A STUDY of SURFACE MOTILITY and BIOFILM FORMATION in Pseudomonas

A STUDY OF SURFACE MOTILITY AND BIOFILM FORMATION IN Pseudomonas

aeruginosa: QUORUM SENSING AND PHOTODYNAMIC ANTIMICROBIAL

CHEMOTHERAPY

Dissertation

Submitted to

The College of Arts and Sciences of the

UNIVERSITY OF DAYTON

In Partial Fulfillment of the Requirements for

The Degree

Doctor of Philosophy in Biology

Tracy Lynn Collins

UNIVERSITY OF DAYTON Dayton, Ohio December 2010

A STUDY OF SURFACE MOTILITY AND BIOFILM FORMATION IN Pseudomonas

aeruginosa: QUORUM SENSING AND PHOTODYNAMIC ANTIMICROBIAL

CHEMOTHERAPY

Name: Collins, Tracy Lynn University of Dayton

APPROVED BY:

______Jayne B. Robinson Ph.D. Major Advisor Chair Department of Biology

______Carissa Krane Ph.D. Committee Member

______Mark Nielsen Ph.D. Committee Member

______John Rowe Ph.D. Committee Member

______Shawn Swavey Ph.D. Committee Member

ABSTRACT

A STUDY OF SURFACE MOTILITY AND BIOFILM FORMATION IN Pseudomonas

aeruginosa: QUORUM SENSING AND PHOTODYNAMIC ANTIMICROBIAL

CHEMOTHERAPY

Name: Collins, Tracy Lynn University of Dayton

Advisor: Dr. Jayne B. Robinson

Pseudomonas aeruginosa is an opportunistic pathogen that commonly causes infection in immunocompromised individuals. This bacterium forms complex communities known as biofilms. Biofilm formation is dependent on motility as well as quorum-sensing. In this study, we show that P. aeruginosa surface motility is inhibited in the presence of the quorum-sensing molecule E,E-farnesol. In the presence of E,E-farnesol, there is an 4- fold increase in rhamnolipid production. Because swarming motility is dependent on rhamnolipid production, this increase could account for the inhibition in swarming motility observed in the presence of E,E-farnesol. In addition, the effect of the cationic porphyrin 5,10,15,20-tetrakis(1-methyl-pyridino)-21H,23H-porphine, tetra-p-tosylate salt

(TMP) on P. aeruginosa biofilms was examined. Exposure to 225 µM TMP and photoactivation resulted in almost complete killing of biofilm associated cell as well as the detachment of wild-type PAO1 biofilms. In contrast, pqsA mutant biofilms that contain less extracellular DNA remained intact. Our results suggest that the action of

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photoactivated TMP on P. aeruginosa biofilms is two-fold: direct killing of individual cells within biofilms and detachment of the biofilm from the substratum. There was no evidence of porphyrin toxicity in the absence of light; however, biofilms pretreated with

TMP without photoactivation were substantially more sensitive to tobramycin than untreated biofilms.

DEDICATION

This is dedicated to my mother, Linda Collins, and to the memory of my grandparents.

ACKNOWLEDGEMENTS

Nothing I ever accomplish is done entirely of my own merit. The Lord has blessed me in my life with the help and support of a number of people.

I would like to first thank the biology department staff: Karen Bahr, Lynda

Routley, Cathy Wolfe, Jay Lee, and Sue Trainum. Each of them does a wonderful job

making sure the department runs smoothly. I’d especially like to acknowledge Karen

Bahr because she has always looked out for me and is my guardian in the department.

I am very thankful to my committee members John Rowe, Mark Nielsen, Carissa

Krane, and Shawn Swavey. I appreciate the time and input each of them has contributed

to my research project.

Because I have been at UD for ten years, there are a number of labmates that I’d

like to thank. From “Robinson Generation I”, I’d to acknowledge Mary Connolly (for

scraping me off the bathroom floor that one time), Paul DeLange (for loaning me $4

when I forgot my purse), Michelle Yingling (for our “complaint” walks), Amy Beumer

(for always listening), Chris Moler (for not drowning at sea), Lisa Kaiser (for your

superlatives), Rachel Ducharme (for being “The Roach”), Bethany Yager (for the pub

story), Kevin Wheeler (for not playing Jazz music), and Ian Kearns (for the techno

music). “Robinson Generation I” was truly made up of a great group of intelligent and

enjoyable individuals who were a pleasure to work with in the lab, even when

Environmental Safety busted us.

I would also like to thank my labmates from “Robinson Generation II”: Sulabha

Chaganaboyana (for being my indian sister), Mike Goodson (for tea and chimney sweeps), Muhamad Shakhatrah (for the baklava), Laura Gueltzow (for your dimples), Jen

Lang (for being “Jen Lange!”), Liz Markus (for being my partner in crime), Liz Raphael

(for the word “ridonculous”), Mariah Roller (for serenading me), Brittany Demmitt (for the “Thursday night cry”), and honorary lab member Alison DesJardins (for Britney

Spears). This group of people has been so much fun to work with and I can’t begin to express the joy that each of them have brought to my life. When I think back to

“Robinson Generation II”, I will always have fonds memories involving tie dying,

dancing to Lady Gaga, and “Pseudomonas in…aeruginosa out.” Thank you especially to

Liz Markus for allowing me to be part of the “tetrakis takeover” team, for our countless

hours in the confocal room, and adventures involving fishnapping, hello kitty, and albino

squirrels. You’re an amazing person and great friend.

In addition, I would like to thank all the grad students for their camaraderie and

friendship, especially Vandana Sharma, Chris Noriega, and Meagan Roddy. Meagan,

thank you for always being a loyal, supportive friend who gets my movie references. It’s

great to be able to talk about “the two Corey’s” or “the Brat Pack” and someone know

precisely who I’m referring to.

Ten years ago, I had no idea what an amazing journey I was about to enter. In

retrospect, I know now that it was part of a divine plan that I was supposed to come to

UD, be surrounded by wonderful people, and most of all work under a truly great mentor.

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The scientist that I am today is a reflection of the dedication and guidance of my advisor

Jayne B. Robinson. Thank you for making me part of your lab and giving me the greatest

education I could ever ask for. I have the highest respect for you, but please know I will

never surrender to you on Pearl Harbor Day.

Most of all, I want to thank my family for their constant love and support. I have

four of the funniest, smart brothers: Paul, Phil, Mike, and Robbie. They have always

been good to me and are great men. Also, I would like to recognize my sister-in-law,

Erica, for being my friend and blessing the family with two beautiful children, Anna and

Sam. And, lastly, I would like to acknowledge my mother, Linda, who deserves the most

credit of all. She is the reason I am a microbiologist. My mother has always stressed the

importance of not settling and striving for the very best. You’ve pushed and disciplined

me when I don’t feel like trying. Thank you for providing for me physically and

emotionally. You are the best mother. I love you.

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TABLE OF CONTENTS

DEDICATION………………………………………………………………………….... v

ACKNOWLEDGEMENTS...... vi

TABLE OF CONTENTS...... ix

LIST OF FIGURES...... xi

LIST OF TABLES...... xv

CHAPTERS

I. Literature Review…………………………………………………………………1

II. E,E-farnesol inhibits surface motility in Pseudomonas aeruginosa through

rhamnolipid production...... 45

a. Abstract...... 46

b. Introduction...... 48

c. Methods...... 53

d. Results...... 60

e. Discussion...... 84

f. Summary...... 93

g. Future Studies...... 95

III. The effect of a cationic porphyrin on Pseudomonas aeruginosa biofilms...... 97

a. Abstract...... 98

b. Introduction...... 100

c. Methods...... 102

d. Results...... 106

e. Discussion...... 116

LITERATURE CITED...... 120

LIST OF FIGURES

Chapter I

Figure 1. The bacterial flagellum...... 29

Figure 2. Model of the structure of type IV pilin monomer in Pseudomonas

aeruginosa...... 30

Figure 3. Model of type IV pili assembly and retraction in Pseudomonas

aeruginosa……………………………………………………………. 31

Figure 4. Model of the main regulatory networks governing pilus biogenesis and

twitching motility in Pseudomonas aeruginosa ...... 32

Figure 5. Structural model for the full-length Tsr dimer based on structures of the

periplasmic and cytoplasmic domains...... 33

Figure 6. Chemotaxis signal transduction pathway in Escherichia coli...... 34

Figure 7. Chemotaxis genes in Pseudomonas aeruginosa...... 35

Figure 8. Quorum sensing system of bioluminescence in Vibrio fischeri...... 36

Figure 9. The LasRI-RhlRI quorum-sensing systems in Pseudomonas

aeruginosa ...... 37

Figure 10. Structure of P. aeruginosa quorum-sensing molecules...... 38

Figure 11. Regulation of Las and Rhl systems quorum-sensing systems in P.

aeruginosa...... 39

Figure 12. C12 compounds causing inhibition of C. albicans filamentation...... 40

Figure 13. Five stages of biofilm development of P. aeruginosa...... 41

Figure 14. Photosensitisation pathways for photosensitizer...... 42

Figure 15. Photosensitizer absorption maxima...... 43

Figure 16. Molecular structure of 5,10,15,20-tetrakis(1-methyl-pyridino)-

21H,23H-porphine (TMPyP)...... 44

Chapter II

Figure 1. Macroscopic twitching motility of wild-type PAO1 in the presence of

E,E-farnesol...... 68

Figure 2. Microscopic twitching motility of wild-type PAO1 in the presence of

E,E-farnesol...... 69

Figure 3. Swarming motility of wild-type PAO1, pilJ (FA6) mutant, and

complemented pilJ mutant on NB (0.5% agar) in the presence of E,E-

farnesol and dodecanol...... 70

Figure 4. Swarming motility of wild-type PAO1 and pilJ (FA6) mutant on

PPGAS agar in the presence of E,E-farnesol...... 71

Figure 5. Swarming motility of wild-type PA14 and sadB mutant in the

presence of E,E-farnesol...... 72

Figure 6. Swimming motility of wild-type PAO1, pilJ (FA6) mutant, and

complemented pilJ mutant in the presence of E,E-farnesol...... 73

Figure 7. Rhamnose production by wild-type PAO1 and pilJ (FA6) mutant cells

harvested from swarm plates supplemented with E,E-farnesol...... 75

xii

Figure 8. Immunoblot analysis of total pilin in the presence of E,E-farnesol..... 76

Figure 9. Immunoblot analysis of flagellin B harvested from wild-type PAO1 and

pilJ (FA6) mutant cells grown in LB broth supplemented with E,E-

farnesol...... 77

Figure 10. Immunoblot analysis of flagellin B harvested from wild-type PAO1

cells grown on swarm plates supplemented with E,E-farnesol...... 78

Figure 11. Immunoblot analysis of PilB in wild-type PAO1 grown in the

presence of E,E-farnesol...... 79

Figure 12. Quantification of PilB immunoprecipitated from wild-type PAO1

following exposure to E,E-farnesol…………………...... 80

Figure 13. In-vivo 3H-methylation of wild-type PAO1 and pilJ mutant (FA6)

cells in response to E,E-farnesol...... 82

Figure 14. Quantification of PilJ methylation in response to E,E-farnesol ...... 83

Chapter III

Figure 1. Confocal scanning laser micrographs of P. aeruginosa biofilms...... 109

Figure 2. Effect of TMP and light irradiation on cell survival of P. aeruginosa

biofilm associated cells...... 110

Figure 3. Effect of TMP and light irradiation on cell survival of P. aeruginosa

biofilm associated cells via LIVE/DEAD BacLight stain...... 111

Figure 4. Gel electrophoresis analysis of plasmid (pUCP18) DNA treated with

TMP and irradiated...... 112

xiii

Figure 5. Quantification of extracellular DNA of wild-type MPAO1 biofilms

treated with TMP and light irradiation...... 113

Figure 6. Quantification of extracellular DNA of wild-type MPAO1 biofilms

treated with TMP with and without light irradiation...... 114

Figure 7. Confocal scanning laser micrographs of P. aeruginosa wild-type PAO1

biofilms treated with tobramycin only and TMP and tobramycin in the

absence of light...... 115

xiv

LIST OF TABLES

Chapter II

Table 1. Bacterial strains used in this study...... 67

Table 2. Surface motility of P. aeruginosa wild-type and mutant strains...... 74

Table 3. Effect of E,E-farnesol on pilJ expression: Alkaline Phosphatase Activity

of pilJ::TnphoA fusion in wild-type MPAO1...... 81

Chapter I

LITERATURE REVIEW

Swimming motility and Swarming motility

Pseudomonas aeruginosa is capable of three forms of motility: swimming, swarming, and twitching motility. Swimming motility is movement through an aqueous environment via the rotation of flagella (Harshey 2003). In a semi-solid environment, the cell switches from swimming to swarming motility (Fraser and Hughes 1999, Harshey

1994). While swimming motility is characterized by individual cell movement, swarming motility relies on the smooth migration of a group of cells. During swarming motility, cells differentiate from vegetative cells into swarmer cells that are elongated and have two polar flagella (Fraser and Hughes 1999, Kohler et al. 2000). This morphological change is induced by nutrient depletion and in response to certain amino acids (i.e. glutamate, aspartate, proline, and histidine) (Kohler et al. 2000, Toutain et al.

2005).

Like swimming motility, swarming motility is also mediated through the rotation of flagella. Flagella can be divided into three sections: the basal body, the hook, and the filament. In gram-negative bacteria, the basal body consists of four stacked rings. The first set of rings are known as the MS rings. The M ring is embedded in the cell membrane followed by the S ring. The S ring is produced through the expression of fliF and is associated with the cell membrane in the periplasmic space. These two rings serve as anchor between the flagellum and the cell (Harshey 2003, Sowa and Berry 2008). The next ring is the P ring which is fixed in the peptidoglycan, and the L ring located in the outer membrane. A central rod passes through all four rings and is attached to the hook.

The flagellum hook is either left handed or right handed and rotates during swimming and swarming motility. The hook is composed of the protein FlgE. There are

two other hook associated proteins HAP1 (FlgK) and HAP3 (FlgL) that form the junction between the hook and filament. HAP2 (FlgD) is responsible for capping the filament protein (Ikeda et al. 1987, Sower and Berry 2008). The filament is helically composed of proteins known as flagellins, FliC. As the flagellum is assembled, flagellin subunits travel through the hollow core of the filament where they are attached to the tip of the flagellum. This attachment is directed by the flagellum cap (Fig. 1) (Yonekura et al.

2000).

The bacterial flagellum is powered by a proton motive force, or in some cases a sodium motive force (Blair 2003, Hirota and Imae 1983, Larsen et al. 1974). The flagellar motor is divided into two components: the rotor and the stator. The rotor is composed of the FliG, FliM, and FliN (Toker and Macnab 1997, Francis et al. 1994,

Khan et al. 1992). These switch proteins direct the rotation of the flagellum either counterclockwise or clockwise. The second component of the motor is the stator. The motor is composed of the Mot proteins MotA and MotB. These intergral proteins exist in a ratio of four MotA to two MotB proteins and are responsible for conducting protons through the inner membrane resulting in the rotation of the motor (Blair 2003, Kojima and Blair 2001). P. aeruginosa also possesses another set of stator proteins known as

MotCD. Both sets of stator proteins, MotAB and MotCD, are functionally redundant for swimming motility; however, under high agar concentrations (i.e. swarming conditions

[>0.5% agar]) ΔmotCD mutants are unable to swarm. These findings suggest that both stators have separate roles during swarming motility (Toutain et al. 2005).

In addition to flagella, it has been suggested that swarming motility requires Type

IV pili. pilA mutants in PAK background have been specifically shown to be deficient in

swarming motility. These findings indicate that Type IV pili are either necessary for

surface propagation or may sense the viscosity of the surface thereby signaling the initiation of swarming (Kohler et al. 2000). However other studies have reported that

Type IV pili are unnecessary for swarming motility in PAO1 background (Rashid and

Kornberg et al. 2000). These conflicting findings may be due to strain variations between

PAK and PAO1.

On swarm plates, cells move away from the point of inoculation and form a

distinct pattern referred to as tendrils. This surface colonization is dependent upon the

production of the biosurfactant rhamnolipid. Rhamnolipid biosynthesis is dependent on

quorum-sensing, specifically the rhl system (Kohler et al. 2000).

Rhamnolipids are glycolipids composed of mono- or di-rhamnose linked to the

fatty acid 3-hydroxylalkanoic acid. RhlG initiates fatty acid synthesis of the lipid

component of rhamnolipids. These fatty acids are converted to the precursor 3-

hydroxyalcanoyl-3-hydroxyalcanoate (HAA) by RhlA (Deziel et al. 2003). The final step

in this biosynthetic pathway involves the sequential transfer of dTDP-L-rhamnose to

HAA resulting in mono-rhamnolipid and the addition of another dTDP-L-rhamnose to mono-rhamnolipid producing di-rhamnolipid. These two reactions are catalyzed by RhlB and RhlC, respectively (Maier and Soberon-Chavez 2000, Rahim et al. 2001).

The exact function of rhamnolipids is still unclear; however, they appear to be necessary for swarming motility and reduce surface tension (Kohler et al. 2000). While rhlA mutants (deficient in HAA and rhamnolipids) are unable to swarm, rhlB mutants

(deficient in mono-and di-rhamnolipids) and rhlC mutants (deficient in di-rhamnolipids) retain their ability to swarm (Caiazza et al. 2005). However, rhlB and rhlC mutants

exhibit altered swarming patterns in wild-type cells. Wild-type cells typically possess long radiating tendrils that consist of large cell-free areas between tendrils. rhlB and rhlC mutants have short irregular tendrils that are colonized with cells between adjacent tendrils. Additionally, swarming motility is completely inhibited when purified rhamnolipids are added to swarm plates (Caiazza et al. 2005). These observations suggest that rhamnolipids and their precursor HAA have different roles in swarming motility. HAA is believed to be the minimal surfactant necessary for swarming motility, while rhamnolipids modulate swarming motility by preserving the cell-free areas between tendrils.

Twitching motility

Pseudomonas aeruginosa possesses the ability to translocate across abiotic and biotic surfaces through a mechanism known as twitching motility. “Twitching motility” was first coined by Lautrop in 1961 (Lautrop 1961). When examining Acinetobacter calcoaceticus, Lautrop observed that these cells appeared to be moving in a jerking fashion. This movement has also been observed in Neisseria gonorrhea and Myxococcus xanthus. Twitching motility is almost entirely restricted to Gram-negative bacteria, but has been observed in Streptococcus sanguis, a Gram-positive bacterium.

Twitching motility allows cells to rapidly colonize a surface during high nutrient availability and involves cell-to-cell contact (Mattick 2002). Furthermore, this type of motility facilitates colonial behavior involved in biofilm formation and fruiting bodies

(O’Toole and Kolter 1998). Twitching motility occurs on surfaces with low water content as well as hydrated surfaces (Mattick 2002, Bradley 1980). It is distinct from other forms of motility, such as swimming and swarming, because it is mediated through

the extension and retraction of Type IV pili rather than the rotation of flagella (Bradley

1980, Skerker and Berg 2001).

In P. aeruginosa, approximately 40 genes have been identified whose products

are required for twitching motility and are directly involved in pili biogenesis (Alm and

Mattick 1997, Beatson et al. 2002). Pili are 5-7 nm in diameter, extending several µm in length and are composed of a single protein subunit, pilin, which is encoded by the pilA

gene (Mattick 2002). Pilin is arranged in a helical conformation consisting of five

subunits per turn. The primary structure of pilin is approximately 145-160 amino acids

long with a positively charged leader sequence and a highly conserved, hydrophobic

amino terminal domain (Keizer et al. 2001). This domain forms the core of the pilus

fiber, a coiled-coil of ∝-helices. Beyond the conserved amino-terminal region, there is a

hydrophilic central and carboxy-terminal domain (Forest and Tainer 1997). These

domains surround the core region as a scaffold of β-sheets (Fig. 2) (Mattick 2002).

Furthermore, the majority of structural and antigenic variation of pilin between species

resides in the central and carboxy-terminal domains (Tennet and Mattick 1994, Mattick

2002).

In addition to PilA, minor pilin-like proteins have been identified in P. aeruginosa

(PilE, PilV, PilW, PilX, FimT, FimU). These pilin-like proteins resemble PilA by possessing the hydrophobic amino-terminal ∝-helical regions (Alm et al 1996, Alm and

Mattick 1995, Alm and Mattick 1996). The exact function of these proteins is not entirely known; however, they are required for twitching motility and are believed to form the base of the pilus fiber.

During biogenesis and assembly of pili, the PilA subunit is modified by PilD.

This protein has dual functions, acting first as a peptidase cleaving the pilin leader

sequence and then as a transmethylase adding a methyl group to the resulting N-terminal residue (Pepe and Lory 1998, Lory and Strom 1997, Strom and Lory 1993). Following pre-pilin modification, pilin is assembled on a base of the minor pilins through the actions of PilB and PilC (Nunn et al. 1990). PilB is an ATPase required for pili extension while PilC is believed to be required to cap or stabilize pili. The pilus is assembled through a gated pore in the outer membrane, composed of a dodecameric doughnut-shape

complex known as PilQ (Martin et al. 1993).

Retraction of the pilus is mediated through the action of PilT and PilU, PilB

homologs (Fig. 3) (Hobbs and Mattick 1993, Kaiser 2000, Mattick 2002). These

ATPases are responsible for the depolymerization of the pilus into its pilin subunits.

In P. aeruginosa, production of Type IV pili is controlled by a sensor-regulator

pair, PilS and PilR (Hobbs et al. 1993, Strom and Lory 1993). PilS acts as the sensor

protein autophosphorylating in response to a particular environmental stimulus. This

histidine kinase is localized to the poles (Boyd 2000). Following autophosphorylation,

PilS transfers its phosphate group to PilR, the response regulator. In its phosphorylated

form, PilR activates transcription of pilA by interacting with RNA polymerase and RpoN

(Fig. 4) (Darzins and Russell 1997). The specific stimulus responsible for turning on this

system, however, is unknown.

Chemotaxis and MCPs

Pseudomonas aeruginosa is a motile bacterium exhibiting chemotaxis to a wide

range of chemical stimuli (Hamilton and Sheely 1971, Kato et al. 1992, Kelly-

Wintenberg and Montie 1994, Moench and Konetzka 1978, Kearns et al. 2001). P. aeruginosa’s ability to chemotax towards, or away from, these chemical stimuli is mediated through chemotactic transducers known as MCPs, or methyl-accepting chemotaxis proteins. Through sequence homology, P. aeruginosa MCPs are believed to be transmembrane proteins resembling E. coli MCPs (Ferrandez et al. 2002, Stover et al.

2000). X-ray crystallography of E. coli MCPs has revealed that they are dimers consisting of a periplasmic and cytoplasmic domain (Fig. 5) (Kim et al. 1999, Milburn et al. 1991, Lefman et al. 2004). The periplasmic ligand binding domain of MCPs is responsible for detecting chemoeffector levels, while the cytoplasmic region is a signaling domain that undergoes reversible methylation of 4-6 glutamic acid residues

(Hazelbauer et al. 2008, Milburn et al. 1991, Springer et al. 1977). The methylation state of the MCP acts as molecular memory of the chemical environment.

MCPs of E. coli belong to a “two component” signal transduction pathway directing chemotactic responses. Within this pathway, six additional chemotaxis proteins act in concert with the MCP to direct cell movement (Fig. 6) (Madigan et al. 1997). The

MCP acts as a receptor binding the ligand. Upon binding the ligand, this protein undergoes a conformational change transducing a signal that controls two interrelated processes, excitation and sensory adaptation. Excitation is a process controlling the direction of cell movement in response to a chemical stimulus (Schulmeister et al. 2008,

Hess et al. 1988). During excitation in Escherichia coli, the MCP forms a ternary complex with CheA (histidine kinase) and CheW (linker protein). The transduced signal alters CheA activity by stimulating autophosphorylation. CheA-P acts as a phosphodonor to the response regulator protein, CheY (Tindall et al. 2010, Hess et al. 1988).

In E. coli and S. enterica, the flagella motor has been shown to have a default counterclockwise rotation. During counterclockwise rotation, peritrichous flagella form a bundle sheath causing the cell to swim in a single direction called a “smooth run”.

Unphosphorylated CheY directs counterclockwise rotation of the flagella. When

phosphorylated, CheY-P interacts with FliM, the flagella motor. This interaction triggers

the flagellum to rotate clockwise. Clockwise rotation of the flagella motor causes the

flagellar bundle to come apart causing the cell to “tumble”, thus changing direction. In

the presence of a chemoattractant, a bias is generated such that cells exhibit long runs

punctuated by periodic tumbles, thus allowing the cell to migrate in response to a

chemical gradient.

Sensory adaptation counteracts the excitation process by allowing the bacteria to

adapt to a sustained stimulus. This process is regulated by methylation and

demethylation of the MCP. In the presence of a positive stimulus, MCPs are methylated;

whereas, in the presence of a negative stimulus or absence of a positive stimulus MCPs

are demethylated. In E. coli, the methylation state of MCPs is controlled by two proteins,

CheR and CheB (Springer and Koshland 1997). CheR is a methyltransferase protein using an S-adenosylmethionine as a methyl source to methylate glutamate residues in the cytoplasmic region of the MCP (Springer and Koshland 1997). CheB is a methylesterase

that cleaves the methyl groups from the glutamate residues of the MCP (Bray 2002,

Stock and Koshland 1978). This protein is active when phosphorylated by CheA-P. In other words, methylation of the MCP favors the kinase-on signaling state, while demethylation of the MCP favors the kinase-off state.

In comparison to the signal transduction system in E. coli, there appears to be a

high level of diversity among motile bacteria. Most motile bacteria possess multiple

homologues of che-like genes such as the Rhodobacter sphaeroides, Sinorhizobium

meliloti, and Caulobacter crescentus. Pseudomonas aeruginosa has che-like genes

arranged in five clusters (Fig. 7) (Ferrandez et al. 2002). Cluster I, II, III, and V have been shown to be involved in swimming motility chemotaxis and cluster IV is directly related to twitching motility (Darzin 1994, Ferrandez et al. 2002, Kato et al. 1992, Kearns

et al. 2001, Masduki et al. 1995).

A comparative analysis of E. coli and P. aeruginosa has led to a model of the signal transduction pathway controlling twitching motility in P. aeruginosa (Darzins and

Russell 1997). In this model, an environmental signal stimulates the formation a ternary complex consisting of PilJ (MCP), PilL (CheA homologue), and PilI (CheW homologue)

(Fig. 4). This complex activates the kinase activity of PilL transferring a phosphate group to the response regulators, PilG and/or PilH (CheY homologues). These response regulators are believed to regulate pili biogenesis and twitching motility. In this model, sensory adaption is controlled by PilK and ChpB. PilK, a CheR homologue, is thought to act as a methyltransferase adding methyl groups to the MCP, while ChpB, a CheB homologue, is the methylesterase removing these methyl groups (Mattick 2002, Faulke

1997).

Studies have demonstrated there to be diversity between E. coli methyl-accepting chemotaxis genes. E. coli has five genes encoding the MCPs Tar, Tap, Trg, Tsr, and Aer.

Each of these MCPs binds and detects specific ligands: Tsr (serine), Tar

(aspartate/maltose), Tap (dipeptides), Trg (galactose/ribose), and Aer (oxygen).

Currently, 26 genes in P. aeruginosa have been identified as E. coli mcp homologue

genes; however, these genes have not been as extensively characterized (Croft et al. 2000,

Ferrandez et al. 2002, Kuroda et al. 1995, Stover et al. 2000, Wu et al. 2000).

Previous studies have demonstrated that P. aeruginosa exhibits chemotactic responses to L-amino acids, sugars, thiocyanic, isothiocyanic esters, and inorganic phosphate (Craven and Montie 1985, Kato et al. 1992, Moench and Konetzka 1978,

Moulton and Montie 1979). In addition, MCP methylation assays of P. aeruginosa have shown that an approximately 73 kDa protein is labeled with L-[methyl-3H]methionine in

response to L-serine, L-arginine, and ∝-aminoisobutyrate (Craven and Montie 1983).

This study demonstrated that chemotaxis towards these amino acids is mediated through

MCPs.

Studies using a P. aeruginosa mutant defective in chemotaxis to L-serine led to the identification of a MCP transducer gene designated pctA. This gene encodes for a

629 amino acid polypeptide with a calculated mass of 68,042 daltons. PctA also possesses structural features typical of MCPs. These features include two hydrophobic transmembrane domains, a hydrophilic periplasmic region, and a hydrophilic cytoplasmic domain. In addition, PctA has two potential methylation sites within the cytoplasmic domain. Significant homology to enteric MCPs was observed in the highly conserved domain (HCD) of this protein (Kuroda et al. 1995).

In Pseudomonas aeruginosa, PctA is directly involved in chemotaxis to L-amino acids. PctA has been shown to be specifically necessary for chemotaxis to L-glycine, L- serine, L-threonine, and L-valine. There also appears to be a reduction in chemotaxis to other L-amino acids in the pctA mutant (Kuroda et al. 1995).

In addition to pctA, two other che-like genes, mcpA and mcpB, have been

identified in P. aeruginosa as genes encoding for MCPs (Ferrandez et al. 2002). These

genes belong to cluster II genes and are involved in swimming/swarming motility. Both

mcpA and mcpB mutants demonstrate a general chemotaxis defect towards amino acids,

sugars, organic acids, and aromatic compounds in tryptone and LB soft agar plates. This

defect is complemented when mcpA and mcpB are expressed in trans in their respective mutants (Ferrandez et al. 2002). However, mcpA mutants were not deficient in chemotaxis and behaved liked wild-type cells in minimal-medium soft agar plates.

Like PctA, McpA and McpB possess certain structural features resembling E.

coli–like MCP architecture. McpA contains a HCD, but only one transmembrane

domain. McpB also contains a HCD, but, in contrast to McpA, does not have any

transmembrane domains commonly found in MCPs. This suggests that McpB exist as a

cytoplasmic MCP. Additionally, McpA and McpB are the only P. aeruginosa MCPs that

have C-terminal pentapeptides such as those associated with the high abundance

receptors of E. coli (Ferrandez et al. 2002). This structural analysis, along with

chemotaxis assays, supports the idea that McpA and McpB play a role in chemotactic

signal transduction in P. aeruginosa.

Pseudomonas aeruginosa not only possesses E. coli MCP homologues necessary

for chemotactic responses in swimming and swarming conditions, but also has one

required for twitching motility. This homologue, pilJ, belongs to the pilGHIJKL, cluster

IV, chemotactic system (Darzins 1994). Comparative analysis of PilJ with other MCPs

shows that this protein is most similar to E. coli MCPs. Further studies revealed that PilJ

is 26% identical to Tsr (Darzins 1994).

Like enteric MCPs (e.g. Tsr), PilJ consists of two hydrophobic amino acid

sequence acting as transmembrane domains. The first transmembrane region is located at

the N-terminus of the protein and the second transmembrane region is located near the

middle of the protein. In addition, PilJ and Tsr share similar Kyte-Doolittle

hydrophilicity-hydrophobicity plots. These plots show two large stretches of

hydrophobic amino acids, TM1 and TM2. TM1 is 25 amino acids long found near the N-

terminus and TM2 is 27 amino acids long found near the middle of the protein (Darzins

1994).

The strongest degree of homology between PilJ and Tsr is located in the C-

terminal domains, especially within the highly conserved domain (HCD). HCDs are

found in all MCPs and located near the middle of the cytoplasmic signaling domain

(Alley et al. 1992). Genetic evidence suggests that CheW directly interacts with this

domain and CheA to form a ternary complex during chemotaxis (Bhatnagar et al. 2010,

Lui and Parkinson 1991). In addition to the HCD, two methylation regions, K1 and R1,

are positioned in the cytoplasmic domain.

In contrast to the cytoplasmic domain, PilJ displays little similarity in the periplasmic domain to Tsr (Darzins 1994). Because the periplasmic domain is involved in ligand binding, this implies that Tsr and PilJ bind different ligands. Additional studies have shown that PilJ also does not possess a NIT domain present in several bacterial chemotaxis receptors, such as in E. coli receptors (Shu et al. 2003). NIT domain is a nitrate- and nitrite-sensing domain detected in various receptor components of signal transduction pathways. Even though the exact ligand(s) detected by PilJ has not been

determined, comparative analysis of PilJ has revealed that its ligand binding domain is most similar to E. coli canonical MCPs (I. Zhulin, personal communication).

Again, pilJ belongs to the pilGHIJKL operon and under the control of a promoter located upstream of pilG. Little is known about the regulation of pilJ; however, microarray analysis of P. aeruginosa revealed that pilJ is QS repressed (Wagner et al.

2003). In addition, studies have shown that genes containing a las box were usually negatively regulated by quorum-sensing. Sequence analysis of the pilJ gene has revealed the presence of a putative las box, although further tests are needed to confirm the function of this box (Wagner, personal correspondence).

PilJ varies from other MCPs in P. aeruginosa in its function. PilJ has been shown to be necessary for twitching motility, but is not required for swarming motility.

Macroscopic twitching motility assays showed that pilJ mutant cells do not exhibit a twitch zone. Also, microscopic observations of these cells revealed that they are non- motile and grow as compact colonies (Darzins 1994). In contrast to the wild-type PAO1, pilJ mutant cells do not fully extend functional pili and are present at both poles rather than one (DeLange et al. 2007). The presence of these shortened pili is also evident by the fact that pilJ mutant cells exhibit wild-type sensitivity to some pili specific phage

(PO4, B3, and F116) (Darzins 1994, Whitchurch et al. 2004). The presence of these shorten pili may also account for the existence of certain strains of P. aeruginosa (i.e.

PAK) in which pilJ mutant cells are able to twitch, but demonstrate aberrant twitching motility (Whitchurch et al. 2004). Interestingly, E. coli MCP mutants do not behave like

P. aeruginosa MCP mutants and remain motile. P. aeuginosa MCP’s have dual function and are involved not only in chemotaxis, but motility in general.

More recently it has been revealed that PilJ also negatively regulates swarming

motility (Caiazza et al. 2007). In frame deletion mutants of pilJ exhibit hyperswarming and increased flagellar reversals. These phenotypes are identical to those of a sadB

mutant strain. Epistasis analysis indicates sadB is genetically upstream of pilJ

demonstrating its ability to control swarming motility through the CheIV chemosensory

system.

Polar localization of chemoreceptor complexes in E. coli have shown that Tsr

receptors are located at the polar region of the cells (Maddock and Shapiro 1993). These

receptors are organized into groups of three, with extensive interaction between the dimer

interfaces. This clustering of receptors, referred to as a “trimer of dimers”, is thought to

be responsible for the cell’s high sensitivity to changes in the chemical environment and

wide dynamic range (Kim et al. 2002).

Furthermore, three dimensional electron microscopic imagining of the polar

region of E. coli revealed the presence of membrane invaginations. These invaginations

are believed to facilitate protein-protein interaction of chemoreceptors, such as Tsr

(Lefman 2004). Also, this three-dimensional network of membranes might stabilize the formation of CheA and CheW with the receptor. Such membrane invaginations, as well as, clustering of chemoreceptors has not been identified in P. aeruginosa. However, we

have shown PilJ to localize in the polar regions of the cell (DeLange et al. 2007). In addition, the pili and the pili assembly ATPases (PilB, PilT, and PilU) localize to cellular poles suggesting that there is clustering of chemoreceptor molecules and the presence of membrane invaginations in P. aeruginosa (Chiang et al. 2005).

Quorum-sensing in Gram-negative bacteria.

Quorum-sensing (QS) is a type of cell to cell communication used by bacteria to

coordinate population behavior through the use of quorum-sensing molecules (QSM)

known as autoinducers (Bassler 2002). These molecules are excreted by individual cells and increase in the environment with a corresponding increase in cell population. When a critical threshold concentration is achieved, specific target genes are expressed or repressed. This is known as population density dependent gene expression.

In Gram-negative bacteria, the main quorum-sensing molecules are known as acyl-homoserine lactones (AHL). This was first discovered in the marine bacterium

Vibrio fischeri in which quorum-sensing is responsible for the regulation of bioluminescence (Nealson et al. 1970). Gram-negative quorum-sensing circuits are composed of two regulatory proteins: LuxI and LuxR (Fig. 8) (Miller and Bassler 2001).

LuxI proteins are responsible for the synthesis of AHL while the LuxR proteins act as the transcriptional regulator. AHL’s directly interact with LuxR, which in turn binds to specific promoter regions of QS regulated genes. Pseudomonas aeruginosa is composed of two pairs of LuxRI homologues, LasRI and RhlRI (Fig. 9) (Miller and Bassler 2001).

LasI and RhlI are autoinducer synthases that catalyze the formation of 3-oxo-C12HSL and

C4HSL, respectively (Fig. 10) (Gambello and Iglewski 1991, Passador et al. 1993,

Lazdunski et al. 2004, Juhas et al. 2005). At high cell density, LasR binds its QSM

cognate 3-oxo-C12HSL. Together this complex binds to the las box in the promoter

region preceeding genes encoding for the expression of virulence factors. This LasR-

C12HSL complex induces the expression of lasI creating a positive feedback loop (Seed

et al. 1995).

LasR-C12HSL also positively regulates the expression of rhlR (Ochsner and

Reiser 1995). This gene encodes for the transcriptional regulator, RhlR, which binds to

its cognate C4HSL and activates the expression of several target genes. Similar to the

LasR/I system, RhlR-C4HSL induces the expression of rhlI producing a positive feedback loop. These two systems work in tandem with one another to control the expression of

several virulence factors (i.e. alkaline protease, elastase, pyocyanin) and coordinate

bacterial behavior (i.e. surface motility and biofilm formation) (Miller and Bassler 2001).

Besides activating the expression of RhlR, C12HSL has been shown to inhibit the binding

of C4HSL to RhlR (Pesci et al. 1997). This regulation allows for the sequential activation

of each signaling system.

In addition to the LasRI and RhlRI systems, P. aeruginosa also possesses another

signaling molecule known as 2-heptyl-3-hydroxy-4-quinolone (PQS) (Fig. 10) (Pesci et al. 1999, Juhas et al. 2005). PQS, or the Pseudomonas quinolone signal, belongs to a group of molecules called 4-hydroxy-2-alkylquinoline (HAQs) known for their antimicrobial activity (Deziel et al. 2004). This system has been shown to regulate the production of several virulence factors such as pyocyanin, elastase, and rhamnolipids

(Pesci et al. 1999, Calfee et al. 2001, Deziel et al. 2003, Deziel 2004). PQS has

specifically been detected in the lungs of cystic fibrosis patients (Collier et al. 2002). The

PQS system is under the control of both the LasRI and RhlRI systems (Fig. 11) (Venturi

2006).

PQS is synthesized from anthranilate. Initially, the cell converts chorismate to this precursor of PQS through the phnAB operon (Cao et al. 2001, Deziel et al 2004).

This operon is responsible for the production of anthranilate synthase. Anthranilate is

converted to 4-hydroxy-2-heptyl-quinoline (HHQ) via the action of pqsABCD. HHQ is

an intercellular signaling molecule released by the cell and taken up by another cell

where it is converted into PQS by the gene product of pqsH (Deziel et al 2004).

The PQS system is under the control of PqsR. This protein directs transcription

of the phnAB and pqsABCD operons (Gallagher et al. 2002, Deziel et al. 2004). As

previously stated, this system is also governed by the Las and Rhl systems. LasR

positively regulates pqsH and pqsR (Gallagher et al. 2002, Wade et al. 2005). In terms of

the Rhl system, PQS has been shown to directly interact with RhlR to control the

transcription of specific genes such as one responsible for the production of elastase, lasB

(Pesci et al. 1999).

Several factors control quorum-sensing in P. aeruginosa (Fig. 11) (Venturi 2006).

A further layer of complexity is added to P. aeruginosa quorum-sensing with the existence of two orphan cognate LuxR-like regulators, QscR and VsqR. These two LuxR regulators do not possess cognate QSM. QscR has been shown to regulate LasR/RhlR, whereas VqsR positively regulates quorum-sensing through the Las system (Venturi

2006). At low cell densities there is a corresponding low concentration of the QSM’s 3- oxo-C12HSL and C4HSL. QscR acts on the Las and Rhl systems by binding to the transcriptional regulators LasR and RhlR. This binding prevents the interaction of the transcriptional regulator with its cognate QSM and inhibits its ability to bind to the promoter regions of quorum-sensing controlled genes. As the concentration of AHLs

increase, QscR-AHL interactions occur, promoting the dissociation of the heterodimers

QscR/LasR and QscR/RhlR, thus promoting LasR/3-oxo-C12HSL and RhlR/ C4HSL

interactions. The competitive binding of QscR to LasR and RhlR prevents the premature

production of several QS controlled products such as elastase, pyocyanin, and HCN

(Chugani et al. 2001, Ledgham et al. 2003).

VqsR positively controls quorum-sensing in P. aeruginosa through the expression of the autoinducer synthase, LasI (Juhas et al. 2004, 2005). vqsR knock-out mutants do not produce AHLs and, therefore, produce fewer virulence factors. In addition, these mutants demonstrate a reduced pathogenicity in a nematode infection model system

(Juhas et al. 2004).

Farnesol

Candida albicans is an opportunistic pathogen that is a part of the normal microflora of humans. This fungus has two distinct morphologies: yeast and filamentous

(mycelium). Yeast cells are typical involved in commensal interactions while the filamentous form is important in opportunistic infections (Gow et al. 2002). This is evident by the presence of the predominance of the filamentous form in deep areas of infected tissue and the reduction in host mortality when infected with mutants impaired in their ability to filament in the mouse model (Lo et al. 1997).

Several environmental factors can trigger this yeast to mycelium dimorphism including change in pH, temperature, nutrient availability, inoculum size, and the presence of chemicals that stimulate or repress that filamentous pathway (Brown 2002).

Among those chemicals responsible for repressing the filamentous pathway is E,E- farnesol. This volatile sesquiterpene is produced by C. albicans and was the first quorum-sensing molecule identified in eukaryotes (Hornby et al 2001). In C. albicans,

E,E-farnesol has been shown to inhibit the conversion of yeast to mycelium, as well as affect biofilm formation (Hornby et al. 2001, Mosel et al. 2005).

It has been shown that there is a positive correlation between the presence of P.

aeruginosa and C. albicans in infected hosts (Hermann et al. 1999). In addition, P. aeruginosa has been shown to limit the growth of C. albicans (Kerr et al. 1994).

Interestingly, antibiotic treatment and irradication of P. aeruginosa infections in CF patients results in an increase in C. albicans population number in infected tissue (Burns et al. 1999). Research examining this interspecies interaction have shown that P. aeruginosa QSM 3-oxo-C12HSL mimics E,E-farnesol by repressing C. albicans filamentation without affecting cell growth (Hogan et al. 2004). Furthermore, when filamentous C. albicans is exposed to 3-oxo-C12HSL, it reverts back to its yeast

morphology. 3-oxo-C12HSL’s ability to functionally mimic E,E-farnesol has been linked

to the structural similarity between these two compounds (Hogan et al. 2004). Both 3-

oxo-C12HSL and E,E-farnesol possess a 12-carbon chain length moiety in common (Fig.

12). Other C12 compounds such as dodecanol have similar antagonistic effects on filamentous C. albicans while acylated homoserine lactones of varying carbon backbones lengths do not.

Recent studies have indicated that P. aeruginosa is not only capable of communicating with C. albicans through 3-oxo-C12 HSL, but that this communication is

bi-directional with E,E-farnesol affecting P. aeruginosa virulence and community

behavior. In the presence of E,E-farnesol, production of QSM PQS and the PQS-

controlled virulence factor, pyocyanin are reduced (Cugini et al. 2007). E,E-farnesol

stimulates a non-productive interaction of PqsR to the promoter region of pqsA, the first

gene in the biosynthesis pathway of PQS. This interaction results in reduced transcript

levels of pqsA.

In addition to affecting the virulence factor pyocyanin, E,E-farnesol inhibits swarming motility in P. aeruginosa (McAlester et al. 2008). This inhibition was demonstrated in a number of CF isolates as well as the laboratory strain PAO1; however, the mechanism has yet to be determined. Despite the ability of E,E-farnesol to prevent swarming motility, it does not alter the cells ability to attach to surfaces (McAlester et al.

2008). In contrast to these results, E,E-farnesol inhibits biofilm formation of mixed cultures of P. aeruginosa, Enterococcus faecalis, and Staphylococcus aereus (Dowd et al.

2009).

Additional studies have been conducted examining the interspecies interaction of

C. albicans with several fungal and bacterial species through its QSM, E,E-farnesol.

E,E-farnesol has specifically been shown to increase susceptibility of Escherichia coli and Staphylococcus aereus to antibiotics by increasing cell permeability (Brehm-Stecher and Johnson, 2003; Inoue et al., 2004; Jabra-Rizk et al., 2006). In addition, E,E-farnesol induces apoptosis in the fungus Aspergillus nidulans (Semighini et al. 2006).

Biofilm formation

Biofilms are complex communities of bacterial cells within a hydrated matrix.

Several bacteria possess the ability to form these communities. Among those species are

Escherichia coli, Pseudomonas aeruginosa, Staphylococcus aureus, Staphylococcus epidermidis, Neisseria gonorrhoeae, Streptococcus mutans, and Burkholderia cepacia.

Biofilms are both environmentally and medically significant. Within the environment, biofilms are found on rocks in streams and rivers, as well as, in extreme environments

(e.g. hot springs and frozen glaciers). Biofilms are also beneficial for bioremediation processes in terms of eliminating oil spills and sewage treatment; however, they can be

harmful in food preparation and the clogging and corrosion of water and oil pipelines.

Biofilms are medically significant by attaching to abiotic (e.g. stints and catheters) and biotic (e.g. urinary tracts and tissues) surfaces (Costerton et al. 1999, Donlan and

Costerton 2002, Speer et al. 1988).

There are five distinct stages of biofilm formation in P. aeruginosa (Fig. 13)

(Monroe 2007). The first stage of biofilm development is initial attachment. During

initial attachment, planktonic cells swim towards the intended surface through the

rotation of their flagella and attach themselves. This attachment is dependent on several

factors such as hydrophobicity between the cell and surface, electrostatic interactions,

temperature, and steric hindrances (Dunne 2002). Following initial attachment, the cell

transitions to irreversible attachment. Irreversible attachment is mediated through the gene sadB and is dependent upon Type IV pili and other adhesins (Caiazza and O’Toole

2004).

After attachment, the next stage of biofilm formation is maturation I or microcolony formation. During maturation I, cells begin to replicate and spread across the surface through the extension and retraction of Type IV pili. The genes gacA and pilB are required for microcolony formation (Parkins et al. 2001, Sauer et al. 2002). gacA encodes for a response regulator belonging to the two component regulatory system

GacA/GacS and, as previously discussed, pilB encodes for an ATPase responsible for

Type IV pili extension.

Microcolonies eventually differentiate into mature biofilms. This stage is known as maturation II. These macrocolonies are characterized by a mushroom shaped architecture and consists of multiple channels through which nutrients and waste are

transported (Sauer et al. 2002). The final stage of biofilm development is dispersion.

During this stage, cells are released from the mature biofilm either passively or actively.

These cells can then swim and recolonize another surface thus forming a new biofilm.

Mature biofilms are composed of exopolysaccharides (EPS), proteins, and extracellular DNA. EPS constitutes 50-90% of the total organic carbon of biofilms and is highly hydrated (Flemming et al. 2000). EPS is primarily composed of polysaccharides.

For Gram-negative bacteria, these polysaccharides can be either neutral or anionic.

Anionic polysaccharides such as uronic acids (e.g. D-glucuronic, D-galacturonic, and mannuronic acids) or ketal-linked pryruvates can bind the divalent cations calcium or magnesium. This cross-linking between polysaccharides and cations facilitate better binding force for the biofilm (Flemming et al. 2000).

The loci psl, pel, and alg, are responsible for the production of EPS in P. aeruginosa (Evans and Linker 1973, Friedman and Kolter 2004, Matsukawa and

Greenberg 2004). The psl genes pslA and pslB are involved in attachment and biofilm formation (Friedman and Kolter 2004). Genes belonging to the alg loci are directly involved in the production of capsular polysaccharide alginate. Alginate is a major component of mucoid strains and is studied extensively because of its importance in the

CF lung (Gacesa 1998, Govan and Deretic 1996, Govan and Fyfe 1978). P. aeruginosa strains isolated from CF lungs produce high quantities of this polysaccharide. Because most natural occurring isolates are non-mucoid and produce little alginate, conversion to the mucoid state is believed to be important in the pathogenesis of CF patients.

As previously mentioned, EPS is highly hydrated and is able to incorporate large quantities of water through hydrogen bonding. This characteristic protects the biofilm by

preventing dessication and inhibits the diffusion of antibiotics across the matrix (Donlan

2000).

Extracellular DNA is also a significant component of the biofilm structure.

Extracellular DNA was previously believed to be released during cellular lysis; however, it has been shown that this extracellular DNA is acquired through a process independent of this mechanism. In P. aeruginosa, this mechanism appears to involve the release of

membrane vesicles from the outer membrane (Schooling et al. 2009).

Studies examining the function of extracellular DNA in biofilm formation

demonstrated that when grown in the presence of DNase I, P. aeruginosa was unable to form biofilms. In addition, 12, 36, and 60 hour old biofilms treated with DNase I were dissolved, whereas 84 hour old biofilms were affected to a minor degree. These results indicate that extracellular DNA is required for the initial establishment of P. aeruginosa biofilms (Whitchurch et al. 2002).

Recent studies have also shown that extracellular DNA contributes to the overall architecture of the biofilm (Allesen-Holm et al. 2006). In comparison to the wild-type strain, lasIrhlI, pqsA, and fliMpilA mutant biofilms contain reduced amounts of extracellular DNA and have an altered biofilm structure. pqsA mutant biofilms do not exhibit the mushroom shaped pattern of wild-type biofilms and are thin and flat. In

addition, fliMpilA mutant biofilms form irregular microcolonies.

Besides contributing to the establishment and architecture of biofilms,

extracellular DNA also induces the host immune response. Studies have shown that

extracellular DNA activates neutrophils. Also, P. aeruginosa biofilms treated with

DNase I are reduced in their ability to stimulate the release of cytokines IL-8 and IL-

1beta and exhibit a reduction in the upregulation of neutrophil activation markers CD18,

CD11b, and CD66b from the host immune system (Fuxman Bass et al. 2010). In addition, lasIrhlI mutant strain biofilms, known for having a low content of extracellular

DNA, are inhibited in their capacity to stimulate the release of proinflammatory cytokines by neutrophils. These findings indicate that extracellular DNA is a significant proinflammatory component of P. aeruginosa biofilms.

Photodynamic Therapy

Photodynamic therapy (PDT) is a type of therapy that combines the use of visible light and a photosensitizer (PS) to generate cytotoxic reactive oxygen species and free radicals. PDT has previously been used in the treatment of certain cancers and other diseases such as macular degeneration. In the last decade, there has been growing interest in the use of this type of therapy to treat bacterial infections. The application of this therapy in terms of microorganisms is known as photodynamic antimicrobial chemotherapy (PACT).

During PACT, the PS absorbs light and is excited from ground state to an excited singlet state. In this state, the excited PS experiences an electron spin change and is

converted to an excited triplet state (Wainwright 2008). In the presence of oxygen, the

excited PS can undergo two different reactions: Type I and Type II reaction. During

Type I reaction, electrons are transferred from the excited PS to oxygen in the immediate

environment resulting in the production of reactive oxygen species (ROS) (i.e. hydroxyl

radicals and superoxide). In contrast, there is a direct transfer of energy from the triplet

PS to oxygen in Type II reaction. This transfer of energy results in the formation of

1 singlet oxygen ( O2) (Fig. 14) (Wainwright and Crossley 2004, Robertson et al. 2009).

Reactive oxygen species produced by Type I and II reaction are responsible for

photodamage of cellular components. Due to morphological differences between

microbial species, the mode of action of a photosensitizer towards cellular components

varies from species to species. Type I reaction has specifically been shown to affect

membrane permeability through lipid peroxidation. Lipid peroxidation leads to loss of

fluidity and ion permeability (Korytowski 1992). Singlet oxygen produced by Type II

reaction has also been shown to target cell wall/cell membrane components such as

lipids, sterols, and peptides. Products from these reactions vary. For instance, during

Type I reaction cholesterol is converted to cholesterol-7α and 7β-hydroperoxide;

1 whereas, cholesterol is converted to 5α-isomer by O2 produced by Type II reaction

(Girotti 1990).

In addition to targeting cell wall/membrane components, nucleic acids are also

targeted during PACT. Hydroxyl radicals created by Type I processes attack the sugar

1 moiety of a nucleic acid. In contrast, O2 produced by Type II processes target the guanine base (Foote 1990). These cytotoxic events lead to base substitution, strand

cleavage, mutation, and inhibition of replication.

The photodynamic mode of action is usually determined by the class of

compound. Photosensitizers are typically aromatic molecules capable of forming the long-lived triplet state. The amount of energy required for each photosensitizer to

1 produce O2 is dependent of the molecular structure of that compound (Fig. 15)

(Wainwright 1998).

There are several different classes of photosensitizers such as cationic azine

photosensitizers (e.g. methylene blue, acridine orange, toluidine blue O); cyanines and

merocyanine 540 (e.g. pyrvinium and stilbazium); and, macrocyclic photosensitizers (e.g.

porphyrins and phthalocyanines). Among macrocyclic photosensitizers is a group of compounds known as porphyrins. Porphyrins are heterocyclic macrocycles distinguished by four pyrrole subunits. Because these molecules are highly conjugated, they have intense absorption between the wavelengths 600-650nm (Wainwright 1998).

Porphyrins are characterized by their ability to bind free O2. This family of

compounds includes naturally-derived porphyrins such as chlorophyll and heme. These

naturally occurring porphyrins have been shown to have bactericidal effects particularly

on a range of anaerobic bacteria. In contrast to chlorophylls, haemin demonstrates

reactivity in the absence of light (Nitzan et al. 1994).

In addition to naturally-derived porphyrins, the effects of synthetic porphyrins on bacterial viability have also been examined. Studies involving meso-substituted porphyrins, have shown cationic derivatives to be more successful than anionic derivatives at photoinactivating Gram-negative bacteria (Merchat et al. 1996). For instance, the anionic haematoporphyrin only photoinactivates Gram-negative following permeabilization of the outer membrane (Malik et al. 1992). In contrast, the cationic photosensitizer 5-phenyl-10,15,20-tris(N-methyl-4-pyridyl)porphyrin (TriP[4]) has been

demonstrated to successfully inactivate Gram-positive, Gram-negative, and fungi in the

absence of membrane permeabilizers (Lambrechts et al. 2005, Merchat et al. 1996).

Also, studies examining the effects of the cationic porphyrin 5,10,15,20-tetrakis(1- methyl-pyridino)-21H,23H-porphine, tetra-p-tosylate salt (TMP) have shown it to photoinactivate Staphylococcus aureus and Pseudomonas aeruginosa (Fig. 16) (Di Poto et al. 2009, Donnelly et al. 2007).

Besides exhibiting cytotoxic effects on planktonic cells, various photosensitizers have been determined to photoinactivate biofilms. The phenothiazinium photosensitizer new methylene blue specifically targets P. aeruginosa biofilms by photoinactivating bacterial cells and through the degradation of exopolysaccharides (Wainwright et al.

2002). Studies in our lab, as well as other labs, have shown P. aeruginosa biofilms to be disrupted by TMP and have suggested that in addition to photoinactivating biofilm associated cells, extracellular DNA is a target of this porphyrin (Donnelly et al. 2007,

Collins et al. 2010). This is further supported by studies showing TMP’s ability to intercalate in between DNA base pairs, causing photoinduced strand breakage when irradiated (Kelly and Murphy 1985, Pasternack and Gibbs 1996, Collins et al. 2010).

Additionally, TMP is known to significantly reduce S. aureus survival and, when combined with antibiotics, disrupt established biofilms (Di Poto et al. 2009). Interaction of these photosensitizers with bacterial cells and biofilm components indicate the potential clinical application of them in the treatment of bacterial infections.

Figure 1. The bacterial flagellum (Yonekura et al. 2000). HAP1 and HAP3: junction between the hook and filament, HAP2: filament cap, OM: outer membrane, PG: peptidoglycan layer, IM: cytoplasmic membrane, FliF: S-ring.

Figure 2. Model of the structure of type IV pilin monomer in Pseudomonas aeruginosa (Mattick 2002). Blue α-helices: highly conserved, hydrophobic amino terminal domain; Green and pink β-sheets: hydrophilic central and carboxy-terminal domain.

Figure 3. Model of type IV pili assembly and retraction in Pseudomonas aeruginosa

(Mattick 2002). PilA: pilin subunit; PilE, PilV, PilW, PilX, FimU: minor pilin subunits;

PilD: pilin peptidase and transmethylase; PilB: ATPase responsible for pili extension;

PilC: pili cap; PilQ: gated pore; PilT: ATPase responsible for pili retraction.

Figure 4. Model of the main regulatory networks governing pilus biogenesis and twitching motility in Pseudomonas aeruginosa (Darzins and Russell 1997). PilJ:

MCP; PilK: transmethylase; PilI: linker protein; PilL: histidine kinase; PilG and PilH: response regulators; PilS: environmental stimuli sensor protein; PilR: environmental stimuli response regulator; RpoN: alternative sigma factor; pilD: gene encoding pilin peptidase and transmethylase; pilC: gene encoding pili cap; pilB: gene encoding ATPase responsible for extension; pilA: gene encoding pilin subunit.

Figure 5. Structural model for the full-length Tsr dimer based on structures of the

periplasmic and cytoplasmic domains (Lefman et al. 2004). X-ray crystallography of

E. coli MCPs has revealed that they are dimers consisting of a periplasmic and cytoplasmic domain. The periplasmic domain is responsible for detecting the chemoeffector levels, while the cytoplasmic domain is a signaling domain that undergoes reversible methylation of 4-6 glutamic acid residues. The methylation state of the MCP acts as molecular memory of the chemical environment.

Figure 6. Chemotaxis signal transduction pathway in Escherichia coli (Madigan et

al. 1997). MCP: methyl-accepting chemotaxis protein; CheR: methyltransferase; CheB: methylesterase; CheW: linker protein; CheA: histidine kinase; CheY: response regulator;

CheZ: phosphatase.

Figure 7. Chemotaxis genes in Pseudomonas aeruginosa (Ferrandez et al. 2002).

Chemotaxis genes are arranged in five gene clusters. pilJ, a gene encoding for a methyl- accepting chemotaxis protein (MCP), belongs to Cluster IV genes. Cluster IV genes are associated with twitching motility.

Figure 8. Quorum-sensing system of bioluminescence in Vibrio fischeri (Miller and

Bassler 2001). LuxI (square) is the AHL synthase responsible for the production of the autoinducer N-(3-oxohexanoyl)-homoserine lactone (hexagons). At a critical population density, the autoinducer binds to the transcriptional activator, LuxR (circles). The LuxR- autoinducer complex activates transcription of the luxICDABE operon involved in

bioluminescence.

Figure 9. The LasRI-RhlRI quorum-sensing systems in Pseudomonas aeruginosa

(Miller and Bassler 2001). LasI is the AHL synthase (square) responsible for the

production of the autoinducer N-(3-oxododecanoyl)-homoserine lactone (triangles). At a critical cell population, N-(3-oxododecanoyl)-homoserine lactone binds to LasR (circles),

the transcriptional activator. This LasR-autoinducer complex positively regulates the

transcription of several virulence genes, as well as, rhlR. RhlI is the AHL synthase

(square) responsible for the production of N-(butryl)-homoserine lactone (pentagons). At a critical cell concentration, N-(butryl)-homoserine lactone binds the transcriptional regulator RhlR. This RhlR-autoinducer complex also positively regulates the transcription of several virulence and target genes.

Figure 10. Structure of P. aeruginosa quorum-sensing molecules (Juhas et al.

2005). 3-oxo C12-HSL: quorum-sensing molecule of the LasRI circuit; C4-HSL: quorum-

sensing molecule of the RhlRI circuit; PQS, or 2-heptyl-3-hydroxy-4-quinolone:

Pseudomonas quinolone signal known for controlling several genes associated with

antimicrobial activity.

Figure 11. Regulation of Las and Rhl systems quorum-sensing systems in P. aeruginosa (Venturi 2006). The Las system is at the top of the hierarchy controlling the

Rhl and PQS systems. Arrows indicate positive gene regulation; whereas, short parallel lines indicate negative gene regulation.

Figure 12. C12 compounds causing inhibition of C. albicans filamentation (Hogan et al. 2004). 3-oxo-C12 homoserine lactone (3OC12HSL), C12 homoserine lactone

(C12HSL), dodecanol, and farnesol inhibit filamentation of C. albicans at concentrations

<200 µM.

Figure 13. Five stages of biofilm development of P. aeruginosa (Monroe 2007).

Stage 1: initial attachment; Stage 2: irreversible attachment; Stage 3: maturation I; Stage

4: maturation II; and, Stage 5: dispersion.

Figure 14. Photosensitisation pathways for photosensitizer (Wainwright and

Crossley 2004). Photosensitisation of a photosensitizer (PS) can result either in the transfer of electrons to oxygen (Type I reaction) resulting in reactive oxygen species

(ROS) or the direct transfer of energy to oxygen (Type II reaction) resulting in the

1 formation of singlet oxygen ( O2).

Figure 15. Photosensitizer absorption maxima (Wainwright 1998).

Figure 16. Molecular structure of 5,10,15,20-tetrakis(1-methyl-pyridino)-21H,23H- porphine (TMPyP) (Snyder et al 2006). TMP is a porphyrin composed of four pyrole subunits and four phenyl groups.

Chapter II

E,E-farnesol inhibits surface motility in P. aeruginosa through rhamnolipid production

(Portion to be submitted for publication)

Abstract

E,E-FARNESOL INHIBITS SURFACE MOTILITY IN P. aeruginosa THROUGH RHAMNOLIPID PRODUCTION

Name: Collins, Tracy Lynn University of Dayton

Advisor: Dr. Jayne B. Robinson

Pseudomonas aeruginosa and Candida albicans both exhibit cell-to-cell communication

through the use of quorum-sensing molecules (QSM) known as autoinducers. Because

there is a positive correlation between the presence of P. aeruginosa and C. albicans in

opportunistic infections, we examined whether the QSM of one organism can affect the

other. Previous research has shown that P. aeruginosa QSM cognate, 3-oxo-C12 HSL,

mimics C. albicans QSM cognate E,E-farnesol by preventing the conversion of yeast to

mycelium. These results suggest that P. aeruginosa is capable of communicating with C.

albicans through 3-oxo-C12 HSL. Previous research in our lab examining the effects of

E,E-farnesol on P. aeuginosa revealed that E,E-farnesol substantially inhibited

production of QSMs. Because twitching and swarming motility are both quorum-sensing

controlled in P. aeruginosa, we examined the effect of E,E-farnesol on each. Twitching

and swarming motility were both decreased when wild-type PAO1 cells were exposed to

25, 100, and 250 µM E,E-farnesol. Interestingly, pilJ mutant cells retained there ability to swarm in the presence of E,E-farnesol. Expression studies showed that E,E-farnesol did not affect pilJ expression and in-vivo 3H-methylation of PilJ in the presence of E,E-

farnesol was inconsistent. In addition, there was an increase in rhamnolipid production when cells were grown in the presense of E,E-farnesol. These biosurfactants are known to regulate swarming motility in P. aeruginosa. In response to 250 µM E,E-farnesol,

there was a 4.0-fold and a 23.2–fold increase in rhamnolipid production of wild-type

PAO1 and pilJ mutant cells, respectively. Changes in the rhamnolipid concentrations

could account for the inhibition of swarming motility observed in the presence of E,E-

farnesol.

Introduction

Pseudomonas aeruginosa is a gram-negative, rod-shaped bacterium. This

organism is ubiquitous, commonly inhabiting soil and aqueous environments, as well as

in animal and plant tissues (Wilson and Dowling 1998). Because it possesses the ability

to utilize a wide range of organic compounds as a sole source of carbon, P. aeruginosa

can adapt and thrive in a variety of nutrient environments. P. aeruginosa is an

ecologically significant bacterium and is important to the nitrogen cycle, specifically

denitrification and nitrate assimilation (Zumft 1997). This bacterium has also proven to

be beneficial in bioremediation by degrading specific contaminants such as chlorinated

pesticides and crude oil (Freedman et al. 2004, Zhang et al. 2005).

P. aeruginosa is not only an ecologically important bacterium, but is classified as

an opportunistic pathogen and is responsible for a high percentage of nosocomial

infections (Emori and Gaynes 1993). As an opportunistic pathogen, P. aeruginosa

exploits the host defenses to initiate infection and has been implicated in infections of the

urinary tract, respiratory system, soft tissue, gastrointestinal, dermatitis, bacteremia, and a

variety of other systemic infections. Pseudomonas aeruginosa is equipped with a

substantial number of virulence factors that disrupt the host immune system and can

cause extensive tissue damage (Kharazmi 1991, Woods and Iglewski 1983).

Immunocompromised patients, such as individuals suffering from neutropenic cancer or HIV, as well as those with severe burns, are highly susceptible to P. aeruginosa

infections. Additionally, P. aeruginosa causes severe lung infections in cystic fibrosis patients and is a leading cause of mortality in this population (Wagner et al. 2003). In addition to humans, P. aeruginosa is a pathogen of a wide range of animal and plant species, including mice, fruit flies, nematode worms, and mustard plants (D’Argenio et al. 2002, Hahn 1997, Rashid and Kornberg 2000, Tennet and Mattick. 1994).

Pseudomonas aeruginosa is distinguished by its ability to attach to both abiotic

(e.g. medical devices) and biotic (e.g. eye epithelial cells) surfaces. Following attachment to the surface, cells have the potential to colonize and form mature biofilms.

Biofilms are encased, microbial colonies attached to a surface and are considered a major contributor to infection. Within these biofilms, bacteria are protected from the effects of antibiotics and attack by the host immune system (Costerton 2001). Mucus blocking the airway passages of cystic fibrosis patients creates an anaerobic, nutrient rich environment ideal for the propagation of bacterial growth and biofilm formation (Lyczak et al. 2000,

Hassett et al. 2009). Because of the medical implications of P. aeruginosa infections, it is important to understand the mechanism of biofilm formation and function.

Two cellular components, flagella and Type IV pili, are necessary for the formation of biofilms (O’Toole and Kolter 1998). Flagella and Type IV pili are extracellular filaments that mediate both cell motility and adhesion. During biofilm formation, bacterial cells are propelled towards a surface by the rotation of the flagella in a liquid environment. Once at the surface, the bacterial cell will attach itself to the surface via its flagella and Type IV pili, as shown in Figure 1 (O’Toole and Kolter 1998).

Through the repeated extension and retraction of the pili, cells translocate across the surface and aggregate into microcolonies. Microcolonies eventually differentiate into

mature biofilms (O’Toole and Kolter 1998). The process by which a bacterium

translocates across a solid surface through the extension and retraction of pili is known as

twitching motility. Strains deficient in genes required for the synthesis of Type IV pili are unable to form microcolonies. Also, flgK mutant cells are unable to attach to the

surface (O’Toole and Kolter 1998). These results indicate that Type IV pili and flagella

play a role in initial attachment and early biofilm formation.

Many bacteria demonstrate the ability to communicate using quorum-sensing

systems. P. aeruginosa possesses one of the most well understood quorum-sensing

models which is composed of the las and rhl systems (Pesci et al. 1997). During

quorum-sensing the bacterial species is able to sense critical population density through

the use of quorum-sensing molecules (QSM) known as autoinducers or, in the case of P.

aeruginosa, acyl-homoserine lactones (AHLs). These molecules accumulate in the

environment during high cell density and diffuse into surrounding cells. When an

intracellular threshold of autoinducers is reached, these molecules activate certain target

genes. Several of these target genes encode for virulence factors, such as exoproteases

and pyocyanin (de Kievit and Iglewski 2000). There is evidence that quorum-sensing

influences biofilm formation (Davies et al. 1998), as well as twitching and swarming

motility (Wagner et al. 2003).

Swarming motility is movement across a semi-solid surface via the rotation of flagella. Swarming cells of P. aeruginosa are usually elongated and possess two polar

flagella (Kohler et al. 2000). During swarming motility, P. aeruginosa cells migrate as

defined groups forming tendril patterns. This type of surface motility is dependent on

biosurfactants known as rhamnolipids. Rhamnolipid production is quorum-sensing

controlled and regulated through the rhl system (Kohler et al. 2000). rhlI and rhlR mutants are unable to swarm.

Recently, an extracellular molecule was discovered to mediate quorum-sensing systems in the polymorphic fungus, Candida albicans (Hornby et al. 2001). This QSM,

E,E-farnesol, prevents the yeast-to-mycelium conversion, resulting in actively budding yeasts. C. albicans is an opportunistic pathogen whose cellular morphology is associated with its virulence (Gow et al. 2002). The yeast form is predominantly involved in commensal interactions, whereas, the mycelium form is common in opportunistic infections. Because there is a positive correlation between the presence of P. aeruginosa and C. albicans within biological fluids of human origin, it is important to determine how and when these two organisms communicate and interact with each other (Hermann et al.

1999).

A recent study demonstrated that the P. aeruginosa QSM cognate, 3-oxo-C12

homoserine lactone, mimics E,E-farnesol by preventing the conversion of yeast to

mycelium (Hogan et al. 2004). These results suggest that P. aeruginosa is capable of

communicating with C. albicans through 3-oxo-C12 HSL. In order to determine if the

reverse is true, the effects of E,E-farnesol on P. aeruginosa AHL production were

assessed in our lab by separation using thin layer chromatography (TLC) and detection

with the Agrobacterium tumefaciens, NTL4 lacZ-based AHL reporter strain. TLC

analysis revealed that E,E-farnesol substantially inhibited production of QSMs by P.

aeruginosa cells. Because twitching and swarming motility are both quorum-sensing

controlled in P. aeruginosa, we examined the effect of E,E-farnesol on each. In this

study, we demonstrated that E,E-farnesol inhibits both twitching and swarming motility to varying degrees and examine the mechanism of that inhibition in P. aeruginosa.

Methods

Bacterial strains and growth conditions. Strains and plasmids used in this study are listed in Table 1. For broth cultures, P. aeruginosa was either grown in Luria-Bertani

(LB) medium (Difco, Detroit, Michigan) or PPGAS minimal medium (Wild et al. 1997).

PPGAS medium consists of NH4Cl (20 mM), KCl (20 mM), Tris-HCl (pH 7.2) (120

3 mM), MgSO4 (1.6 mM), glucose (0.5%), and peptone (1.0%). For in vivo H- methylation, cells were grown overnight in Minimal Salts medium (40 mM K2HPO4, 20

-3 mM KH2PO4, 7.6 mM [NH4]2SO4, 0.2 mM MgSO4 · 7 H2O, 9.2 x 10 mM FeCl3 · 6

H2O, adjusted to pH 7.0) supplemented with 0.04% (w/v) succinate and 0.5 mM L-

methionine. Antibiotics were added as necessary at the following concentrations: 60

µg/ml tetracyclin or 60 µg/ml carbenicillin. E,E-farnesol was purchased from Sigma-

Aldrich. Fresh stocks were prepared in methanol for every use.

Twitching motility assays. (i) Macroscopic. Twitching motility assays were

performed as previously described with modifications (Darzins 1994). LB (1% agar)

plates were stab inoculated with a needle to the bottom of the Petri dish with an overnight

culture of plate grown P. aeruginosa cells and incubated for 24 h at 37ºC. E,E-farnesol

was dissolved in methanol and added to agar media at the final concentrations of 25, 100,

or 250 µM. An equal amount of methanol only was added to plates to serve as solvent

control.

(ii) Microscopic. Strains were inoculated onto blocks of LB (1% agar) and covered with

glass cover slips. E,E-farnesol was dissolved in methanol and added to agar media at the final concentrations of 25, 100, or 250 µM. An equal amount of methanol only was

added to plates to serve as solvent control. Following 5 h incubation at 37ºC, the

outermost regions of the motile zone were examined using Olympus BX51 microscope at

a magnification of 600X under DIC.

Swarming motility assay. Swarming motility assays were performed as previously

described with modifications (Rashid and Kornberg 2000). Swarm agar plates were

either composed of nutrient broth (NB) (EMD Chemicals, Darmstadt, Germany)

supplemented with 0.5% (w/v) glucose and solidified with 0.5% agar or PPGAS medium

and solidified with 0.5% (w/v) agar. Plates were allowed to solidify for 3 h at room

temperature and were stab inoculated using 2 µl of a culture grown overnight in LB at

37°C. Plates were incubated for 24 h at 37ºC. As in the twitching motility assay, plates

were supplemented with E,E-farnesol to a final concentration of either 25, 100, or 250

µM. An equal amount of methanol only was added to swarm plates to serve as a solvent

control and equal concentrations of dodecanol were added to swarm plates to serve as a

hydrophobicity control.

Swimming motility assay. Swimming motility assays were performed as previously

described with modifications (O’Toole and Kolter 1998). Swim plates were composed of

LB solidified with 0.3% (w/v) agar. Plates were allowed to solidify for 3 h and were stab

inoculated using 2 µl of a culture grown overnight in LB at 37°C. Plates were incubated

for 24 h at 37°C. Plates were supplemented with E,E-farnesol to final concentration of

either 25, 100, or 250µM. An equal amount of methanol only was added to plates to serve as a solvent control.

Rhamnolipid quantification assay. Orcinol assays were performed as previously reported with modifications (Wilhem et al. 2007). For quantification of rhamnolipids from swarm plates, cells were resuspended from NB (0.5% agar) plates to an OD590nm of

2.0 in PBS. NB (0.5% agar) plates were supplemented with specified concentration of

E,E-farnesol. Negative controls received methanol only. Cells were removed by

centrifugation (10 min at 6,000 x g). 50 µl of the supernatant was diluted with water to a

volume of 300 µl and extracted twice with 600 µl of diethylether. Pooled fractions were

evaporated to dryness and resuspended in 100 µl of distilled water. Dissolved extracts

were mixed with 100 µl 1.6% orcinol and 800 µl of 60% sulfuric acid and heated to 80°C

for 30 min. The absorbance at 421nm was measured. The amount of rhamnose produced

was quantified by comparison to rhamnose standards of known concentrations assuming

1 µg of rhamnose was equivalent to 2.5 µg of rhamnolipids.

Isolation of and immunoblot analysis of pilin, PilB, and flagellin. (i) Isolation of

pilin, PilB, and flagellin. For isolation of pilin and PilB, P. aeruginosa was grown

overnight in LB broth at 37°C. The following day, 0.1 ml of the overnight culture was

subcultured in to 10 ml of fresh LB broth and grown at 37°C to an OD590nm= 0.7. 0.1 ml was subcultured into 10 ml of fresh LB broth supplemented with either E,E-farnesol or an equal volume of methanol serving as a negative control. Cells were grown to an

OD590nm= 0.7 and washed 3X in CTX buffer. Cells were solubilized by boiling for 10 min in Laemmli sample buffer. Whole cell fractions of PilB were also harvested from NB

(0.5% agar) plates supplemented with E,E-farnesol. Flagellin was harvested from whole cells grown either on NB (0.5% agar) plates or in LB broth supplemented with E,E- farnesol and solubilized by boiling for 10 min in Laemmli sample buffer. (ii)

Immunoblot analysis. Immunoblotting was conducted as previously described with modifications (Sambrook and Russell 2001). Equivalent amounts of cell protein were loaded and separated by SDS-PAGE on a 12% Tris-HCl gel. Proteins were transferred at

100V for 15 min to a nitrocellulose membrane by electroblotting. Purified whole cell pilin protein was detected using rabbit anti-pilin antiserum (a gift of Randy Irvin) as the primary antibody followed by the secondary antibody (anti-rabbit immunoglobulin G conjugated to alkaline phosphatase) (Promega, Madison,WI). Purified PilB protein was detected using rabbit anti-PilB antibody as the primary antibody followed by the secondary antibody (anti-rabbit immunoglobulin G conjugated to alkaline phosphatase).

And, purified flagellin protein was detected using rabbit anti-flagellin B antibody as the primary antibody followed by the secondary antibody (anti-rabbit immunoglobulin G conjugated to alkaline phosphatase).

Immunoprecipitation of PilB and protein quantification. (i) Whole cell fractions of

PilB. For isolation of PilB from whole cells, P. aeruginosa was grown overnight in LB broth at 37°C. The following day, 0.1 ml of the overnight culture was subcultured into

10 ml of fresh LB broth supplemented with either E,E-farnesol or an equal volume of methanol serving as a negative control. Cells were grown at 37°C to an OD590nm= 0.7.

0.1 ml was subcultured into 10 ml of fresh LB broth. Cells were grown to an OD590nm=

0.7 and washed 3X in CTX buffer and resuspended in 75 µl CTX buffer and 75 µl

Laemmli Sample Buffer. Cells were solubilized by boiling for 10 min in Laemmli sample buffer. (ii) Immunoprecipitation of PilB. 25 µl of Pansorbin cells was added to

200 µl of each protein extract and incubated on ice for 30 min. Pansorbin cells were removed by centrifugation for 1 min at 8,000 rpm and the supernatant transferred to sterile microcentrifuge tubes. 1µl of rabbit anti-PilB antibodies was added to each sample and incubated overnight at 4°C. The following day, 25 µl of Pansorbin cells was added to each sample and incubated for 30 min on ice. Each sample was washed 3X in wash buffer (50 mM Tris pH=8.4, 450 mM NaCl, 0.5% Triton X-100), resuspended in 60

µl of 2X Laemmli Sample buffer, and boiled for 2 min. Pansorbin cell debris was removed by centrifugation for 1 min at 8,000 rpm and the supernatant transferred to sterile microcentrifuge tubes. Immunoprecipitated protein was measured using standard

BSA protein assay (Pierce, Rockford, IL).

pilJ expression. P. aeruginosa (strain 31801) carrying a chromosomal pilJ::TnphoA fusion was assayed for alkaline phosphatase activity as previously described with modifications (Poole and Braun 1988). Cells were grown overnight in LB broth with or without E,E-farnesol at 37°C. The following morning, 300 µl of cells were harvested by centrifugation for 5 min at 12,000 x g and resuspended in 600 µl of 0.1M Tris-HCl (pH

8.0). Cells were permeabilized with 20 µl of chloroform and vortexed for 10 s.

Following permeabilization, cells were centrifuged for 1 min at 12,000 x g and 500µl of the supernatant was transferred to 500 µl of p-nitrophenyl phosphate (2 mg/ml in 0.1M

Tris-HCl [pH 8.0]). Each sample was incubated at room temperature for 15 min. Each

reaction was stopped by the addition of 60 µl of 1N NaOH to achieve a final

concentration 0.1N. Alkaline phosphatase-mediated release of p-nitrophenyl was measure at 405 nm and reported relative to the quantity of cells used in the assay

(OD600nm). Expression of pilJ in log phase cells was measured by subculturing 0.1 ml of

overnight cells into 10 ml of fresh LB supplemented with or without E,E-farnesol and

grown at 37°C. Cells were grown to an OD600nm of 0.3 and alkaline phosphatase activity

measured as described above.

In vivo 3H-methylation assay. P. aeruginosa was grown overnight in MSS as described

above. In vivo 3H-methylation was conducted as previously described with modifications

(Craven and Montie 1983). The following day, 1 ml volumes of cells were washed 3X in

CTX buffer. Chloramphenicol was added to 1ml of washed cells at a concentration of

200 µg/ml to inhibit de novo protein synthesis. 10 µCi of L-[methyl-3H]methionine was added to arrested cells and incubated for 30 min at 37ºC. Following the 30 min incubation with L-[methyl-3H]methionine, cells were exposed to either 25, 100, or 250

µM E,E-farnesol for 15 min. Cells were pelleted by centrifugation for 5 min at 12,000 x g. Following centrifugation, cells were solubilized by boiling 10 min in Laemmli sample buffer and proteins separated by SDS-PAGE on a 12% Tris-HCl gel. Following separation, the gel was stained in coomasie blue for 30 min and destained for 2 h. The stained gel was soaked in Amplify (Amersham Pharmacia, Piscataway, NJ) for 30 min to enhance 3H labeled proteins and dried for 55 min at 60ºC. Hyperfilm was exposed to the

dried gel for 3 days at -70°C and developed.

Immunoprecipitation of PilJ:YFP using anti-YFP antibodies. Following in vivo 3H-

methylation of FA6 + pJY, 25 µl of Pansorbin cells was added to 200 µl of protein

extract and incubated on ice for 30 min. Pansorbin cells were removed by centrifugation

for 1 min at 8,000 rpm and the supernatant transferred to sterile microcentrifuge tubes.

10 µl of Living Color A.v. monoclonal antibodies (JL-8) (i.e. anti-YFP antibodies) was

added to each sample and incubated overnight at 4°C. The following day, 25 µl of

Pansorbin cells was added to each sample and incubated for 30 min on ice. Each sample was washed 3X in wash buffer (50 mM Tris pH=8.4, 450mM NaCl, 0.5% Triton X-100), resuspended in 60 µl of 2X Laemmli Sample buffer, and boiled for 2 min. Pansorbin cell debris was removed by centrifugation for 1 min at 8,000 rpm and the supernatant transferred to sterile microcentrifuge tubes. For scintillation counting, 10 µl of the supernatant was transferred to 2 ml of cold 10% Trichloroacetic acid (TCA) in glass tubes and incubated on ice for 30 min. A vacuum manifold containing glass filters was used to filter each sample. Glass tubes were washed 2X with 5% TCA to collect any residual sample. Glass filters were washed with 10 ml of 95% ethanol and allowed to air dry. Each filter was placed into 10 ml of BioSafe Scintillation fluid and counts per minute (CPMs) were measured using a scintillation counter.

Results

Effect of E,E-farnesol on twitching motility. In the absence of E,E-farnesol, wild-type

PAO1 exhibited normal twitching motility patterns characterized by a defused twitch

zone between the agar plate interface (Fig. 1A). When inoculated into plates containing

E,E-farnesol, there was a substantial reduction in twitching motility (Fig. 1B-1D). This

reduction increased with increasing concentrations of E,E-farnesol. In addition, these

reduced twitch zones were serrated and did not exhibit a smooth circular pattern as

typical twitch zones. Wild-type PAO1 was inoculated onto plates containing dodecanol

as a hydrophobicity control. Twitching motility was unaffected in the presence of the

dodecanol control (data not shown).

Twitch zones were also examined microscopically. Wild-type PAO1 was

inoculated onto small LB (1.0% agar) slabs and covered with a coverslip. In the absence

of E,E-farnesol, spearhead-like aggregates were observed moving away from the point of

inoculation (Fig. 2A). When inoculated onto plates with E,E-farnesol, the cells exhibited

altered twitching motility characterized by circular patterns at the edge of the twitch zone

(Fig. 2B).

Effect of E,E-farnesol on swarming motility. When inoculated onto NB (0.5% agar) plates containing methanol only, wild-type PAO1 demonstrated normal swarming

motility characterized by the typical tendril pattern customarily observed (Fig. 3). When

inoculated onto plates containing 25 and 100 µM E,E-farnesol, wild-type PAO1

swarming motility was severely inhibited (Fig. 3).

In contrast to wild-type PAO1 cells, pilJ (FA6) mutant cells exhibited a

hyperswarming phenotype which lacked the distinct tendrils normally observed in wild- type PAO1 (Fig. 3). In the presence of 25 and 100 µM E,E-farnesol, pilJ (FA6) mutant

cells were slightly inhibited in their ability swarm. However, after 48 h, pilJ (FA6) mutant cells were able to overcome the inhibiting effects of E,E-farnesol and were restored in their swarming capacity.

The complemented pilJ mutant demonstrated swarming motility comparable to wild-type PAO1, and like the wild-type was inhibited in its ability to swarm in the presence of E,E-farnesol (Fig. 3).

All three strains were inoculated onto swarm plates containing dodecanol. Plates containing dodecanol served as a hydrophobicity control. In the presence of dodecanol, wild-type PAO1, pilJ (FA6) mutant, and complemented pilJ mutant displayed a hyperswarming phenotype (Fig 3D-E).

When inoculated onto PPGAS (0.5% agar) plates supplemented methanol, wild- type PAO1 and pilJ (FA6) mutant cells exhibited swarming motility characterized by a tendril pattern. This pattern was more pronounced on this media than on NB (0.5% agar)

(Fig. 4A). In the presence of 25, 100, and 250 µM E,E-farnesol, wild-type PAO1 swarming motility was inhibited. This inhibition was also more prominent on this media than NB (0.5% agar) plates (Fig. 4). In contrast, the ability of the pilJ (FA6) mutant to swarm was unaffected in the presence of E,E-farnesol (Fig. 4).

The effect of E,E-farnesol on sadB mutant swarming motility was also examined.

In contrast to wild-type PA14, the sadB mutant exhibited a hyperswarming phenotype

similar to the pilJ (FA6) mutant (Fig. 5). In the presence of 25 µM E,E-farnesol, sadB mutant cells were slightly inhibited in their ability to swarm; however, in the presence of

100 and 250 µM E,E-farnesol, swarming motility was unaffected (Fig. 5).

In order to further determine the mechanism by which E,E-farnesol inhibits surface motility, several mutant strains were examined for their ability to twitch and

swarm (Table 2). A pilB mutant (B), pqsH mutant, and groEL mutant were all inhibited

in their ability to twitch and swarm. Because these strains exhibited twitching and

swarming phenotypes resembling wild-type cells exposed to E,E-farnesol, this suggests

that these genes may be involved in the mechanism of E,E-farnesol inhibition of surface

motility.

Effect of E,E-farnesol on swimming motility. The effect of E,E-farnesol on wild-type

PAO1, pilJ (FA6) mutant, and complemented pilJ mutant swimming motility was

determined by inoculating LB (0.3% agar) supplemented with E,E-farnesol. When

inoculated onto plates without E,E-farnesol, wild-type PAO1, pilJ (FA6) mutant, and

complemented pilJ mutant exhibited typical swimming motility characterized by concentric rings of surface migration (Fig 6A). Wild-type PAO1, pilJ (FA6) mutant, and

complemented pilJ mutant swimming motility was unaltered in the presence of 25 and

100 µM E,E-farnesol (Fig 6B-C). In addition, all three strains were inoculated onto swim plates containing dodecanol as a hydrophobicity control. In the presence of

dodecanol, wild-type PAO1, pilJ (FA6) mutant, and complemented pilJ mutant demonstrated normal swimming motility (Fig. 6D-E).

Effect of E,E-farnesol on rhamnose production by wild-type PAO1 and pilJ (FA6) mutant. Wild-type PAO1 cells produced 11.82 mg/L of rhamnose. In the presence of 25

µM of E,E-farnesol, there was a 3.8-fold increase (45.01 mg/L) in rhamnose production.

There was also a 3.0-fold (35 mg/L) and a 4.0-fold (47 mg/L) increase in rhamnose production in the presence of 100 or 250 µM E,E-farnesol, respectively (Fig. 7).

In contrast to wild-type PAO1, pilJ (FA6) mutant cells produced substantially less rhamnose (2.1 mg/L). When inoculated onto swarm plates containing 25, 100, and 250

µM E,E-farnesol, there was a 20.1-fold (47.22 mg/L) , 23.6-fold (49.49 mg/L), and 23.2- fold (48.65 mg/L) increase in rhamnose production of the pilJ (FA6) mutant, respectively

(Fig. 7).

Effect of E,E-farnesol on pilin production. Using rabbit anti-pilin antibodies, a distinct band was observed at approximately 18 kDa in each lane (Fig. 8). This molecular weight corresponds to the predicted molecular weight of P. aeruginosa purified pilin monomer.

In the presence of 25 and 250µM E,E-farnesol, the intensity of this band did not

substantially vary in comparison to the negative control.

Effect of E,E-farnesol on flagellin B production. Because swarming motility is

dependent of flagella, flagellin B production was examined. Flagellin B production was

also examined because proteomic data indicated that there was a 2.01 decrease in the

production of flagellin-related proteins in the presence of E,E-farnesol (Pierce personal

communication). When P.aeruginosa cells were either grown in LB broth or NB (0.5% agar) there was a distinct band observed at approximately 53 kDa, the predicted molecular weight of P. aeruginosa flagellin monomer (Fig. 9 and 10). This band did not vary in intensity between wild-type PAO1 and pilJ (FA6) mutant cells (Fig. 9: lanes 1

and 5). In addition, the intensity of this band appeared to be equivalent between those

cells exposed to E,E-farnesol and those unexposed (Fig. 9 and 10).

Effect of E,E-farnesol on PilB production. Because proteomic data showed attenuated

levels of PilB production in the presence of E,E-farnesol, we examined the PilB

production in response to E,E-farnesol. A distinct band was observed at approximately

80 kDa for both broth and plate grown whole cell lysates (Fig. 11A and 11B). This band

corresponds to the predicted molecular weight of P. aeruginosa PilB protein. In broth

conditions, there was a decrease in the intensity of this band when cells were exposed to

250 µM E,E-farnesol (Fig. 11A: lane 3). When grown on NB (1.5% agar) plates, this

band did not vary in intensity in the presence of E,E-farnesol (Fig. 11B: lane 3).

The effects of E,E-farnesol on PilB levels was further examined by

immunoprecipitating PilB with rabbit anti-PilB antibodies from wild-type PAO1 cells

that were exposed to various concentrations of E,E-farnesol. Protein concentrations were

measured using standard BSA protein assay kit. Multiple trials were conducted showing

a small reduction in PilB levels in the presence of E,E-farnesol. For Trial 1, the greatest

reduction (1.58-fold) in PilB production was observed at a concentration of 100 µM (Fig.

12). In Trial 2 and 3, there was a 1.35-fold and 1.13-fold decrease in PilB production in

25 µM E,E-farnesol; however, the standard deviation was high indicating these

differences were not statistically significant (Fig. 12).

Effect of E,E-farnesol of pilJ expression. To determine the effect of E,E-farnesol on pilJ expression, a P. aeruginosa (31801) carrying a chromosomal pilJ::TnphoA fusion was grown in the presence of E,E-farnesol and alkaline phosphatase activity measured. pilJ expression was greater during late stationary phase (0.19±0.01) than early log phase

(0.04±0.01) (Table 3). During late stationary phase, there was a slight decrease in pilJ expression in the presence of 25 and 250 µM E,E-farnesol; however, this reduction was not significant. Likewise, pilJ expression was unaffected in the presence of 25 and 250

µM E,E-farnesol during early log phase.

In vivo 3H-methylation of wild-type PAO1 (AD) and pilJ isogenic mutant in response

to E,E-farnesol. Post translational modication of PilJ in wild-type PAO1 (AD) cells in

the presence of E,E-farnesol was examined through in vivo 3H-methylation. Several

methylated protein bands were observed in the lanes containing unstimulated and

stimulated cells in the MW region of 72 kDa, the predicted molecular weight of PilJ and other P. aeruginosa MCPs. The intensity of these bands increased slightly with the addition of E,E-farnesol. Methylation was greatest in response to E,E-farnesol for cells

exposed to 25 µM for 30 min (Fig. 13: lane 3). In addition to wild-type PAO1 (AD)

cells, pilJ mutant (FA6) cells were also labeled with L-[methyl-3H] methionine and

exposed to E,E-farnesol. Several methylated protein bands were observed for both

unstimulated and stimulated cells. The intensity of these methylated proteins; however,

did not vary upon exposure to E,E-farnesol for 5 or 30 min (Fig. 13: lane 4-6).

Immunoprecipitation of PilJ:YFP in response to E,E-farnesol. To quantitate the methylation of PilJ in response to E,E-farnesol, FA6 + pJY was labeled with L-[methyl-

3H] methionine and immunoprecipitated with Living Color A.v. monoclonal antibodies

(JL-8) (i.e. anti-YFP antibodies). Several trials were conducted producing varying

results. For trial 1 and trial 3, there was a 1.4-fold and a 1.2-fold increase in methylation

in response to 25 µM of E,E-farnesol, respectively (Fig. 14). However, this increase was

not significant for trial 3. In contrast to trial 1 and trial 3, there was a decrease in PilJ

methylation in response to E,E-farnesol in trial 2 (Fig. 14). This decrease was

proportional to increasing concentrations of E,E-farnesol. The greatest decrease (2.1-

fold) was observed at 250 µM E,E-farnesol.

Table 1. Bacterial strains used in this study Strains Characteristics Source P. aeruginosa PAO1 (AD) wild-type A. Darzins PAO1 (P) wild-type E. Pesci PAO1 (H) wild-type D. Hassett MPAO1 wild-type Seattle PAK wild-type L. Burrows PA14 Wild-type G. O’Toole pilJ (FA6) mutant PAO1 (pilJ::TcR) A. Darzins complemented pilJ mutant FA6 (pUCP19::pilJ CbR) A. Darzins FA6 + pJY FA6 (pUCP19::pilJ:YFP CbR) P. DeLange 31801 MPAO1 (pilJ::TnphoA TcR) Seattle pilB mutant (B) PAK (Δ pilB) L. Burrows pilB mutant (S) MPAO1 (pilB::TnphoA TcR) Seattle oprF mutant PAO1 (Δ oprF) D. Hassett pqsA mutant PAO1 (Δ pqsA) E. Pesci pqsH mutant MPAO1 (pqsH::Tn TcR ) E. Pesci pqsR mutant PAO1 (Δ pqsR) E. Pesci qscR mutant PAO1 (qscR::GmR) P. Greenberg sadB mutant PA14 (sadB::Tn5B21 TcR) G. O’Toole groEL mutant MPAO1 (groEL::TnphoA TcR) Seattle

A B C D

Figure 1. Macroscopic twitching motility of wild-type PAO1 in the presence of E,E- farnesol. LB (1.0% agar) plates supplemented E,E-farnesol were stab inoculated with wild-type PAO1 (AD) and incubated for 24 h at 37°C. The smaller white zones represent cells growing on the agar surface. Diffuse zones (twitch zones) represent cells moving at the agar-petri dish interface. (A) methanol only; (B) 25 µM farnesol; (C) 100 µM farnesol; (D) 250 µM farnesol.

A B

Figure 2. Microscopic twitching motility of wild-type PAO1 in the presence of E,E- farnesol. Strains were inoculated onto blocks of LB (1% agar) plates supplemented with

E,E-farnesol and covered with glass coverslips. Following 5 h incubation at 37ºC, the outermost regions of the motile zone were photographed using bright-field microscopy at a magnification of 600X. (A) methanol only; (B) 25 µM farnesol.

A B C D E

Wild-type PAO1

pilJ (FA6) mutant

complemented pilJ mutant

Figure 3. Swarming motility of wild-type PAO1, pilJ (FA6) mutant, and complemented pilJ mutant on NB (0.5% agar) in the presence of E,E-farnesol and

dodecanol. NB (0.5% agar) plates supplemented with either E,E-farnesol or dodecanol

were stab inoculated with P. aeruginosa and incubated for 24 h at 37°C. (A) methanol

only; (B) 25 µM farnesol; (C) 100 µM farnesol; (D) 25 µM dodecanol; (E) 100 µM

dodecanol.

A B C D

Wild-type PAO1

pilJ (FA6) mutant

Figure 4. Swarming motility of wild-type PAO1 and pilJ (FA6) mutant on PPGAS agar in the presence of E,E-farnesol. PPGAS (0.5% agar) plates with E,E-farnesol were stab inoculated with P. aeruginosa and incubated for 24 h at 37ºC. (A) methanol

only; (B) 25 µM farnesol; (C) 100 µM farnesol; (D) 250 µM farnesol.

A B C D

PA14

sadB mutant

Figure 5. Swarming motility of wild-type PA14 and sadB mutant in the presence of

E,E-farnesol. NB (0.5% agar) plates supplemented with E,E-farnesol were stab inoculated with P. aeruginosa and incubated for 24 h at 37°C. (A) methanol only; (B) 25

µM farnesol; (C) 100 µM farnesol; (D) 250 µM farnesol.

A B C D E

Wild-type PAO1

pilJ (FA6) mutant

complemented pilJ mutant

Figure 6. Swimming motility of wild-type PAO1, pilJ (FA6) mutant, and

complemented pilJ mutant in the presence of E,E-farnesol. LB (1.0% agar)

supplemented with E,E-farnesol was stab inoculated with wild-type PAO1 (AD) and

incubated for 24 h at 37°C. (A) methanol only; (B) 25 µM farnesol; (C) 100 µM farnesol; (D) 25 µM dodecanol; (E) 100 µM dodecanol .

Table 2. Surface motility of P. aeruginosa wild-type and mutant strains Strain Swarming motility Twitching motility MPAO1 + + PAO1 (P) + + PAO1 (H) + + PAK + +/- PA14 + + oprF mutant - + pilB mutant (S) +++ - pilB mutant (B) - - pqsA mutant + + pqsH mutant - - pqsR mutant + + qscR mutant - + sadB mutant +++ NM groEL mutant - - Note* NM= Not measured, (+) = motility observed, (-) = no motility observed, (+/-) = reduced motility.

PAO1 30 PilJ rhamnose (mg/L) 20

0 0 25 100 250 Farnesol (uM)

Figure 7. Rhamnose production by wild-type PAO1 and pilJ (FA6) mutant cells

harvested from swarm plates supplemented with E,E-farnesol. Whole cell

suspensions were collected from NB (0.5% agar) plates supplemented with E,E-farnesol

and rhamnolipids extracted with diethylether. Rhamnose levels were detected using the

orcinol method and A421nm was measured. The amount rhamnose in the samples was quantified by comparing to rhamnose standards of known concentrations.

1 2 3 4 5

19.7 kDa

Figure 8. Immunoblot analysis of total pilin in the presence of E,E-farnesol. Whole

cell suspensions were collected from LB broth and solubilized by boiling for 10 min in

Laemmli sample buffer prior to SDS-PAGE analysis on a 12% Tris-HCl gel. Bands were

detected with P. aeruginosa anti-pilin antibody. Equivalent amounts of cell protein (33

µg) were loaded for each sample. Lane 1 and 5: molecular weight marker; Lane 2: methanol only; Lane 3: 25 µM farnesol; Lane 4: 250 µM farnesol.

1 2 3 4 5 6 7 8

53 kDa

Figure 9. Immunoblot analysis of flagellin B harvested from wild-type PAO1 and pilJ (FA6) mutant cells grown in LB broth supplemented with E,E-farnesol. Whole

cell suspensions were collected from LB broth supplemented with E,E-farnesol and

solubilized by boiling for 10 min in Laemmli sample buffer prior to SDS-PAGE analysis

on a 12% Tris-HCl gel. Bands were detected with P. aeruginosa anti-flagellin B

antibody. Equivalent amounts of cell protein (19.4 µg) were loaded for each sample.

Lane 1: wild-type PAO1 (methanol only); Lane: wild-type PAO1 (25 µM farnesol); Lane

3: wild-type PAO1 (100 µM farnesol); Lane 4: wild-type PAO1 (250 µM farnesol); Lane

5: pilJ (FA6) mutant (methanol only); Lane 6: pilJ (FA6) mutant (25 µM farnesol); Lane

7: pilJ (FA6) mutant (100 µM farnesol; Lane 8: pilJ (FA6) mutant (250 µM farnesol).

1 2 3

53 kDa

Figure 10. Immunoblot analysis of flagellin B harvested from wild-type PAO1 cells grown on swarm plates supplemented with E,E-farnesol. Whole cell suspensions were collected from NB (0.5% agar) plates supplemented with E,E-farnesol and solubilized by boiling for 10 min in Laemmli sample buffer prior to SDS-PAGE analysis on a 12% Tris-HCl gel. Bands were detected with P. aeruginosa antiflagellin B

antibody. Equivalent amounts of cell protein (18.6 µg) were loaded for each sample.

Lane 1: methanol only; Lane 2: 25 µM farnesol; Lane 3: 250 µM farnesol.

1 2 3 1 2 3

80 kDa 68kd

A B

Figure 11. Immunoblot analysis of PilB in wild-type PAO1 grown in the presence of

E,E-farnesol. Whole cell suspension were collected from either LB broth or NB (0.5% agar) plates supplemented with E,E-farnesol and solubilized by boiling for 10 min in

Laemmli Sample buffer prior to SDS-PAGE analysis on a 12% Tris-HCl gel. Bands were detected with P. aeruginosa anti-PilB antibody. Equivalent amounts of cell protein were loaded for each sample. A = broth grown cells, B = swarm plate grown cells. Lane

1: no methanol; Lane 2: 25 µM farnesol; Lane 3: 250 µM farnesol.

Trial 1 Trial 2 Trial 3 control 1316.0±93.4 882.6±349.5 674.1±154.3 25 µM farnesol 1113.8±121.9 656.0±108.3 597.7±55.6 100 µM farnesol 834.5±235.9 776.9±237.1 599.9±82.8 250 µM farnesol 1175.7±150.5 764.0±70.6 743.4±48.0

Figure 12. Quantification of PilB immunoprecipitated from wild-type PAO1 following exposure to E,E-farnesol. Whole cells were exposed to E,E-farnesol and PilB

was precipitated using P. aeruginosa anti-PilB antibodies. Precipitated protein was measured using standard BSA protein assay.

Table 3. Effect of E,E-farnesol on pilJ expression: Alkaline Phosphatase

Activity of pilJ::TnphoA fusion in wild-type MPAO1

Sample Growth phase Farnesol Conc. (µM) Alkaline Phosphatase Activity (A405/A600) 1 Overnight - 0 0.19±0.01 2 Overnight + 25 0.16±0.03 3 Overnight + 250 0.14±0.01 4 Log -/- 0 0.04±0.01 5 Log -/+ 25 0.04±0.01 6 Log -/+ 250 0.05±0.03 7 Log +/+ 25 0.05±0.01 8 Log +/+ 250 0.05±0.01 Note* Overnight cells: (+) = grown in media with E,E-farnesol, (-) = grown in media

without E,E-farnesol. Log phase cells (OD600nm= 0.3): (-/-) = grown overnight without

E,E-farnesol, subcultured in media without E,E-farnesol, (-/+) = grown overnight without

E,E-farnesol, subcultured into media with E,E-farnesol, (+/+) = grown overnight with

E,E-farnesol, subcultured into media with E,E-farnesol.

1 2 3 4 5 6 1 2 3 4 5 6

72 kDa

Figure 13. In-vivo 3H-methylation of wild-type PAO1 and pilJ mutant (FA6) cells in

response to E,E-farnesol. Cells were labeled with L-[methyl-3H]methionine and

exposed to 25 µM E,E-farnesol for either 5 or 30 min. The coomasie stained gel appears

on the left and the fluorogram appears on the right. Lane 1: wild-type PAO1 (methanol

only); Lane 2: wild-type PAO1, 5 min; Lane 3: wild-type PAO1, 30 min; Lane 4: pilJ

(FA6) mutant (methanol only); Lane 5: pilJ (FA6) mutant, 5 min; Lane 6: pilJ (FA6)

mutant, 30 min.

Figure 14. Quantification of PilJ methylation in response to E,E-farnesol. FA6 +

pJY cells were labeled with L-[methyl-3H]methionine and exposed to either 25, 100, or

250 µM E,E-farnesol for 15 min. Following in-vivo 3H-methylation, PilJ:YFP was precipitated using anti-YFP antibodies and PilJ methylation was measured using a scintillation counter.

Discussion

In this study we examined the effect of E,E-farnesol on surface motility of P.

aeruginosa. In the presence of E,E-farnesol, there was a substantial decrease in twitching

motility (Fig. 1). This inhibition was dose dependent with the greatest reduction observed at 250 µM E,E-farnesol. Interestingly, microscopic examination of wild-type

PAO1 exposed to E,E-farnesol revealed that these cells exhibited circular patterns at the

edge of the twitch zone (Fig. 2B). This altered pattern is similar to that of a pilH mutant

(Darzins 1994). pilH belongs to the Type IV pili biogenesis gene cluster and encodes for

a CheY homologue implicated in the regulation of Type IV pili biogenesis and twitching

motility. These results indicate that E,E-farnesol may act on P. aeruginosa through the

Type IV signal transduction pathway.

In addition to twitching motility, E,E-farnesol inhibited swarming motility in

wild-type PAO1 (Fig. 3 and Fig. 4). Previous research has demonstrated the ability of

E,E-farnesol to inhibit swarming motility of P. aeruginosa CF isolates; however, this

previous study did not provide a mechanism for this inhibition (McAlester et al. 2007).

Because the MCP PilJ has been shown to negatively regulate swarming motility,

we also examined the effects of E,E-farnesol on swarming motility of a pilJ mutant

(Caiazza et al. 2007). pilJ is also a part of the Type IV pili biogenesis signal transduction pathway (Darzins 1994). In the presence of E,E-farnesol, pilJ (FA6) mutant cells’ initial

ability to swarm was slightly reduced. After 48 h these pilJ (FA6) mutant cells were able

to swarm like pilJ (FA6) mutant cells unexposed to E,E-farnesol (Fig. 3). Because pilJ

(FA6) mutant cells maintained their ability to swarm in the presence of E,E-farnesol, this

suggested to us that E,E-farnesol acts on P. aeuginosa Type IV signal transduction

pathway through PilJ. The slight decrease in pilJ (FA6) mutant cells swarming motility

in the presence of E,E-farnesol may indicate that E,E-farnesol binds to MCPs other than

PilJ.

Due to the pilJ (FA6) mutant’s ability to swarm in the presence of E,E-farnesol,

we examined pilJ expression and methylation response following exposure to this QSM.

pilJ expression was unaffected in the presence of E,E-farnesol (Table 3). PilJ

methylation was also assessed to determine if E,E-farnesol acts on PilJ through post-

translational modification. In-vivo 3H-methylation of wild-type PAO1 showed an

increase in the level of methylation of proteins migrating at 72 kDa, the molecular weight

of PilJ, when E,E-farnesol was present (Fig. 13). This increase was not observed when pilJ (FA6) mutant cells were exposed to E,E-farnesol. These results suggest that E,E- farnesol acts on P. aeruginosa by binding to PilJ; however, P. aeruginosa possesses numerous MCPs that have an approximate molecular of 72 kDa.

In order to determine if PilJ was specifically methylated in response to E,E- farnesol, PilJ was immunoprecipitated following in-vivo 3H-methylaton. The results of

these experiments were variable (Fig. 14). In trials 1 and 3, there was an increase PilJ

methylation in response to 25 µM of E,E-farnesol; however, in trial 2 there was a

decrease in PilJ methylation following exposure to increasing concentrations of E,E-

farnesol. In the presence of a positive stimulus (attractant), MCPs are methylated;

whereas, in the presence of a negative stimulus (repellent) or absence of a positive

stimulus MCPs are demethylated. Because all three trials were conducted using the same

conditions, it is difficult to conclude whether E,E-farnesol acts as an attractant or

repellent. This variation could be due to the unstable bond between the methyl group and

the tritiated methionine of L-[methyl-3H]methionine used to measure methylation.

According to the manufacturer, this bond may begin to degrade within a month

(Amersham Pharmacia, personal communication). If degradation occurs and the methyl

groups are released from the tritiated methionine, quantification of PilJ methylation could

appear less than its actual measurement.

Previous studies in our lab investigating the methylation pattern of PilJ in

response to multiple chemicals demonstrated an increase in methylation towards

molecules consisting of a C12 backbone (i.e. 3-oxo-C12-HSL and L-∝-

phosphatidylethanolamine [PE] dilauryl [C12:0]). E,E-farnesol is structurally similar to

3-oxo-C12-HSL and also consists of a C12 backbone. 3-oxo-C12-HSL is a QSM that has

been shown to be involved in surface motility of P. aeruginosa (Glessner et al. 1999).

Although we were unnsuccessful in establishing the methylation pattern of PilJ in the

presence of E,E-farnesol, any potential alteration in that pattern may be attributed to the structural similarity of E,E-farnesol to 3-oxo-C12-HSL. This similarity may allow E,E-

farnesol to compete with 3-oxo-C12-HSL for the ligand binding domain of PilJ and inhibit

swarming and twitching motility through the Type IV signal transduction pathway.

To further delineate the mechanism by which E,E-farnesol inhibits surface motility in P. aeruginosa, proteomic analysis was conducted by a collaborating lab.

Approximately 60 proteins, were differentially expressed in the presence of 25 µM E,E- farnesol (Pierce, personal communication). This data was examined for any proteins

involved in surface motility. In the presence of 25 µM E,E-farnesol, there was a 2.0-fold

decrease in PilB and flagellin-related proteins. PilB is involved in Type IV pili biogenesis and flagellin B is the protein monomer that composes the flagellum filament.

In addition, there was a 1.97-fold decrease in OprF, an outer membrane protein, levels

and a 10.01-fold decrease in chaperone GroEL levels in the presence of 25 µM E,E-

farnesol.

Because PilB is required for Type IV pili extention, a decrease in its production

could possibly account for the inhibition of twitching motility observed in the presence of

E,E-farnesol. In order to confirm the proteomic data, the effect of E,E-farnesol on PilB

levels was examined through immunoblot analysis using anti-PilB antibodies. When

cells were exposed to E,E-farnesol in broth conditions, there was a slight decrease in PilB

production in the presence of 250 µM E,E-farnesol (Fig. 11A). In contrast, PilB levels

were unaffected when cells were grown on swarm plates supplemented with E,E-farnesol

(Fig. 11B). In addition, PilB production was also examined by exposing wild-type PAO1

cells to E,E-farnesol and then immunoprecipitating PilB using anti-PilB antibodies.

Varying results were observed in multiple trials. A slight reduction was observed in PilB

levels when cells were exposed to E,E-farnesol; however, the only significant decrease

was observed in Trial 1 at a concentration of 100 µM E,E-farnesol (Fig. 12). The

variation observed in all three trials could be attributed to the fact that the anti-PilB

antibodies used were specific for PilB in a wild-type PAK background rather than the

strain we used, wild-type PAO1. These antibodies may have cross-reacted with proteins

other than PilB in wild-type PAO1. Due to variability in these results, it cannot be

determined whether E,E-farnesol affects twitching motility through PilB production.

In terms of swarming motility, PilB has never been shown to be involved in this type of motility. To determine if PilB is necessary for swarming motility, two pilB mutant strains were examined for their ability to swarm. One of the pilB mutant strains was unable to swarm while the other was unaffected (Table 2). This difference in swarm phenotype could be due to the fact each were made in different wild-type backgrounds,

MPAO1 and PAK. The pilB strain incapable of swarming was constructed from wild- type PAK, a strain of P. aeruginosa known to have abnormal motility patterns.

PilA production was also assessed using immunoblot analysis. There was no observable difference in whole cell PilA production in the presence of 25 or 250 µM E,E- farnesol (Fig. 8). These results indicate that the ability of E,E-farnesol to inhibit P. aeruginosa twitching motility cannot be attributed to levels of the pilin subunit. While

E,E-farnesol does not affect PilB or PilA levels it may affect extension and/or retraction of Type IV pili through any of the other 40 genes required for twitching motility in P. aeruginosa (Alm and Mattick 1997, Beatson et al. 2002).

As previously stated, proteomic analysis of wild-type PAO1 has also revealed a

2.0-fold and a 1.9-fold decrease in flagellin related proteins and OprF levels, respectively

(Pierce, personal communication). Among the potential flagellin related proteins affected is flagellin B, the protein subunit making up the flagellum filament. Because swarming motility is dependent on flagella, a decrease in flagellin B production could potentially account for the inhibition observed in the presence of E,E-farnesol. Immunblot analysis of wild-type PAO1 cells harvested from broth and swarm plates showed no difference in flagellin B production in the presence of E,E-farnesol (Fig. 9 and Fig. 10). These results indicate that E,E-farnesol does not inhibit swarming motility through flagellin B

production. Our inability to confirm the proteomic data may indicate that either we were not successful in replicating the exact conditions in which it was conducted or that another flagellin related protein, such as a flagellin related hook protein, was down regulated rather than flagellin B.

The mechanism by which E,E-farnesol inhibits swarming motility was further studied by examining the swarming phenotype of an oprF mutant. OprF is an outer membrane protein of P. aeruginosa that functions as a transmembrane pore involved in regulating cell shape and growth in low-osmolarity media through altering cell permeability (Brinkman et al 2000, Saint et al. 2000, El Hamel et al. 2000, Freulet-

Marriere et al. 2000). The oprF mutant was completely deficient in its ability to swarm

(Table 2). Presently, no published literature has implicated OprF in swarming motility in

P. aeruginosa. These results along with the proteomic data suggest that E,E-farnesol may inhibit swarming motility by decreasing cell permeability. By altering the permeability of the cell membrane, E,E-farnesol could potentially hinder the cells ability to conduct H+ and, therefore, affect the proton motive force needed to power the flagellum motor. Other studies have also indicated that cell permeability plays a role in regulating cell motility. In a study examining the effects of the two-component regulatory system PprB and PprA on global gene expression, a mutation in the response regulator gene pprB resulted in a substantial reduction of swimming and swarming motility (Dong et al. 2005). This two-component regulatory system known for controlling cell permeability as well as the expression of several quorum-sensing controlled genes (Wang et al. 2004, Dong et al. 2005).

As previously stated, there was a 10.01-fold decrease in chaperone GroEL levels

in the presence of 25 µM E,E-farnesol. GroEL is a heat shock protein that is responsible

for the folding/and or assembly of certain polypeptides, as well as the transport of some

secretory proteins across the cell membrane (Fujita et al. 1998). Presently, there is no

published literature indicating that this protein is involved in surface motility in P.

aeruginosa; however, we demonstrated that a groEL mutant is deficient in swarming and

twitching motility (Table 2). These results suggest that GroEL may directly or indirectly

control surface motility. If this is true, then E,E-farnesol may partially regulate surface

motility by regulating GroEL protein levels.

In this study, other genes were also examined as possible targets of E,E-farnesol

(Table 2). The pqsH mutant was the only mutant strain exhibiting a swarm and twitch

negative phenotype consistent with that of wild-type cells exposed to E,E-farnesol. PqsH

is responsible for converting the precursor 4-hydroxy-2-heptyl-quinoline (HHQ) to the

quorum-sensing molecule (QSM) PQS. Previous research has shown that E,E-farnesol

affects the PQS system by stimulating a non-productive interaction of PqsR to the

promoter region of pqsA (Cugini et al. 2007). This interaction results in reduced

transcript levels of pqsA and, consequently, a decrease in PQS production. PQS is known to regulate rhamnolipid production, a biosurfactant involved in swarming motility. This

suggests that E,E-farnesol may inhibit swarming motility through rhamnolipid production.

In order to determine the effects of E,E-farnesol on rhamnolipid production, wild-

type PAO1 and pilJ (FA6) mutant cells were harvested from swarm plates supplemented

with E,E-farnesol and rhamnose measured by orcinol assay (Wilhem et al. 2007). In the

presence of 25, 100, and 250 µM E,E-farnesol, there was a 3.8-fold, 3.0-fold, and 4.0- fold increase in rhamnose levels, respectively (Fig. 7). Recent studies have shown that rhamnolipids and their precursor HAA are crucial in regulating swarming motility.

Rhamnolipids regulate swarming patterns by maintaining the cell-free areas between tendrils and HAA functions solely as a wetting agent (Caiazza et al. 2005). This study also demonstrated that swarming motility is completely inhibited when purified rhamnolipids were added to swarm plates (Caiazza et al. 2005). This indicates that swarming motility is highly regulated by the levels of rhamnolipids and HAA present and that E,E-farnesol may inhibit swarming motility by altering these concentrations. In this study, the ability of E,E-farnesol to inhibit swarming motility was more pronounced when cells were inoculated onto PPGAS plates (Fig. 4). These results further support the idea that E,E-farnesol affects swarming motility through rhamnolipid levels because this media is typically used to promote the production of these biosurfactants.

Interestingly, the pilJ (FA6) mutant produced substantially less rhamnolipids than the wild-type PAO1; however, when exposed to E,E-farnesol they produced equivalent amounts of rhamnolipids. Because pilJ (FA6) mutant cells retain their ability to swarm in the presence of increased levels of rhamnolipids, this suggests that either the inhibition in swarming motility is independent of this increase or that pilJ mutant cells are unable to sense and respond to rhamnolipids. Evidence supporting the latter hypothesis can be found in recent studies examining the function of the gene sadB in swarming motility and biofilm formation. Epistasis analysis of SadB indicated that it functions upstream of the

CheIV chemotaxis cluster and that a sadB mutant is phenotypically similar to PilJ

(Caiazza et al. 2007). Mutations in sadB and pilJ resulted in hyperswarming, increased

flagellar reversals, and decreased biofilm formation. In addition, sadB mutants were unable to sense and/or respond to rhamnolipids (Caiazza et al. 2005). If this also is true

for the pilJ (FA6) mutant, then this explains how the pilJ (FA6) mutant is able to swarm

in the presence of increased rhamnolipid levels. Like the pilJ (FA6) mutant, the sadB

mutant also maintained its ability to swarm in the presence of E,E-farnesol (Fig. 5).

Overall, the mechanism by which E,E-farnesol inhibits swarming and twitching

motility in P. aeruginosa is complex, involving several proteins and pathways. Because

swarming and twitching motility is involved in biofilm formation, E,E-farnesol could

potentially be used in the inhibition of biofilm formation and the treatment of established

biofilms. In addition to previous research, this study indicates that the interaction

between P. aeruginosa and C. albicans is complicated. Understanding this interplay

between these two organisms could lead to manipulation and disruption of the microbial

community of CF patients.

Summary

Quorum-sensing is a way for bacterial cells to communicate within a population.

Because several quorum-sensing species occupy the same natural environment, it is important to determine whether there are not only intraspecies, but interspecies interactions. This study was conducted in order to determine whether the yeast Candida albicans can communicate with Pseudomonas aeruginosa through its quorum-sensing molecule, E,E-farnesol, and the effect of that communication on community behavior.

E,E-farnesol was shown to inhibit swarming and twitching motility. Interestingly, pilJ

(FA6) mutant cells were only slightly inhibited in their ability to swarm in the presence of E,E-farnesol. After 48 h, pilJ (FA6) mutant cells were completely restored in their ability to swarm. These results indicated that E,E-farnesol may act on P. aeruginosa through PilJ; however, this could not be verified through pilJ expression nor PilJ methylation.

We were able to show that there was an increase in rhamnolipid production when

P. aeruginosa was exposed to E,E-farnesol. We also demonstrated that pilJ (FA6) mutant cells produce less rhamnolipids than the wild-type PAO1 and that in the presence of E,E-farnesol there is also an increase in rhamnolipid production equivalent to wild-

type PAO1. Because swarming motility is dependent on rhamnolipid concentrations, this

increase in rhamnolipid production could account for the inhibition we observe. pilJ

(FA6) mutant’s ability to retain swarming motility in the presence of E,E-farnesol could

indicate that PilJ is a receptor for rhamnolipids and, in its absence, the mutant is

incapable of sensing and responding to increased levels of this surfactant.

In additon to increasing rhamnolipid levels, proteomic analysis showed that OprF

levels are downregulated in the presence E,E-farnesol. Also, oprF mutant cells are impaired in their ability to swarm. These results suggest that E,E-farnesol may affect swarming motility by altering cell permeability. Overall, we demonstrated that the method by which E,E-farnesol inhibits surface motility in P. aeruginosa is complex, involving multiple mechanisms. These findings are of great importance because surface motility plays a key role in biofilm formation in P. aeruginosa. By affecting P. aeruginosa’s capacity to properly form biofilms, we could interfer with its ability to successfully colonize a host.

Future Studies

In order to confirm that E,E-farnesol inhibits swarming motility through

rhamnolipid production, additional experiments must be conducted. This could be

accomplished by adding rhamnolipid extracts isolated from cells exposed to E,E-farnesol

or purified rhamnolipids to swarm plates and examinining the swarming phenotype of wild-type PAO1 and pilJ (FA6) mutant. If an increase in rhamnolipid levels is

responsible for inhibiting swarming motility, this should be apparent on the plates

inoculated with wild-type PAO1. Also, if PilJ senses rhamnolipids, then swarming

motility of the pilJ (FA6) mutant would be unaffected in the presence of these additional

rhamnolipids.

To further clarify that PilJ senses and responds to rhamnolipids, in-vivo 3H-

methylation of PilJ in the presence of rhamnolipids should be conducted. If PilJ senses

rhamnolipids there will be a change in the methylation pattern of this protein. In

addition, previous studies have indicated that PilJ controls swarming motility through cell

reversals. It would be interesting to see if E,E-farnesol affects cell reversals. Preliminary

tests in our lab were unsuccessful in determining the effects of E,E-farnesol on cell

reversals using published protocols (Caiazza and O’Toole 2004). For instance, published

literature suggests using 15% ficoll to emulate swarming motility; however, no

movement was observed under the microscope using these conditions. Additional work

is needed to modify and optimize these methods. Nevertheless, establishing this

information could lead to a model where in the presence of E,E-farnesol itself or

increased rhamnolipids levels, PilJ senses and responds to these levels by controlling cell

reversals.

In addition, the effect of E,E-farnesol on OprF levels and its relationship to

swarming motility should be further investigated. Immunoblot analysis of OprF

production in the presence of E,E-farnesol should be conducted in order to confirm the

proteomic data. Also, more studies need to be performed in order to establish the

connection between cell permeability and surface motility. If by overexpressing oprF we observe a hyperswarming phenotype, then this lends evidence to our theory. In addition, the permeability of P. aeruginosa following exposure to E,E-farnesol could be examined using fluorescent dyes. These results could lead to an additional model where in the presence of E,E-farnesol, OprF levels are down regulated leading to a reduction in cell permeability. This reduction could affect the cells ability to conduct H+ and, therefore,

affect the proton motive force necessary to drive flagella rotation.

Lastly, the greatest effect of E,E-farnesol on the P. aeruginosa proteome was

observed in terms of GroEL levels. Because there was a 10-fold decrease in GroEL

levels in the presence of E,E-farnesol, we examined swarming motility of a groEL

mutant. In comparison to its parent strain MPAO1, this mutant was unable to swarm.

Complementation of this mutant should be conducted in order to see if swarming motility

is restored; thus, verifying that GroEL levels affect this phenotype. Also, it would be interesting to see whether rhamnolipid production is altered in the groEL mutant. If there

is an increase in rhamnolipid production in the groEL mutant, this would indicate how

GroEL levels affect swarming motility in P. aeruginosa.

Chapter III

The effect of a cationic porphyrin on Pseudomonas aeruginosa biofilms Collins, T.L., Markus, E.A., Hassett, D.J., and J.B. Robinson. 2010. Curr. Microbiol. 61: 411-416.

Abstract

THE EFFECT OF A CATIONIC PORPHYRIN ON Pseudomonas aeruginosa BIOFILMS

Name: Collins, Tracy Lynn University of Dayton

Advisor: Dr. Jayne B. Robinson

Current studies have indicated the utility of photodynamic therapy using porphyrins in

the treatment of bacterial infections. Photoactivation of porphyrins results in the

1 production of singlet oxygen ( O2) that damages biomolecules associated with cells and

biofilms, e.g., proteins, polysaccharides, and DNA. The effect of a cationic porphryin on

P. aeruginosa PAO1 biofilms was assessed by exposing static biofilms to 5,10,15,20-

tetrakis(1-methyl-pyridino)-21H,23H-porphine, tetra-p-tosylate salt (TMP) followed by

irradiation. Biofilms were visualized using confocal scanning laser microscopy (CSLM) and cell viability determined using the LIVE/DEAD BacLight viability assay and standard plate counts. At a concentration of 100 µM TMP, there was substantial killing of cells within P. aeruginosa PAO1 wild-type and pqsA mutant biofilms with little

disruption of the biofilm matrix or structure. Exposure to 225 µM TMP resulted in

almost complete killing as well as the detachment of wild-type PAO1 biofilms. In

contrast, pqsA mutant biofilms that contain less extracellular DNA remained intact.

Standard plate counts of cells recovered from attached biofilms revealed a 4.1-log10 and a

3.9-log10 reduction in viable cells of wild-type PAO1 and pqsA mutant strains,

respectively. Our results suggest that the action of photoactivated TMP on P. aeruginosa biofilms is two-fold: direct killing of individual cells within biofilms and detachment of the biofilm from the substratum. There was no evidence of porphyrin toxicity in the absence of light; however, biofilms pretreated with TMP without photoactivation were substantially more sensitive to tobramycin than untreated biofilms.

Introduction

Pseudomonas aeruginosa is an opportunistic pathogen known to cause infections

in immunocompromised individuals and is the leading cause of mortality among cystic

fibrosis (CF) patients (Govan and Deretic 1996). The organism possesses a number of virulence factors that contribute to its ability to invade and colonize its host (van Delden and Iglewski 1998, van Delden 2004). In addition, it forms complex communities known as biofilms; hydrated matrices of cells consisting of polysaccharides, extracellular DNA, and proteases (D’Argenio et al. 2002, Freidman and Kolter 2004, Matsukawa and

Greenberg 2004, Nemoto et al. 2004, Whitchurch et al. 2002). P. aeruginosa has been shown to form biofilms on abiotic (e.g. catheters and stents) as well as biotic (e.g. urinary tract and lung tissue) surfaces (Costerton et al. 1999, Donlan and Costerton 2002, Speer et al. 1988). Biofilms are of significant medical importance because they confer the ability to evade the host immune system and render the cells more resistant to antimicrobial agents (Costerton 2001, Nickel et al. 1985). These common characteristics lead to persistent and chronic infections (Costerton et al. 1999).

Photodynamic therapy (PDT) has been useful in the treatment of certain cancers and other diseases such as macular degeneration. In recent years, there has been increased interest in using PDT as a means to treat bacterial infections (Wainwright

2009). PDT requires three components: light, oxygen, and a photosensitizer. Light activated cationic porphyrins transfer energy to molecular oxygen resulting in the

1 production of singlet oxygen ( O2). This mechanism is known as the Type II reaction.

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1 O2 reacts with different components (e.g. phospholipids, peptides, and sterols) of the cell

wall and cell membranes and also mediates DNA damage and cell death (Wainwright

1998). The cationic porphyrin 5,10,15,20-tetrakis(1-methyl-pyridino)-21H,23H-

porphine, tetra-p-tosylate salt (TMP) specifically causes DNA damage by intercalating

between DNA base pairs, causing photoinduced strand breakage when irradiated (Kelly

and Murphy 1985, Pasternack and Gibbs 1996).

Previous studies have demonstrated the ability of cationic porphyrins to successfully photoinactivate Gram-positive and Gram-negative bacteria, as well as fungi

(Hamblin and Hasan 2004). In this study, we examined the effectiveness of TMP against cultures of P. aeruginosa biofilms. TMP at a concentration of 2.5 mg ml-1 has been

shown to reduce P. aeruginosa PAO1 planktonic cell populations by >102 cfu ml-1

(Donnelly et al. 2007). In the same study, it was demonstrated that higher concentrations

(5.0 mg ml-1) of TMP were necessary to achieve the same level of killing in bacteria

enmeshed within biofilms; however, they did not examine the effect of TMP on biofilm

structure. Additionally, this porphyrin is known to significantly reduce S. aureus survival

and, when combined with antibiotics, disrupt established biofilms (Di Poto et al. 2009).

In this study, we investigated the effects of photoactivated TMP on P. aeruginosa

biofilms using two strains: a wild-type PAO1 strain and its isogenic pqsA mutant that has

previously been shown to produce biofilms with highly reduced levels of extracellular

DNA (Allesen-Holm et al. 2006). Additionally, we investigated the ability of TMP to

affect biofilms in the absence of photoactivation.

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Methods

Bacterial strains, growth conditions and chemicals. The P. aeruginosa PAO1 wild-

type and isogenic pqsA mutant strain were obtained from Eb Pesci (East Carolina

University School of Medicine). P. aeruginosa strains were grown aerobically with

shaking in Minimal Salts medium (40 mM K2HPO4, 20 mM KH2PO4, 7.6 mM

-3 [NH4]2SO4, 0.2 mM MgSO4 · 7 H2O, 9.2 x 10 mM FeCl3 · 6 H2O, 0.2% [wt/vol]

glucose; adjusted pH 7.0) at 37°C (Craven and Montie 1983, Sambrook et al. 1989). For

static biofilms, P. aeruginosa strains were grown overnight in Minimal Salts medium at

37°C with shaking. The following day, bacteria were diluted in fresh media to an

OD590nm of 0.15. Five hundred microliters of the standardized culture was added to

sterile polystyrene cuvettes and incubated statically for 24 h at 37°C. For examination of

static biofilms using CSLM, sterile microscope slides were submerged in standardized

cell suspensions and incubated statically at 37°C for 24 h.

Photosensitizer. 5,10,15,20-tetrakis(1-methyl-pyridino)-21H,23H-porphine, tetra-p- tosylate salt (TMP) was purchased through Sigma-Aldrich. A 12.5 mg ml-1 TMP stock

solution was prepared in dH2O and filter sterilized. TMP was added to cell suspensions

and biofilms at various concentrations. TMP concentrations of 100 (0.14 mg ml-1) and

225 µM (0.14 mg ml-1) were chosen based on their effectiveness in preliminary trials.

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Photoactivation. TMP was activated using a 100-Watt mercury vapor lamp fitted with a colored glass filter (Newport FSR-GG420) blocking wavelengths shorter than 400 nm.

Samples were irradiated for various exposure times at an intensity of 220-240 Joules/cm2.

CSLM of static biofilms. Overnight biofilms formed on slides were rinsed in phosphate

buffered saline (PBS), pH 7.0, and transferred to 50 ml tubes containing PBS

supplemented with TMP at a concentration of either 100 or 225 µM. Negative control

slides were transferred to 50 ml tubes containing only PBS. Following pre-exposure to

TMP, biofilms were irradiated for 10 min and washed briefly in PBS. Bacterial viability

was assessed in biofilm cultures using LIVE/DEAD BacLight bacterial viability assay

(Molecular Probes Inc., Eugene, OR), containing SYTO9 and propidium iodide dyes.

Biofilms were visualized with an Olympus FV1000 CSLM (Olympus America, Center

Valley, PA) using a 60X oil immersion objective. The excitation and emission

wavelengths used for visualizing SYTO9 were 488nm and 520nm, respectively.

Propidium iodide was visualized using excitation and emission wavelengths of 543nm

and 580nm, respectively. Biofilm images were acquired in 0.4 µm optical sections for

the entire thickness of the biofilm.

Effect of TMP on viability of biofilm associated cells. Static biofilms formed in sterile

polystyrene cuvettes, as described above, were used to quantify cell survival.

Supernatants from 24 h biofilms were removed and replaced with PBS containing TMP

and irradiated for 10 min. Cells released from the biofilm following treatment were

collected from the supernatant by centrifugation and resuspended in PBS. The remaining

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attached biofilm was washed once with PBS and attached cells were released from the

surface using mechanical shearing by repeated pipeting. The number of viable cells

present in the supernatants and biofilms following release by mechanical shearing were

determined by plating on LB (1.5% agar) plates. Plates were incubated for 24 h at 37°C.

Measurement of LIVE/DEAD cells in static biofilms. Static biofilms were grown in

sterile polystyrene cuvettes as described above. Supernatant from 24 h biofilms were

removed and replaced with PBS containing TMP and irradiated for 10 min. Negative

controls were exposed to PBS only. Biofilms were resuspended in PBS and added in a

ratio of 1:1 to LIVE/DEAD BacLight stain in a 96-well black microtitre plates.

Fluorescence was measured using a Wallac Victor2 1420 Multilabel Counter.

Measurement of extracellular DNA in biofilms using PicoGreen. Static biofilms were grown in sterile polystyrene cuvettes as described above. Supernatant from 24 h biofilms were removed and replaced with PBS containing TMP and irradiated for 5 min. Attached biofilms were resuspended in PBS. Resuspended cells were centrifuged for 2 min at

10,000 x g and 100µl of supernatant was transferred to a polystyrene microtitre plate containing 20µl/well of Pico Green Reagent (200-fold diluted dimethylsulfoxide, DMSO in TE buffer) (Molecular Probes Inc., Eugene, OR). The mixture was incubated at room temperature for 3 min in the dark. Fluorescence was measured using a Wallac Victor2

1420 Multilabel Counter.

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Combined TMP and tobramycin treatment. Biofilms were grown on glass slides as described above. The 24 h biofilms were incubated for 10 min in 225 µM TMP. Slides were rinsed in PBS and transferred to tubes containing MSG or MSG supplemented with

100µg ml-1 of tobramycin. Slides were incubated in tobramycin for 2 h at 37°C. All incubations were performed in the dark. Cell viability within biofilms was assessed using the LIVE/DEAD BacLight bacterial viability kit and visualized with an Olympus

FV1000 CSLM as described above.

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Results

TMP effects on biofilm structure and cell viability. The effect of TMP on 24 h P.

aeruginosa PAO1 biofilms was assessed using CSLM and viable plate counts. In the

absence of TMP, wild-type PAO1 cells formed dense biofilms on glass slides (Fig. 1A)

and in polystyrene cuvettes (data not shown). When wild-type PAO1 biofilms were

exposed to 100 µM TMP and irradiated for 10 min, there was a decrease in biofilm

density and the majority of cells within the biofilm were non-viable based on

LIVE/DEAD staining (Fig. 1C and Fig. 3). Exposure to 225 µM TMP and 10 min of

irradiation resulted in a nearly complete disruption and clearance of established wild-type

PAO1 biofilms (Fig. 1E). The few remaining attached cells were nonviable. Shorter

periods of light exposure or lower concentrations of TMP resulted in less clearance of the

biofilms (data not shown). Interestingly, biofilms exposed to TMP but not irradiated

appeared to be expanded in volume without a loss of cell viability (Fig. 1G).

Standard plate counts of cells recovered from biofilms formed in polystyrene

cuvettes were used to quantify the effects of photoactivated TMP. Wild-type biofilms exposed to 225 µM TMP and 10 min of irradiation exhibited a 4.1-log10 decrease in

viable cells in the attached biofilm population (Fig. 2). There was a 4.5-log10 reduction in the number of viable cells in the recovered supernatants of wild-type PAO1 biofilms following the same TMP treatment (Fig. 2). The recovered supernatants contained the cells sloughed off as the result of TPM exposure and irradiation.

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In contrast to wild-type biofilms, pqsA mutant biofilms were significantly

different in overall structure. As previously noted, these biofilms are not confluent

(Allesen-Holm et al. 2006) (Fig. 1B). When exposed to 100 or 225 µM TMP and

irradiated for 10 min there was a decrease in cell viability of attached cells (Fig. 1D and

1F, Fig. 3). Standard plate counts of attached cells showed a 3.9-log10 decrease in cell

viability at TMP concentrations of 225 µM (Fig. 2). Similarly, there was a 4.2-log10

reduction in cell viability of pqsA cells collected from supernatants of irradiated biofilms

treated with 225 µM TMP (Fig. 2). Although exposure to TMP and irradiation resulted

in cell death, this treatment did not lead to the disruption or clearance of the pqsA mutant

biofilms observed with wild-type cells. Without photoactivation, TMP did not affect cell

viability or disrupt the architecture of pqsA mutant biofilms (Fig. 1H)

DNA degradation in the presence of TMP. To determine the effect of TMP on DNA, pUCP18 plasmid DNA was exposed to TMP and irradiated. The untreated control samples had the three expected forms of plasmid DNA: covalently closed circles, relaxed circular and linear cut (Fig. 4: lanes 1 and 7). Plasmid DNA exposed to light only appeared similar to the control (Fig. 4: lanes 2 and 8).

Plasmid DNA treated with 100 or 225 µM TMP without subsequent photoactivation resulted in retarded mobility of DNA (Fig 4: lanes 3, 5, 9, and 11) as expected due to its ability to intercalate into DNA (12, 16). The combination of TMP and irradiation for either 5 or 30 min resulted in the complete degradation of pUCP18 plasmid

DNA at concentrations of 100 or 225 µM TMP (Fig. 4: lanes 4, 6, 10, and 12).

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To determine the effect of TMP on extracellular DNA, wild-type MPAO1

biofilms were exposed to TMP and irradiated. Extracellular DNA was collected from

biofilm supernatants and quantified using Pico Green. The amount of fluorescence is

directly proportional to the quantity of extracellular DNA present. When exposed to

TMP and irradiated, there was a decrease in fluorescence in comparison to untreated

biofilms (Fig.5). This decrease was more pronounced when biofilms were exposed to

higher concentrations of TMP.

In order to determine if the decrease in fluorescence was due to degradation of

extracellular DNA by TMP or TMP competing with Pico Green dye for DNA binding,

the assay was repeated using dark controls. There was a decrease in fluorescence when

biofilms were exposed to TMP in the dark (Fig. 6).

Effect of tobramycin on TMP treated biofilms. Wild-type PAO1 biofilms were

exposed to TMP for 10 min followed by exposure to 100 µg ml-1 of tobramycin for 2 h

with all steps performed in the dark. As noted above, TMP treatment without

photoactivation resulted in an expansion and loss of biofilm density with no observable

reduction in cell viability. In biofilms treated with tobramycin there was a reduction in

cell viability which was limited to cells near the surface of the biofilms, where oxygen is

most plentiful (Fig. 7A). Treatment with TMP and subsequent exposure to tobramycin resulted in substantial clearance of the biofilms and greater loss of cell viability throughout the biofilms than with either single treatment (Fig. 7B).

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wild-type PAO1 pqsA mutant A B A B

C D C D

EE F F

GG H

Figure 1. Confocal scanning laser micrographs of P. aeruginosa biofilms. Biofilms

were grown on glass slides for 24 h under static conditions in MSM and then exposed to

specified concentrations of TMP. Following exposure to TMP, biofilms were either

irradiated (A-F) with a 100-Watt mercury vapor lamp for 10 min or incubated in the dark

(G-H). Bacterial viability was determined using the LIVE/DEAD BacLight Bacterial

Viability assay. Cells staining red are considered dead while cells staining green are

alive. The images show horizontal optical sections from the midpoint of the biofilms

flanked by vertical optical sections in biofilms treated with: (A-B), No TMP, light; (C-D),

100 µM TMP, light; (E-F), 225 µM TMP, light; and (G-H) 225 µM TMP, dark.

109

1.00E+01

1.00E+00

1.00E-01

1.00E-02

1.00E-03

1.00E-04 Surviving Fraction Surviving

1.00E-05

1.00E-06

1.00E-07

1.00E-08 wild-type PAO1 pqsA mutant

Figure 2. Effect of TMP and light irradiation on cell survival of P. aeruginosa

biofilm associated cells. Established biofilms of wild-type PAO1 and the pqsA mutant were treated with TMP and irradiated 10 min with a 100-Watt mercury vapor lamp. Cells were collected from the supernatant of treated biofilms, as well as from the remaining attached biofilm. Cell suspensions were diluted and plated onto LB (1.5% agar) plates and incubated 24 h at 37°C. CFU were used to determine the surviving fraction.

Attached cells: no TMP , 225 µM TMP ; Supernatant cells: no TMP , 225 µM TMP

110

Green/Red Fluorescence Green/Red 15

0 no TMP 100uM TMP 225uM TMP

Figure 3. Effect of TMP and light irradiation on cell survival of P. aeruginosa biofilm associated cells via LIVE/DEAD BacLight stain. Established biofilms of wild- type PAO1 and the pqsA mutant were treated with TMP and irradiated 10 min with a

100-Watt mercury vapor lamp. Cells were collected from the supernatant of treated biofilms and from the remaining attached biofilm. Cells suspensions were mixed in 1:1 ratio with LIVE/DEAD BacLight stain and incubated in the dark for 15 min.

Fluorescence was measured using a Wallac Victor2 1420 Multilabel Counter. PAO1

(attached) , PAO1 (supernatant) , pqsA mutant (attached) , pqsA mutant

(supernatant) .

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1kb 1 2 3 4 5 6 7 8 9 10 11 12

Contributed by Elizabeth Anne Markus

Figure 4. Gel electrophoresis analysis of plasmid (pUCP18) DNA treated with TMP and irradiated. Purified plasmid (pUCP18) DNA (100 ng ml-1) was exposed to either

0 µM, 100 µM or 225 µM TMP and irradiated with a 100-Watt mercury vapor lamp for

0, 5, or 30 min. Lane 1: 0 TMP, nonirradiated control DNA; Lane 2: 0 TMP, irradiated

5 min, Lane 3: 100 µM TMP, nonirradiated, Lane 4: 100 µM TMP, irradiated 5 min,

Lane 5: 225 µM TMP, nonirradiated, Lane 6: 225 µM TMP, irradiated 5 min, Lane 7: 0

TMP, nonirradiated, Lane 8: 0 TMP, irradiated 30 min, Lane 9: 100 µM TMP, nonirradiated, Lane 10: 100 µM TMP, irradiated 30 min, Lane 11: 225 µM TMP, nonirradiated, and Lane 12: 225 µM TMP, irradiated 30 min.

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Figure 5. Quantification of extracellular DNA of wild-type MPAO1 biofilms treated with TMP and light irradiation. Wild-type MPA01 biofilms were treated with 10 µM,

100 µM, and 225 µM of TMP and irradiated 5 min with a 100-Watt mercury vapor lamp.

Attached biofilms were resuspended in PBS. Resuspended cells were centrifuged for 2 min at 10,000 x g and 100 µl of supernatant was transferred to a polystyrene microtitre plate containing 20 µl/well of Pico Green Reagent (200-fold diluted dimethylsulfoxide,

DMSO in TE buffer). The mixture was incubated at room temperature for 3 min in the dark. Fluorescence was measured using a Wallac Victor2 1420 Multilabel Counter.

MPA01 No TMP , MPA01 10 µM TMP ; MPA01 100 µM TMP , MPA01 225 µM

TMP .

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16000

14000

12000

10000

8000 Fluorescence

6000

4000

2000

0 light dark

Figure 6. Quantification of extracellular DNA of wild-type MPAO1 biofilms treated with TMP with and without light irradiation. Wild-type MPA01 biofilms were treated with 10 µM of TMP and irradiated 1 h with a 100-Watt mercury vapor lamp. A dark control was also treated with 10 µM of TMP for 5 min in PBS. Attached biofilms were resuspended in PBS. Resuspended cells were centrifuged for 2 min at 10,000 x g and

100 µl of supernatant was transferred to a polystyrene microtitre plate containing

20µl/well of Pico Green Reagent (diluted 200-fold dimethylsulfoxide (DMSO) in TE buffer). The mixture was incubated at room temperature for 3 min in the dark.

Fluorescence was measured using a Wallac Victor2 1420 Multilabel Counter. MPA01 no

TMP ; MPA01 with 10 µM TMP .

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A B

Figure 7. Confocal scanning laser micrographs of P. aeruginosa wild-type PAO1

biofilms treated with tobramycin only and TMP and tobramycin in the absence of light. Biofilms were grown on glass slides for 24 h at 37°C and then exposed to TMP

(225 µM) for 10 min in the dark. Following exposure to TMP, biofilms were incubated with tobramycin (100 µg ml-1) for 2 h at 37°C. Bacterial viability was determined using the LIVE/DEAD BacLight Bacterial Viability assay. Cells staining red are considered dead while cells staining green are alive. The images show horizontal optical sections from the midpoint of the biofilms flanked by vertical optical sections in biofilms treated with: (A) tobramycin; and (B) TMP + tobramycin.

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Discussion

In this study, we examined the effects of the cationic porphyrin, TMP, on

established P. aeruginosa biofilms. TMP exposure plus photoactivation resulted in a

substantial reduction in the numbers of viable bacteria within established wild-type P.

aeruginosa biofilms as shown by viability staining (Fig. 1C and 1E, Fig. 3) and standard

plate counts (Fig. 2). Bacterial killing required photoactivation, indicating that there was

no dark toxicity associated with TMP (Fig. 1G).

Previous studies have demonstrated that TMP at higher concentrations (5.0 mg

-1 ml ) than used in this study resulted in a 1.2-log10 reduction of wild-type PAO1 isolates

grown in biofilms when irradiated for 5 min (Donnelly et al. 2007). We were able to

achieve higher rates of killing (4.1-log10 reduction) of biofilm associated wild-type PAO1

cells using concentrations of TMP as low as 0.32 mg ml-1 (225 µM) (Fig. 2). This difference in killing rates can be attributed to the different conditions under which the established biofilms were grown and treated.

Previous studies did not evaluate the change in P. aeruginosa biofilm structure following treatment with TMP and light. In addition to killing biofilm associated

bacteria, treatment with TMP followed by irradiation resulted in substantial disruption

and clearance of wild-type PAO1 biofilms (Fig. 1C and 1E). At a concentration of 225

µM TMP, wild-type biofilms were completely disrupted with few cells remaining

attached (Fig. 1E). Without photoactivation, TMP did not lead to clearance of wild-type

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biofilms; however, there was a noticeable expansion in the biofilm and loss of density

(Fig. 1G). These results indicate that TMP photoactivation affects biofilms in two ways: direct killing of cells and the disruption of the biofilm architecture. Additionally, while

TMP toxicity is dependent on photoactivation, it is able to alter biofilm architecture in the absence of light by an unknown mechanism.

To determine if disruption of established P. aeruginosa wild-type biofilms was

solely due to inactivation of biofilm associated cells or also involved the extracellular

matrix of the biofilm, we examined the effects of TMP on extracellular DNA.

Extracellular DNA has previously been shown to be necessary for normal biofilm

formation and contributes to the overall architecture (Whitchurch et al. 2002). We

attempted to quantify extracellular DNA in biofilms following treatment with TMP and

photoactivation compared with untreated biofilms using various DNA stains, such as Pico

Green. We observed a decrease in extracellular DNA of biofilms treated with TMP and

light, but there was also a decrease in extracellular DNA of biofilms that were treated

with TMP in the absence of light (Fig. 5 and 6). Because both TMP and Pico Green

intercalate between base pairs, we were unable to determine if this decrease in

fluorescence was proportional to a reduction in extracellular biofilm matrix DNA or due

to competitive inhibition, i.e., the intercalation of TMP preventing binding of Pico Green.

Due to the difficulty in quantifying extracellular biofilm DNA following TMP

exposure using established staining techniques, we assessed the effects of TMP on

purified pUCP18 DNA and a pqsA mutant. Prior to irradiation, pUCP18 plasmid DNA mobility was retarded following exposure to TMP, indicating intercalation of TMP (Fig.

4, lanes 3, 5, 9, and 11). Exposure to TMP and subsequent photoactivation led to

117

complete degradation of pUCP18 plasmid DNA (Fig. 4, lanes 4, 6, 10, and 12). These results coincide with previous studies that demonstrated that TMP intercalates between

DNA base pairs, causing photocleavage of DNA (Kelly and Murphy 1985, Pasternack and Gibbs 1996). Thus, TMP is expected to intercalate into available extracellular DNA within biofilms and, upon irradiation, lead to disruption of the DNA. Degradation of extracellular DNA in the biofilm matrix using DNase has been previously shown to disrupt biofilm architecture and lead to the dissolution of the biofilm (Whitchurch et al.

2002). TMP photocleavage of DNA would similarly result in the disruption of biofilms.

The pqsA mutant, defective in a late portion of the P. aeruginosa quorum-sensing system, has been shown to produce biofilms with substantially lower levels of extracellular DNA (Allesen-Holm et al. 2006). In the presence of TMP and light, high levels of killing were observed in pqsA mutant biofilms (Fig. 1D and F, Fig. 3).

However, in contrast to wild-type biofilms, the biofilms formed by the pqsA mutant were not disrupted by this treatment. The inability of TMP photoactivation to disrupt pqsA biofilms could be attributed, in part, to the lack of extracellular DNA in these biofilms.

We conclude that disruption of P. aeruginosa PAO1 wild-type biofilms by TMP and light treatment is partially due to its effect on the extracellular DNA matrix and not just photoinactivation of the cells within the matrix. However, we acknowledge that the lack of dissolution of pqsA mutant biofilms by TMP photoactivation may not be solely due to differences in DNA content as these biofilm differ from wild-type biofilms in a number of important ways.

The ability of TMP to intercalate into DNA, leading to an unwinding and expansion of the DNA volume, could explain the expansion of wild-type PAO1 biofilms

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treated with TMP but not exposed to light. This change in the architecture of P.

aeruginosa biofilms treated with TMP in the absence of photoactivation led us to explore

how this might affect the ability of antibiotics to kill bacteria within biofilms. We

examined the combined effects of TMP and the antibiotic tobramycin in the dark on

established biofilms. Treatment of wild-type PAO1 biofilms with tobramycin, the major

front-line antibiotic used in the treatment of CF lung disease, did not result in substantial

biofilm clearance and led to minimal killing of biofilm associated cells (Fig. 7A). Killing

was primarily localized to the top layer of the biofilm. In contrast, exposure of wild-type

PAO1 biofilms to TMP prior to treatment with tobramycin resulted in significant biofilm

clearance and enhanced killing of cells (Fig. 7B). One of the limitations of photodynamic therapy is the delivery of light to infections in deep tissue. These findings are especially important because they show that TMP can act to disrupt biofilm structure when activated by light and also by a light independent mechanism that enhances killing when combined with tobramycin. A light independent treatment has the potential to be applied when trying to eradicate P. aeruginosa biofilms that are not easily accessible to irradiation such as those associated with cystic fibrosis patients.

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Literature Cited

1. Allesen-Holm, M., Barken, K.B., Yang, L., Klausen, M., Webb, J.S., Kjelleberg, S., Molin, S., Givskov, M., and T. Tolker-Nielsen. 2006. A characterization of DNA release in Pseudomonas aeruginosa cultures and biofilms. Mol. Microbiol. 59: 1114-1128.

2. Alley, M.R.K., Maddock, J.R., and L. Shapiro. 1992. Polar localization of a bacterial chemoreceptor. Genes Devel 6: 825-836.

3. Alm, R.A., Hallinan, J.P., Watson, A.A., and J.S. Mattick. 1996. Fimbrial biogenesis genes of Pseudomonas aeruginosa: pilW and pilX increase the similarity of type IV fimbriae to the GSP protein-secretion systems and pilY1 encodes a gonococcal PilC homologue. Mol. Microbiol. 22: 161-173.

4. Alm, R.A., and J.S. Mattick. 1995. Identification of a gene, pilV, required for type IV fimbrial biogenesis in Pseudomonas aeruginosa, whose product possesses a pre-pilin-like leader sequence. Mol. Microbiol. 16: 485-496.

5. Alm, R.A., and J.S. Mattick. 1996. Identification of two genes with prepilin-like leader sequences involved in type IV fimbrial biogenesis in Pseudomonas aeruginosa. J. Bacteriol. 178: 3809-3817.

6. Alm, R.A., and J.S. Mattick. 1997. Genes involved in the biogenesis and function of type IV fimbriae in Pseudomonas aeruginosa. Gene. 192: 89-98.

7. Bhatnagar, J., Borbat, P.P., Pollard, A.M., Bilwes, A.M., Freed, J.H., and B.R. Crane. 2010. Structure of the ternary complex formed by a chemotaxis receptor signaling domain, the CheA histidine kinase, and the coupling protein CheW as determined by pulsed dipolar ESR spectroscopy. Biochemistry 49: 3824-3841.

8. Bassler, B.L.. 2002. Small talk. Cell-to-cell communication in bacteria. Cell 109: 421-424.

9. Beatson, S.A., Whitchurch, C.B., Sargent, J.L., Levesque, R.C., and J.S. Mattick. 2002. Differential regulation of twitching motility and elastase production by Vfr in Pseudomonas aeruginosa. J. Bacteriol. 184: 3605-3613.

10. Blair, D.F. 2003. Flagellar movement driven by proton translocation. FEBS Lett.

120

545: 86-95.

11. Boyd, J.M. 2000. Localization of the histidine kinase PilS to the poles of Pseudomonas aeruginosa and identification of a localization domain. Mol. Microbiol. 2000: 153-162.

12. Bradley, D.E. 1980. A function of Pseudomonas aeruginosa PAO pili: twitching motility. Can. J. Microbiol. 26: 146-154.

13. Bray, D. 2002. Bacterial chemotaxis and the question of gain. Proc. Natl. Acad. Sci. USA 99: 7-9.

14. Brehm-Stecher, B.F., and E.A. Johnson. 2003. Sensitization of Staphylococcus aureus and Escherichia coli to antibiotics by the sesquiterpenoids nerolidol, farnesol, bisabolol, and apritone. Antimicrob. Agents Chemother. 47: 3357–3360.

15. Brinkman, F.S.L., Bains, M., and R.E.W. Hancock. 2000. The amino terminus of Pseudomonas aeruginosa outer membrane protein OprF forms channels in lipid bilayer membranes: correlation with a three dimensional model. J. Bacteriol. 182: 5251–5255.

16. Brown, A.J. 2002. Morphogenetic signaling pathways in Candida albicans in Candida and Candidiasis. Calderone, R.A. (edn.). Washington, DC: American Society for Microbiology Press: 95-106.

17. Burns, J., Van Dalfsen, J., Shawar, R., Otto, K., Garber, R., Quan, J., Montgomery, A.B., Albers, G.M., Ramsey, B.W., and A.L. Smith. 1999. Effect of intermittent administration of inhaled tobramycin on respiratory microbial flora in patients with cystic fibrosis. J. Infect. Dis. 179: 1190–1196.

18. Caiazza, N.C. and G.A. O’Toole. 2004. SadB is required for the transition from reversible to irreversible attachment during biofilm formation by Pseudomonas aeruginosa PA14. J. Bacteriol. 186: 4476-4485.

19. Caiazza, N.C., Shanks, R.M., and G.A. O’Toole. 2005. Rhamnolipids modulate swarming motility patterns in Pseudomonas aeruginosa. J. Bacteriol. 187: 7351- 7361.

20. Caiazza, N.C., Merritt, J.H., Brothers, K.M., and G.A. O'Toole. 2007. Inverse regulation of biofilm formation and swarming motility by Pseudomonas aeruginosa PA14.J. Bacteriol. 189: 3603-3612.

21. Calfee, M.W., Coleman, J.P., and E.C Pesci. 2001. Interference with Pseudomonas quinolone signal synthesis inhibits virulence factor expression by Pseudomonas aeruginosa. Proc. Natl. Acad. Sci. USA 98: 11633-11637.

121

22. Cao, H., Krishnan, G., Goumnerov, B., Tsongalis, J., Tompkins, R., and L.G. Rahme. 2001. A quorum-sensing-associated virulence gene of Pseudomonas aeruginosa encodes a LysR-like transcription regulator with a unique self- regulatory mechanism. Proc. Natl. Acad. Sci. USA 98: 14613-14618.

23. Chiang, P., Habash, M., and L.L. Burrows. 2005. Disparate subcellular localization patterns of Pseudomonas aeruginosa Type IV pilus ATPases involved in twitching motility. J. Bacteriol. 187: 829-839.

24. Chugani, S.A., Whiteley, M., Lee, K.M., D’Argenio, D., Manoil, C., and E.P. Greenberg. 2001. QscR, a modulator of quorum-sensing signal synthesis and virulence in Pseudomonas aeruginosa. Proc. Natl. Acad. Sci. USA 98: 2752- 2757.

25. Collier, D.N., Anderson, L., McKnight, S.L., Noah, T.L., Knowles, M., Boucher, R., Schwab, U., Gilligan, P., and E.C. Pesci. 2002. A bacterial cell to cell signal in the lungs of cystic fibrosis patients. FEMS Microbiol. Lett. 215: 41-46.

26. Collins, T.L., Markus, E.A., Hassett, D.J., and J.B. Robinson. 2010. The effects of a cationic porphyrin of P. aeruginosa biofilms. Curr. Microbiol. 61: 411-416.

27. Costerton, J.W., Stewart, P.S., and E.P. Greenberg. 1999. Bacterial biofilms: a common cause of persistent infections. Science 284: 1318-1322.

28. Costerton, J.W. 2001. Cystic fibrosis pathogenesis and the role of biofilms in persistent infection. Trends Microbiol. 9: 50-52.

29. Craven, R.C., and T.C. Montie. 1983. Chemotaxis of Pseudomonas aeruginosa: involvement of methylation. J Bacteriol. 154: 780-786.

30. Craven, R.C., and T.C. Montie. 1985. Regulation of Pseudomonas aeruginosa chemotaxis by nitrogen source. J. Bacteriol. 164: 544-549.

31. Croft, L., Beatson, S. A., Whitchurch, C. B., Huang, B., Blakeley, R. L., and J.S. 2000. An interactive web-based Pseudomonas aeruginosa genome database: discovery of new genes, pathways and structures. Microbiology 146: 2351-2364.

32. Cugini, C., Worth Calfee, M., Farrow, J.M, Morales, D.K., Pesci, E.C., and D.A. Hogan. 2007. Farnesol, a common sesquiterpene, inhibits PQS production in Pseudomonas aeruginosa. Mol. Microbiol. 65: 896-906.

33. D’Argenio, D.A., Calfee, M.W., Rainey, P.B., and E.C. Pesci. 2002. Autolysis and autoagggregation in Pseudomonas aeruginosa colony morphology mutants. J. Bacteriol. 184: 6481-6489.

122

34. Darzins, A. 1994. Characterization of a Pseudomonas aeruginosa gene cluster involved in pilus biosynthesis and twitching motility: sequence similarity to the chemotaxis protein of enterics and the gliding bacterium Myxococcus xanthus. Mol. Microbiol. 11: 137-153.

35. Darzins, A., and M.A. Russell. 1997. Molecular genetic analysis of type-4 pilus biogenesis and twitching motility using Pseudomonas aeruginosa as a model system. Gene 192: 109-115.

36. Davies, D.G., Parsek, M.R., Pearson, J.P., Iglewski, B.H, Costerton, J.W., and E.P. Greenberg. 1998. The involvement of cell-to-cell signals in the development of a biofilm. Science 280: 295-298.

37. de Kievit, T.R., and B.H. Iglewski. 2000. Bacterial quorum-sensing in the pathogenic relationships. Infect. Immun. 68: 4839-4849.

38. DeLange, P.A., Collins, T.L., Pierce, G.E., and J.B Robinson. 2007. PilJ localizes to cell poles and is required for type IV pilus extension in Pseudomonas aeruginosa. Curr. Microbiol. 55: 389-395.

39. Deziel, E., Lepine, F., Milot, S., and R. Villemur. 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 149: 2005–2013.

40. Deziel, E., Lepine, F., Milot, S., He, J., Mindrinos, M.N., Tompkins, R.G., and L.G. Rahme. 2004. Analysis of Pseudomonas aeruginosa 4-hydroxy-2- alkylquinolines (HAQs) reveals a role for 4-hydroxy-2-heptylquinoline in cell-to- cell communication. Proc. Natl. Acad. Sci. USA 101: 1339-1344.

41. Di Poto, A., Sbarra, M., Provenza, G., Visai, L., and P. Speziale. 2009. The effect of photodynamic treatment combined with antibiotic action or host defence mechanisms on Staphylococcus aureus biofilms. Biomaterials 30: 3158-3166.

42. Dong, Y.H., Zhang, X.F., Soo, H.M., Greenberg, E.P., and L.H. Zang. 2005. The two-component response regulator PprB modulates quorum-sensing signal production and global gene expression in Pseudomonas aeruginosa. Mol. Microbiol. 56: 1287-1301.

43. Donlan R.M. 2000. Role of biofilms in antimicrobial resistance. ASAIO J. 46: 47–52.

44. Donlan, R.M., and J.W. Costerton. 2002. Biofilms: Survival Mechanisms of Clinically Relevant Microorganisms. Clin. Microbiol. Rev. 15:167–193.

123

45. Donnelly, R., McCarron, P., Cassidy, C., Elborn, J.S., and M. Tunney. 2007. Delivery of photosensitisers and light through mucus: Investigations into the potential use of photodynamic therapy for treatment of Pseudomonas aeruginosa cystic fibrosis pulmonary infection. J. Controlled Release 117: 217-226.

46. Dowd, S.E., Sun, Y., Smith, E., Kennedy, J.P., Jones, C.E., and R. Wolcott. 2009. Effects of biofilm treatments on the multi-species Lubbock chronic wound biofilm model. J. Wound Care 18: 508, 510-512.

47. Dunne, W.M. 2002. Bacterial adhesion: seen any good biofilms lately? Clin. Microbiol. Rev. 15: 155-166.

48. El Hamel, C., Freulet, M.A., Jaquinod, M., De, E., Molle, G., and N. Orange. 2000. Involvement of the C-terminal part of Pseudomonas fluorescens OprF in the modulation of its pore-forming properties. Biochem. Biophys. Acta 1509: 237–244.

49. Emori, T.G., and R.P. Gaynes. 1993. An overview of nosocomial infections, including the role of the microbiology laboratory. Microbiol. Rev. 6: 428-442.

50. Evans, L.R. and A. Linker. 1973. Production and characterization of the slime polysaccharide of Pseudomonas aeruginosa. J. Bacteriol. 116: 915-924.

51. Faulke, J.J, Bass, R.B., Butler, S.L., Chervitz, S.A., and M.A. Danielson. 1997. The two-component signaling pathway of bacterial chemotaxis: a molecular view of signal transduction by receptors, kinases, and adaptation enzymes. Annu. Rev. Cell Dev. Biol. 13: 457-512.

52. Ferrandez, A., Hawkins, A.C., Summerfield, D.T., and C.S. Harwood. 2002. Cluster II che genes from Pseudomonas aeruginosa are required for an optimal chemotactic response. J Bacteriol. 184: 4374-4383.

53. Flemming, H-C, Wingender, J., Griegbe, and C. Mayer. 2000. Physico-chemical properties of biofilms. In: Evans LV, editor. Biofilms: recent advances in their study and control. Amsterdam: Harwood Academic Publishers: 19–34.

54. Foote, C. S. 1990. Future directions and applications in photodynamic theory. SPIE Institute Series 1S6: 115–126.

55. Forest, K.T. and J.A. Tainer. 1997. Type IV pilus structure: outside to inside and top and bottom. Gene 192: 165-169.

56. Francis, N.R., Sosinsky, G.E., Thomas, D., and D.J. DeRosier. 1994. Isolation, characterization and structure of bacterial flagellar motors containing the switch complex. J. Mol. Biol. 235: 1261-1270.

124

57. Fraser, G.M., and C. Hughes. 1999. Swarming motility. Curr. Opin. Microbiol. 2: 630-635.

58. Freedman, D.L., Swamy, M., Bell, N.C., and M.F. Verce. 2004. Biodegradation of chloromethane by Pseudomonas aeruginosa strain NB1 under nitrate-reducing and aerobic conditions. Appl. Environ. Microbiol. 70: 4629-4634.

59. Freulet-Marriere, M.A., El Hamel, C., Chevalier, S., De, E., Molle, G., and N. Orange. 2000. Evidence for association of lipopolysaccharide with Pseudomonas fluorescens strain MF0 porin OprF. Res. Microbiol. 151: 873–876.

60. Friedman, L. and R. Kolter. 2004. Two genetic loci produce distinct carbohydrate-rich structural components of the Pseudomonas aeruginosa biofilm matrix. J. Bacteriol. 186: 4457-4465.

61. Friedman, L. and R. Kolter. 2004. Genes involved in matrix formation in Pseudomonas aeruginosa PA14 biofilms. Mol. Microbiol. 51: 675-690.

62. Fujita, M., Amemura, A., and H. Aramaki. 1998. Transcription of the groESL operon in Pseudomonas aeruginosa PAO1. FEMS Microbiol. Lett. 163: 237-242.

63. Fuxman Bass, J.I., Russo, D.M., Gabelloni, M.L., Geffner, J.R., Giordano, M., Catalano, M., Zorreguieta, A., Trevani, A.S. 2010. Extracellular DNA: a major proinflammatory component of Pseudomonas aeruginosa biofilms. J. Immunol. 184: 6386-6395.

64. Gacesa P. 1998. Bacterial alginate biosynthesis: recent progress and future prospects. Microbiology 144:1133–1143.

65. Gallagher, L.A., McKnight, S.L., Kuznetsova, M.S., Pesci, E.C., and C. Manoil. 2002. Functions required for extracellular quinolone signaling by Pseudomonas aeruginosa. J. Bacteriol. 184: 6472-6480.

66. Gambello, M.J., and B.H. Iglewski. 1991. Cloning and characterization of the Pseudomonas aeruginosa lasR gene, a transcriptional activator of elastase expression. J. Bacteriol. 173: 3000–3009.

67. Girotti, A. W. 1990. Photodynamic lipid peroxidation in biological systems. J. Photochem. and Photobiol. 51: 497–509.

68. Glessner, A., Smith, R.S., Iglewski, B.H., and J.B. Robinson. 1999. Roles of Pseudomonas aeruginosa las and rhl quorum-sensing systems in control of twitching motility. J. Bacteriol. 181: 1623-1629.

69. Govan, J., and J. Fyfe. 1978. Mucoid Pseudomonas aeruginosa and cystic fibrosis: resistance of the mucoid form to carbenicillin, flucloxacillin, and

125

tobramycin and the isolation of mucoid variants in vitro. J. Antimicrob. Chemother. 4: 233–240.

70. Govan, J., and V. Deretic. 1996. Microbial pathogenesis in cystic fibrosis: mucoid Pseudomonas aeruginosa and Burkholderia cepacia. Microbiol. Rev. 60: 539– 574.

71. Gow, N.A., Brown, A.J., and F.C. Odds. 2002. Fungal morphogenesis and host invasion. Curr. Opin. Microbiol. 5: 366–371.

72. Hahn, H.P. 1997. The type IV pilus is the major virulence-associated adhesion of Pseudomonas aeruginosa-a review. Gene 192: 99-108.

73. Hamblin, M.R., and T. Hasan. 2004. Photodynamic therapy: a new antimicrobial approach to infectious disease. Photochem. Photobiol. Sci. 3: 436-450.

74. Hamilton, P.B., and G. Sheely. 1971. Chemotactic response to amino acids by Pseudomonas aeruginosa in a semisolid nitrate medium. J. Bacteriol. 108: 596- 598.

75. Harshey, R.M. 1994. Bee’s aren’t the only ones: swarming in gram-negative bacteria. Mol. Microbiol. 13: 389-394.

76. Harshey, R.M. 2003. Bacterial motility on a surface: many ways to a common goal. Annu. Rev. Microbiol. 57: 249-273.

77. Hassett, D.J., Sutton, M.D., Schurr, M.J., Herr, A.B., Caldwell, C.C., and J.O. Matu. 2009. Pseudomonas aeruginosa hypoxic or anaerobic biofilm infections within cystic fibrosis airways. Trends Microbiol. 17: 130-138.

78. Hazelbauer, G.L., Falke, J.J., and J.S. Parkinson. 2008. Bacterial chemoreceptors: high performance signaling in networked arrays. Trends Biochem. Sci. 33: 9-19.

79. Hermann, C., Hermann, J., Munzel, U., and Ruchel, R. 1999. Bacterial flora accompanying Candida yeasts in clinical specimens. Mycoses 42: 619–627.

80. Hess, J.F., Oosawa, K., Kaplan, N., and M.I. Simon. 1988. Phosphorylation of three proteins in the signaling pathway of bacterial chemotaxis. Cell 53: 79-87.

81. Hirota, N., and Y. Imae. 1983. Na+-driven flagellar motors of an alkalophilic Bacillus strain YN-1. J. Biol. Chem. 258: 10577-10581.

82. Hobbs, M., Collie, E.S., Free, P.D., Livingston, S.P., and J.S. Mattick. 1993. PilS and PilR, a two component transcriptional regulatory system controlling

126

expression of type IV fimbriae in Pseudomonas aeruginosa. Mol. Microbiol. 7: 669-682.

83. Hobbs, M., and J.S. Mattick. 1993. Common components in the assembly of type IV fimbriae, DNA transfer systems, filamentous phage and protein-secretion apparatus: a general system for the formation of surface-associated protein complexes. Mol. Microbiol. 10: 233-243.

84. Hogan, D.A., Vik, A., and R. Kolter. 2004. A Pseudomonas aeruginosa quorum- sensing molecule influences Candida albicans morphology. Mol. Microbiol. 54: 1212-1223.

85. Hornby, J. M., Jensen, E.C., Lisec, A.D., Tasto, J.J., Jahnke, B., Shoemaker, R., Dussault, P., and K.W. Nickerson. 2001. Quorum-sensing in the dimorphic fungus Candida albicans is mediated by farnesol. Appl. Environ. Microbiol. 67: 2982–2992.

86. Ikeda, T., Homma, M., Iino, T., Asakura, S., and R. Kamiya. 1987. Localization and stoichiometry of hook-associated proteins within Salmonella typhimurium flagella. J. Bacteriol. 169: 1168–1173.

87. Inoue, Y., Shiraishi, A., Hada, T., Hirose, K., Hamashima, H., and J. Shimada. 2004. The antibacterial effects of terpene alcohols on Staphylococcus aureus and their mode of action. FEMS Microbiol. Lett. 237: 325–331.

88. Jabra-Rizk, M.A., Meiller, T.F., James, C.E., and M.E. Shirtliff. 2006. Effect of farnesol on Staphylococcus aureus biofilm formation and antimicrobial susceptibility. Antimicrob. Agents Chemother. 50: 1463–1469.

89. Juhas, M, Wiehlmann, L., and B. Huber. 2004. Global regulation of quorum- sensing and virulence by VqsR in Pseudomonas aeruginosa. Microbiology 150: 831-841.

90. Juhas, M., Eberl, L., and B. Tummler. 2005. Quorum-sensing: the power of cooperation in the world of Pseudomonas. Environ. Microbiol. 7: 459-471.

91. Kaiser, D. 2000. Bacterial motility: How do pili pull? Curr. Biol. 10: 777-780.

92. Kato, J., Ito, A., Nikata, T., and H. Ohtake. 1992. Phosphate taxis in Pseudomonas aeruginosa. J. Bacteriol. 174: 5149-5151.

93. Kearns, D.B., Robinson, J., and L.J. Skimkets. 2001. Pseudomonas aeruginosa exhibits directed twitiching motility up phosphatidylethanolamine gradients. J. Bacteriol. 183: 763-767.

127

94. Keizer, D.W., Slupsky, C.M., Kalisiak, M., Campbell, A.P., Crump, M.P., Sastry, P.A., Hazes, B., Irvin, R.T., and B.D. Sykes. 2001. Structure of a pilin monomer from Pseudomonas aeruginosa: implications for the assembly of pili. J. Biol. Chem. 276: 24186-24193.

95. Kelly, J.M., and M.J. Murphy. 1985. A comparative study of the interaction of 5,10,15,20-tetrakis(N-methylpyridinum-4-yl)porphyrin and its zinc complex with DNA using fluorescence spectroscopy and topoisomerisation. Nucleic Acids Res. 13: 167-184.

96. Kelly-Wintenberg, K., and T.C. Montie. 1994. Chemotaxis to oligopeptides by Pseudomonas aeruginosa. Appl. Environ. Microbiol. 60: 363-367.

97. Kerr, J.R. 1994. Suppression of fungal growth exhibited by Pseudomonas aeruginosa. J Clin. Microbiol. 32: 525–527.

98. Khan, I.H., Reese, T.S., and S. Khan. 1992. The cytoplasmic component of the bacterial flagellar motor. Proc. Natl. Acad. Sci. USA 89: 5956-5960.

99. Kharazmi, A. 1991. Mechanisms involved in the evasion of the host immune defence by Pseudomonas aeruginosa. 30: 201-205.

100. Kim, K.K., Yokota, H., and S.H. Kim. 1999. Four-helical-bundle structure of the cytoplasmic domain of a serine chemotaxis receptor. Nature 400:787-792.

101. Kim, S., Wang, W., and K.K. Kim. 2002. Dynamic and clustering model of bacterial chemotaxis receptors: Structural basis for signaling and high sensitivity. Proc. Natl. Acad. Sci. U.S.A. 99:11611-11615.

102. Kohler, T., Kocjancic Curty, L., Barja, F., van Delden, C., and J. Pechere. 2000. Swarming motility of Pseudomonas aeruginosa is dependent on cell-to-cell signaling and requires flagella and pili. J. Bacteriol. 182: 5990-5996.

103. Kojima, S., and D.F. Blair. 2001. Conformational change in the stator of the bacterial motor. Biochemistry 40: 13041-13050.

104. Korytowski, W., Bachowski, G. J., and Girotti, A. W. 1992. Photoperoxidation of cholesterol in homogeneous solution, isolated membranes, and cells: comparison of the 5 - and 6 - hydroperoxides as indicators of singlet oxygen intermediacy. J. Photochemist and Photobiol. 56: 1–8.

105. Kuroda, A., Kumano, T., Taguchi, K., Nikata, T., Kato, J., and H. Ohtake. 1995. Molecular cloning and characterization of a chemotactic transducer gene in Pseudomonas aeruginosa. J Bacteriol. 177:7019-7025.

128

106. Lambrechts, S., Aalders, M., Verbraak, F., Lagerberg, J., Dankert, J., and J. Schuitmaker. 2005. Effect of albumin on the photodynamic inactivation of microorganisms by a cationic porphyrin. J. Photochem. Photobiol. 79: 51-57.

107. Larsen, S.H., Adler, J., Gargus, J.J., and R.W. Hogg. 1974. Chemomechanical coupling without ATP: the source of energy for motility and chemotaxis in bacteria. Proc. Natl. Acad. Sci. USA 71: 1239-1243.

108. Lautrop, H. 1961. Bacterium anitratum transferred to the genus Cytophaga. Int. Bull. Bacteriol. Nomencl. 11:107-108.

109. Lazdunski, A.M., Ventre, I., and J.N. Sturgis. 2004. Regulatory circuits and communication in gram-negative bacteria. Nat. Rev. Microbiol. 7: 581–592.

110. Lefman, J., Zhang, P., Hirai, T., Weis, R.M., Juliani, J., Bliss, D., Kessel, M., Bos, E., Peters, P., and S. Subramaniam. 2004. Three-dimensional electron microscopic imaging of membrane invaginations in Escherichia coli overproducing the chemotaxis Receptor Tsr. J. Bacteriol. 186:5052-5061.

111. Ledgham, F., Ventre, I., Soscia, C., Foglino, M., Sturgis, J.N., and A. Lazdunski. 2003. Interactions of the quorum-sensing regulator QscR: interaction with itself and the other regulators of Pseudomonas aeruginosa LasR and RhlR. Mol. Microbiol. 48: 199-210.

112. Lo, H.J., Kohler, J.R., DiDomenico, B., Loebenberg, D., Cacciapuoti, A., and G.R. Fink. 1997. Nonfilamentous Candida albicans mutants are avirulent. Cell 90: 939–949.

113. Lory, S., and M.S. Strom. 1997. Structure-function relationship of type IV prepilin peptidase of Pseudomonas aeruginosa-a review. Gene 192:117-121.

114. Lui, J.D., and J.S. Parkinson. 1991. Genetic evidence for interaction between the CheW and Tsr proteins during chemoreceptor signaling by Escherichia coli. J. Bacteriol. 173: 4941-4951.

115. Lyczak, J.B., Cannon, C.L., and G.B. Pier. 2000. Establishment of Pseudomonas aeruginosa infection: lessons from a versatile opportunist. Microbes Infect. 2: 1051-1060.

116. McAlester G, O'Gara F, and J.P. Morrisey. 2008. Signal-mediated interactions between Pseudomonas aeruginosa and Candida albicans. J. Med. Microbiol. 57: 563-569.

117. Maddock, J. R., and L. Shapiro. 1993. Polar location of the chemoreceptor complex in the Escherichia coli cell. Science 259: 1717-1723.

129

118. Madigan, M.T., Martinko, J.M, and J. Parker. 1997. Biology of microorganisms. Prentice Hall, Upper Saddle River, NJ: 244.

119. Maier, R.M., and G. Soberon-Chavez. 2000. Pseudomonas aeruginosa rhamnolipids: biosynthesis and potential applications. Appl. Microbiol. Biotechnol. 54: 625-633.

120. Malik, Z., Ladan, H., and Y. Nitzan. 1992. Photodynamic inactivation of Gram- negative bacteria: problems and possible solutions. J. Photochem. Photobiol. 14: 262-266.

121. Martin, P.R., Hobbs, M., Free, P.D., Jeske, Y., and J.S. Mattick. 1993. Characterization of pilQ, a new gene required for the biogenesis of type IV fimbriae in Pseudomonas aeruginosa. Mol. Microbiol. 9: 857-868.

122. Masduki, A., Nakamura, J., Ohga, T., and Umezaki, R., Kato, J., and H. Ohtake. 1995. Isolation and characterization of chemotaxis mutants and genes of Pseudomonas aeruginosa. J. Bacteriol. 177: 948-952.

123. Matsukawa, M., and E.P. Greenberg. 2004. Putative exopolysaccharide synthesis genes influence Pseudomonas aeruginosa biofilm development. J. Bacteriol. 186: 4449-4456.

124. Mattick, J.S. 2002. Type IV pili and twitching motility. Annu. Rev. Microbiol. 56: 289-314.

125. Merchat, M., Bertolini, G., Giacomini, P., Villanueva, A., and G. Jori. 1996. Meso-substituted cationic porphyrins as efficient photosensitizers of Gram- positive and Gram-negative bacteria. J. Photochem. Photobiol. 32: 153-157.

126. Milburn, M.V., Prive, G.G., Milligan, D.L., Scott, W.G., Yeh, J., Jancarik, J., Koshland, D.E. Jr, and S.H. Kim. 1991. Three-dimensional structures of the ligand-binding domain of the bacterial aspartate receptor with and without a ligand. Science 254: 1342-1347.

127. Miller, M.B., and B.L. Bassler. 2001. Quorum-sensing in bacteria. Annu. Rev. Microbiol. 55: 165-199.

128. Moench, T. T., and W.A. Konetzka. 1978. Chemotaxis in Pseudomonas aeruginosa. J. Bacteriol. 133: 427-429.

129. Monroe, D. 2007. Looking for Chinks in the Armor of Bacterial Biofilms. PLoS Biology 5: 2458-2461.

130. Mosel, D.D., Dumitru, R., Hornby, J.M., Atkin, A.L., and K. W. Nickerson. 2005. Farnesol concentrations required to block germ tube formation in Candida

130

albicans in the presence and absence of serum. Appl. Environ. Microbiol. 71: 4938–4940.

131. Moulton, R.C., and T.C. Montie. 1979. Chemotaxis by Pseudomonas aeruginosa. J. Bacteriol. 137: 427-429.

132. Nealson, K.H., Platt, T., and J.W. Hastings. 1970. Cellular control of the synthesis and activity of the bacterial luminescent system. J. Bacteriol. 104: 313– 322.

133. Nemoto, K., Hirota, K., Mukarami, K., Taniguti, K., Murata, H., Viducic, D, and Y. Miyake. 2003. Effect of Varidase (streptodornase) on biofilms formed by Pseudomonas aeruginosa. Chemotherapy 49: 121-125.

134. Nickel, J.C., Ruseska, I., Wright, J.B., J.W. Costerton. 1985. Tobramycin resistance of Pseudomonas aeruginosa cells growing as a biofilm on urinary tract catheter. Antimicrob. Agents Chemother. 27: 619-624.

135. Nitzan, Y., Wexler, H. M., and S.M. Finegold. 1994. Inactivation of anaerobic bacteria by various photosensitized porphyrins or by hemin. Curr. Microbiol. 29: 125–131.

136. Nunn, D., Bergman, S., and S. Lory. 1990. Products of three accessory genes, pilB, pilC, and pilD, are required for biogenesis of Pseudomonas aeruginosa pili. J. Bacteriol. 172: 2911-2919.

137. Ochsner, U.A., and J. Reiser. 1995. Autoinducer-mediated regulation of rhamnolipid biosurfactant synthesis in Pseudomonas aeruginosa. Proc. Natl. Acad. Sci. USA 92: 6424-6428.

138. O’Toole, G.A., and R. Kolter. 1998. Flagellar and twitching motility are necessary for Pseudomonas aeruginosa biofilm development. Mol. Microbiol. 30: 295-304.

139. Parkins, M.D., Ceri, H., and D.G. Storey. 2001. Pseudomonas aeruginosa GacA, a factor in multihost virulence, is also essential for biofilm formation. Mol. Microbiol. 40: 1215-1226.

140. Passador, L., Cook, J.M., Gambello, M.J., Rust, L., and B.H. Iglewski. 1993. Expression of Pseudomonas aeruginosa virulence genes requires cell-to-cell communication. Science 260: 1127–1130.

141. Pasternack, R.F., and E.J Gibbs. 1996. Porphyrin and metalloporphyrin interactions with nucleic acids. Met. Ions Biol. Syst. 33: 367-397.

131

142. Pepe, J.C., and S. Lory. 1998. Amino acid substitutions in PilD, a bifunctional enzyme of Pseudomonas aeruginosa. Effect on leader peptidase and N- methyltransferase activities in vitro and in vivo. 273: 19120-19129.

143. Pesci, E.C., Pearson, J.P., Seed, P.C., and B.H. Iglewski. 1997. Regulation of las and rhl quorum-sensing in Pseudomonas aeruginosa. J. Bacteriol. 179: 3127- 3132.

144. Pesci, E.C., Milbank, J.B., Pearson, J.P., McKnight, S., Kende, A.S, Greenberg, E.P., and B.H. Iglewski. 1999. Quinolone signaling in the cell-to-cell communication system of Pseudomonas aeuginosa. Proc. Natl. Acad. Sci. USA 96: 11229-11234.

145. Rahim, R., Ochsner, U.A., Olvera, C., Graninger, M., Messner, P., Lam, J.S., and G. Soberon-Chavez. 2001. Cloning and functional characterization of Pseudomonas aeruginosa rhlC gene that encodes rhamnosyltransferase 2, an enzyme responsible for di-rhamnolipid biosynthesis. Mol. Microbiol. 40: 708- 718.

146. Rashid, M.H., and A. Kornberg. 2000. Inorganic polyphosphate is needed for swimming, swarming, and twitching motilities of Pseudomonas aeruginosa. Proc. Natl. Acad. Sci. USA 97: 4885-4890.

147. Robertson, C.A., Hawkins-Evans, D., and H. Abrahamse. 2009. Photodynamic therapy (PDT): A short review on cellular mechanisms and cancer research applications for PDT. J. Photochem. Photobiol. 96: 1-8.

148. Saint, N., El Hamel, C., De, E., and G. Molle. 2000. Ion channel formation by N-terminal domain: a common feature of OprFs of Pseudomonas and OmpA of Escherichia coli. FEMS Microbiol. Lett. 190: 261–265.

149. Sambrook, J., Fritsch, E.F., and T. Maniatis. 1989. Molecular cloning: a laboratory manual. 2nd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY

150. Sambrook, J., and Russell, D. 2001. Molecular cloning: a laboratory manual. 3rd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.

151. Sauer, K., Camper, A.K., Ehrlich, G.D., Costerton, J.W., and D.G. Davies. 2002. Pseudomonas aeruginosa displays multiple phenotypes during development as a biofilm. J. Bacteriol. 184: 1140-1154.

152. Schooling, S.R., Hubley, A. and T.J. Beveridge. 2009. Interactions of DNA with biofilm-derived membrane vesicles. J. Bacteriol. 191: 4097-4102.

132

153. Schulmeister, S., Ruttorf, M., Thiem, S., Kentner, D., Lebiedz, D., and V. Sourjik. 2008. Protein exchange dynamics at chemoreceptor clusters in Escherichia coli. Proc. Natl. Acad. Sci. USA 105: 6403-6408.

154. Seed, P.C., Passador, L., and B.H. Iglewski. 1995. Activation of the Pseudomonas aeruginosa lasI gene by LasR and the Pseudomonas autoinducer PAI: an autoinduction regulatory heirarchy. J. Bacteriol. 177: 654-659.

155. Semighini, C., Hornby, J., Dumitru, R., Nickerson, K., and S. Harris. 2006. Farnesol-induced apoptosis in Aspergillus nidulans reveals a possible mechanism for antagonistic interactions between fungi. Mol. Microbiol. 59: 753–764.

156. Shu, C.T., Ulrich, L.E., and I.B. Zhulin. 2003. The NIT domain: a predicted nitrate-responsive module in bacterial sensory receptors. Trends Biochem. Sci. 28: 121-124.

157. Skerker, J.M., and H.C. Berg. 2001. Direct obervation of extention and retraction of type IV pili. Proc. Natl. Acad. Sci. 98: 6901-6904.

158. Snyder, J., Lambert, L., and P. Ogilby. 2006. 5,10,15,20-tetrakis(N-methyl-4- pyridyl)-21H,23H-porphine (TMPyP) as a sensitizer for singlet oxygen imaging in cells: characterizing the irradiation-dependent behavior of TMPyP in a single cell. Photochem. Photobiol. 82: 177-184.

159. Sowa, Y., and R.M. Berry. 2008. Bacterial flagellar motor. Q. Rev. Biophys. 41: 103-132.

160. Speer, A.G., Cotton, P.B., Rode, J., Seddon, A.M., Neal, C.R., Holton, J., and J.W. Costerton. 1988. Biliary stent blockage with bacterial biofilm. A light and electron microscopy study. Ann. Intern. Med. 108: 546-553.

161. Springer, M.S., Goy, M.F., and J. Adler. 1977. Sensory transduction in Escherichia coli: two complementary pathways of information processing that involve methylated proteins. Proc. Natl. Acad. Sci. U.S.A. 74: 279-284.

162. Springer, W.R, and D.E. Koshland. 1997. Identification of a protein methyltransferase as the cheR gene product in the bacterial sensing system. Proc. Natl. Acad. Sci. U.S.A. 74: 533-537.

163. Stock, J.B., and D.E. Koshland. 1978. A protein methylesterase involved in bacterial sensing. Proc. Natl. Acad. Sci. U.S.A. 75: 3659-3663.

164. Stover, C.K., Pham, X.Q., Erwin, A.L., Mizoguchi, S.D., Warrener, P., Hickey, M.J., Brinkman, F.S., Hufnagle, W.O., Kowalik, D.J., Lagrou, M., Garber, R.L., Goltry, L., Tolentino, E., Westbrock-Wadman, S., Yuan, Y., Brody, L.L., Coulter, S.N., Folger, K.R., Kas, A., Larbig, K., Lim, R., Smith, K., Spencer, D.,

133

Wong, G.K., Wu, Z., Paulsen, I.T., Reizer, J., Saier, M.H., Hancock, R.E., Lory, S., and M.V. Olson. 2000. Complete genome sequence of Pseudomonas aeruginosa PA01, an opportunistic pathogen. Nature 406: 959-964.

165. Strom, M.S., and S. Lory. 1993. Structure-function and biogenesis of the type IV pili. Annu. Rev. Microbiol. 47: 565-596.

166. Tennet, J.M., and J.S. Mattick. 1994. Type IV fimbriae. In Fimbriae: Aspects of adhesion, genetics, biogenesis, and vaccines, ed. P Klemm: 127-146. Boca Raton: CRC.

167. Tindall, M.J., Porter, S.L., Maini, P.K., and J.P. Armitage. 2010. Modeling chemotaxis reveals the role of reversed phosphotransfer and a bi-functional kinase-phosphatase. PLos Comput. Biol. 6: 1-10.

168. Toker, A.S., and R.M. Macnab. 1997. Distinct regions of bacterial flagellar switch protein FliM interact with FliG, FliN and CheY. J. Mol. Biol. 273: 623- 634.

169. Toutain, C.M., Zegans, M.E.T., and G.A. O’Toole. 2005. Evidence for two flagellar stators and their role in the motility of Pseudomonas aeruginosa. J. Bacteriol. 187: 771-777.

170. van Delden, C., and B.H. Iglewski. 1998. Cell-to-cell signaling and Pseudomonas aeruginosa infections. Infect. Dis. 4: 551-560.

171. van Delden, C. 2004. Virulence factors in Pseudomonas aeruginosa. In: Ramos J-L, edn. Pseudomonas: virulence and gene regulation. New York, Kluwer Academic/Plenum Publishers: 3-45.

172. Venturi, V. 2006. Regulation of quorum-sensing in Pseudomonas. FEMS Microbiol. Rev. 30: 274-291.

173. Wade, D.S., Calfee, M.W., Rocha, E.R., Ling, E.A., Engstrom, E., Coleman, J.P., and E.C. Pesci. 2005. Regulation of Pseudomonas quinolone signal synthesis in Pseudomonas aeruginosa. J. Bacteriol. 187: 4372-4380.

174. Wagner, V.E. Bushnell, D., Passador, L., Brooks, A.I., and B.H. Iglewski. 2003. Microarray analysis of the Pseudomonas aeruginosa quorum-sensing regulons: effects of growth phase and environment. J. Bacteriol. 185: 2080-2095.

175. Wainwright, M. 1998. Photodynamic antimicrobial chemotherapy. J. Antimicrobial Chemotherapy. 42: 13-28.

134

176. Wainwright, M., Phoenix, D.A., Nickson, P.B., Morton, L.H.G. 2002. The use of new methylene blue in Pseudomonas aeruginosa biofilm destruction. Biofouling 18: 247–249.

177. Wainwright, M. and K.B. Crossley. 2004. Photosensitising agents – circumventing resistance and breaking down biofilms: a review. International Biodeterioration and Biodegradation. 53: 119-126.

178. Wainwright, M. 2008. Photodynamic Therapy: The Development of New Photosensitisers. Anti-Cancer Agents in Medicinal Chemistry. 8: 280-291.

179. Wainwright, M. 2009. Photoantimicrobials – So what’s stopping us? Photodiag. Photodyn. Ther. 6: 167-169.

180. Wang, L.H., Weng, L.X., Dong, Y.H., and L.H. Zhang. 2004. Specificity and enzyme kinetics of the quorum-quenching N-acyl homoserine lactone lactonase (AHL-lactonase). J. Biol. Chem. 279: 13645–13651.

181. Whitchurch, C.B., Tolker-Nielsen, T., Ragas, P.C., and J.S. Mattick. 2002. Extracellular DNA required for bacterial biofilm formation. Microbiology 295: 1487.

182. Whitchurch, C.B., Leech, A.J., Young, M.D., Kennedy, D., Sargent, J.L., Bertrand, J.J., Semmler, A.B., Mellick, A.S., Martin, P.R., Alm, R.A., Hobbs, M., Beatson, S.A., Huang. B., Nguyen, L., Commolli, J.C., Engel, J.N., Darzins, A., and J.S. Mattick. 2004. Characterization of a complex chemosensory signal transduction system which controls twitching motility in Pseudomonas aeruginosa. Mol. Microbiol. 52: 873-893.

183. Wild, M. Caro, A.D., Hernández, A.L., Miller, R.M., and G. sóberon-Chávez. 1997. Selection and partial characterization of Pseudomonas aeruginosa mono- rhamnolipid deficient mutant. FEMS Microbiol. Lett. 153: 279-285.

184. Wilhem, S., Gdynia, A., Tielen, P., Rosenau, F., and K.E. Jaeger. 2007. The autotransporter esterase EstA of Pseudomonas aeruginosa is required for rhamnolipid production, cell motility, and biofilm formation. J. Bacteriol. 18: 6695-6703.

185. Wilson, R., and R.B. Dowling. 1998. Lung Infections. 3. Pseudomonas aeruginosa and other related species. Thorax 53: 213-219.

186. Woods, D.E., and B.H. Iglewski. 1983. Toxins of Pseudomonas aeruginosa: new perspectives. Rev. Infect. Dis. 5: 715-722.

135

187. Wu, H., Kato, J., Kuroda, A., Ikeda, T., Takiguchi, N., and H. Ohtake. 2000. Identification and characterization of two chemotactic transducers for inorganic phosphate in Pseudomonas aeruginosa. J. Bacteriol. 182: 3400–3404.

188. Yonekura, K., Maki, S., Morgan, D.G., DeRosier, D.J., Vonderviszt, F., Imada, K., and K. Namba, 2000. The bacterial flagellar cap as the rotary promoter of flagellin self-assembly. Science 290: 2148-2152.

189. Zhang, G., Wu, Y., Qian, X., and Q. Meng. 2005. Biodegradation of crude oil by Pseudomonas aeruginosa in the presence of rhamnolipids. J. Zhejiang Univ. Sci. 6: 725-730.

190. Zumft, W.G. 1997. Cell biology and the molecular basis of denitrification. Microbiol. Mol. Biol. Rev. 533-616.

136