of

multiple processes. Well-studied targets include those involved in exopolysaccharide production (alginate, Psl and Pel), flagella, twitching motility and the metabolism of bis-

(3’,5’)-cyclic diguanylate (P. J. Baynham et al., 2006; P. J. Baynham & Wozniak, 1996;

C. J. Jones et al., 2014, 2013; Martínez-Granero et al., 2014; Ramsey et al., 2005; Tart et al., 2006). AmrZ is composed of three domains, including a flexible N terminus (residues

1-11), a DNA binding ribbon-helix-helix domain (residues 12-66) and a C-terminal domain

(CTD, residues 67-108). The ribbon-helix-helix domain is highly similar to other members in the Arc superfamily such as Arc, CopG and MetJ (Knight et al., 1989; Schreiter &

Drennan, 2007; Vershons et al., 1985; Waligora et al., 2010). AmrZ tetramerizes in solution and its CTD truncation variant AmrZ1-66 binds to DNA as a dimer of dimers, suggesting that AmrZ interacts with its DNA targets as oligomers, likely as tetramers

(Pryor et al., 2012; Waligora et al., 2010).

This study sought to dissect AmrZ-mediated activation mechanism of PalgD. We demonstrate the presence of four AmrZ binding sites (ZBSs) in PalgD, each of which is required for promoter activation. In addition, correct phasing between these ZBSs is crucial for PalgD activation and AmrZ induces the formation of trans DNA-AmrZ complexes when bound to a PalgD fragment in vitro. These results suggest DNA looping during PalgD activation.

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Materials & Methods

Bacterial strains, plasmids, oligonucleotides and culture conditions.

All bacterial strains, plasmids and DNA oligonucleotides are listed in Table 4.1.

Strains of Escherichia coli and Pseudomonas aeruginosa were incubated as previously

described (C. J. Jones et al., 2013). All E. coli strains were cultured in LB (1 liter LB contains 10 g tryptone, 5 g yeast extract, 5 g sodium chloride) or on LA (LB with 1.5%

(w/v) agar). Where necessary, 15 µg/ml tetracycline was used to maintain plasmids in E. coli. All P. aeruginosa strains were cultured in LBNS or on LANS (LB or LA without sodium chloride, respectively). In P. aeruginosa, 100 µg/ml tetracycline was used if needed. All strains were grown at 37oC unless stated otherwise. All oligonucleotides were

ordered desalted from Sigma Aldrich with the exception of DNA primers for FRET

experiments, which were ordered with PAGE purification from Midland Certified Reagent

Company.

Nucleotide manipulations.

PalgD variants with mutations or truncations were generated by overlap extension

PCR as previously described (Heckman & Pease, 2007). For scrambling of nucleotide

sequences, the CLC sequence viewer software was used to randomize the order of

nucleotide sequences. In the DNA phasing experiments, sequences of 5bp and 10bp

insertions are “CATGC” and “CATGCATGCA”, respectively.

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Transcriptional reporter assays.

All transcriptional reporters were constructed in the mini-CTX-lacZ plasmid

(Becher & Schweizer, 2000). PalgD fragments (-633 - +368 relative to the transcription start site) were cloned into mini-CTX-lacZ after digestion by HindIII and BamHI (New

England Biolabs). Transcriptional reporters were then transferred and integrated into chromosomes of specified P. aeruginosa strains. A 96-well plate reader (Molecular

Devices SpectraMax M2) was used to measure absorbance at 420 nm and 550 nm, and β- galactosidase activities were determined as previously described (J. H. Miller, 1992).

DNA binding studies. We used EMSA (electrophoretic mobility shift assays) modified from a previously described protocol (C. J. Jones et al., 2014) to study DNA- protein interactions. 5’-[6FAM] labeled DNA used for EMSA was amplified using Quick- load® Taq 2X Mastermix (New England Biolabs), [6FAM]-labeled forward primers and

non-labeled reverse primers, and PAO1 genomic DNA as the template. Each EMSA

reaction contained 20 mM Bis-Tris pH 6.5, 4 mM MgCl2, 200 µg/ml acetylated BSA, 130

mM NaCl, 1 mM DTT, 5% glycerol, 150 ng/µl Poly d[(I-C)] (Roche), 10 nM [6FAM] labeled DNA, and a specified concentration of recombinant AmrZ or its variants. Protein-

DNA binding was equilibrated at room temperature (25°C) for 20 min after the addition of all reagents. 10 µl of each reaction was loaded onto a cold pre-equilibrated 4% non- denaturing acrylamide gel. Electrophoresis was conducted for 25 min at 200 V in 0.5%

TBE on ice in the dark. A Typhoon scanner (GE Lifescience) (Settings: fluorescence,

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PMT800, 520BP 40 CY2 Blue FAM, 100 µm pixel size) was used to detect FAM fluorescence within gels.

Fluorescence Resonance Energy Transfer (FRET)

DNA preparation: 5’ termini of oligonucleotides (algD157 and algD160) (Midland

Certified Reagent Company) were conjugated with Cyanine5 NHS ester and Cyanine3

NHS ester (Lumiprobe) respectively, as recommended by the manufacturer. Labeled oligonucleotides were purified by HPLC using a Poroshell 120 (50 x 4.6 mm) RP column with an EC-C18 stationary phase (Agilent). Samples were dissolved in 0.1 M triethylammonium acetate (TEAA) pH 7.0 and eluted with a 3-30% acetonitrile (ACN) gradient at 1% ACN/min. Labeled fractions were collected and acetonitrile removed by rotary vacuum. Oligonucleotides were concentrated and buffer exchanged to TE with

Amicon Ultra-4 3kDa centrifugal filter units. Oligonucleotides were used as primers in a

PCR reaction using pBX1 as the template to generate the DNA fragment (Quick-Load®

Taq 2X Master mix, New England Biolabs). PCR products were PAGE purified on a 4% acrylamide gel and concentrated using Amicon Ultra-4 10kDa centrifugal filter units.

FRET measurement: The composition of each final FRET reaction mixture was the same as EMSA, with the exception of a different glycerol concentration (2%), and the addition of 1.5 mM Trolox and 4% PEG8000 (polyethylene Glycol 8000). Trolox and

PEG8000 were added as a triplet state quencher and the crowding agent, respectively (Ge,

Luo, & Xu, 2011; Paudel & Rueda, 2014). The mixture was placed in a quartz cuvette

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(Starna, 16.12F-Q-1.5/Z8.5) and measured for fluorescent response with AmrZ

concentrations up to 10 µM. Fluorescence measurements were performed on a Fluoromax-

4 (Horiba Jobin Yvon), with attached circulating water bath set to 25oC. Samples were

excited at 548 nm (Cy3) or 646 nm (Cy5), and emission spectra were scanned from 560

nm to 700 nm. Apparent FRET values were calculated by dividing the emission intensity

at 666nm by the combined intensities at 562nm and 666nm.

Results

Identification of four AmrZ binding sites in PalgD

A previous study identified one AmrZ binding site centered at -289 bp relative to the transcription start site and its importance for PalgD activation (P. J. Baynham &

Wozniak, 1996). In addition, we utilized ChIP-Seq to uncover AmrZ binding sites throughout the P. aeruginosa genome and discovered a second AmrZ binding region in

PalgD (Figure 4.1B; (C. J. Jones et al., 2014)). Two ZBSs were therefore anticipated in

this promoter based on sequence similarity to the consensus. They are designated as ZBS2

(-289bp relative to the transcription start site, the one identified previously) and ZBS3 (-

130bp) respectively (Figure 4.1A). This nomenclature will be explained later in this

section. EMSAs (electrophoretic mobility shift assays) were then used to verify AmrZ

binding to ZBS3. Three [6FAM]-labeled DNA fragments were amplified such that they

contain ZBS2, ZBS3, or neither ZBS, respectively. AmrZ bound to each fragment that

contains a ZBS but showed no binding to the negative control DNA (Figure 4.1C). The

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DNA binding-deficient variant AmrZR22A was also included in this assay and did not

exhibit binding to any DNA fragment (Figure 4.1C). Overall, these results demonstrate that

AmrZ specifically recognizes both AmrZ binding sites within PalgD.

We next sought to understand the significance of both ZBSs during PalgD

activation. One or both ZBSs were disrupted by randomly scrambling their nucleotide

sequences and their effects on AmrZ binding and PalgD activation were determined.

EMSA studies showed that disruption of both ZBSs impaired, but did not eliminate, AmrZ

binding to the DNA (Figure 4.2A). To study their effects on algD transcription in vivo, four algD-lacZ transcriptional fusions were constructed, including PalgD wild type (WT) and

PalgD with scrambled ZBS2, ZBS3, or both, respectively. These fusions were integrated into the chromosome of the parental FRD1 or its isogenic ΔamrZ mutant. In FRD1, ZBS2 scrambling caused dramatic reduction of PalgD activity (57%) compared to WT while the loss of intact ZBS3 had a modest effect (11% reduction). Mutations of both ZBSs resulted in further reduction (74%). However, the PalgD activity was completely eliminated in the

FRD1 ΔamrZ strain (Figure 4.2B), validating that amrZ is absolutely required for algD transcription activation (P. J. Baynham & Wozniak, 1996). Since disrupting the AmrZ binding to both ZBSs did not recapitulate the results seen in the ΔamrZ strain, we explore additional mechanisms of regulation.

We propose two possible explanations for different outcomes of scrambling both

ZBSs and amrZ deletion. 1.) AmrZ activates PalgD indirectly via interacting with or regulating expression of other proteins that are required for PalgD activation. 2.) AmrZ still binds to PalgD despite the scrambling of both ZBSs. The first scenario is unlikely

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since neither RNA-Seq nor DNA microarrays revealed an AmrZ-dependent alginate

regulator (C. J. Jones et al., 2014; Xu & Wozniak, 2015). In addition, co-

immunoprecipitation experiments did not demonstrate AmrZ binding to other TFs (C. J.

Jones and S. W. Harshman, unpublished data). Secondly, observed partial AmrZ binding

could be conferred by ineffective scrambling or additional ZBS(s) in the DNA fragments.

To test if residual AmrZ binding is due to ineffective scrambling, we determined AmrZ

binding to a ZBS2-containing DNA fragment after ZBS2 deletion or replacement by a

sequence known not to bind to AmrZ (Ramsey et al., 2005). Surprisingly, AmrZ still bound

to this fragment after either deletion or replacement (Figure 4.3), suggesting the presence

of another AmrZ binding site. ChIP-Seq analyses indicate additional ZBS(s) is not likely

to be outside of the two regions (Figure 4.1B). Therefore, we concluded that within each

AmrZ binding region (~100bp) there is more than one ZBS. In consideration of the length

of a ZBS (~18bp) and the sequence similarity constraint to the consensus (C. J. Jones et

al., 2014; Pryor et al., 2012), we proposed two additional ZBSs (ZBS1 and ZBS4) within each region, that is total four ZBSs in PalgD. This hypothesis was tested via EMSA assays

(Figure 4.4). The DNA fragment containing only ZBS1 or ZBS4 still retained AmrZ binding while no binding was seen with any ZBS-free DNA fragment (Figure 4.4B).

Therefore, our DNA binding studies demonstrate four AmrZ binding sites within PalgD.

Proposed positions and sequences of these four ZBSs are shown in Figure 4.5, with ZBS1

and ZBS4 located at 321bp and 74bp upstream of the transcription start site, respectively.

In order to evaluate the significance of these ZBSs during algD transcription, we

scrambled DNA sequences of ZBSs and measured effects on AmrZ binding and PalgD

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activation. Since effects of losing ZBS2 and ZBS3 have been determined (Figure 4.2), we

scrambled ZBSs in pairs (ZBS1/2 and ZBS3/4) (Figure 4.6A). Scrambling of ZBS1/2 or

ZBS3/4 eliminated AmrZ binding of their respective DNA fragment (Figure 4.6B).

Importantly, the PalgD activity suffered dramatic reduction from disruption of either ZBS

pair while losing all four had a similar effect as an amrZ deletion (Figure 4.6C). Taken

together, above results validate the significance of four ZBSs during PalgD activation.

Correct phasing between ZBS2 and ZBS3 is critical during PalgD activation

Since all four ZBSs are necessary for PalgD activation and AmrZ forms oligomers,

we investigated whether AmrZ oligomers bound to these ZBSs form higher order

complexes and thus induce PalgD DNA looping. If true, the relative phasing between ZBSs would be critical. We therefore inserted 5 bp between ZBS2 and ZBS3 (Figure 4.7A), cloned this promoter variant into the lacZ transcriptional fusion and integrated it into the

FRD1 chromosome. The algD promoter activity reduced significantly (70%) with this insertion (Figure 4.7B). This reduction was not due to the insertion event itself, since a 10 bp insertion, which restores DNA phasing, did not affect PalgD activity (Figure 4.7B). We therefore conclude that proper DNA phasing between these ZBSs is crucial for PalgD activation.

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AmrZ binding induces trans DNA-AmrZ complex formation

The DNA phase-dependent activation of PalgD suggests that transcription activation of this promoter requires contacts between upstream and downstream factors.

These interactions are likely to occur via looping out of the intervening DNA. We therefore hypothesize that AmrZ induces DNA looping when binding to four ZBSs. If true, the interaction between AmrZ oligomers may result in proximity between ZBSs. Fluorescence resonance energy transfer (FRET) is widely used to measure proximity (<10 nm) during protein-protein and protein-DNA interactions and was therefore used to test this hypothesis. The DNA fragment used for FRET experiments was amplified from PalgD

with ZBS1 and ZBS4 at two ends respectively and underwent end-labeling by two

fluorophores (Cy3 and Cy5) (Figure 4.8A). The FRET efficiency peaked at 1µM AmrZ

while an excessive amount of AmrZ inhibited FRET (Figure 4.8B), which is intriguing and

will be addressed further in Discussion. Successful FRET depends on the AmrZ DNA

binding ability, as the DNA binding deficient variant AmrZR22A failed to induce FRET

(Figure 4.8D). These FRET observations indicate that AmrZ-DNA interactions result in

proximity between Cy3 and Cy5 fluorophores, suggesting complex formation between

AmrZ and DNA. These complexes can be formed in cis (within same DNA molecule) or

in trans (between two separate DNA molecules). The cis complex is formed when AmrZ

binding induces DNA bending/looping and in this case ZBSs from two ends of the same

DNA molecule are very close (<10 nm). On the other hand, the formation of trans

complexes suggests proximity between two DNA molecules when they are bound and held

together by AmrZ. To differentiate between these two possibilities, we constructed a DNA

fragment with a 105bp (10 DNA turns) truncation (-250bp to -146bp) between ZBS2 and

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ZBS3 (Figure 4.8C), in which the intervening DNA is shortened to 47bp. Considering the

steric effects of AmrZ oligomers on the DNA, the length of flexible DNA is expected to

be even shorter, which would be problematic for DNA looping due to high bending energy

(Allemand, Cocco, Douarche, & Lia, 2006). Moreover, the distance between two operator

sites is well over a hundred base pairs in most DNA looping examples (Allemand et al.,

2006; Cournac & Plumbridge, 2013). Therefore, the intervening DNA is likely to be too

short, if not impossible, to bend, preventing cis complex formation. This truncation

however, does not affect the formation of trans complexes. No reduction of the FRET

efficiency was observed after this truncation (Figure 4.8D), suggesting that AmrZ binding to PalgD induces trans DNA-AmrZ complex formation.

Discussion

Alginate production in P. aeruginosa is under tight control by a number of regulators. In this study, we identified four ZBSs in PalgD and showed that each is necessary for transcription activation. Our results also highlight the significance of proper phasing between ZBSs and reveal the formation of trans DNA-AmrZ complexes.

Alginate is a key virulence factor responsible for P. aeruginosa persistence in CF lung infections. Since activation of alginate production is frequently observed during late stages of CF infections when the gain of fitness exceeds the cost of alginate production, it provides an excellent example of bacterial patho-adaptability. Alginate production requires at least 13 biosynthesis enzymes, whose expression is under the control by numerous

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regulators (Ramsey et al., 2005). All biosynthesis genes, except algC, reside within the

alginate biosynthesis operon. The promoter of this operon is particularly complex. So far

it is known to contain binding sites for 7 different TFs, and strikingly, many of these sites

are ≥250 bp upstream of the transcription start site (Figure 4.1A). In this study, we

discovered three additional AmrZ binding sites (ZBS1, ZBS3 and ZBS4) within this

promoter. It is interesting that ZBS4 overlaps with IHF1.2 (Figure 4.1A; (D J Wozniak,

1994)). However, since IHF is dispensable for alginate production under our growth condition (Delic-Attree, Toussaint, Froger, Willison, & Vignais, 1996), the observed effect

of scrambling ZBS3/4 is therefore likely solely due to the loss of AmrZ binding (Figure

4.6).

It is common for a promoter to contain two or more binding sites of one TF, and

~75% of negatively and ~25% of positively regulated promoters are bound by a TF at

several locations in E. coli (Neidhardt & Curtiss, 1996). Multiple binding sites have distinct functions in different promoters during transcription regulation. For instance, the arginine

biosynthesis regulator ArgR interacts with the ARG box of its target promoters and this

ARG box contains two ArgR binding sites separated by 3 bp. Removal of one site results

in 100-fold reduction of the ArgR binding affinity to the other one, indicative of cooperative binding between two sites (Tian, Lim, Carey, & Maas, 1992). Based on our

EMSA data, there does not seem to be cooperative binding between ZBS1 and ZBS2 or

ZBS3 and ZBS4 (Figure 4.4).

The presence of multiple binding sites is often required for DNA loop formation, which may be induced by oligomerization of TFs from two or more binding sites. DNA

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looping is commonly seen and plays a central role during transcriptional control in both eukaryotic and prokaryotic organisms (Allemand et al., 2006). DNA looping was first demonstrated in the araBAD system in E. coli (Dunn, Hahn, Ogdent, & Schleif, 1984;

Lobell & Schleif, 1990). The araBAD operon encodes proteins necessary for arabinose catabolism, whose expression is tightly controlled by AraC. AraC functions as an activator/repressor depending on the presence/absence of arabinose. Without arabinose,

AraC dimers bind to two sites --araO2 and a half site of araI (araI1). AraC induces DNA loop formation when binding to these two sites, preventing transcription initiation.

However, arabinose-bound AraC alters conformation and shifts its position from araO2 to araI2, leading to a broken DNA loop and subsequent transcription initiation of this operon

(Lobell & Schleif, 1990).

DNA loop formation requires proper phasing between protein binding sites. Since each DNA turn is approximately 10.5 bp, an insertion/deletion of 5 bp between protein binding sites would place two proteins in an opposite orientation, reducing protein-protein interactions and subsequent DNA looping. This phase requirement of DNA looping has been employed to support DNA loop formation induced by AraC and the lac repressor

(Dunn et al., 1984; Han et al., 2009; Krämer et al., 1987). In this study, our results demonstrate that a correct phase between ZBS2 and ZBS3 is crucial, suggesting interactions between upstream and downstream factors, likely achieved by DNA looping, are required for PalgD activation.

In order to provide direct evidence for PalgD DNA looping, we designed FRET experiments in which two pairing fluorophores (Cy5 and Cy3) are placed 5bp away from

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ZBS1 and ZBS4, respectively. This 5bp spacing is designed to reduce protein-induced

fluorescence enhancement, which occurs when a protein is very close to a fluorophore

(Hwang & Myong, 2014). When incubated with this DNA fragment, AmrZ, but not a DNA

binding-deficient variant AmrZR22A, induces energy transfer from Cy3 to Cy5, indicating

proximity (<10nm) between these two fluorophores. As this energy transfer occurs

between fluorophores from two DNA molecules (Figure 4.8), we speculate that each trans

complex is composed of two AmrZ tetramers and two DNA fragments. These interactions

are likely to be stable, as the FRET efficiency did not fluctuate over time (up to 20min)

(data not shown). Additionally, a temperature increase from 25°C to 37°C did not affect

FRET efficiency (data not shown), suggesting that these interactions are likely to be similar

during human infections. The FRET efficiency is calculated as the proportion of the Cy3

energy transferred to Cy5 and is the average of the whole solution. Since other conformations of trans DNA-AmrZ complexes, such as Cy3 DNA-AmrZ-Cy3 DNA, are present with the same probability but do not elicit FRET, actual FRET of Cy3 DNA-AmrZ-

Cy5 DNA is expected to more efficient, indicating even a shorter distance between ends of

two DNA molecules. It is intriguing that excessive AmrZ inhibits FRET (Figure 4.8D), which might be explained by oligomerization of all DNA-bound AmrZ with a saturated amount of free AmrZ in solution. Although we are unable to provide a direct evidence of

AmrZ-induced DNA looping, this study reveals that AmrZ holds multiple ZBSs together in vitro, which may be critical for PalgD DNA loop formation in vivo. Additional factors, such as the DNA bending protein IHF and the nucleotide-associated protein Hp-1, may

contribute to looping of this promoter in the cell. This is corroborated by the in vivo

observation that removing the 105bp region used in the truncation experiment (Figure

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4.8C&D) resulted in abolished PalgD transcription (Figure 4.9). Further investigations are

required to validate this theory.

With data obtained here and from previous studies, we propose a model of PalgD

activation. A DNA loop forms when PalgD is bound by AmrZ, IHF, and Hp-1. Formation

of this DNA loop brings remote elements closer to the transcription start site and allows

interactions between remote proteins and RNA polymerase. Specifically, two remote AlgR

dimers interact with the 3rd AlgR dimer that is bound to a proximal site, which further optimizes the DNA conformation and allows direct contacts by Vfr and the NtrC homolog

AlgB to RNA polymerase. Both proper DNA conformation and interactions with RNA polymerase stabilize promoter binding by the polymerase and transcription is then initiated.

Of note, this complex activation mechanism allows the integration of different activation signals during activation and the loss of any activating signal often prevents transcription initiation. This is important for bacteria as alginate production is an energy-exhausting process. In this model, a high intracellular AmrZ level is necessary for PalgD activation because DNA looping requires all four ZBSs bound by AmrZ. Therefore, the intracellular

AmrZ concentration may serve as an additional means of regulation, and it was previously shown that mucoid P. aeruginosa produces significantly more AmrZ than non-mucoid strains (C. J. Jones et al., 2013).

Overall, in this study we have identified four AmrZ binding sites in PalgD and provided support for the DNA looping model of this complex promoter. Further understanding of this promoter may allow exploitation of this activation mechanism and

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development of novel means to prevent or disrupt alginate production during P. aeruginosa

infections.

Acknowledgement

Public Health Service awards AI097511 and NR013898 (DJW), an NSF instrument development grant DBI0923551 (VHW) and funds provided from the Public Health

Preparedness for Infectious Diseases (PHPID) program (phpid.osu.edu) supported this work. BX was partially supported by the student trainingship grant from the cystic fibrosis foundation (XU13H0).

We thank Dr. Thomas Hollis for kindly providing purified AmrZ and AmrZR22A, and Dr. Joe J. Harrison for providing the E. coli S17λpir strain. We are grateful to Sheri

Dellos-Nolan and Dr. Meenu Mishra for critical reading of this manuscript as well as Dr.

Gayan Senavirathne for valuable scientific discussions. We also acknowledge Dr.

Nrusingh Mohapatra for assistance with the β-galactosidase assay.

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Figure 4.1. Identification of a second AmrZ binding site in PalgD.

(A.) Schematic of PalgD and transcription factor binding sites. Note: the AlgB binding site

(~-274bp to -224bp), the CysB binding site (~+90 to +116) and two Hp-1 (AlgP) binding sites (~-432 to -332, ~-116 to +111) are not shown here due to their unprecise locations

(Michael J Schurr, 2013). Designed algD fragments used in (C.) are also shown here.

Numbers indicate the distance relative to the transcription start site. (B.) An example image of ChIP-Seq shows two AmrZ binding peaks in PalgD. “Before IP” and “After IP”

represent DNA samples before and after chromatin immuno-precipitation (IP) with AmrZ

(C. J. Jones et al., 2014). (C.) EMSA experiments were performed to determine AmrZ

binding to above four DNA fragments. “-” indicates free DNA. “+” means the addition of

50 nM AmrZ or a DNA-binding deficient variant AmrZR22A. Arrows point to retardation

of DNA mobility.

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Figure 4.2. Scrambling of ZBS2 and ZBS3 results in reduced DNA binding and impaired PalgD activation.

(A.) Sequences of ZBS2 and ZBS3 were randomly scrambled and resulting affinities AmrZ were measured by EMSA. “-” indicates no protein, while “+” lanes each contained 40 nM

AmrZ. Arrows point to retardation of DNA mobility. “F”: Free DNA. (B.) Scrambling of

ZBS2 and ZBS3 led to impaired PalgD activities. “No” represents promoterless lacZ as the negative control, and “WT” represents wild type PalgD. “ZBS2”, “ZBS3” and “ZBS23” refer to corresponding AmrZ binding sites which were randomly scrambled in the algD- lacZ transcriptional fusion constructs. Y axis is the relative PalgD activity as the percentage

of WT. ***: P<0.001 using unpaired two-tailed student t-test analyses of three independent

experiments.

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Figure 4.3. Deletion of ZBS2 or replacement by an algB sequence does not eliminate

AmrZ binding.

(A.) Wild type ZBS2 sequence and the selected algB sequence which has the same GC%.

(B.) EMSA assays to determine AmrZ binding to DNA fragments containing mutated

ZBS2. “F”: Free DNA. “+” represents the addition of 40nM AmrZ. Arrows point to the shifts of DNA on the gel.

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Figure 4.4. Identification of two additional ZBSs in PalgD.

(A.) Proposal of two additional ZBSs (ZBS1 and ZBS4) and design of DNA fragments used for EMSA assays in (B.). The control DNA fragment “4” was used as the negative control. (B.) EMSA assays to determine the presence of two additional ZBSs. Arrows point to retardation of DNA mobility. Addition of 50 nM AmrZ was labeled as “+”. “F”: Free

DNA.

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Figure 4.5. Proposed positions and nucleotide sequences of four ZBSs.

The PalgD DNA sequence was examined, and positions (B.) and nucleotide sequences (C.) of four ZBSs were proposed based on EMSA results and their sequence similarities to the

AmrZ binding consensus sequence (A.). Numbers in (B.) represent the distance from the transcription start site. (A.) is adapted from (C. J. Jones et al., 2014).

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Figure 4.6. Scrambling of all ZBSs results in abolished PalgD activation.

(A.) ZBSs were randomly scrambled in pairs as shown in grey boxes. Design of DNA fragments used for EMSA experiments (B.) is also shown here. (B.) AmrZ binding affinities of WT or scrambled DNA fragments were determined by EMSA. Each DNA was tested for its binding affinity to 50nM AmrZ. Arrows represent DNA shifts caused by

AmrZ binding. “F”: Free DNA. (C.) PalgD activities were determined after scrambling two or four ZBSs. Y axis is the relative PalgD activity as percentage compared to WT.

***: P<0.001 using unpaired two-tailed student t-test analyses of three independent experiments.

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Figure 4.7. Correct DNA phasing between ZBSs is critical for PalgD activation.

(A.) The lightning symbol indicates the insertion location and insertion sizes (5bp or 10bp).

(B.) After a 5bp or 10bp insertion, the PalgD activity was measured. “No” is the lacZ fusion without a promoter. ***: P<0.001; ns: not significant. Unpaired, two-tailed student t-tests

were performed based on data from three independent experiments.

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Figure 4.8. AmrZ binding results in formation of trans DNA-AmrZ complexes.

(A.) & (C.) PalgD DNA fragments designed for FRET experiments contains Cy5 and Cy3

fluorophores on each end. (A.) WT PalgD. (C.) PalgD with 105bp truncation (-250bp to -

146bp). (B.) Effects of different AmrZ concentrations on observed FRET when the WT

DNA fragment (A.) was tested. (D.) AmrZR22A failed to induce FRET, and AmrZ induced a comparable FRET level of the DNA with 105bp truncation between ZBS2 and ZBS3.

1µM AmrZ or AmrZR22A was used here. “ns”: not significant; ***: P<0.001. Unpaired, two-tailed student t-tests were performed from three independent experiments.

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*** 100

50 %WT ACTIVITY

No WT ∆105bp

Figure 4.9. The 105bp truncation between ZBS2 and ZBS3 disrupts PalgD activation.

“No” is the lacZ fusion without a promoter. ***: P<0.001. Unpaired, two-tailed student t- tests were performed based on data from three independent experiments.

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Table 4.1. Strains, plasmids, and oligonucleotides used in Chapter 4

Strains/Plasmids Relevant genotype/description/sequence Sources Strains Escherichia coli NEB5α fhuA2 Δ(argF-lacZ)U169 phoA glnV44 Φ80 Δ(lacZ)M15 New gyrA96 recA1 relA1 endA1 thi-1 hsdR17 England Biolabs S17λpir pro thi hsdR+ TpR SmR; chromosome::RP4-2 Tc::Mu- (Simon et Kan::Tn7/λpir al., 1983) P. aeruginosa FRD1 mucA22 (Ohman & Chakrabarty , 1981) FRD1200 amrZ::xylE-aacC1 in FRD1 (Wozniak et al., 2003) OSUPA211 algD-lacZ transcriptional fusion at the attB site in FRD1 This study (FRD1/PalgD- lacZ) OSUPA213 Promoterless lacZ at the attB site in FRD1 This study OSUPA226 amrZ deletion; WT algD-lacZ transcriptional fusion at This study the attB site in FRD1 OSUPA260 ZBS2-scrambled algD-lacZ transcriptional fusion at the This study attB site in FRD1 OSUPA261 ZBS3-scrambled algD-lacZ transcriptional fusion at the This study attB site in FRD1 OSUPA263 ZBS2/3-scrambled algD-lacZ transcriptional fusion at This study the attB site in FRD1 OSUPA283 algD-lacZ transcriptional fusion with a 5bp insertion This study between ZBS2 and ZBS3 in FRD1 OSUPA285 algD-lacZ transcriptional fusion with a 10bp insertion This study between ZBS2 and ZBS3 in FRD1 OSUPA32 algD-lacZ transcriptional fusion with a 105bp insertion between ZBS2 and ZBS3 in FRD1 Plasmids mini-CTX-lacZ Vector for transcriptional fusion analyses. TetR (Hoang TT Becher A, Schweizer HP., 2000) pBX1 mini-CTX-PalgD-lacZ. Two primers (algD65, algD66) This study were used to amplify the PalgD pBX16 mini-CTX-PalgD-lacZ with the ZBS2 This study (CCATTGGCCATTACCAGCCTCCC) replaced by a same-length and same-GC% sequence from the algB CDS (GCCTCGAAGACGAAGGCTACAGC). Generated using four primers: algD65, 66, 85 and 86

Continued

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Table 4.1. Continued pBX17 mini-CTX-PalgD-lacZ with a ZBS2-containing sequence This study (CCATTGGCCATTACCAGCCTCCC) deleted. Generated using four primers: algD65, 66, 87 and 88 pBX34 mini-CTX-PalgD-lacZ with the ZBS2 This study (CCATTGGCCATTACCAGCCTCCC) randomly scrambled to (CCGATTTCCTCCCTAGACCCCGA). Generated using four primers: algD65, 66, 104 and 105 pBX35 mini-CTX-PalgD-lacZ with the ZBS3 This study (CTAGTGGCCATTGGCAGGCA) randomly scrambled to (AGAGATCCGCTCGTTGACGG). Generated using four primers: algD65, 66, 106 and 107 pBX37 mini-CTX-PalgD-lacZ with both ZBS2 and ZBS3 This study scrambled pBX46 mini-CTX-PalgD-lacZ with a 5bp insertion (CAGTC) This study between ZBS2 and ZBS3. Generated using four primers: algD65, 66, 122 and 123 pBX48 mini-CTX-PalgD-lacZ with a 10bp insertion This study (CAGTCAGTCA) between ZBS2 and ZBS3. Generated using four primers: algD65, 66, 128 and 129 pBX66 mini-CTX-PalgD-lacZ with both ZBS1 and ZBS2 This study scrambled (total 63bp). Generated using four primers: algD65, 66, 144 and 145 pBX67 mini-CTX-PalgD-lacZ with both ZBS3 and ZBS4 This study scrambled (total 67bp). Generated using four primers: algD65, 66, 152 and 153 pBX68 mini-CTX-PalgD-lacZ with all four ZBSs scrambled. This study pBX71 mini-CTX-PalgD-lacZ with a 105bp deletion between This study ZBS2 and ZBS3. Generated using four primers: algD65, 66, 164 and 165 Oligonucleotides algD65 CCCCAAGCTTCTCTTTCGGCACGCCGAC This study algD66 CCGGGATCCCCGACATAGCCCAAACCAAAG This study algD71 [6FAM] ACCGTTCGTCTGCAAGTCATG This study algD73 ACCAACTTGATGGCCTTTCCG This study algD85 GCTGTAGCCTTCGTCTTCGAGGCATTACGCCGGA This study GATGGCATTTC algD86 GCCTCGAAGACGAAGGCTACAGCGCCATTACAT This study GCAAATTACGATTGC algD87 GCAATCGTAATTTGCATGTAATGGCATTACGCCG This study GAGATGGCATTTC algD88 GAAATGCCATCTCCGGCGTAATGCCATTACATGC This study AAATTACGATTGC algD104 TCGGGGTCTAGGGAGGAAATCGGATTACGCCGG This study AGATGGCATTTC algD105 CCGATTTCCTCCCTAGACCCCGAGCCATTACATG This study CAAATTACGATTGC Continued

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Table 4.1. Continued algD106 CCGTCAACGAGCGGATCTCTTTGCAGAATGCAT This study GGCGGCTG algD107 AGAGATCCGCTCGTTGACGGTTTAACGGAAAGG This study CCATCAAGTT algD108 CTTAATCTTCGACCCATGCAC This study algD109 [6FAM] GGAATCCTTAAGGTTTGCTTAAG This study algD110 TCCCGCTCGCGAAATATTTG This study algD111 [6FAM] CAAACGGCCGGAACTTCCCT This study algD112 TAGTTCGGTCCATAGAATTCAAG This study algD122 CCGGAATGAAACAGAgactgAGCCGCTTTTACCGC This study algD123 GCGGTAAAAGCGGCTcatgcTCTGTTTCATTCCGG This study algD128 CCGGAATGAAACAGAtgactgactgAGCCGCTTTTAC This study CGC algD129 GCGGTAAAAGCGGCTcagtcagtcaTCTGTTTCATTCC This study GG algD132 GAGACGGTTTCAAGTTGCTCTG This study algD144 GCGATTCGCGGATGGCCTCTCTATAACTGAGGC This study GAGAT TGGGGAAAAGTCTGTGGGCTG algD145 AGGCCATCCGCGAATCGCCGAGCTACGTCTCAA This study CATTCCACCTATGCAAATTACGATTGCAAAG algD146 ATTACGCCGGAGATGGCATTT This study algD147 TTGAATTGGGGAAAAGTCTG This study algD148 [6FAM] TTTAACGGAAAGGCCATCAAG This study algD152 GATGATAAGTGTGAAACTCCCATTTGCATCTTAT This study ATCCCCTCGGGTTGCAGAATGCATGGCGGCTG algD153 AATGGGAGTTTCACACTTATCATCGTACGCTAGT This study AAGATGAGACGT ATTTCCAAATATTTCGCGAG algD156 [6FAM] TATTTCCAAATATTTCGCGAGC This study algD157 (5’-amino modifier C6) This study TCAAGGCGGAAATGCCATCT algD160 (5’-amino modifier C6) This study ATATCCGTTTGATATCACTTGATAC algD164 TGGCCACTAGTTGCAGAATGTCGTAATTTGCATG This study TAATGG algD165 CCATTACATGCAAATTACGACATTCTGCAACTAG This study TGGCCA algD168 GATACCAACTTGATGGCCTTTC This study CTX1 CCTCGTTCCCAGTTTGTTCC (Tart et al., 2006) attB2 GTCGCCGCCGGCGATGC (Tart et al., 2006) attB4 CGCCCTATAGTGAGTCG (Tart et al., 2006) attB5 CGCCCCAACCTCGCTGG (Tart et al., 2006)

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Chapter 5. Overall discussion and synopsis

In his book “Adaptability: the art of winning in an age of uncertainty”, Max

McKeown, a writer and consultant in strategy and leadership, stated that “all failure is

failure to adapt, all success is successful adaptation”. This quote is widely applicable to other disciplines such as evolution biology. All living organisms have adapted to adjust themselves upon the entrance into a new “home”.

The Gram-negative bacterium Pseudomonas aeruginosa is an excellent example of an organism that can prosper in diverse environments. Prevalent in natural sites such as fresh water and soil, this bacterium is also readily found on/in medical devices and at various hospital sites. To make it worse, P. aeruginosa exhibits more recalcitrance to disinfectants and other antimicrobial treatments compared with most other bacterial species. A large selection of virulence factors produced by this pathogen aids its survival against the host immune defense. Although typically a healthy person is not susceptible to its infections, P. aeruginosa poses a significant threat to hospitalized patients and those with compromised immunity or other complications. Importantly, P. aeruginosa is capable of causing both acute and chronic infections. As discussed in detail in Chapter 1, this

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bacterium adjusts the expression of different virulence factors during the switch between acute and chronic infections. This switch is a rather complicated process requiring multiple factors such as c-di-GMP signaling and the Gac/Rsm system (Coggan & Wolfgang, 2012;

Hengge, 2009). Full understanding of the explicit mechanism during this switch is vital to the ultimate control of P. aeruginosa infections.

P. aeruginosa contains an unusually large genome (6.2 Mbp in the type strain

PAO1), compared to ~4.6 Mbp in Escherichia coli and ~2.0 Mbp in Streptococcus pneumoniae (Blattner et al., 1997; Hoskins et al., 2001). 12% of P. aeruginosa genes participate in expression control and the regulatory network in P. aeruginosa contains 1020 interactions, demonstrating a complex nature of regulation in this organism (Galán-

Vásquez, Luna, & Martínez-Antonio, 2011). Within this regulatory network, the transcription factor AmrZ plays a central role in coordinating the expression of a variety of virulence factors. Previous studies in the Wozniak laboratory have shown that the outcome of AmrZ-mediated regulation can be activation (alginate and twitching motility) or repression (flagella-mediated motility, Psl and c-di-GMP) (P. J. Baynham et al., 2006; P.

J. Baynham & Wozniak, 1996; C. J. Jones et al., 2014, 2013; Tart et al., 2006). We also determined the crystal structure of AmrZ1-66 when bound by DNA (Pryor et al., 2012).

These studies have significantly deepened our understanding of AmrZ-mediated regulation and demonstrated the global role of AmrZ during P. aeruginosa pathogenesis. However, multiple questions need to be addressed. First, the AmrZ-mediated activation mechanism of twitching motility remains elusive. In addition, albeit a number of transcription factor binding sites have been identified in the algD promoter, the activation mechanism of this

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promoter is largely unknown. Furthermore, the solved crystal structure only provides

information of the AmrZ truncation variant AmrZ1-66, and roles of the C-terminal domain during AmrZ regulation require further investigation. This thesis has been aimed at addressing these questions.

It is well known that organisms adjust themselves upon environmental changes via corresponding cellular responses. For instance, P. aeruginosa cells exhibit substantially different gene expression profiles when oxygen is suddenly depleted or between two growth modes (e.g., in static biofilm versus planktonic growth) (Dötsch et al., 2012; Trunk et al., 2010). In recognition of this, we sought to focus specifically on surface-grown bacteria during the study of twitching motility. Distinct activities of pilA transcription under various growth conditions demonstrate the remarkable adaptability of this bacterium

(Figure 2.1). We designed the cellophane-based twitching motility assay to facilitate the harvest of the bacterial subpopulation with active twitching motility. Transcription profiling experiments were then conducted in order to define the AmrZ regulon and identify AmrZ-dependent gene(s) that is required for twitching motility. The AmrZ regulon was also defined in two separate studies yet both focused on planktonic bacteria. These two studies were carried out by DNA microarray (Elizabeth A. Waligora, unpublished data) and by RNA-Seq (C. J. Jones et al., 2014), respectively. The comparison between the current study with two previous reports reveals strikingly little overlap. Two microarray results only share two common targets (gcbA and PA2081) while there are five AmrZ- dependent genes/operons (PA0102, PA3235, PA4033, gcbA and the PA5167 operon) shared by the current microarray study and RNA-Seq. The diguanylate cyclase-encoding

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gene gcbA appeared in all three studies, validating its repression by AmrZ. Many other

AmrZ-dependent processes revealed in the current study however, were not revealed in either previous report with planktonic bacteria. Certain genes identified exclusively in this

thesis encode proteins that are involved in processes such as denitrification as well as

production of rhamnolipid and pyocin. In addition, AmrZ targets (algD and fleQ) did not

show differential expression in the current study, which is anticipated owing to a

significantly lower AmrZ level in PAO1 compared to mucoid strains (C. J. Jones et al.,

2013). Besides growth conditions, other factors such as analysis methods and the strain

background may also contribute to these differences. The AmrZ regulon defined in the

current study contains 112 genes with none known to be required for TM. One possibility

is that these AmrZ-dependent tfp gene(s) are yet identified in previous studies such as

transposon mutant screening (Jacobs et al., 2003). Identification of this gene(s) may require

further understanding of the TFP regulatory circuit. As an alternative approach,

spontaneous suppressor screening was conducted to identify mutations that bypass AmrZ

regulation and thereby restore TM. However, mutations identified by whole genome

sequencing do not appear to have a direct link to the TFP system (Appendix A). We are

therefore unable to define the molecular mechanism of AmrZ-mediated activation of TM.

To decipher this mechanism, other approaches such as a screen using the overexpression

vector library should be given special consideration. Meanwhile, as mentioned in Appendix

A, AmrZ might regulate TM indirectly such as by affecting intracellular cAMP signaling.

Therefore, it will be instrumental to determine the effect of amrZ inactivation on the

intracellular cAMP concentration.

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Findings in Chapter 3 and Chapter 4 have brought novel understandings of how

AmrZ executes regulation. Many members within the Arc protein superfamily such as Mnt

and NikR form tetramers, while others exist exclusively as dimers in solution (Schreiter &

Drennan, 2007). As shown in Chapter 3, we demonstrate that the AmrZ CTD mediates

tetramerization. In addition, our nano-ESI MS results showed that each AmrZ-DNA

complex contains four AmrZ monomers, likely an AmrZ tetramer. Further biochemical

investigations are required to determine which residues in the CTD are involved in

monomer-monomer interactions that are necessary for tetramerization. A good starting

approach would be alanine scanning, which can be used to screen for amino acid

substitutions that negatively impact AmrZ tetramerization. Since the CTD deletion results

in reduced DNA binding by AmrZ, it is interesting to determine how the CTD contributes

to DNA binding. This can be explored by resolving the crystal structure of the full length

AmrZ-DNA complex.

The discovery of AmrZ tetramerization also facilitates our understanding of the

algD promoter activation mechanism. Before the current study, many transcription factor

binding sites have been identified in this complex promoter with half within the far

upstream region (Figure 4.1). Nevertheless, a detailed understanding of the activation

mechanism is lacking. A DNA looping model was proposed with little supporting evidence

(Mohr et al., 1992). In Chapter 4, we identified four AmrZ binding sites within the

promoter, and both DNA phasing and FRET experiments support the DNA looping model.

We speculate that AmrZ oligomerization between four AmrZ binding sites directly participates in looping out of the intervening DNA, allowing remote transcription factors

145

such as AlgR, Vfr and AlgB to contact proximal transcriptional components. Although

DNA looping has been observed in multiple bacterial transcription control systems including the lac repressor, AraC, and NtrC (Krämer et al., 1987; Lobell & Schleif, 1990;

Su, Porter, Kustu, & Echols, 1990), the algD promoter may be one of the most complicated examples in bacteria that involve multiple transcription factors during DNA loop formation. This is a significant advancement in our understanding of transcription activation. If our DNA looping model stands true, the loss of AmrZ tetramerization is anticipated to result in failed DNA looping, abrogating transcription initiation. It is therefore puzzling that the CTD truncation mutant AmrZ1-66 partially activates this promoter when overexpressed (Chapter 3). Nano-ESI MS experiments showed that the majority of AmrZ1-66 exist as dimers or monomers while a small fraction correspond to tetramers (Chapter 3), and AmrZ1-66 binds to DNA as a dimer of dimers (Pryor et al.,

2012). It is therefore important to determine if there is any interaction between two dimer pairs of AmrZ1-66 from two binding sites.

Collectively, here we are able to further enhance our understanding of AmrZ- mediated regulation. It is clear that efficacious regulation relies on a proper level of AmrZ.

Therefore, different AmrZ intracellular concentrations allow AmrZ to achieve its regulation differentially under diverse conditions. A high concentration of AmrZ in a mucoid strain results in activation of alginate production and the repression of the flagella machinery and the exopolysaccharide Psl, while in non-mucoid strains, the AmrZ level is significantly lower so that alginate production is ceased while the expression of flagella and Psl becomes de-repressed (P. J. Baynham & Wozniak, 1996; C. J. Jones et al., 2013;

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Tart et al., 2006). Therefore, AmrZ plays a vital role for P. aeruginosa in choosing an appropriate lifestyle when entering into different environments, such as in the CF airway versus the bloodstream. AmrZ is thereby a key contributor for successful adaptation by this pathogen. As Max McKeown stated, “Success is successful adaptation”, if we can exploit

AmrZ-mediated regulation, we may find a new approach to intervene and create the

“failure to adapt” scenario, making this pathogen more sensitive to antibiotics and other antimicrobials, in order to control P. aeruginosa infections.

147

References

Ahlgren, H. G., Benedetti, A., Landry, J. S., Bernier, J., Matouk, E., Radzioch, D., … Nguyen, D. (2015). Clinical outcomes associated with Staphylococcus aureus and Pseudomonas aeruginosa airway infections in adult cystic fibrosis patients. BMC Pulmonary Medicine, 15(1), 67. Aim, R. A., Hallinan, J. P., Watson, A. A., & Mattick, J. S. (1996). Fimbrial biogenesis genes of Pseudomonas aeruginosa: pilW and pilX increase the similarity of type 4 fimbriae to the GSP protein-secretion systems and pilY1 encodes a gonococcal PilC homologue. Molecular Microbiology, 22(1), 161–173. Ali, M. H., & Imperiali, B. (2005). Protein oligomerization: How and why. Bioorganic and Medicinal Chemistry, 13(17), 5013–5020. Allemand, J. F., Cocco, S., Douarche, N., & Lia, G. (2006). Loops in DNA: An overview of experimental and theoretical approaches. European Physical Journal E, 19(3), 293–302. Alm, R. A., Bodero, A. J., Free, P. D., & Mattick, J. S. (1996). Identification of a novel gene, pilZ, essential for type 4 fimbrial biogenesis in Pseudomonas aeruginosa. Journal of Bacteriology, 178(1), 46–53. Alm, R. A., & Mattick, J. S. (1997). Genes involved in the biogenesis and function of type- 4 fimbriae in Pseudomonas aeruginosa. Gene, 192(1), 89–98. Amiel, E., Lovewell, R. R., O’Toole, G. a, Hogan, D. a, & Berwin, B. (2010). Pseudomonas aeruginosa evasion of phagocytosis is mediated by loss of swimming motility and is independent of flagellum expression. Infection and Immunity, 78(7), 2937–45. Amikam, D., & Galperin, M. Y. (2006). PilZ domain is part of the bacterial c-di-GMP binding protein. Bioinformatics, 22(1), 3–6. Amouyal, M., Mortensen, L., Buc, H., & Hammer, K. (1989). Single and double loop 148

formation when deoR repressor binds to its natural operator sites. Cell, 58, 545– 551. Ansari, A. Z., & Mapp, A. K. (2002). Modular design of artificial transcription factors. Current Opinion in Chemical Biology, 6(6), 765–772. Anyan, M. E., Amiri, A., Harvey, C. W., Tierra, G., Morales-Soto, N., Driscoll, C. M., … Shrout, J. D. (2014). Type IV pili interactions promote intercellular association and moderate swarming of Pseudomonas aeruginosa. Proceedings of the National Academy of Sciences, 201414661. Artimo, P., Jonnalagedda, M., Arnold, K., Baratin, D., Csardi, G., De Castro, E., … Stockinger, H. (2012). ExPASy: SIB bioinformatics resource portal. Nucleic Acids Research, 40(W1), 597–603. Balasubramanian, D., Kumari, H., & Mathee, K. (2014). Pseudomonas aeruginosa AmpR: an acute-chronic switch regulator. Pathogens and Disease, (May 2014), n/a–n/a. Balleza, E., López-Bojorquez, L. N., Martínez-Antonio, A., Resendis-Antonio, O., Lozada-Chávez, I., Balderas-Martínez, Y. I., … Collado-Vides, J. (2009). Regulation by transcription factors in bacteria: beyond description. FEMS Microbiology Reviews, 33(1), 133–151. Banin, E., Brady, K. M., & Greenberg, E. P. (2006). Chelator-induced dispersal and killing of Pseudomonas aeruginosa cells in a biofilm. Applied and Environment Microbiology, 72(3), 2064–2069. Baraquet, C., Murakami, K., Parsek, M. R., & Harwood, C. S. (2012). The FleQ protein from Pseudomonas aeruginosa functions as both a repressor and an activator to control gene expression from the pel operon promoter in response to c-di-GMP. Nucleic Acids Research. Barraud, N., Hassett, D. J., Hwang, S. H., Rice, S. A., Kjelleberg, S., & Webb, J. S. (2006). Involvement of nitric oxide in biofilm dispersal of Pseudomonas aeruginosa. Journal of Bacteriology, 188(21), 7344–7353. Baynham, P., Brown, A. L., Hall, L. L., & Wozniak, D. J. (1999). Pseudomonas aeruginosa AlgZ, a ribbon-helix-helix DNA-binding protein, is essential for alginate synthesis and algD transcriptional activation. Molecular Microbiology, 33(5), 1069–1080. Baynham, P. J., Ramsey, D. M., Gvozdyev, B. V., Cordonnier, E. M., & Wozniak, D. J. (2006). The Pseudomonas aeruginosa ribbon-helix-helix DNA-binding protein AlgZ (AmrZ) controls twitching motility and biogenesis of type IV pili. Journal of

149

Bacteriology, 188(1), 132–140. Baynham, P. J., & Wozniak, D. J. (1996). Identification and characterization of AlgZ, an AlgT-dependent DNA-binding protein required for Pseudomonas aeruginosa algD transcription. Molecular Microbiology, 22(1), 97–108. Beatson, S. A., Whitchurch, C. B., Annalese, B. T., Mattick, J. S., & Semmler, A. B. T. (2002). Quorum sensing is not required for twitching motility in Pseudomonas aeruginosa, 184(13), 3598–3604. Beatson, S. A., Whitchurch, C. B., Jennifer, L., Levesque, R. C., Mattick, J. S., & Sargent, J. L. (2002). Differential regulation of twitching motility and elastase production by Vfr in Pseudomonas aeruginosa. Journal of Bacteriology, 184(13), 3605–3613. Becher, A., & Schweizer, H. P. (2000). Integration-proficient Pseudomonas aeruginosa vectors for isolation of single-copy chromosomal lacZ and lux gene fusions. BioTechniques, 29, 948–952. Beck, L. L., Smith, T. G., & Hoover, T. R. (2007). Look, no hands! Unconventional transcriptional activators in bacteria. Trends in Microbiology, 15(12), 530–537. Belete, B., Lu, H., & Wozniak, D. (2008). Pseudomonas aeruginosa AlgR regulates type IV pilus biosynthesis by activating transcription of the fimU-pilVWXY1Y2E operon. Journal of Bacteriology, 190(6), 2023–2030. Bell, C. E., Frescura, P., Hochschild, A., & Lewis, M. (2000). Crystal structure of the λ repressor C-terminal domain provides a model for cooperative operator binding. Cell, 101(7), 801–811. Berek, D. (2010). Size exclusion chromatography--a blessing and a curse of science and technology of synthetic polymers. Journal of Separation Science, 33(3), 315–335. Bertrand, J. J., West, J. T., & Engel, J. N. (2010). Genetic analysis of the regulation of type IV pilus function by the Chp chemosensory system of Pseudomonas aeruginosa. Journal of Bacteriology, 192(4), 994–1010. Betz, J. L., & Fall, M. Z. (1988). Effects of dominant-negative lac repressor mutations on operator specificity and protein stability. Gene, 67(2), 147–158. Bjarnsholt, T., Jensen, P. Ø., Burmølle, M., Hentzer, M., Haagensen, J. A. J., Hougen, H. P., … Givskov, M. (2005). Pseudomonas aeruginosa tolerance to tobramycin, hydrogen peroxide and polymorphonuclear leukocytes is quorum-sensing dependent. Microbiology, 151(2), 373–383.

150

Blair, R., Goodrich, J., & Kugel, J. (2013). Using FRET to Monitor Protein-Induced DNA Bending: The TBP-TATA Complex as a Model System. In M. Bina (Ed.), Gene Regulation SE - 16 (Vol. 977, pp. 203–215). Humana Press. Blattner, F. R., Plunkett, G. 3rd, Bloch, C. A., Perna, N. T., Burland, V., Riley, M., … Shao, Y. (1997). The complete genome sequence of Escherichia coli K-12. Science (New York, N.Y.), 277(5331), 1453–1462. Bobadilla Fazzini, R. A., Skindersoe, M. E., Bielecki, P., Puchałka, J., Givskov, M., & Martins dos Santos, V. A. P. (2013). Protoanemonin: a natural quorum sensing inhibitor that selectively activates iron starvation response. Environmental Microbiology, 15(1), 111–120. Bogan, A. A., & Thorn, K. S. (1998). Anatomy of hot spots in protein interfaces. Journal of Molecular Biology, 280, 1–9. Boles, B. R., Thoendel, M., & Singh, P. K. (2005). Rhamnolipids mediate detachment of Pseudomonas aeruginosa from biofilms. Molecular Microbiology, 57(5), 1210–23. Borlee, B. R., Goldman, A. D., Murakami, K., Samudrala, R., Wozniak, D. J., & Parsek, M. R. (2010). Pseudomonas aeruginosa uses a cyclic-di-GMP-regulated adhesin to reinforce the biofilm extracellular matrix. Molecular Microbiology, 75(4), 827– 842. Borukhov, S., & Nudler, E. (2008). RNA polymerase: the vehicle of transcription. Trends in Microbiology, 16(3), 126–134. Boucher, J. C., Schurr, M. J., & Deretic, V. (2000). Dual regulation of mucoidy in Pseudomonas aeruginosa and sigma factor antagonism. Molecular Microbiology, 36(2), 341–51. Breitwieser, G. E. (2004). G protein-coupled receptor oligomerization: implications for G protein activation and cell signaling. Circulation Research, 94(1), 17–27. Brigitte E. Raumann Carl O. Pabo & Robert T. Sauer, M. A. R. (1994). DNA recognition by beta-sheets in the Arc repressor-operator crystal structure. Nature, 367(754 - 757). Broad Institute. (n.d.). Picard. Retrieved from http://broadinstitute.github.io/picard/ Brockson, M. E., Novotny, L. a., Mokrzan, E. M., Malhotra, S., Jurcisek, J. a., Akbar, R., … Bakaletz, L. O. (2014). Evaluation of the kinetics and mechanism of action of anti-integration host factor-mediated disruption of bacterial biofilms. Molecular Microbiology, 93(August), n/a–n/a.

151

Browning, D. F., & Busby, S. J. (2004). The regulation of bacterial transcription initiation. Nature Reviews. Microbiology, 2(1), 57–65. Bullock, B. N., Jochim, A. L., & Arora, P. S. (2011). Assessing helical protein interfaces for inhibitor design. Journal of the American Chemical Society, 133(36), 14220– 14223. Burrows, L. L. (2012). Pseudomonas aeruginosa twitching motility: type IV pili in action. Annual Review of Microbiology, 66, 493–520. Byrd, M. S., Pang, B., Hong, W., Waligora, E. a., Juneau, R. a., Armbruster, C. E., … Swords, W. E. (2011). Direct evaluation of Pseudomonas aeruginosa biofilm mediators in a chronic infection model. Infection and Immunity, 79(8), 3087–3095. Campa, M., Bendinelli, M., & Friedman, H. (1993). Pseudomonas aeruginosa as an opportunistic pathogen. New York: Plenum Press. Chang, W.-S., van de Mortel, M., Nielsen, L., Nino de Guzman, G., Li, X., & Halverson, L. J. (2007). Alginate production by Pseudomonas putida creates a hydrated microenvironment and contributes to biofilm architecture and stress tolerance under water-limiting conditions. Journal of Bacteriology, 189(22), 8290–8299. Chemani, C., Imberty, A., de Bentzmann, S., Pierre, M., Wimmerová, M., Guery, B. P., & Faure, K. (2009). Role of LecA and LecB lectins in Pseudomonas aeruginosa- induced lung injury and effect of carbohydrate ligands. Infection and Immunity, 77(5), 2065–2075. Chew, S. C., Kundukad, B., Seviour, T., van der Maarel, J. R. C., Yang, L., Rice, S. A., … Kjelleberg, S. (2014). Dynamic remodeling of microbial biofilms by functionally distinct exopolysaccharides. mBio, 5(4), e01536–14. Chivers, P. T., & Sauer, R. T. (1999). NikR is a ribbon-helix-helix DNA-binding protein. Protein Science : A Publication of the Protein Society, 8(11), 2494–2500. Chivers, P. T., & Sauer, R. T. (2002). NikR repressor: High-affinity nickel binding to the C-terminal domain regulates binding to operator DNA. Chemistry and Biology, 9(10), 1141–1148. Chua, S. L., Liu, Y., Yam, J. K. H., Chen, Y., Vejborg, R. M., Tan, B. G. C., … Yang, L. (2014). Dispersed cells represent a distinct stage in the transition from bacterial biofilm to planktonic lifestyles. Nature Communications, 5, 4462. Clementi, F. (1997). Alginate production by Azotobacter vinelandii. Critical Reviews in Biotechnology, 17(4), 327–361.

152

Cobb, L. M. L., Mychaleckyj, J. J. C., Wozniak, D. J., & López-Boado, Y. S. (2004). Pseudomonas aeruginosa flagellin and alginate elicit very distinct gene expression patterns in airway epithelial cells: implications for cystic fibrosis disease. The Journal of Immunology, 173(9), 5659–5670. Coburn, B., Wang, P. W., Diaz Caballero, J., Clark, S. T., Brahma, V., Donaldson, S., … Guttman, D. S. (2015). Lung microbiota across age and disease stage in cystic fibrosis. Scientific Reports, 5, 10241. Coggan, K. a., & Wolfgang, M. C. (2012). Global regulatory pathways and cross-talk control Pseudomonas aeruginosa environmental lifestyle and virulence phenotype. Current Issues in Molecular Biology, 14(2), 47–70. Comer, J. E., Marshall, M. A., Blanch, V. J., Deal, C. D., & Castric, P. (2002). Identification of the Pseudomonas aeruginosa 1244 pilin glycosylation site. Infection and Immunity, 70(6), 2837–2845. Comolli, J. C., Hauser, A. R., Waite, L., Whitchurch, C. B., Mattick, J. S., & Engel, J. N. (1999). Pseudomonas aeruginosa gene products PilT and PilU are required for cytotoxicity in vitro and virulence in a mouse model of acute pneumonia. Infect. Immun., 67(7), 3625–3630. Cooper GM. (2000). The cell: a molecular approach. 2nd edition. Retrieved from http://www.ncbi.nlm.nih.gov/books/NBK9850/ Cornelis, P., & Dingemans, J. (2013). Pseudomonas aeruginosa adapts its iron uptake strategies in function of the type of infections. Frontiers in Cellular and Infection Microbiology, 3(November), 75. Cournac, A., & Plumbridge, J. (2013). DNA looping in prokaryotes: experimental and theoretical approaches. Journal of Bacteriology, 195(6), 1109–19. Crick, F. (1970). Cendral dogma of molecular biology. Nature. Cullen, L., & McClean, S. (2015). Bacterial adaptation during chronic respiratory infections. Pathogens, 4(1), 66–89. Cystic Fibrosis Foundation. (2013). Patient registry annual data report. Dasgupta, N., Ferrell, E. P., Kanack, K. J., West, E. H., Ramphal, R., & West, S. E. H. (2002). fleQ, the gene encoding the major flagellar regulator of Pseudomonas aeruginosa, is sigma70 dependent and is downregulated by Vfr, a homolog of Escherichia coli cyclic AMP receptor Protein. Journal of Bacteriology, 184(19), 5240–5250.

153

Dasgupta, N., Wolfgang, M. C., Goodman, A. L., Arora, S. K., Jyot, J., Lory, S., & Ramphal, R. (2003). A four-tiered transcriptional regulatory circuit controls flagellar biogenesis in Pseudomonas aeruginosa. Molecular Microbiology, 50(3), 809–824. Davies, D. G., & Marques, C. N. H. (2009). A fatty acid messenger is responsible for inducing dispersion in microbial biofilms. Journal of Bacteriology, 191(5), 1393– 1403. Davies, D. G., Parsek, M. R., Pearson, J. P., Iglewski, B. H., Costerton, J. W., Greenberg, E. P., & David G. Davies James P. Pearson, Barbara H. Iglewski, J. W. Costerton, E. P. Greenberg, M. R. P. (1998). The involvement of cell-to-cell signals in the development of a bacterial biofilm. Science, 280(1998), 295–298. de Bentzmann, S., Roger, P., Dupuit, F., Bajolet-Laudinat, O., Fuchey, C., Plotkowski, M. C., … Puchelle, E. (1996). Asialo GM1 is a receptor for Pseudomonas aeruginosa adherence to regenerating respiratory epithelial cells. Infection and Immunity, 64(5), 1582–1588. Delic-Attree, I., Toussaint, B., Froger, A., Willison, J. C., & Vignais, P. M. (1996). Isolation of an IHF-deficient mutant of a Pseudomonas aeruginosa mucoid isolate and evaluation of the role of IHF in algD gene expression. Microbiology, 142(10), 2785–2793. Deretic, V., & Konyecsni, W. M. (1990). A procaryotic regulatory factor with a histone H1-like carboxy-terminal domain: clonal variation of repeats within algP, a gene involved in regulation of mucoidy in Pseudomonas aeruginosa. Journal of Bacteriology, 172(10), 5544–54. Devaraj, A., Justice, S. S., Bakaletz, L. O., & Goodman, S. D. (2015). DNABII proteins play a central role in UPEC biofilm structure. Molecular Microbiology, 96(6), 1119–1135. Déziel, E., Lépine, F., Milot, S., & Villemur, R. (2003). rhlA is required for the production of a novel biosurfactant promoting swarming motility in Pseudomonas aeruginosa: 3-(3-hydroxyalkanoyloxy) alkanoic acids (HAAs), the precursors of rhamnolipids. Microbiology, (2003), 2005–2013. Doggett, R. G. (1969). Incidence of mucoid Pseudomonas aeruginosa from clinical sources. Applied Microbiology, 18(5), 936–937. Doig, P., Todd, T., Sastry, P. A., Lee, K. K., Hodges, R. S., Paranchych, W., & Irvin, R. T. (1988). Role of pili in adhesion of Pseudomonas aeruginosa to human 154

respiratory epithelial cells. Infect. Immun., 56(6), 1641–1646. Donlan, R. M., & Costerton, J. W. (2002). Biofilms : survival mechanisms of clinically relevant microorganisms. Clinical Microbiology Reviews, 15(2), 167–193. Döring, G., Conway, S. P., Heijerman, H. G., Hodson, M. E., Høiby, N., Smyth, A., & Touw, D. J. (2000). Antibiotic therapy against Pseudomonas aeruginosa in cystic fibrosis: a European consensus. The European Respiratory Journal, 16(4), 749–67. Dötsch, A., Eckweiler, D., Schniederjans, M., Zimmermann, A., Jensen, V., Scharfe, M., … Häussler, S. (2012). The Pseudomonas aeruginosa transcriptome in planktonic cultures and static Biofilms using RNA sequencing. PLoS ONE, 7(2), e31092. Drake, D., & Montie, T. C. (1988). Flagella, motility and invasive virulence of Pseudomonas aeruginosa. Journal of General Microbiology, 134(1), 43–52. Drake, J. W. (1991). A constant rate of spontaneous mutation in DNA-based microbes. Proceedings of the National Academy of Sciences, 88(16), 7160–7164. Drozdetskiy, A., Cole, C., Procter, J., & Barton, G. J. (2015). JPred4: a protein secondary structure prediction server. Nucleic Acids Research, 1–6. Dunn, T. M., Hahn, S., Ogdent, S., & Schleif, R. F. (1984). An operator at -280 base pairs that is required for repression of araBAD operon promoter : Addition of DNA helical turns between the operator and promoter cyclically hinders repression. Proceedings of the National Academy of Sciences, 81(August), 5017–5020. Eloe-fadrosh, E. A., Brady, A., Crabtree, J., Drabek, E. F., Ma, B., Mahurkar, A., … Fraser, M. (2015). Functional dynamics of the gut microbiome in elderly people during probiotic consumption. mBio, 6(2), 1–12. Fic, E., Bonarek, P., Gorecki, A., Kedracka-Krok, S., Mikolajczak, J., Polit, A., … Wasylewski, Z. (2009). cAMP receptor protein from Escherichia coli as a model of signal transduction in proteins - A review. Journal of Molecular Microbiology and Biotechnology, 17(1), 1–11. Flemming, H. C., Neu, T. R., & Wozniak, D. J. (2007). The EPS matrix: The “House of Biofilm Cells.” Journal of Bacteriology. Folkesson, A., Jelsbak, L., Yang, L., Johansen, H. K., Ciofu, O., Høiby, N., & Molin, S. (2012). Adaptation of Pseudomonas aeruginosa to the cystic fibrosis airway: an evolutionary perspective. Nature Reviews Microbiology, 10(12), 841–51. Friedrich, C., Bulyha, I., & Søgaard-Andersen, L. (2013). Outside-in assembly pathway of

155

the type IV pili system in Myxococcus xanthus. Journal of Bacteriology, 196(2). Fuchs, E. L., Brutinel, E. D., Jones, A. K., Fulcher, B., Urbanowski, M. L., Yahr, T. L., … Fulcher, N. B. (2010). The Pseudomonas aeruginosa Vfr regulator controls global virulence factor expression through cyclic AMP-dependent and -independent mechanisms. Journal of Bacteriology, 192(14), 3553–3564. Fulcher, N. B., Holliday, P. M., Klem, E., Cann, M. J., & Wolfgang, M. C. (2010). The Pseudomonas aeruginosa Chp chemosensory system regulates intracellular cAMP levels by modulating adenylate cyclase activity. Molecular Microbiology, 76(4), 889–904. Gacesa, P. (1998). Bacterial alginate biosynthesis--recent progress and future prospects. Microbiology (Reading, England), 144 ( Pt 5(1 998), 1133–43. Galán-Vásquez, E., Luna, B., & Martínez-Antonio, A. (2011). The regulatory network of Pseudomonas aeruginosa. Microbial Informatics and Experimentation, 1(1), 3. Gasteiger, E., Hoogland, C., Gattiker, A., Duvaud, S., Wilkins, M., Appel, R., & Bairoch, A. (2005). Protein identification and analysis tools on the ExPASy server. In J. Walker (Ed.), The Proteomics Protocols Handbook SE - 52 (pp. 571–607). Humana Press. Gautier, L., Cope, L., Bolstad, B. M., & Irizarry, R. A. (2004). affy—analysis of Affymetrix GeneChip data at the probe level. Bioinformatics, 20(3), 307–315. Ge, X., Luo, D., & Xu, J. (2011). Cell-free protein expression under macromolecular crowding conditions. PLoS ONE, 6(12). Gellatly, S. L., & Hancock, R. E. W. (2013). Pseudomonas aeruginosa: new insights into pathogenesis and host defenses. Pathogens and Disease, 67(3), 159–173. Gentleman, R. C., Carey, V. J., Bates, D. M., Bolstad, B., Dettling, M., Dudoit, S., … Zhang, J. (2004). Bioconductor: open software development for computational biology and bioinformatics. Genome Biology, 5(10), R80. Gibiansky, M. L., Conrad, J. C., Jin, F., Gordon, V. D., Motto, D. A., Mathewson, M. a, … Wong, G. C. L. (2010). Bacteria use type IV pili to walk upright and detach from surfaces. Science, 330(6001), 197. Giltner, C. L., Nguyen, Y., & Burrows, L. L. (2012). Type IV pilin proteins: versatile molecular modules. Microbiology and Molecular Biology Reviews, 76(4), 740– 772.

156

Gloag, E. S., Turnbull, L., Huang, A., Vallotton, P., Wang, H., Nolan, L. M., … Whitchurch, C. B. (2013). Self-organization of bacterial biofilms is facilitated by extracellular DNA. Proceedings of the National Academy of Sciences, 110(28), 11541–6. Gomila, M., Peña, A., Mulet, M., Lalucat, J., & García-Valdés, E. (2015). Phylogenomics and systematics in Pseudomonas. Frontiers in Microbiology, 6(March), 1–13. Govan, J. R. W., & Deretic, V. (1996). Microbial pathogenesis in cystic fibrosis: mucoid Pseudomonas aeruginosa and Burkholderia cepacia. Microbiological Reviews, 60(3), 539–574. Gunnelius, L., Hakkila, K., Kurkela, J., Wada, H., Tyystjärvi, E., & Tyystjärvi, T. (2014). The omega subunit of the RNA polymerase core directs transcription efficiency in cyanobacteria. Nucleic Acids Research, 42(7), 4606–4614. Güvener, Z. T., & Harwood, C. S. (2007). Subcellular location characteristics of the Pseudomonas aeruginosa GGDEF protein, WspR, indicate that it produces cyclic- di-GMP in response to growth on surfaces. Molecular Microbiology, 66(November), 1459–1473. Hall, B., McLoughlin, A. J., Leung, K. T., Trevors, J. T., & Lee, H. (1998). Transport and survival of alginate-encapsulated and free lux-lac marked Pseudomonas aeruginosa UG2Lr cells in soil. FEMS Microbiology Ecology, 26(1), 51–61. Hall-Stoodley, L., Costerton, J. W., & Stoodley, P. (2004). Bacterial biofilms: from the natural environment to infectious diseases. Nature Reviews. Microbiology, 2(2), 95–108. Han, L., Garcia, H. G., Blumberg, S., Towles, K. B., Beausang, J. F., Nelson, P. C., & Phillips, R. (2009). Concentration and length dependence of DNA looping in transcriptional regulation. PloS One, 4(5), e5621. Hauser, A. R. (2009a). The type III secretion system of Pseudomonas aeruginosa: infection by injection. Nat Rev Microbiol, 7(9), 654–665. Hauser, A. R. (2009b). The type III secretion system of Pseudomonas aeruginosa: infection by injection. Nat Rev Micro, 7(9), 654–665. Hauser, A. R., & Rello, J. (2003). Severe infections caused by Pseudomonas aeruginosa. Boston: Kluwer Academic Publishers. Heckman, K. L., & Pease, L. R. (2007). Gene splicing and mutagenesis by PCR-driven overlap extension. Nat. Protocols, 2(4), 924–932.

157

Hegge, F. T., Hitchen, P. G., Aas, F. E., Kristiansen, H., Løvold, C., Egge-Jacobsen, W., … Koomey, M. (2004). Unique modifications with phosphocholine and phosphoethanolamine define alternate antigenic forms of Neisseria gonorrhoeae type IV pili. Proceedings of the National Academy of Sciences of the United States of America, 101(29), 10798–10803. Hengge, R. (2009). Principles of c-di-GMP signalling in bacteria. Nat Rev Micro, 7(4), 263–273. Henrichsen, J. (1983). Twitching motility. Annual Review Microbiology, 37(27), 81–93. Hickman, J., & Harwood, C. (2008). Identification of FleQ from Pseudomonas aeruginosa as a c-di-GMP-responsive transcription factor. Molecular Microbiology, 69(May), 376–389. Hickman, J. W., Tifrea, D. F., & Harwood, C. S. (2005). A chemosensory system that regulates biofilm formation through modulation of cyclic diguanylate levels. Proceedings of the National Academy of Sciences of the United States of America, 102(40), 14422–14427. Hoang, T. T., Karkhoff-Schweizer, R. R., Kutchma, A. J., & Schweizer, H. P. (1998). A broad-host-range Flp-FRT recombination system for site-specific excision of chromosomally-located DNA sequences: application for isolation of unmarked Pseudomonas aeruginosa mutants. Gene, 212(1), 77–86. Hoang TT Becher A, Schweizer HP., K. A. J. (2000). Integration-proficient plasmids for Pseudomonas aeruginosa: site-specific integration and use for engineering of reporter and expression strains. Plasmid, 43. Hobbs, M., Collie, E. S. R., Free, P. D., Livingston, S. P., & Mattick, J. S. (1993). PilS and PilR, a two-component transcriptional regulatory system controlling expression of type 4 fimbriae in Pseudomonas aeruginosa. Molecular Microbiology, 7(5), 669– 682. Hoiby, N. (1982). Microbiology of lung infections in cystic fibrosis patients. Acta Paediatrica Scandinavica, 71(Suppl. 301), 33–54. Hoskins, J., Alborn, W. E., Arnold, J., Blaszczak, L. C., Burgett, S., DeHoff, B. S., … States, U. (2001). Genome of the bacterium Streptococcus pneumoniae strain R6. Journal of Bacteriology, 183(19), 5709–17. Hoyne, P. a, Haas, R., Meyer, T. F., Davies, J. K., & Elleman, T. C. (1992). Production of Neisseria gonorrhoeae pili (fimbriae) in Pseudomonas aeruginosa. Journal of

158

Bacteriology, 174(22), 7321–7. Huang, B., Whitchurch, C. B., & Mattick, J. S. (2003). FimX, a multidomain protein connecting environmental signals to twitching motility in Pseudomonas aeruginosa. Journal of Bacteriology, 185(24), 7068–7076. Huhulescu, S., Simon, M., Lubnow, M., Kaase, M., Wewalka, G., Pietzka, A. T., … Allerberger, F. (2011). Fatal Pseudomonas aeruginosa pneumonia in a previously healthy woman was most likely associated with a contaminated hot tub. Infection, 39(3), 265–269. Hwang, H., & Myong, S. (2014). Protein induced fluorescence enhancement (PIFE) for probing protein-nucleic acid interactions. Chemical Society Reviews, 43(4), 1221– 9. Ihaka, R., & Gentleman, R. (1996). R: a language for data analysis and graphics. Journal of Computational and Graphical Statistics, 5(3), 299–314. Inclan, Y. F., Huseby, M. J., & Engel, J. N. (2011). FimL regulates cAMP synthesis in Pseudomonas aeruginosa. PloS One, 6(1), e15867. Ishikawa-Ankerhold, H. C., Ankerhold, R., & Drummen, G. P. C. (2012). Advanced fluorescence microscopy techniques--FRAP, FLIP, FLAP, FRET and FLIM. Molecules (Basel, Switzerland), 17(4), 4047–132. Jacobs, M. A., Alwood, A., Thaipisuttikul, I., Spencer, D., Haugen, E., Ernst, S., … Manoil, C. (2003). Comprehensive transposon mutant library of Pseudomonas aeruginosa. Proceedings of the National Academy of Sciences, 100(24), 14339– 14344. Jain, R., Behrens, A.-J., Kaever, V., & Kazmierczak, B. I. (2012). Type IV pilus assembly in Pseudomonas aeruginosa over a broad range of cyclic di-GMP concentrations. Journal of Bacteriology, 194(16), 4285–4294. Jarrell, K. F., & McBride, M. J. (2008). The surprisingly diverse ways that prokaryotes move. Nature Reviews. Microbiology, 6(6), 466–476. Jenkins, a T. a, Buckling, A., McGhee, M., & Ffrench-Constant, R. H. (2005). Surface plasmon resonance shows that type IV pili are important in surface attachment by Pseudomonas aeruginosa. Journal of the Royal Society, Interface / the Royal Society, 2(3), 255–259. Johnson, M. D. L., Garrett, C. K., Bond, J. E., Coggan, K. A., Wolfgang, M. C., & Redinbo, M. R. (2011). Pseudomonas aeruginosa PilY1 binds integrin in an RGD- and

159

calcium-dependent manner. PLoS ONE, 6(12), e29629. Jones, A. K., Fulcher, N. B., Balzer, G. J., Urbanowski, M. L., Pritchett, C. L., Schurr, M. J., … Wolfgang, M. C. (2010). Activation of the Pseudomonas aeruginosa AlgU regulon through mucA mutation inhibits cyclic AMP/Vfr signaling. Journal of Bacteriology, 192(21), 5709–17. Jones, C. J., Newsom, D., Kelly, B., Irie, Y., Jennings, L. K., Xu, B., … Wozniak, D. J. (2014). ChIP-Seq and RNA-Seq reveal an AmrZ-mediated mechanism for cyclic di-GMP synthesis and biofilm development by Pseudomonas aeruginosa. PLoS Pathogens, 10(3), e1003984. Jones, C. J., Ryder, C. R., Mann, E. E., & Wozniak, D. J. (2013). AmrZ modulates Pseudomonas aeruginosa biofilm architecture by directly repressing transcription of the psl operon. Journal of Bacteriology, 195(8), 1637–1644. Jones, D. T. (1999). Protein secondary structure prediction based on position-specific scoring matrices. Journal of Molecular Biology, 292(2), 195–202. Kai, T., Tateda, K., Kimura, S., Ishii, Y., Ito, H., Yoshida, H., … Yamaguchi, K. (2009). A low concentration of azithromycin inhibits the mRNA expression of N-acyl homoserine lactone synthesis enzymes, upstream of lasI or rhlI, in Pseudomonas aeruginosa. Pulmonary Pharmacology & Therapeutics, 22(6), 483–6. Karas, M., Bahr, U., & Dülcks, T. (2000). Nano-electrospray ionization mass spectrometry: addressing analytical problems beyond routine. Fresenius’ Journal of Analytical Chemistry, 366, 669–676. Kato, J., Misra, T. K., & Chakrabarty, a M. (1990). AlgR3, a protein resembling eukaryotic histone H1, regulates alginate synthesis in Pseudomonas aeruginosa. Proceedings of the National Academy of Sciences of the United States of America, 87(8), 2887– 91. Kazmierczak, B. I., Lebron, M. B., & Murray, T. S. (2006). Analysis of FimX, a phosphodiesterase that governs twitching motility in Pseudomonas aeruginosa. Molecular Microbiology, 60(4), 1026–43. Kearns, D. B., Robinson, J., Lawrence, J., Kearns, D. B., & Robinson, J. (2001). Pseudomonas aeruginosa exhibits directed twitching motility up phosphatidylethanolamine gradients. Journal of Bacteriology, 183(2), 763–767. Kelly, B. J., Fitch, J. R., Hu, Y., Corsmeier, D. J., Zhong, H., Wetzel, A. N., … White, P. (2015). Churchill: an ultra-fast, deterministic, highly scalable and balanced

160

parallelization strategy for the discovery of human genetic variation in clinical and population-scale genomics. Genome Biology, 16(1), 1–14. Klausen, M., Aaes-Jørgensen, A., Molin, S., & Tolker-Nielsen, T. (2003). Involvement of bacterial migration in the development of complex multicellular structures in Pseudomonas aeruginosa biofilms. Molecular Microbiology, 50(1), 61–68. Klausen, M., Heydorn, A., Ragas, P., Lambertsen, L., Aaes-Jørgensen, A., Molin, S., & Tolker-Nielsen, T. (2003). Biofilm formation by Pseudomonas aeruginosa wild type, flagella and type IV pili mutants. Molecular Microbiology, 48(6), 1511–1524. Kluger, R., & Alagic, A. (2004). Chemical cross-linking and protein-protein interactions- a review with illustrative protocols. Bioorganic Chemistry, 32(6), 451–72. Knight, K. L., Bowie, J. U., Vershon, A. K., Kelley, R. D., Sauer, R. T., Vershong, A. K., & Sauer, R. T. (1989). The Arc and Mnt repressors. A new class of sequence- specific DNA-binding protein. Journal of Biological Chemistry, 264(7), 3639– 3642. Kohler, T., Curty, L. K., Barja, F., Van Delden, C., Pechere, J. C., Köhler, T., … Pechère, J.-C. (2000). Swarming of Pseudomonas aeruginosa is dependent on cell-to-cell signaling and requires flagella and pili. Journal of Bacteriology, 182(21), 5990– 5996. Konyecsni, W. M., & Deretic, V. (1990). DNA sequence and expression analysis of algP and algQ, components of the multigene system transcriptionally regulating mucoidy in Pseudomonas aeruginosa: algP contains multiple direct repeats. Journal of Bacteriology, 172(5), 2511–20. Krämer, H., Niemöller, M., Amouyal, M., Revet, B., von Wilcken-Bergmann, B., & Müller-Hill, B. (1987). lac repressor forms loops with linear DNA carrying two suitably spaced lac operators. The EMBO Journal, 6(5), 1481–1491. Kuchma, S. L., Griffin, E. F., & O’Toole, G. A. (2012). Minor pilins of the type IV pilus system participate in the negative regulation of swarming motility. Journal of Bacteriology, 194(19), 5388–5403. Kyte, J., & Doolittle, R. F. (1982). A simple method for displaying the hydropathic character of a protein. Journal of Molecular Biology, 157(1), 105–132. Lau, G. W., Hassett, D. J., Ran, H., & Kong, F. (2004). The role of pyocyanin in Pseudomonas aeruginosa infection. Trends in Molecular Medicine, 10(12), 599– 606.

161

Leech, A. J., Sprinkle, A., Wood, L., Wozniak, D. J., & Ohman, D. E. (2008). The NtrC family regulator AlgB, which controls alginate biosynthesis in mucoid Pseudomonas aeruginosa binds directly to the algD promoter. J. Bacteriol., 190(2), 581–589. Leid, J. G., Willson, C. J., Shirtliff, M. E., Hassett, D. J., Parsek, M. R., & Jeffers, a. K. (2005). The exopolysaccharide alginate protects Pseudomonas aeruginosa biofilm bacteria from IFN-gamma-mediated macrophage killing. The Journal of Immunology, 175(11), 7512–7518. Létoffé, S., Redeker, V., & Wandersman, C. (1998). Isolation and characterization of an extracellular haem-binding protein from Pseudomonas aeruginosa that shares function and sequence similarities with the Serratia marcescens HasA haemophore. Molecular Microbiology, 28(6), 1223–1234. Levine, M., & Tjian, R. (2003). Transcription regulation and animal diversity. Nature, 424(6945), 147–151. Lewis, M., Chang, G., Horton, N. C., Kercher, M. A., Pace, H. C., Schumacher, M. A., … Lu, P. (1996). Crystal structure of the lactose operon repressor and its complexes with DNA and inducer. Science (New York, N.Y.), 271(5253), 1247–1254. Li, H., & Durbin, R. (2009). Fast and accurate short read alignment with Burrows-Wheeler transform. Bioinformatics (Oxford, England), 25(14), 1754–60. Li, W., Cowley, a., Uludag, M., Gur, T., McWilliam, H., Squizzato, S., … Lopez, R. (2015). The EMBL-EBI bioinformatics web and programmatic tools framework. Nucleic Acids Research, 1–5. Lister, P. D., Wolter, D. J., & Hanson, N. D. (2009). Antibacterial-resistant Pseudomonas aeruginosa: Clinical impact and complex regulation of chromosomally encoded resistance mechanisms. Clinical Microbiology Reviews, 22(4), 582–610. Livak, K. J., & Schmittgen, T. D. (2001). Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) method. Methods (San Diego, Calif.), 25, 402–408. Lobell, R. B., & Schleif, R. F. (1990). DNA looping and unlooping by AraC protein. Science (New York, N.Y.), 250(4980), 528–532. Loris, R., Tielker, D., Jaeger, K.-E., & Wyns, L. (2003). Structural basis of carbohydrate recognition by the lectin LecB from Pseudomonas aeruginosa. Journal of Molecular Biology, 331(4), 861–870.

162

Lovewell, R. R., Patankar, Y. R., & Berwin, B. (2014). Mechanisms of phagocytosis and host clearance of Pseudomonas aeruginosa. American Journal of Physiology. Lung Cellular and Molecular Physiology, 306(7), L591–603. Luo, Y., Zhao, K., Baker, A. E., Kuchma, S. L., Coggan, K. a, Wolfgang, M. C., … O’Toole, G. A. (2015). A hierarchical cascade of second messengers regulates Pseudomonas aeruginosa surface behaviors. mBio, 6(1), 1–11. Ma, L., Conover, M., Lu, H., Parsek, M. R., Bayles, K., & Wozniak, D. J. (2009). Assembly and development of the Pseudomonas aeruginosa biofilm matrix. PLoS Pathogens, 5(3). Ma, L., Jackson, K. D., Landry, R. M., Parsek, M. R., & Wozniak, D. J. (2006). Analysis of Pseudomonas aeruginosa conditional psl variants reveals roles for the Psl polysaccharide in adhesion and maintaining biofilm structure postattachment. Journal of Bacteriology, 188(23), 8213–8221. Ma, S., Selvaraj, U., Ohman, D. E., Quarless, R., Hassett, D. J., & Wozniak, D. J. (1998). Phosphorylation-independent activity of the response regulators AlgB and AlgR in promoting alginate biosynthesis in mucoid Pseudomonas aeruginosa. Journal of Bacteriology, 180(4), 956–968. Ma, X., Lai, L. B., Lai, S. M., Tanimoto, A., Foster, M. P., Wysocki, V. H., & Gopalan, V. (2014). Uncovering the stoichiometry of Pyrococcus furiosus RNase P, a multi- subunit catalytic ribonucleoprotein complex, by surface-induced dissociation and ion mobility mass spectrometry. Angewandte Chemie International Edition, 53(43), 11483–11487. Madl, T., Van Melderen, L., Mine, N., Respondek, M., Oberer, M., Keller, W., … Zangger, K. (2006). Structural basis for nucleic acid and toxin recognition of the bacterial antitoxin CcdA. Journal of Molecular Biology, 364(2), 170–185. Mann, E. E., & Wozniak, D. J. (2012). Pseudomonas biofilm matrix composition and niche biology. FEMS Microbiology Reviews, 36(4), 893–916. Marchler-Bauer, A., Derbyshire, M. K., Gonzales, N. R., Lu, S., Chitsaz, F., Geer, L. Y., … Bryant, S. H. (2014). CDD: NCBI’s conserved domain database. Nucleic Acids Research, 43(D1), D222–D226. Mardis, E. R. (2013). Next-generation sequencing platforms. Annual Review of Analytical Chemistry (Palo Alto, Calif.), 6, 287–303. Margulies, M., Egholm, M., Altman, W. E., Attiya, S., Bader, J. S., Bemben, L. a, …

163

Rothberg, J. M. (2005). Genome sequencing in microfabricated high-density picolitre reactors. Nature, 437(7057), 376–380. Markussen, T., Marvig, R. L., Gomez-Lozano, M., Aanaes, K., Burleigh, a. E., Hoiby, N., … Jelsbak, L. (2014). Environmental heterogeneity drives within-host diversification and evolution of Pseudomonas aeruginosa. mBio, 5(5), e01592–14– e01592–14. Martínez-Antonio, A. (2011). Escherichia coli transcriptional regulatory network. Network Biology, 1(1), 21–33. Martínez-Granero, F., Redondo-Nieto, M., Vesga, P., Martín, M., & Rivilla, R. (2014). AmrZ is a global transcriptional regulator implicated in iron uptake and environmental adaption in P. fluorescens F113. BMC Genomics, 15, 237. Marvig, R. L., Khademi, H., Evolution, W., Adaptation, R., Acquisition, I., Feeds, R. S. S., & Journal, a S. M. (2014). Within-host evolution of Pseudomonas aeruginosa reveals adaptation. mBio, 5(3), 1–8. Matsui, H., Sano, Y., Ishihara, H., & Shinomiya, T. (1993). Regulation of pyocin genes in Pseudomonas aeruginosa by positive (prtN) and negative (prtR) regulatory genes. Journal of Bacteriology, 175(5), 1257–1263. Matthews, J., Sunde, M., Gell, D., & Grant, R. (2012). Protein dimerization and oligomerization in biology. Mattick, J. (2002). Type IV pilus and twitching motility. Annu. Rev. Microbiol., 56, 289– 314. Mattick, J. S., Whitchurch, C. B., & Alm, R. a. (1996, November 7). The molecular genetics of type-4 fimbriae in Pseudomonas aeruginosa--a review. Gene. McKenna, A., Hanna, M., Banks, E., Sivachenko, A., Cibulskis, K., Kernytsky, A., … DePristo, M. a. (2010). The Genome Analysis Toolkit: a MapReduce framework for analyzing next-generation DNA sequencing data. Genome Research, 20(9), 1297–303. Melville, S., & Craig, L. (2013). Type IV pili in Gram-positive bacteria. Microbiology and Molecular Biology Reviews : MMBR, 77(3), 323–41. Merritt, J. H., Ha, D.-G., Cowles, K. N., Lu, W., Morales, D. K., Rabinowitz, J., … O’Toole, G. A. (2010). Specific control of Pseudomonas aeruginosa surface- associated behaviors by two c-di-GMP diguanylate cyclases. mBio, 1(4).

164

Migula, W. (1894). Ueber ein neues System der Bakterien. Arb. Bakteriol. Inst. Karlsruhe, 1, 235–238. Miller, J. H. (1992). A short course in bacterial genetics : a laboratory manual and handbook for Escherichia coli and related bacteria. Plainview, N.Y.: Cold Spring Harbor Laboratory Press. Miller, R. M., Tomaras, A. P., Barker, A. P., Voelker, R., Chan, E. D., Vasil, A. I., … Vasil, M. L. (2008). Pseudomonas aeruginosa twitching motility-mediated chemotaxis towards phospholipids and fatty Acids: specificity and metabolic requirements. Journal of Bacteriology, 190(11), 4038–4049. Mittal, R., Sharma, S., Chhibber, S., & Harjai, K. (2006). Contribution of quorum-sensing systems to virulence of Pseudomonas aeruginosa in an experimental pyelonephritis model. Journal of Microbiology, Immunology, and Infection = Wei Mian Yu Gan Ran Za Zhi, 39(4), 302–309. Mohr, C. D., Leveau, J. H. J., Krieg, D. P., Hibler, N. S., Deretic, V., & Mohr CD Krieg DP, Hibler NS, Deretic V., L. J. H. (1992). AlgR-binding sites within the algD promoter make up a set of inverted repeats separated by a large intervening segment of DNA. J. Bacteriol, 174. Mohr, C. D., Leveau, J. H., Krieg, D. P., Hibler, N. S., Deretic, V., Mohr CD Krieg DP, Hibler NS, Deretic V., L. J. H., & N, H. (1991). AlgR, a response regulator controlling mucoidy in Pseudomonas aeruginosa, binds to the FUS sites of the algD promoter located unusually far upstream from the mRNA start site. J Bacteriol, 173. Moncalián, G., & De La Cruz, F. (2004). DNA binding properties of protein TrwA, a possible structural variant of the Arc repressor superfamily. Biochimica et Biophysica Acta - Proteins and Proteomics, 1701(1-2), 15–23. Muller, C., Bahlawane, C., Aubert, S., Delay, C. M., Schauer, K., Michaud-Soret, I., & De Reuse, H. (2011). Hierarchical regulation of the NikR-mediated nickel response in Helicobacter pylori. Nucleic Acids Research, 39(17), 7564–7575. Murray, T. S., & Kazmierczak, B. I. (2006). FlhF is required for swimming and swarming in Pseudomonas aeruginosa. Journal of Bacteriology, 188(19), 6995–7004. Murray, T. S., & Kazmierczak, B. I. (2008). Pseudomonas aeruginosa exhibits sliding motility in the absence of type IV pili and flagella. Journal of Bacteriology, 190(8), 2700–2708.

165

Neidhardt, F. C., & Curtiss, R. (1996). Escherichia coli and Salmonella: cellular and molecular biology. Washington, D.C.: ASM Press. Nguyen, Y., Sugiman-Marangos, S., Harvey, H., Bell, S. D., Charlton, C. L., Junop, M. S., & Burrows, L. L. (2015). Pseudomonas aeruginosa minor pilins prime type IVa pilus assembly and promote surface display of the PilY1 adhesin. Journal of Biological Chemistry, 290(1), 601–611. Noble, W. C., & White, P. M. (1969). Pseudomonads and man. Transactions of the St. John’s Hostpital Dermatological Society, 55(2), 202–208. Nolan, L. M., Beatson, S. a, Croft, L., Jones, P. M., George, A. M., Mattick, J. S., … Whitchurch, C. B. (2012). Extragenic suppressor mutations that restore twitching motility to fimL mutants of Pseudomonas aeruginosa are associated with elevated intracellular cyclic AMP levels. MicrobiologyOpen, 1(4), 490–501. O’Connor, J. R., Kuwada, N. J., Huangyutitham, V., Wiggins, P. A., & Harwood, C. S. (2012). Surface sensing and lateral subcellular localization of WspA, the receptor in a chemosensory-like system leading to c-di-GMP production. Molecular Microbiology, 86(September), 720–729. O’Sullivan, D. J., & O’Gara, F. (1992). Traits of fluorescent Pseudomonas spp. involved in suppression of plant root pathogens. Microbiological Reviews, 56(4), 662–676. O’Toole, G. A., Gibbs, K. A., Hager, P. W., Phibbs, P. V, & Kolter, R. (2000). The global carbon metabolism regulator Crc is a component of a signal transduction pathway required for biofilm development by Pseudomonas aeruginosa. Journal of Bacteriology, 182(2), 425–431. O’Toole, G. A., & Kolter, R. (1998). Flagellar and twitching motility are necessary for Pseudomonas aeruginosa biofilm development. Molecular Microbiology, 30(2), 295–304. Ohman, D. E., & Chakrabarty, A. M. (1981). Genetic mapping of chromosomal determinants for the production of the exopolysaccharide alginate in a Pseudomonas aeruginosa cystic fibrosis isolate. Infection and Immunity, 33(1), 142–148. Okkotsu, Y., Tieku, P., Fitzsimmons, L. F., Churchill, M. E., & Schurr, M. J. (2013). Pseudomonas aeruginosa AlgR phosphorylation modulates rhamnolipid production and motility. Journal of Bacteriology, 195(24), 5499–515. Orans, J., Johnson, M. D. L., Coggan, K. a, Sperlazza, J. R., Heiniger, R. W., Wolfgang,

166

M. C., & Redinbo, M. R. (2010). Crystal structure analysis reveals Pseudomonas PilY1 as an essential calcium-dependent regulator of bacterial surface motility. Proceedings of the National Academy of Sciences of the United States of America, 107(3), 1065–70. Pabo, C. O., & Sauer, R. T. (1992). Transcription factors: structural families and principles of DNA recognition. Annual Review of Biochemistry, 61, 1053–1095. Paudel, B. P., & Rueda, D. (2014). Molecular crowding accelerates ribozyme docking and catalysis. Journal of the American Chemical Society, 136, 16700–16703. Pelicic, V. (2008). Type IV pili: e pluribus unum? Molecular Microbiology, 68(4), 827– 37. Persat, A., Inclan, Y. F., Engel, J. N., Stone, H. a., & Gitai, Z. (2015). Type IV pili mechanochemically regulate virulence factors in Pseudomonas aeruginosa. Proceedings of the National Academy of Sciences, 201502025. Petrova, O. E., Cherny, K. E., & Sauer, K. (2014). The Pseudomonas aeruginosa diguanylate cyclase GcbA, a homolog of P. fluorescens GcbA, promotes initial attachment to surfaces, but not biofilm formation, via regulation of motility. Journal of Bacteriology, 196(15), 2827–2841. Porter, M. E., & Dorman, C. J. (2002). In vivo DNA-binding and oligomerization properties of the Shigella flexneri AraC-like transcriptional regulator VirF as identified by random and site-specific mutagenesis. Journal of Bacteriology, 184(2), 531–539. Pryor, E. E., Waligora, E. A., Xu, B., Dellos-Nolan, S., Wozniak, D. J., & Hollis, T. (2012). The transcription factor AmrZ utilizes multiple DNA binding modes to recognize activator and repressor sequences of Pseudomonas aeruginosa virulence genes. PLoS Pathogens, 8(4), e1002648. Qi, Y., Xu, L., Dong, X., Yau, Y. H., Ho, C. L., Koh, S. L., … Liang, Z.-X. (2012). Functional Divergence of FimX in PilZ Binding and Type IV Pilus Regulation. Journal of Bacteriology, 194(21), 5922–5931. Qiu, D., Damron, F. H., Mima, T., Schweizer, H. P., & Yu, H. D. (2008). PBAD-based shuttle vectors for functional analysis of toxic and highly regulated genes in Pseudomonas and Burkholderia spp. and other bacteria. Applied and Environmental Microbiology, 74(23), 7422–7426. Rafferty, J. B., Somers, W. S., Saint-Girons, I., & Phillips, S. E. (1989). Three-dimensional crystal structures of Escherichia coli met repressor with and without corepressor.

167

Nature, 341(6244), 705–710. Ramsey, D. M., Baynham, P. J., & Wozniak, D. J. (2005). Binding of Pseudomonas aeruginosa AlgZ to sites upstream of the algZ promoter leads to repression of transcription. J. Bacteriol., 187(13), 4430–4443. Ramsey, D. M., & Wozniak, D. J. (2005). Understanding the control of Pseudomonas aeruginosa alginate synthesis and the prospects for management of chronic infections in cystic fibrosis. Molecular Microbiology, 56(2), 309–322. Rashid, M. H., & Kornberg, A. (2000). Inorganic polyphosphate is needed for swimming, swarming, and twitching motilities of Pseudomonas aeruginosa. Proceedings of the National Academy of Sciences, 97(9), 4885–4890. Rice, S. A., Tan, C. H., Mikkelsen, P. J., Kung, V., Woo, J., Tay, M., … Kjelleberg, S. (2009). The biofilm life cycle and virulence of Pseudomonas aeruginosa are dependent on a filamentous prophage. The ISME Journal, 3(3), 271–282. Ross, W., Vrentas, C. E., Sanchez-Vazquez, P., Gaal, T., & Gourse, R. L. (2013). The magic spot: A ppGpp binding site on E. coli RNA polymerase responsible for regulation of transcription initiation. Molecular Cell, 50(3), 420–429. Rutherford, S. T., & Bassler, B. L. (2012). Bacterial quorum sensing: its role in virulence and possibilities for its control. Cold Spring Harbor Perspectives in Medicine, 2(11), 1–26. Ryder, C., Ryder, C., Byrd, M., Byrd, M., Wozniak, D. J., & Wozniak, D. J. (2007). Role of polysaccharides in Pseudomonas aeruginosa biofilm development. Current Opinion in Microbiology, 10(6), 644–8. Sano, Y., Matsui, H., Kobayashi, M., & Kageyama, M. (1993). Molecular structures and functions of pyocins S1 and S2 in Pseudomonas aeruginosa. Journal of Bacteriology, 175(10), 2907–2916. Sauer, K., Camper, A. K., Ehrlich, G. D., Costerton, J. W., & Davies, D. G. (2002). Pseudomonas aeruginosa displays multiple phenotypes during development as a biofilm. Journal of Bacteriology, 184(4), 1140–1154. Sauer, K., Cullen, M. C., Rickard, A. H., Zeef, L. A. H., Gilbert, P., & Davies, D. G. (2004). Characterization of nutrient-induced dispersion in Pseudomonas aeruginosa PAO1 biofilm. Journal of Bacteriology, 186(21), 7312–7326. Schaik, E. J. Van, Giltner, C. L., Audette, G. F., Keizer, D. W., Bautista, D. L., Slupsky, C. M., … Irvin, R. T. (2005). DNA binding : a novel function of Pseudomonas

168

aeruginosa type IV pili. Journal of Bacteriology, 187(4), 1455–1464. Schena, M., Shalon, D., Davis, R. W., & Brown, P. O. (1995). Quantitative monitoring of gene expression patterns with a complementary DNA microarray. Science, 270(5235), 467–470. Schobert, M., & Jahn, D. (2002). Regulation of heme biosynthesis in non-phototrophic bacteria. Journal of Molecular Microbiology and Biotechnology, 4, 287–294. Schreiter, E. R., & Drennan, C. L. (2007). Ribbon-helix-helix transcription factors: variations on a theme. Nat Rev Micro, 5(9), 710–720. Schreiter, E. R., Sintchak, M. D., Guo, Y., Chivers, P. T., Sauer, R. T., & Drennan, C. L. (2003). Crystal structure of the nickel-responsive transcription factor NikR. Nature Structural Biology, 10(10), 794–799. Schurr, M. J. (2013). Which bacterial biofilm exopolysaccharide is preferred, Psl or alginate? Journal of Bacteriology, 195(8), 1623–6. Schurr, M. J., Martin, D. W., Mudd, M. H., Hibler, N. S., Boucher, J. C., & Deretic, V. (1993). The algD promoter: regulation of alginate production by Pseudomonas aeruginosa in cystic fibrosis. Cellular & Molecular Biology Research, 39(4), 371– 6. Semmler, A. B. T., Whitchurch, C. B., & Mattick, J. S. (1999). A re-examination of twitching motility in Pseudomonas aeruginosa. Microbiology, 145(10), 2863– 2873. Seo, J., & Darwin, A. J. (2013). The Pseudomonas aeruginosa periplasmic protease CtpA can affect systems that impact its ability to mount both acute and chronic infections. Infection and Immunity, 81(12), 4561–4570. Sera, T. (2009). Zinc-finger-based artificial transcription factors and their applications. Advanced Drug Delivery Reviews, 61(7-8), 513–526. Shendure, J., & Ji, H. (2008). Next-generation DNA sequencing. Nature Biotechnology, 26(10), 1135–1145. Sheth, H. B., Lee, K. K., Wong, W. Y., Srivastava, G., Hindsgaul, O., Hodges, R. S., … Irvin, R. T. (1994). The pili of Pseudomonas aeruginosa strains PAK and PAO bind specifically to the carbohydrate sequence βGalNAc(1–4)βGal found in glycosphingolipids asialo-GM1 and asialo-GM2. Molecular Microbiology, 11(4), 715–723.

169

Shrout, J. D., Chopp, D. L., Just, C. L., Hentzer, M., Givskov, M., & Parsek, M. R. (2006). The impact of quorum sensing and swarming motility on Pseudomonas aeruginosa biofilm formation is nutritionally conditional. Molecular Microbiology, 62(5), 1264–1277. Simon, R., Priefer, U., & Pühler, A. (1983). A broad host re-mobilization system for in vivo genetic-engineering transposon mutagenesis in Gram-negative bacteria. Bio- Technology, 1(9), 784–791. Singh, P. K., Schaefer, a L., Parsek, M. R., Moninger, T. O., Welsh, M. J., & Greenberg, E. P. (2000). Quorum-sensing signals indicate that cystic fibrosis lungs are infected with bacterial biofilms. Nature, 407(6805), 762–764. Siryaporn, A., Kuchma, S. L., O’Toole, G. a., & Gitai, Z. (2014). Surface attachment induces Pseudomonas aeruginosa virulence. Proceedings of the National Academy of Sciences, (20141009). Smyth, G. K. (2005). Limma: linear models for microarray data. In R. Gentleman and V. Carey and S. Dudoit and R. Irizarry and W. Huber (Ed.), Bioinformatics and Computational Biology Solutions Using {R} and Bioconductor (pp. 397–242). New York: Springer. Snitkin, E. S., & Segre, J. a. (2014). Pseudomonas aeruginosa adaptation to human hosts. Nature Genetics, 47(1), 2–3. Soberón-Chávez, G., Lépine, F., & Déziel, E. (2005). Production of rhamnolipids by Pseudomonas aeruginosa. Applied Microbiology and Biotechnology. Sokatch, J. R., Gunsalus, I. C., & Ornston, L. N. (1986). The bacteria : a treatise on structure and function Volume X. Orlando: Academic press. Sonawane, A., Jyot, J., & Ramphal, R. (2006). Pseudomonas aeruginosa LecB is involved in pilus biogenesis and protease IV activity but not in adhesion to respiratory mucins. Infection and Immunity, 74(12), 7035–7039. Sorensen, R. U., & Klinger, J. D. (1987). Biological effects of Pseudomonas aeruginosa phenazine pigments. Antibiot Chemother, 39, 113–124. Sousa, A., & Pereira, M. (2014). Pseudomonas aeruginosa diversification during infection development in cystic fibrosis lungs—a review. Pathogens, 3(3), 680–703. Stanier, R. Y., Palleroni, N. J., & Doudoroff, M. (1966). The aerobic pseudomonads: a taxonomic study. Journal of General Microbiology, 43(2), 159–271.

170

Stapper, A. P. A., Narasimhan, G., Ohman, D. E., Barakat, J., Hentzer, M., Molin, S., … Mathee, K. (2004). Alginate production affects Pseudomonas aeruginosa biofilm development and architecture, but is not essential for biofilm formation. Journal of Medical Microbiology, 53(Pt 7), 679–690. Stoodley, P., Sauer, K., Davies, D. G., & Costerton, J. W. (2002). Biofilms as complex differentiated communities. Annual Review of Microbiology, 56, 187–209. Su, W., Porter, S., Kustu, S., & Echols, H. (1990). DNA-looping and enhancer activity: association between DNA-bound NtrC activator and RNA polymerase at the bacterial glnA promoter. Proceedings of the National Academy of Sciences, 87(14), 5504–5508. Sunya, S., Gorret, N., Delvigne, F., Uribelarrea, J.-L., & Molina-Jouve, C. (2012). Real- time monitoring of metabolic shift and transcriptional induction of yciG::luxCDABE E. coli reporter strain to a glucose pulse of different concentrations. Journal of Biotechnology, 157(3), 379–90. Tart, A. H., Blanks, M. J., & Wozniak, D. J. (2006). The AlgT-dependent transcriptional regulator AmrZ (AlgZ) inhibits flagellum biosynthesis in mucoid, nonmotile Pseudomonas aeruginosa cystic fibrosis isolates. J. Bacteriol., 188(18), 6483– 6489. Tian, G., Lim, D., Carey, J., & Maas, W. K. (1992). Binding of the arginine repressor of Escherichia coli K12 to its operator sites. Journal of Molecular Biology, 226(2), 387–397. Tielker, D., Hacker, S., Loris, R., Strathmann, M., Wingender, J., Wilhelm, S., … Jaeger, K.-E. (2005). Pseudomonas aeruginosa lectin LecB is located in the outer membrane and is involved in biofilm formation. Microbiology, 151(5), 1313–1323. Trevino, V., Falciani, F., & Barrera-Saldana, H. A. (2007). DNA microarrays: a powerful genomic tool for biomedical and clinical research. Molecular Medicine, 13(October), 9–10. Trunk, K., Benkert, B., Quäck, N., Münch, R., Scheer, M., Garbe, J., … Jahn, D. (2010). Anaerobic adaptation in Pseudomonas aeruginosa: definition of the Anr and Dnr regulons. Environmental Microbiology, 12(6), 1719–1733. Turnbull, L., & Whitchurch, C. (2014). Motility assay: twitching motility. In A. Filloux & J.-L. Ramos (Eds.), Pseudomonas Methods and Protocols SE - 9 (Vol. 1149, pp. 73–86). Springer New York.

171

Van Dijk, E. L., Auger, H., Jaszczyszyn, Y., & Thermes, C. (2014). Ten years of next- generation sequencing technology. Trends in Genetics, 30(9). Vershons, A. K., Youderian, P., Susskindli, M., & Sauers, R. T. (1985). The bacteriophage P22 Arc and Mnt repressors. The Journal of Biological Chemistry, 260(22), 12124– 12129. Vert, M., Doi, Y., Hellwich, K.-H., Hess, M., Hodge, P., Kubisa, P., … Schué, F. (2012). Terminology for biorelated polymers and applications (IUPAC Recommendations 2012). Pure and Applied Chemistry, 84(2), 1. Wagner, V. E., & Iglewski, B. H. (2008). P. aeruginosa biofilms in CF infection. Clinical Reviews in Allergy & Immunology, (May), 1–11. Waldburger, C. D., & Sauer, R. T. (1995). Domains of Mnt repressor: roles in tetramer formation, protein stability, and operator DNA binding. Biochemistry, 34, 13109– 13116. Waligora, E. A., Ramsey, D. M., Pryor, E. E., Lu, H., Hollis, T., Sloan, G. P., … Pryor Jr., E. E. (2010). AmrZ beta-sheet residues are essential for DNA binding and transcriptional control of Pseudomonas aeruginosa virulence genes. J. Bacteriol., 192(20), 5390–5401. Wang, Q., Suzuki, A., Mariconda, S., Porwollik, S., & Harshey, R. M. (2005). Sensing wetness : a new role for the bacterial flagellum, 24(11), 2034–2042. Wang, S., Parsek, M. R., Wozniak, D. J., & Ma, L. Z. (2013). A spider web strategy of type IV pili-mediated migration to build a fibre-like Psl polysaccharide matrix in Pseudomonas aeruginosa biofilms. Environmental Microbiology, 15, 2238–2253. Wang, Z., Gerstein, M., & Snyder, M. (2009). RNA-Seq: a revolutionary tool for transcriptomics. Nature Reviews. Genetics, 10(1), 57–63. Weiss, A., & Shaw, L. N. (2015). Small things considered: the small accessory subunits of RNA polymerase in Gram-positive bacteria. FEMS Microbiology Reviews, 39(4), 541–554. West, S. E. H., Sample, A. K., & Runyenjanecky, L. J. (1994). The vfr gene product, required for Pseudomonas aeruginosa exotoxin A and protease production, belongs to the cyclic AMP receptor protein family. Journal of Bacteriology, 176(24), 7532– 7542. Wetterstrand, K. (2015). DNA Sequencing Costs: Data from the NHGRI Genome Sequencing Program (GSP). Retrieved July 7, 2015, from

172

www.genome.gov/sequencingcosts Whitchurch, C. B., Beatson, S. A., Comolli, J. C., Jakobsen, T., Sargent, J. L., Bertrand, J. J., … Engel, J. N. (2005). Pseudomonas aeruginosa fimL regulates multiple virulence functions by intersecting with Vfr-modulated pathways. Molecular Microbiology, 55(5), 1357–1378. Whitchurch, C. B., Erova, T. E., Emery, J. A., Sargent, J. L., Harris, J. M., Semmler, A. B. T., … Wozniak, D. J. (2002). Phosphorylation of the Pseudomonas aeruginosa response regulator AlgR is essential for type IV fimbria-mediated twitching motility. Journal of Bacteriology, 184(16), 4544–4554. Whitchurch, C. B., & Mattick, J. S. (1994). Characterization of a gene, pilU, required for twitching motility but not phage sensitivity in Pseudomonas aeruginosa. Molecular Microbiology, 13(6), 1079–1091. Whitchurch, C. B., Tolker-Nielsen, T., Ragas, P. C., & Mattick, J. S. (2002). Extracellular DNA required for bacterial biofilm formation. Science (New York, N.Y.), 295(5559), 1487. Whiteley, M., Bangera, M. G., Bumgarner, R., Parsek, M., Teitzel, G., Lory, S., & Greenberg, E. P. (2001). Gene expression in Pseudomonas aeruginosa biofilms. Nature, 413(October), 860–864. Winsor, G. L., Lam, D. K. W., Fleming, L., Lo, R., Whiteside, M. D., Yu, N. Y., … Brinkman, F. S. L. (2011). Pseudomonas Genome Database: improved comparative analysis and population genomics capability for Pseudomonas genomes. Nucleic Acids Research, 39(Database issue), D596–600. Wolfgang, M. C., Lee, V. T., Gilmore, M. E., & Lory, S. (2003). Coordinate regulation of bacterial virulence genes by a novel adenylate cyclase-dependent signaling pathway. Developmental Cell, 4(2), 253–63. Wood, T. K., Knabel, S. J., & Kwan, B. W. (2013). Bacterial persister cell formation and dormancy. Applied and Environmental Microbiology, 79(23), 7116–7121. Wozniak, D. J. (1994). Integration host factor and sequences downstream of the Pseudomonas aeruginosa algD transcription start site are required for expression. J. Bacteriol., 176(16), 5068–5076. Wozniak, D. J., & Ohman, D. E. (1994). Transcriptional analysis of the Pseudomonas aeruginosa genes algR, algB, and algD reveals a hierarchy of alginate gene expression which is modulated by algT. J. Bacteriol., 176(19), 6007–6014.

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Wozniak, D. J., Sprinkle, A. B., & Baynham, P. J. (2003). Control of Pseudomonas aeruginosa algZ expression by the alternative sigma factor AlgT. J. Bacteriol., 185(24), 7297–7300. Wurtzel, O., Yoder-Himes, D. R., Han, K., Dandekar, A. a, Edelheit, S., Greenberg, E. P., … Lory, S. (2012). The single-nucleotide resolution transcriptome of Pseudomonas aeruginosa grown in body temperature. PLoS Pathogens, 8(9), e1002945. Wyckoff, T. J., & Wozniak, D. J. (2001). Transcriptional analysis of genes involved in Pseudomonas aeruginosa biofilms. Methods in Enzymology, 336(1993), 144–151. Xu, B., & Wozniak, D. J. (2015). Development of a novel method for analyzing Pseudomonas aeruginosa twitching motility and its application to define the AmrZ regulon. PLoS One, 10(8). Yabuuchi, E., Kosako, Y., Oyaizu, H., Yano, I., Hotta, H., Hashimoto, Y., … Arakawa, M. (1992). Microbiology and immunology: Proposal of Burkholderia gen. nov. and transfer of seven species of the genus Pseudomonas homology group II to the new genus, with the type species Burkholderia cepacia (Palleroni and Holmes 1981) comb. no. Microbiology and Immunology, 36(12), 1251–1275. Zadran, S., Standley, S., Wong, K., Otiniano, E., Amighi, A., & Baudry, M. (2012). Fluorescence resonance energy transfer (FRET)-based biosensors: visualizing cellular dynamics and bioenergetics. Applied Microbiology and Biotechnology, 96(4), 895–902. Zolfaghar, I., Evans, D. J., & Fleiszig, S. M. J. (2003). Twitching motility contributes to the role of pili in corneal infection caused by Pseudomonas aeruginosa. Infection and Immunity, 71(9), 5389–5393.

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Chapter 5

Appendix A. Spontaneous suppressor screening

A1. Results

In Chapter 2, we designed a novel TM approach that provides a convenient method to isolate TM-active subpopulations, and microarray analyses were performed comparing transcriptome profiles of WT and amrZ mutants. However, this approach failed to identify a tfp gene within the AmrZ regulon. We therefore performed spontaneous suppressor screening as an alternative approach to identify the AmrZ-dependent tfp target(s). A relatively large-scale spontaneous mutant screening of the AmrZV20A strain, which expresses a DNA binding deficient AmrZ variant AmrZV20A, was conducted to identify suppressors that adopted genetic changes restoring the TM phenotype. The procedure is shown as a flowchart in Figure A1. We look for suppressors in which mutations outside of amrZ allow TM to bypass AmrZ-mediated regulation, resulting in restored TM in the absence of amrZ. To avoid intragenic revertants, the amrZ gene in each suppressor mutant was sequenced to ensure it encodes the same AmrZV20A variant as the parental strain

(Figure A2B).

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Total three independent spontaneous suppressors (named M1, M2 and M3) have been isolated, all of which have restored TM significantly (Figure A2A). Genomes of these three suppressors were subsequently sequenced and compared to that of their parental strain AmrZV20A, in order to determine which genetic changes have allowed them to restore TM. A limited number of point mutations were identified (Table A1). In the suppressor M1, mutations are found in genes encoding the housekeeping sigma factor

RpoD, major facilitator superfamily transporters, type III secretion proteins and PvdL (a pyoverdine biosynthesis protein). The suppressor M3 contains two mutations, one in the hypothetical gene PA1279 and the other one in kdpB, which encodes the B chain of a potassium transporter. None of the mutated genes identified in M1 and M3 is known to be related to TM and it is mysterious how these two suppressors have restored their TM. Two mutations were identified in the suppressor M2, one in the T6SS (type VI secretion system) gene PA2363 and the other one in pilA. The major pilin PilA is the main subunit of the

TFP filament and required for TM (J. S. Mattick, Whitchurch, & Alm, 1996). The pilA point mutation in suppressor M2 leads to a serine-to-alanine mutation at the 73rd amino acid. Serine contains a hydroxyl group and thus is a polar amino acid. This amino acid plays important roles in catalytic functions of many enzymes and frequently undergoes modifications such as glycosylation and phosphorylation. In P. aeruginosa strain 1244 and

Neisseria gonnorrhoeae, certain serine residues in type IV major pilins are important glycosylation sites (Comer, Marshall, Blanch, Deal, & Castric, 2002; Hegge et al., 2004).

Since this S73A mutation correlates with restored TM in suppressor M2, we hypothesized that Ser73 in PilA represses TFP function, possibly by glycosylation, which is suppressed by functional AmrZ.

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We thereby attempted to recapitulate this pilA mutation in the ΔamrZ mutant. The arabinose-inducible plasmid harboring PilAS73A was placed into a ΔpilA or ΔpilA&ΔamrZ double mutant, and TM measured after inducing PilAS73A expression by arabinose. The

PilAS73A plasmid restored TM in the ΔpilA mutant at a similar level as the one carrying

WT pilA, suggesting this point mutation variant was produced and interchangeable with its wild type counterpart (Figure A3A). However, in the ΔpilA&ΔamrZ mutant, no TM was observed when the PilAS73A plasmid was induced (Figure A3B), suggesting the S73A mutation in pilA alone is unable to suppress the TM defect in the ΔamrZ mutant. These results do not support our hypothesis and we fail to confirm the role of PilAS73A during

AmrZ-mediated regulation of TM.

AmrZ was shown to repress c-di-GMP synthesis (C. J. Jones et al., 2014) and there have been a few studies showing the crosstalk between c-di-GMP and the TFP system (Alm et al., 1996; Amikam & Galperin, 2006; Jain et al., 2012; Qi et al., 2012). We therefore probed the possibility that AmrZ mediates its TM regulation indirectly through modulating the level of this second messenger. Intracellular concentrations of c-di-GMP were measured in these three suppressors and compared to their parental strain AmrZV20A. In suppressors M2 and M3, no change of the c-di-GMP level was observed compared to their parental strain. In contrast, the c-di-GMP level in M1 decreased to the WT level (Figure

A4A), and we thereby suspect that this alteration of the c-di-GMP concentration might correlate with restored TM. Therefore, we attempted to elevate the c-di-GMP level in M2 by overexpressing a diguanylate cyclase-encoding gene gcbA, but no significant reduction of TM was observed (Figure A4B). To determine if a reduced c-di-GMP level would

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restore TM in the ΔamrZ mutant, a plasmid containing the phosphodiesterase gene PA2133

was placed into this strain and TM was measured. This plasmid restored the smooth colony

phenotype of the small colony variant ΔwspF, indicating PA2133 production reduced the

c-di-GMP level (data not shown). However, PA2133 overexpression failed to restore TM

in the ΔamrZ mutant (Figure A5). Above results indicate that alteration of the c-di-GMP

level alone is unable to affect the outcome of AmrZ-mediated TM regulation.

Collectively, in this project we attempted to use an innovative approach by

combining spontaneous suppressor screening with next-generation sequencing. Large scale

screening of the ΔamrZ mutant was conducted to identify spontaneous mutants that

restored TM, and next-generation sequencing was used to determine mutations in those

suppressors. A limited number of mutations were identified and only one mutation

occurred within a tfp gene (pilA) in the suppressor M2. However, we were unable to

recapitulate the phenotype of this pilA mutation in the ΔamrZ strain. Additional

investigations were performed in order to identify the role of c-di-GMP during AmrZ-

mediated regulation of TM. Our results did not support the hypothesis that perturbing c-di-

GMP levels affects the outcome of AmrZ-mediated regulation of TM.

A2. Discussion

High-fidelity DNA replication is achieved by both accurate dNTP incorporation by the DNA polymerase and multiple proofreading and DNA repair mechanisms.

Nevertheless, mutations still occur naturally and the average mutation rate in E. coli is

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approximately ~5*10-10 per base pair per genome replication (J. W. Drake, 1991).

Scientists have been taking advantage of this replication error to screen for mutations that

cause phenotypic changes of interest. As an example, Nolan et al. used this approach to identify suppressors in which TM was restored in the ΔfimL mutant (Nolan et al., 2012).

Although DNA binding-competent AmrZ is required for TM (P. J. Baynham et al., 2006),

amrZ mutants, especially those carrying point mutation variants, occasionally display small

twitching zones. We therefore screened for spontaneous revertants in an amrZ mutant that restored TM. This method was proven successful and three revertants were identified, all of which have mutations outside of the amrZ gene. In the past, it was extremely difficult to map mutation locations in spontaneous suppressors as no marker or label is available.

This problem has been mostly solved in the next-generation sequencing era. Whole genomes sequencing allows the identification of single-nucleotide polymorphisms (SNPs) and small insertions/deletions. However, due to technical limitations, detectable maximum sizes of insertion/deletions are limited to approximately 50 bp during our Illumina data analyses. Possible large insertions/deletions therefore cannot be revealed by our data analyses, which may potentially be solved by better software design in the future.

In each of the three suppressors, a few mutations are identified, but none could be confirmed to be responsible for restored TM. It is therefore intriguing what TM-pertinent genetic changes have occurred in these strains. Since our transcriptomics (both DNA microarray and RNA-seq) failed to identify TM-related AmrZ targets (Chapter 2 and (C.

J. Jones et al., 2014)), we suspect that AmrZ regulates a previously unidentified gene(s) that is required for TM or indirectly by affecting intracellular signaling such as second

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messengers c-di-GMP and 3’,5’-cyclic adenosine monophosphate (cAMP). Although our perturbations of intracellular c-di-GMP concentrations did not affect TM in the suppressor

M1 (Figure A4) and the ΔamrZ mutant (Figure A5), we cannot rule out the role of c-di-

GMP during AmrZ-mediated regulation. This is because c-di-GMP signaling is

subcellularly localized, and its concentrations in different subcellular compartments may

vary and are critical for differential responses to various stimuli (Merritt et al., 2010).

Another candidate is cAMP, which plays important roles in signal transduction in both

prokaryotic and eukaryotic organisms. This molecule plays a critical role in the glucose-

lactose diauxie in E. coli as activation of the lac promoter requires the binding by cAMP-

CRP (cAMP receptor protein) (Fic et al., 2009). In P. aeruginosa, cAMP and the CRP

homolog Vfr participate in regulation of a number of processes such as T3SS, exotoxins

and proteases (Fuchs et al., 2010; West, Sample, & Runyenjanecky, 1994; Wolfgang, Lee,

Gilmore, & Lory, 2003). In addition, vfr disruption results in a TM defect and cAMP is

implicated in fine control of TM (Beatson, Whitchurch, Jennifer, et al., 2002; Fulcher et

al., 2010; Nolan et al., 2012; Whitchurch et al., 2005). The regulatory network of the TM- necessary histidine-kinase protein FimL also intersects with the Vfr-cAMP pathway in the way that FimL regulates cAMP synthesis and vfr overexpression in trans restores TM in

the ΔfimL mutant (Inclan, Huseby, & Engel, 2011; Whitchurch et al., 2005). Intriguingly,

the PilA production pattern (reduced surface pili amounts but similar in whole cell lysates

relative to wild type) of the ΔamrZ mutant is similar to the Δvfr mutant and the ΔfimL mutant (P. J. Baynham et al., 2006; Beatson, Whitchurch, Annalese, Mattick, & Semmler,

2002; Whitchurch et al., 2005). In combination with the fact that both AmrZ and Vfr activate algD transcription, it is therefore plausible that AmrZ and Vfr might share the

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same mechanism of TM regulation. The specific mechanism of Vfr-mediated activation of

TM remains unclear as well and there does not seem to be a hierarchy between these two

TFs based on transcriptome profiling results (C. J. Jones et al., 2014; Wolfgang et al., 2003;

Xu & Wozniak, 2015). Future work is needed to understand regulatory mechanisms of TM by AmrZ and Vfr, and to determine if AmrZ-mediated regulation intersects with cAMP signaling.

A3. Materials and Methods

Bacterial strains, plasmids, oligonucleotides and growth conditions

Information of bacterial strains, plasmids, and oligonucleotides employed in this study is summarized in Table A2. Bacteria (Escherichia coli and Pseudomonas aeruginosa) were grown as previously (Waligora et al., 2010). Briefly, E. coli strains were grown in LB (each liter LB contains 10 g tryptone, 5 g yeast extract, 5 g sodium chloride) or on LA (LB with 1.5% (w/v) agar). 100 µg/ml ampicillin was used to maintain plasmids in E. coli. All P. aeruginosa strains were grown in LBNS or on LANS (LB or LA without sodium chloride, respectively). In P. aeruginosa, 300 µg/ml carbenicillin was used to maintain plasmids. All strains were grown at 37°C unless stated otherwise. All oligonucleotides were ordered desalted from Sigma Aldrich.

In-frame deletion and point mutation mutants were generated by overlap extension

PCR using the gene replacement vector pEX18Ap (Heckman & Pease, 2007; Hoang et al.,

1998). Multi-copy pHERD20T-based plasmids were transferred into P. aeruginosa strains

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through bi-parental mating with the E. coli helper strain S17-1 (Qiu et al., 2008; Simon et

al., 1983).

Subsurface twitching assays

Subsurface twitching assays were performed as a previously described study (Xu

& Wozniak, 2015). Briefly, a single colony of a strain of interest was resuspended in 400

µl LBNS, and P10 tips were immersed in the above suspension and used to stab through

the agar of 1-day-old 1% LANS plates. Plates were incubated upright in a humid chamber

at room temperature for 5 days, and imaged through white epiluminescence via a

ChemicDoc imaging system (Bio-Rad). Relative twitching activities (twitching ratios)

were calculated by dividing diameters of twitching zones by diameters of central colonies.

Spontaneous suppressor screening

The AmrZV20A strain was used as the parental strain during screening. This strain

underwent prolonged incubation (8 days) at room temperature during subsurface twitching

assays. Occasionally stabbed colonies exhibited small twitching zones, from which

bacteria were streaked out and isolated. These isolates were used for the next cycle of

prolonged twitching assays. When a bacterial isolate displayed significantly and

hereditarily restored TM, it was considered as a spontaneous suppressor. During this

screening process, three independent suppressors were isolated and labeled as M1, M2 and

M3.

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Whole genome sequencing

Strains submitted for next-generation sequencing include PAO1, AmrZV20A

(WFPA513), ΔamrZ (WFPA205) and suppressors M1, M2 and M3. Whole genomic DNA was prepared using the Promega Wizard® Genomic DNA Purification kit (Catalog #:

A1120) following the manufacture protocol. Subsequent DNA sample preparation and data

analyses were performed by our collaborator --the Peter White’s research group at Center

for Microbial Pathogenesis in Nationwide Children’s Hospital, Columbus, OH (Kelly et

al., 2015). Briefly, DNA samples were prepared for Illumina MiSeq following manufacture

protocols. Sequences of these strains were aligned to the PAO1 reference using bwa 0.7.4

(H. Li & Durbin, 2009) and duplicate reads were removed by Picard Tools MarkDuplicates

(Broad Institute, n.d.). Base quality scores were recalibrated by GATK 1.6 to remove false

positives that had poorly scored sequencer qualities (McKenna et al., 2010). Variants were called at locations where a significant number of the reads are different from the reference by GATK 1.6 UnifiedGenotyper and these variants were annotated using known gene information from NCBI.

Intracellular c-di-GMP concentration measurements

Intracellular c-di-GMP concentrations of LANS-grown P. aeruginosa were

measured by Laura Jennings from the Matthews Parsek laboratory, and the procedure is

the same as previously described (C. J. Jones et al., 2014).

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A4. Acknowledgement

This work was completed in collaborations with research groups of Peter White from Center for Microbial Pathogenesis at Nationwide Hospital, Columbus, OH and

Matthews Parsek from University of Washington. More specifically, Benjamin Kelly from the Peter White group and Laura Jennings from Matthews Parsek laboratory are directly involved in this work.

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Figure A1. Flowchart of the suppressor screening process.

Repeated screening cycles were performed initially starting from the AmrZV20A strain, later from those isolates that exhibited slightly increased TM. Suppressors were isolated when they had restored TM hereditarily, and their amrZ genes were sequenced to ensure no intragenic mutation within amrZ.

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Figure A2. Three suppressors exhibit restored TM and no intragenic mutation within amrZ.

(A.) Three suppressors (M1, M2 and M3) were identified and their TM was measured via subsurface twitching assays. (B.) Their amrZ genes were sequenced and confirmed absent of intragenic mutations. Data in (A) represent three independent experiments. Unpaired, two-tailed student t-tests were performed to determine statistical differences between TM of strains. ***: P<0.001.

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Figure A3. PilAS73A fails to restore TM in the ΔpilA&ΔamrZ mutant.

The plasmid encoding an empty vector, PilA or PilAS73A was transferred into ΔpilA or

ΔpilA&ΔamrZ, and TM measured by the subsurface twitching assay in the presence of 1% arabinose. Scale bars: 5 mm.

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Figure A4. The suppressor M1 displays a reduced c-di-GMP level while gcbA

overexpression in this strain fails to inhibit TM.

(A.) LANS-grown bacteria were harvested and lysed. Cell extracts were subject to c-di-

GMP measurements, which were normalized to protein amounts. (B.) The plasmid pgcbA

(pBX22) or the empty vector (pHERD20T) was placed into the suppressor M1, and its TM measured via the subsurface twitching assay. Data represent three independent experiments. Unpaired, two-tailed student t-tests were used for statistical analyses. ns: not significant; *: P<0.05.

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Figure A5. TM of the ΔamrZ mutant after expressing the plasmid-borne phosphodiesterase PA2133.

The plasmid pJN2133 was transferred into PAO1 or the ΔamrZ mutant, and TM was

measured in the absence/presence of 0.5% arabinose via the subsurface twitching assay.

Scale bars: 5 mm.

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Table A1. Point mutations identified in three suppressor mutants

Amino Mutant Base Gene Function Location Coding Change Acid # Change Change nonsynonymous rpoD Sigma factor ORF c.C1760G p.T587S SNV probable major facilitator nonsynonymous PA0811 ORF c.C605A p.A202E superfamily SNV (MFS) transporter translocation nonsynonymous M1 pscQ protein in type ORF c.A339T p.Q113H SNV III secretion type III secretion pscD ORF Stop/gain SNV c.C295T p.Q99X apparatus protein Pyoverdine nonsynonymous pvdL ORF c.C4238G p.A1413G synthesis SNV probable MFS nonsynonymous PA2472 ORF c.T1271C p.I424T transporter SNV Predicted nonsynonymous PA2363 component of ORF c.T728C L243P SNV M2 the T6SS TFP major nonsynonymous pilA ORF c.T217G S73A pilin SNV hypothetical nonsynonymous PA1259 ORF c.A1433G p.H478R protein SNV potassium- M3 transporting nonsynonymous kdpB ORF c.G1789T p.A597S ATPase, B SNV chain

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Table A2. Strains, plasmids, and oligonucleotides used in Appendix A

Plasmids pHERD20T Arabinose-inducible vector (Qiu et al., 2008) pBX9 pHERD20T-WT pilA. Primers pilA_CDS_F2 and This study pilA_CDS_R were used to amplify the pilA coding sequence. pBX41 pHERD20T-PilAS73A. Primers pilA_A, E, F and D were This study used to generate the pilA coding sequence with the S73A mutation.

pBX22 pHERD20T-gcbA. (C. J. Jones et al., 2014)

Continued

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Table A2. Continued pJN105 Arabinose-inducible vector (J. W. Hickman et al., 2005) pJN2133 pJN105-PA2133 (J. W. Hickman et al., 2005) Oligonucleotides pilA_CDS_F2 cgctctagaCAACGGAGAGATTCATGAAAGCTC This study pilA_CDS_R cgcgcatgcCGCCTCACCCTCTGAACGAATC This study pilA_A tgctctagaGCTTTTCGCTGATGGCGTC (Xu & Wozniak, 2015) pilA_E GTACTACTGCTGCTACTGCGAC This study pilA_F GTCGCAGTAGCAGCAGTAGTAC This study pilA_D aactgcagCGCATAGCACCCGGCAAG (Xu & Wozniak, 2015)

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