Curriculum Vitae

Characterization of Heme Transport in Pseudomonas aeruginosa and the Preferential Pathway for Heme Uptake

Item Type dissertation

Authors Smith, Aaron Dennison

Publication Date 2015

Abstract Bacterial pathogens require iron for their survival and virulence and have evolved multiple mechanisms to acquire this scarce micro-nutrient. The Gram-negative opportunistic pathogen Pseudomonas aeruginosa acquires heme as an iron source through the ...

Keywords ABC transporter; iron acquisition; outer membrane receptor; transporter; ATP-Binding Cassette Transporters; Heme; Pseudomonas aeruginosa

Download date 24/09/2021 16:37:31

Link to Item http://hdl.handle.net/10713/4622 Curriculum Vitae

Aaron Dennison Smith

Education:

University of Maryland, Baltimore (2008-present) Department of Pharmaceutical Sciences School of Pharmacy GPA: 3.92 Degree: Ph.D. Expected Date of Degree: May 2015

Dissertation: Characterization of Heme Transport in Pseudomonas aeruginosa and the Preferential Pathway for Heme Uptake

University of Maryland Baltimore County (2002-2007) Department of Biochemistry and Molecular Biology GPA: 3.1 Degree: B.S. Date of Degree: July 2007

Employment:

Graduate Research Assistant (2008-present) Department of Pharmaceutical Sciences School of Pharmacy, University of Maryland, Baltimore

Technical Director (2006-2008) Chemspec, Inc. Baltimore, MD

Teaching Experience:

Teaching Assistant (2008-2009) School of Pharmacy, University of Maryland, Baltimore PharmD courses: Medicinal Chemistry, Microbiology, Pharmacokinetics

Resource Teacher (2006) Vertically Integrated Partnerships K-16 Internship University of Maryland Baltimore County

Leadership Opportunities:

Graduate Recruitment Strategies Taskforce Committee (2012-2013) President: Pharmacy Graduate Student Association (2009-2010) Team Captain: PSC Race for the Cure (2008-2010)

Presentations:

Characterization of the outer membrane heme receptors and the preferential pathway for heme uptake in Pseudomonas aeruginosa. Tetrapyrroles Gordon Research Conference, Newport, Rhode Island. July 2014. Poster presentation.

Characterization of the outer membrane receptor PhuR and the ABC-transporter ShuUV and their contributions to heme utilization. UMB School of Pharmacy Alumni Reunion Day, University of Maryland Baltimore. Fall 2012. Poster presentation.

The DNA Binding Properties of the Cytoplasmic Heme Binding Protein PhuS of Pseudomonas aeruginosa. Graduate Research Day, University of Maryland Baltimore. April 2009. Poster presentation.

The DNA Binding Properties of the Cytoplasmic Heme Binding Protein PhuS of Pseudomonas aeruginosa. Bioinorganic Chemistry Gordon Research Seminar, Ventura, California. January 2009. Poster presentation.

Publications:

Book Chapters:

Smith AD, Wilks A. (2012) Extracellular Heme Uptake and the Challenges of Bacterial Cell Membranes. In S. Lutsenko & J.M. Arguello (Eds.), Metal Transporters (pp. 359-392). Elsevier Inc.: Academic Press.

Submitted Publications:

Smith AD, Modi AR, Sun S, Dawson JH, and Wilks A. (2014) Spectroscopic Determination of Distinct Heme Ligands in the Outer Membrane Receptors PhuR and HasR of Pseudomonas aeruginosa. Biochem. Submitted.

Smith AD and Wilks A (2014) Differential Contributions of the Outer Membrane Receptors PhuR and HasR to Heme Acquisition in Pseudomonas aeruginosa. J Biol Chem. Submitted.

Awards and Honors:

 PSC Departmental Fellowship Research Award Recipient (2012)  PSC Departmental Merit Research Award Recipient (2011)  Rho Chi Induction, Pharmacy Honor Society (2009)  All-America East Conference Academic Honor Roll (4 semesters)  Maryland Saltwater Sportfishermen’s Association (MSSA) Scholarship Award (2002-2006)  NCAA Div. I Athletic Scholarship (2002-2006)

Affiliations:

 American Chemical Society (ACS) Member  Rho Chi, Pharmacy Honors Society

Abstract

Title of Dissertation: Characterization of Heme Transport in Pseudomonas aeruginosa and the Preferential Pathway for Heme Uptake

Aaron Dennison Smith, Doctor of Philosophy, 2015

Dissertation Directed by: Dr. Angela Wilks, Professor, Department of Pharmaceutical Sciences, School of Pharmacy

Bacterial pathogens require iron for their survival and virulence and have evolved multiple mechanisms to acquire this scarce micro-nutrient. The Gram-negative opportunistic pathogen Pseudomonas aeruginosa acquires heme as an iron source through the Phu

(Pseudomonas heme utilization) and Has (Heme assimilation system) systems. The studies herein detail the initial purification and characterization of the outer membrane (OM) HasR and PhuR receptors. A series of site-directed mutagenesis and spectroscopic studies confirmed HasR, in keeping with previously characterized OM receptors, coordinates heme through the conserved N-terminal plug His-221 and His-624 of the surface exposed

FRAP-loop. In contrast PhuR coordinates heme through His-124 and Tyr-519 ligands not previously reported in OM receptors but associated with high affinity heme binding proteins. In vivo studies utilizing a combination of bacterial genetics, isotopic labeling

(13C-heme), and qRT-PCR further revealed that both receptors are required for optimal heme uptake. However, whereas deletion of hasR leads to an inability to regulate heme uptake, loss of PhuR results in decreased efficiency in heme uptake, despite a significant up regulation in HasR protein levels. The results are consistent with PhuR being the major heme uptake receptor, while HasR senses and regulates extracellular heme uptake. Thus

PhuR and HasR represent non-redundant receptors required for accessing and regulating heme uptake across a wide range of physiological conditions found upon infection. The

research presented herein also involved optimization of the ABC-transporter ShuUV along with the soluble periplasmic heme binding ShuT proteins from Shigella dysenteriae, which are involved in the transport of heme across the cytoplasmic membrane and into the cell.

By generating and screening a series of expression constructs we were able to obtain a construct that resulted in increased expression levels of ShuUV homodimer.

Reconstitution of ShuUV in lipososmes with heme loaded ShuT trapped in the interior of the liposome gave a functional system that could transport heme on activation with ATP.

Taken together, the current research lays the foundation for future spectroscopic and structural studies aimed at understanding the molecular mechanisms of membrane bound heme transport proteins.

Characterization of Heme Transport in Pseudomonas aeruginosa and the Preferential Pathway for Heme Uptake

by Aaron Dennison Smith

Dissertation submitted to the Faculty of the Graduate School of the University of Maryland, Baltimore in partial fulfillment of the requirements for the degree of Doctor of Philosophy 2015

Dedicated to My Family

And everyone who has shared in this journey

iii

Acknowledgements

It has been an extremely difficult and challenging journey, full of sacrifices and hardships, to finally have arrived at this point in my graduate career. However, there have also been many rewards and accomplishments that greatly overshadow the many trials and hardships faced. I cannot look back on my graduate career without remembering all of the positive influences that helped me succeed and shape me into the independent scientist I am today. The current work presented within was not the sole accomplishments of a single individual, but a compilation of thoughts and ideas, discussions and guidance, and collaborations and friendships from so many exceptional individuals that made this manuscript possible and greatly influenced my goal of pursuing a PhD.

I must first thank my advisor and mentor Angela Wilks. She has been a constant driving force towards the accomplishment of this goal and could not have hoped for a better mentor to train and learn from. I sincerely appreciate all of her guidance and support, from scientific discussions and bouncing ideas off her, to attending scientific conferences and being able to present my work to the scientific community. She has always been supportive in allowing me to pursue my intellectual curiosities, even so far as allowing me to learn and conduct new research and techniques with another lab in

California. Thank you also for believing in me and not giving up on me when times were rough and seemed quite impossible to overcome, both professionally and personally. I will forever be grateful for everything you have done for me and consider it a privilege and honor to have worked with and learned from your expertise.

I must also thank my committee members Sarah Michel, C.S. Raman, and Peter

Swaan for their guidance and insightful discussions throughout my graduate career, as well as my external member Herschel Wade for accepting to sit on my committee. Their support has been vital to the success of finishing my PhD. There have been many others as well who have greatly helped with the research presented herein. Maureen Kane, Bob

Ernst, and Jace Jones were extremely helpful with the mass spectrometry training and analysis and were always present for discussion and troubleshooting. Our collaborators

John Dawson and his lab members Anuja Modi and Shengfang Sun for all of their work they did for the MCD, as well as Amanda Oglesby-Sherrouse for generously providing bacterial strains utilized in the studies presents in this manuscript. I also would like to thank Tom Poulos and his lab members for bringing me into their lab and making me feel welcome for the time I spent at University of California, Irvine. Sarah Michel, Amanda

Oglesby-Sherrouse, Patrick Wintrode, and all of their lab members were very helpful in our lab group meetings and I was always appreciative of the advice and ideas they shared.

I also want to thank my current lab mates, Kellie, Geoff, Bennet, Susana, Hermes, and

Weiliang, as well as past Wilks’ lab members, Maura, Kylie, Beth, and Katalin. You each had an impact on the success of my graduate career in your own ways, both inside and outside of the lab.

I want to especially thank Maura, Kat, Joe, and Maryanna. Maura was the first person I met when I joined the department and remember her recruiting me to join the lab. Ever since then she has always been there for constant advice and encouragement in lab, to share a good laugh with, or just meet up in the mornings for a cup of coffee or a drink after a stressful day. It was bittersweet to see her graduate and leave the lab, but we

v still have remained very close friends and has been encouraging to see her succeed as an exceptional scientist. Kat and Joe became really good friends who began the program with me and together we experienced and overcame the highs and lows of graduate school. I will always remember the great times we had and all the laughter we shared. I wish you both the best in your future endeavors and know we will always remain friends.

In the short time I’ve known Maryanna she has already made an enormous impact on my life. Thank you for helping get me through some of the hardest of times and being the amazing person you are.

Last but not least, I have been tremendously fortunate to have had such amazing support and encouragement from my family. I couldn’t imagine being who I am and where I am today without their unconditional support and love. Thank you Mom and

Dad for believing in me and encouraging me to pursue my dreams and shaping me into the person I am today.

Table of Contents

Page Dedication ...... iii

Acknowledgements ...... iv

List of Tables ...... xiii

List of Figures ...... xiv

List of Abbreviations ...... xvii

CHAPTER

1. Heme and Iron Acquisition by Pathogenic Bacteria ...... 1

1.1 General Introduction ...... 1

1.1.1 Iron – An Essential Nutrient for Life ...... 1

1.1.2 Iron and Virulence ...... 2

1.2 Host-Pathogen Relationship and the Innate Immune Response ...... 3

1.3 Iron Regulatory Networks ...... 5

1.3.1 Fur-mediated Iron Regulation ...... 5

1.3.2 sRNA-mediated Iron Regulation ...... 7

1.3.3 Regulation of Iron and Heme Uptake by the Extracytoplasmic Function

Sigma (ECF-σ) Factor Signaling Cascade ...... 10

1.4 Iron Acquisition Systems in Pathogenic Bacteria ...... 12

1.5 Heme Acquisition Systems in Pathogenic Bacteria ...... 14

1.5.1 Heme Uptake Across the Gram-Positive Bacterial Cell Wall ...... 14

vii

1.5.2 Active Transport of Heme Across the Outer Membranes of Gram-Negative

Bacteria ...... 21

1.5.3 ATP-Dependent Transport Across the Cytoplasmic Membranes ...... 23

1.5.4 Heme Degradation of Utilization ...... 26

1.5.4.1 Canonical Enzymes of Heme Degradation ...... 27

1.5.4.2 A Paradigm Shift in Heme Degradation and the Identification of the Isd-

like Heme Monooxygenases ...... 30

1.5.5 Heme Utilization in Pseudomonas aeruginosa ...... 32

1.6 Conclusions ...... 34

1.7 References ...... 35

2. Spectroscopic and Biochemical Characterization of the Pseudomonas aeruginosa

Outer Membrane Heme Receptors ...... 57

2.1 Introduction ...... 57

2.2 Materials and Methods ...... 67

2.2.1 General Methods ...... 67

2.2.2 Bacterial Strains and Media ...... 67

2.2.3 Expression Vector Construction and Site-Directed Mutagenesis ...... 68

2.2.4 Expression and Purification of HasAp ...... 69

2.2.5 Expression of PhuR and HasR ...... 70

2.2.6 Outer Membrane Isolation from PhuR or HasR Expressing Cells ...... 70

2.2.7 Detergent Screening of PhuR and HasR ...... 71

2.2.8 Evaluation of Heme in the Presence of Detergent Micelles ...... 72

2.2.9 Solubilization and Purification of PhuR and HasR ...... 72

viii

2.2.10 Determination of Extinction Coefficients ...... 74

2.2.11 Circular Dichroism of PhuR and HasR ...... 74

2.2.12 Magnetic Circular Dichroism ...... 75

2.2.13 Isolation and Characterization of Freshly Isolated Human Hemoglobin ...... 75

2.2.14 Hemin- and Hemoglobin-agarose Pulldowns of PhuR and HasR ...... 77

2.2.15 Heme Transfer from holo-HasA to apo-HasR and Hemoglobin to apo-PhuR 77

2.2.16 Chemical Cross-linking of PhuR and Hemoglobin or HasR and HasA ...... 78

2.2.17 Iron and Heme Regulation of PhuR and HasR by Western Blot Analysis .....79

2.3 Results ...... 80

2.3.1 Expression and Purification of PhuR and HasR ...... 80

2.3.2 Detergent Screenings for PhuR and HasR Solubilization and Purification ...... 80

2.3.3 Spectroscopic Characterization of Wild Type PhuR and HasR ...... 82

2.3.4 Identification of Tyr-519 as the Heme Ligand on the Extracellular

FRAP-loop ...... 86

2.3.5 Characterization of the N-terminal Plug Mutant H124A ...... 90

2.3.6 Heme Binding and Transfer to PhuR and HasR ...... 92

2.3.7 Iron/Heme Regulation of PhuR and HasR ...... 97

2.4 Discussion ...... 99

2.5 References ...... 103

3. Examining the Individual Contributions of PhuR and HasR to Heme Acquisition in Pseudomonas aeruginosa ...... 111

3.1 Introduction ...... 111

3.2 Materials and Methods ...... 116

3.2.1 General Methods ...... 116

3.2.2 Bacterial Strains and Culture Conditions ...... 116

3.2.3 Generation of PAO1 Receptor Mutants ...... 117

3.2.4 Preparation and Isolation of 13C-Heme ...... 119

3.2.5 Extraction of Biliverdin Isomers from P. aeruginosa Supernatants ...... 120

3.2.6 Mass Spectrometry Analysis of Extracted BVIX Isomers ...... 122

3.2.7 PAO1 RNA Extraction and Purification ...... 122

3.2.8 Generation of cDNA and qRT-PCR Analysis ...... 123

3.2.9 SDS-PAGE and Western Blot Analysis of Wild Type and Mutant Strains of

PAO1 ...... 125

3.3 Results ...... 126

3.3.1 Generation of Unmarked PAO1 Receptor Mutants...... 126

3.3.2 Growth Phenotypes of PAO1 Wild Type and Receptor Mutants...... 126

3.3.3 Heme Metabolism Profiles of PAO1 Wild Type and Receptor Mutants ...... 128

3.3.4 qPCR and Western Blot Analysis of the Heme Uptake Genes in the Receptor

Mutant Strains ...... 132

3.4 Discussion ...... 136

3.5 References ...... 141

4. Heme Transport Across the Cytoplasmic Membrane: Reconstitution of the

Shigella dysenteriae ShuUV ABC Transporter in a Proteoliposome System ...... 146

4.1 Introduction ...... 146

4.2 Materials and Methods ...... 156

4.2.1 General Methods ...... 156

4.2.2 Bacterial Strains and Culture Conditions ...... 157

4.2.3 Construction of Over-expression Vectors for Optimizing ShuUV

Expression ...... 157

4.2.4 Test Expressions of Optimized Constructs...... 158

4.2.5 Construction of the Chimeric Over-expression Construct pCh1SdUV ...... 158

4.2.6 In-gel Tryptic Digestion and Extraction of Expressed Proteins ...... 159

4.2.7 Peptide Mass Fingerprinting of Extracted Peptides by MALDI-MS ...... 161

4.2.8 Expression and Purification of ShuUV from the pCh1SdUV Construct ...... 161

4.2.9 Expression and Purification of ShuT ...... 163

4.2.10 Heme Reconstitution of ShuT ...... 164

4.2.11 Expression and Purification of ShuS ...... 164

4.2.12 Preparation of ShuUV Reconstituted in Proteoliposomes...... 165

4.2.13 Reconstitution of ShuT into Proteoliposomes ...... 166

4.2.14 ATPase and Heme Transport Assays ...... 167

4.3 Results ...... 167

4.3.1 Plasmid Construction and Test Expressions of ShuUV Constructs ...... 167

4.3.2 Functional Assay of Chimeric ShuUV ...... 176

4.4 Discussion ...... 176

4.5 References ...... 181

5. Summary and Future Directions ...... 189

5.1 Summary ...... 189

5.2 Future Directions ...... 195

5.3 Conclusions ...... 197

5.4 References ...... 198

List of References ...... 200

xii

List of Tables TABLE Page 3.1 Bacterial strains and plasmids used in the current studies ...... 118

3.2 qPCR primers and probes used in the current studies ...... 124

4.1 List of generated ShuUV expression constructs and description of observed expression profiles ...... 168

4.2 Peptides identified by in-gel tryptic digestion and mass fingerprint analysis by

MALDI-MS ...... 172

xiii

List of Figures FIGURE Page 1.1 Fur-mediated iron regulation ...... 6

1.2 Transcription of the tandem prrF regulatory small RNAs (sRNAs) and proposed read-through of prrH in P. aeruginosa ...... 9

1.3 Transcriptional activation of heme uptake through the extracytoplasmic function sigma (ECF-σ) factor signaling cascade ...... 11

1.4 Heme uptake system of Gram-positive pathogens ...... 15

1.5 Structural properties of the Isd proteins ...... 17

1.6 Proposed “hand clasp” model of heme transfer between IsdA and IsdC ...... 19

1.7 TonB dependent outer membrane receptor architecture of apo-ShuS from S. dysenteriae ...... 22

1.8 Model of the S. dysenteriae ShuUV ABC transporter in complex with its substrate binding protein ShuT ...... 24

1.9 Structures of the heme degrading enzymes from Gram-negative and Gram-positive pathogens ...... 28

1.10 Heme degradation by the canonical heme oxygenases to BVIXα ...... 29

1.11 Heme degradation through HemO, IsdG/I, and MhuD ...... 31

1.12 Heme uptake system from the Gram-negative pathogen P. aeruginosa ...... 33

2.1 Structure of the hemophore holo-HasA from S. marcescens ...... 59

2.2 Structure of the S. marcescens HasA-HasR complex ...... 63

2.3 Sequence alignment of the heme coordinating extracellular loop from known heme binding outer membrane receptors ...... 66

2.4 Predicted signal peptide cleavage sites of PhuR and HasR ...... 81

xiv

2.5 Absorption spectra of heme in various detergents ...... 83

2.6 Purification of HasR and PhuR from cultures supplemented with heme or hemoglobin ...... 85

2.7 Absorption and MCD spectra of the Fe(III) holo-HasR and PhuR receptors ...... 87

2.8 Absorption and MCD spectra of the Fe(III) PhuR Y519A mutant ...... 89

2.9 Absorption and MCD spectra of the Fe(III) PhuR Y519H and H124A mutants .....91

2.10 Heme binding properties of detergent solubilized HasR ...... 93

2.11 Heme binding properties of detergent solubilized PhuR ...... 95

2.12 Evaluation of secondary structural elements of HasR and PhuR by circular dichroism ...... 96

2.13 Western blot analysis of the heme uptake proteins ...... 98

3.1 13C-heme labeling and BVIX isomer fragmentation patterns ...... 121

3.2 PAO1 wild type and receptor mutant growth curves ...... 127

3.3 BVIX isomer fragmentation patters for PAO1 wild type and mutant strains supplemented with 0.5 μM heme ...... 130

3.4 BVIX isomer fragmentation patters for PAO1 wild type and mutant strains supplemented with 5 μM heme ...... 131

3.5 Effect of heme and iron on mRNA transcription levels of the heme utilization systems ...... 134

3.6 Effect on heme and iron on the heme uptake expression levels in PAO1 wild type and receptor mutant strains ...... 135

3.7 Proposed model for the P. aeruginosa heme cell surface signaling system ...... 138

4.1 Structure of the periplasmic heme binding protein ShuT ...... 149

4.2 Structure of holo-ChuS from E. coli ...... 151

4.3 Proposed model of transport for heme through the periplasmic ABC transport system ...... 155

4.4 Sequence analysis and expression profile of predicted ShuUV ...... 170

4.5 Sequence coverage of ShuV from MALDI-MS mass fingerprinting analysis of tryptic digests ...... 173

4.6 Analysis of the expression construct pCh1SdUV ...... 175

4.7 Functional assay for in vitro heme transport in proteoliposomes ...... 177

4.8 Comparison of α-helix 1 from the ABC transporters involved in heme and vitamin

B12 uptake ...... 180

xvi

List of Abbreviations

4-HCCA α-cyano-4-hydroxycinnamic acid

ABC ATP-binding cassette

ADP Adenosine diphosphate

ALA Aminolevulinic acid

Amp Ampicillin

ATP Adenosine triphosphate

BOG β-octylglucoside

BVIX Biliverdin IX

Carb Carbenicillin

CBP Cytoplasmic binding protein

CD Circular dichroism

CM Cytoplasmic membrane

CMC Critical micelle concentration

CV Column volumes

DDM n-dodecyl-β-D-maltoside

DTT Dithiothreitol

ECF Extracytoplamsic function

EDC 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide hydrochloride

ESI Electrospray ionization

FC-15 Fos-Choline-15

Fur Ferric uptake regulator

xvii

Has Heme assimilation system

Hb Hemoglobin

HO Heme oxygenase

HS High spin

IPTG Isopropyl β-D-1-thiogalactopyranoside

Isd Iron surface determinant

LB Luria-Bertani

LC-MS/MS Liquid chromatography tandem mass spectrometry

LDAO Lauryl-N,N-dimethylamine oxide

LS Low spin

MALDI-MS Matrix assisted laser desorption ionization mass spectrometry

Mb Myoglobin

MCD Magnetic circular dichroism

MRM Multiple reaction monitoring

NBD Nucleotide binding domain

NEAT Near iron transport

NMR Nuclear magnetic resonance

OD Optical density

OM Outer membrane

PAGE Polyacrylamide gel electrophoresis

PBP Periplasmic binding protein

PCR Polymerase chain reaction

Phu Pseudomonas heme uptake

xviii

PIA Pseudomonas isolation agar

PrrF Pseudomonas regulatory RNA involving iron (Fe)

PrrH Pseudomonas regulatory RNA involving heme qRT-PCR Quantitative real-time PCR

QS Quorum sensing

RBS Ribosomal binding site

RFQ-RR Rapid freeze-quence resonance Raman

RNAP RNA polymerase

ROS Reactive oxygen species

SD Shine-Dalgarno

SDS Sodium dodecyl sulfate

Sulfo-NHS N-hydroxysulfosuccinimide

TB Terrific broth

TFA Trifluoroacetic acid

TMD Transmembrane domain

TOF Time of flight

TQD Triple quadrupole

TX-100 Triton X-100

WT Wild type

ZW3-14 Zwittergent 3-14

xix

Chapter 1

Heme and Iron Acquisition by Pathogenic Bacteria

1.1 General Introduction

1.1.1 Iron – An Essential Nutrient for Life

Iron is an essential nutrient for most life on earth. It facilitates many biological processes such as aerobic respiration, oxygen transport and storage, DNA synthesis, nitrogen fixation, and facilitates various redox chemistries, among many others.

However, there are several obstacles biological systems must overcome to obtain and satisfy their demands for iron. Despite being the fourth most abundant metal in the earth’s crust, biologically available iron is relatively low. Under aerobic conditions and physiological pH, iron is primarily found in its oxidized ferric form (Fe3+). Fe3+ is extremely insoluble under these conditions, usually forming insoluble iron oxides, hematite, and magnetite. On the other hand reduced ferrous iron (Fe2+) is soluble, but has the ability through Fenton chemistry to generate toxic free radicals (1, 2). These highly reactive oxygen species (ROS) have the propensity to wreak havoc in cells, damaging lipids, carbohydrates, proteins, and nucleic acids (3, 4). Furthermore, within the body the majority of iron is found complexed to protoporphyrin IX as heme. In addition to being

1 able to generate ROS through Fenton chemistry, heme is also very hydrophobic in nature and can bind to lipids resulting in membrane destabilization (5). In an effort to control the toxic side effects of iron as well as ensure it remains biologically available for cellular functions, iron and heme are sequestered by iron and heme binding proteins. For example, the majority of intracellular iron is contained in the iron storage protein ferritin or in iron-sulfur cluster enzymes, such as those involved in the TCA cycle (6-8).

Extracellular iron is tightly bound to transferrin or lactoferrin (9). Heme is generally found in respiratory cytochromes, enzymes involved in oxidative reactions (10, 11), and the oxygen transport and storage proteins hemoglobin (Hb) and myoglobin (Mb) (12, 13).

Heme in particular, serves as an attractive source of iron to invading pathogens, as ~75% of the total iron in the body is complexed as heme, and thus pathogens have evolved sophisticated mechanisms to acquire this vital nutrient (14-17). In fact, it has been demonstrated that the infectious pathogens Staphylococcus aureus and Bordetella pertussis preferentially utilize heme as an iron source during certain stages of infection

(18, 19).

1.1.2 Iron and Virulence

Iron is often the limiting factor for bacterial growth and survival. Additionally, the expression of several virulence factors is dependent upon the pathogens ability to acquire iron. Pathogens produce a variety of virulence factors in response to iron that contribute to and enhance their pathogenicity. These include the secretion of toxins, such as Shiga toxin and hemolysins in pathogenic Escherichia coli and Shigella dysenteriae

(20), or exotoxin A and elastase in Pseudomonas aeruginosa (21), which are responsible

2 for inducing cellular damage and thereby increasing local concentrations of iron. Iron is also intricately involved in the regulation of biofilm formation in many pathogens through the type II and III secretion systems and fimbrial adhesins (22, 23). Quorum sensing (QS), a cell density dependent signaling mechanism, is also regulated by iron and contributes further to the expression of several virulence factors (24). Even iron and heme acquisition systems themselves contribute to the virulence of pathogens by providing a means to hijack heme and iron from the respective host binding proteins (15).

Many of these systems are interconnected and regulated by complex intertwined regulatory networks, yet all are dependent in some fashion upon iron acquisition (25, 26).

1.2 Host-Pathogen Relationship and the Innate Immune Response

Pathogenic bacteria encounter conditions of extreme iron limitation during infection. Free iron is virtually non-existent under these conditions as it is primarily bound to host iron containing proteins. The innate immune response further restricts the invading pathogens ability to access iron by deploying active countermeasures to sequester iron from the invading pathogens. Hepcidin is the primary hormone of iron homeostasis in mammals and has a central role in the innate immune response (27, 28).

Hepcidin was shown to be induced during inflammation and infection and exhibited antimicrobial activity in vitro (29, 30). This activity is a result of decreased iron export from macrophages and enterocytes, effectively inducing a state of hypoferremia thereby limiting the access to serum iron. Hepcidin was shown to directly bind to the iron exporter ferroportin (31-33) causing endocytic internalization and degradation, resulting in decreased iron export and retention of iron within intracellular stores (34, 35).

Lactoferrin and siderocalin (also known as lipocalin-2 or neutrophil gelatinase- associated protein (NGAL)) provide front line defense against microbial infections by binding and sequestering cellular iron at the site of infection (36-38). Both are highly upregulated as an acute response to bacterial infection and are secreted by neutrophils and macrophages at the sites of infection. Siderocalin can directly bind a variety of bacterial siderophores, including the primary siderophore enterobactin from E. coli, effectively inhibiting siderophore mediated iron uptake (38, 39). This effect has been demonstrated by reduced bacterial replication on addition of siderocalin at sites of infection and the fact siderocalin knockout mice exhibit increased susceptibility to bacterial infections (40-42). In a concerted effort with siderocalin the iron-binding protein lactoferrin also prevents microbial iron acquisition by directly sequestering serum iron and preventing siderophore mediated iron uptake (37). Furthermore, it was recognized decades ago that lactoferrin had additional antimicrobial properties, specifically the destabilization of Gram-negative membranes and release of lipopolysaccharide (43). The concerted effort of the regulatory effects of hepcidin to sequester iron, and lactoferrin and siderocalin to scavenge circulating iron, places the invading bacteria at a distinct disadvantage in obtaining iron from the host. To overcome the host response to bacterial infection, pathogens have evolved multiple strategies to circumvent the host innate immune response in order to acquire iron (44). Iron acquisition sits at the interface of the host-pathogen interaction offering a unique opportunity in terms of an antibacterial strategy by tipping the balance in the struggle for iron in favor of the host.

1.3 Iron Regulatory Networks

1.3.1 Fur-mediated Iron Regulation

Bacteria are able to adapt and respond to levels of iron by altering the expression levels of iron responsive genes. The ferric uptake regulator (Fur) protein is fundamental in maintaining the iron homeostasis of almost all bacteria (Figure 1.1). It was first shown in an E. coli Δfur knockout that iron responsive genes, including the enterochelin biosynthesis genes, were constitutively active (45). This led to the proposal that Fur must be an iron-responsive transcriptional suppressor. Later, studies confirmed that Fur binds

Fe2+ as a corepressor through two conserved histidines and a carboxylate ligand in an axial distorted octahedral geometry (46, 47). Upon binding Fe2+, Fur undergoes conformational changes that allow the formation of a homodimer and subsequent binding to a specific DNA sequence with dyad symmetry GATAATGATAATCATTATC, termed the Fur-box, located in the promoter region of iron responsive genes (48). This in turn prevents access to RNA polymerase (RNAP) thereby repressing transcription. However, upon iron depletion intracellular iron levels drop below a threshold where apo-Fur dissociates into monomers with release from the promoter and allowing RNAP to bind and initiate transcription (49). Structurally, Fur consists of an N-terminal DNA recognition domain and a C-terminal domain required for Fur dimerization and Fe2+ binding (50). Additionally, the E. coli Fur protein was shown to have a Zn2+ binding site distinct from the Fe2+ site that was also required for DNA binding (51, 52). However, this site is absent in P. aeruginosa Fur as well as other closely related bacteria (53).

Figure 1.1 Fur-mediated iron regulation. A. Under iron deplete conditions apo-Fur does not bind DNA, allowing transcription of iron-regulated genes. B. In iron replete conditions, apo-Fur binds Fe2+ and forms a homodimer that is active to bind to the Fur Box located upstream of iron-regulated genes, effectively preventing transcription.

Fur has been shown to control the expression of a wide spectrum of iron responsive genes that serve many diverse cellular functions. For example, in E. coli, over

90 genes have been identified to be under the direct control of Fur, including genes involved in siderophore biosynthesis and transport, heme uptake, carbon metabolism, iron metabolism, and oxidative stress (54-57). It was also discovered that Fur can positively regulate genes, although it was not identified until much later that this was indirect post-transcriptional control through regulatory small RNAs as described below in section 1.3.2.

1.3.2 sRNA-mediated Iron Regulation

In E. coli, it was discovered that Fur could positively regulate genes under iron replete conditions, albeit indirectly by controlling the transcription of a regulatory small

RNA (sRNA) RhyB (58). RhyB has sequence complementary (anti-sense) with genes involved in oxidative stress, iron storage, and other non-essential iron containing proteins

(59, 60). A microarray analysis revealed 18 operons encoding up to 56 genes are under the control of RhyB (61). Complementary base pairing with RNA transcripts of these genes with RhyB forms a duplex RNA complex, which is then targeted for rapid degradation thereby effectively preventing translation (62-64). However, when intracellular iron levels are sufficient, Fe2+ bound Fur blocks transcription of RhyB allowing for the positive regulation of RhyB regulated genes. Since the characterization of RhyB, sRNAs have been characterized in many bacteria, including Pseudomonads

(65). Interestingly, a set of tandem duplicate sRNA’s were discovered in P. aeruginosa named PrrF1 and PrrF2 for Pseudomonas regulatory RNA involving iron (Fe) that

7 function in a similar manner to RhyB (66). The PrrF RNAs were shown to modulate virulence by controlling expression of genes involved in the metabolism of anthranilate, a

PQS precursor (67). A third longer transcript has recently been identified that occurs as a result of a read-through of the rho-independent terminator of PrrF1 (Figure 1.2). This longer transcript had been shown to be dependent on extracellular heme uptake and hence termed PrrH for Pseudomonas regulatory RNA involving heme (68). PrrH heme dependent repression leads to increased expression of genes previously identified as being PrrF-regulated as well as several newly identified genes. The regulation of sRNA’s by Fur therefore provides a sophisticated regulatory strategy bacteria employ to up regulate iron storage proteins, such as bacterioferritin, and proteins involved in oxidative stress protection, such as superoxide dismutase, under iron replete conditions, yet rapidly down regulate non-essential iron containing proteins under conditions of iron starvation, sparing scarce iron stores for essential cellular functions.

However, not all positive regulation by Fur can be explained by the down regulation of sRNA’s. For example, in S. typhimurium, it has been shown that Fur can positively regulate proteins involved in the acid shock response (69). Similarly, in P. aeruginosa, the bacterioferritin gene bfrB was shown to be positively regulated by Fur, however, this was independent of PrrF1/2 transcription (66). Clearly, positive regulation by Fur extends beyond sRNA regulation and requires further investigation to completely understand Fur mediated iron regulation.

Figure 1.2 Transcription of the tandem prrF regulatory small RNAs (sRNAs) and proposed read-through of prrH in P. aeruginosa. Under conditions of low iron, Fur- mediated repression of the prrF genes is alleviated. Transcription is terminated by their rho-independent terminators. However, under conditions of low iron and low heme, an anti-termination event at the PrrF1 rho-independent terminator occurs, presumably through binding of an unidentified protein allowing read-through and termination at the PrrF2 rho-independent terminator, resulting in a longer transcript PrrH that includes the intergenic region termed PrrH.

1.3.3 Regulation of Iron and Heme Uptake by the Extracytoplasmic Function Sigma

(ECF-σ) Factor Signaling Cascade

Bacteria must rapidly acclimatize to changes in their environment in order to survive. In an iron-limiting environment, they must respond rapidly and decisively to the nutrient sources available to them. This can be achieved through the signaling cascade of the ECF-σ factors (Figure 1.3). The components of the ECF-σ factor cascade consist of an outer membrane (OM) receptor that binds a specific substrate. Upon substrate binding, a signal is transduced to the periplasmic face of the receptor where it interacts with the anti-sigma factor that then releases the bound ECF-σ factor in the cytoplasm.

The activated ECF-σ factor is then able to bind to and direct the core RNAP complex to the promoters of target genes to initiate transcription (70-72). The ECF-σ genes encoding these components are usually under control of the ECF-σ factor itself, resulting in positive auto-regulation and concurrent amplification of the signal. There are examples of receptor/ECF-σ factors that sense and respond to iron such as the FecA/I in E. coli which senses Fe3+-citrate (73), and HasR/I in Serratia marcescens which senses the heme loaded hemophore HasA (74). Signaling in either cascade results in the up regulation of systems required for import and utilization of the respective substrates. P. aeruginosa has a unique example of an iron responsive ECF-σ factor system where two different

ECF-σ factors (FpvI and PvdS) are regulated by the same anti-sigma factor (FpvR) (75).

The OM receptor FpvA senses Fe3+-pyoverdine which relays the signal through FpvR activating FpvI or PvdS (76, 77). FpvI targets the fpvA gene while PvdS targets genes responsible for pyoverdine biosynthesis. Additionally, it was shown PvdS controlled the regulation of the virulence factors exotoxin A (78) and endoprotease PrlP (79).

Figure 1.3 Transcriptional activation of heme uptake through extracytoplasmic function sigma (ECF-σ) factor signaling cascade. Signaling through the substrate loaded OM receptor induces conformational changes across the membrane to the anti-σ factor, rendering it inactive. This in turn allows for release of the σ factor into the cytoplasm where it can form a transcription initiation complex with RNAP allowing for expression of heme uptake genes.

1.4 Iron Acquisition Systems in Pathogenic Bacteria

The dependence of pathogens to acquire iron is central to their survival and virulence. One of the most effective strategies pathogens employ is the synthesis and secretion of low molecular weight (400-1000 Da) iron chelators termed siderophores.

Siderophores scavenge iron from the extracellular environment with extremely high affinities, ranging from 109 to 1053 M-1 (80). This apparent high affinity for iron allows them to compete with the up regulation of the host iron binding proteins transferrin and lactoferrin during infection (81). Siderophores consist of a peptide backbone that is biosynthesized by the nonribosomal peptide synthases (NRPS) or NRPS-independent pathways followed by addition of catechols, hydroxymates, and/or hydroxycarboxylates to the peptide backbone as the iron chelating ligands (82, 83). Many pathogens produce unique siderophores such as enterobactin in E. coli and Salmonella enterica (84, 85), vibriobactin in Vibrio cholera (86), mycobactin in Mycobacterium tuberculosis (87), and staphyloferrin in S. aureus (88). Additionally, pathogens are capable of biosynthesizing more than one type of siderophore such as pyochelin (89) and pyoverdine (90) in P. aeruginosa. The evolution of bacteria specific siderophores allows for increased fitness within a competitive polymicrobial community, however, some pathogens have further adapted to hijack siderophores synthesized by other microbes (91, 92).

Siderophores are too large to passively diffuse across the membrane or be transported through porins and instead require specific OM receptors on the bacterial cell surface to actively transport the iron-loaded siderophore across the OM. These receptors belong to a large class of TonB-dependent OM receptors specific to Gram-negative bacteria that are involved in the recognition and transport of a wide variety of substrates

(93, 94). They generally consist of a 22-stranded anti-parallel β-barrel that forms the translocation channel along with an N-terminal plug that occludes the channel to prevent non-specific passive diffusion (95). Extracellular loops with varying length extend to the extracellular space that recognize and facilitate binding of their specific substrates. As no suitable energy source is available in the periplasm of Gram-negative pathogens, such as

ATP, the proton motive force of the cytoplasmic membrane provides the energy necessary for active transport into the periplasm (96). The OM receptor is coupled to the cytoplasmic membrane (CM) across the periplasmic space (PM) by the energy transducing complex TonB-ExbBD through a specific interaction with the so called

TonB-box of the receptor (97, 98). In order for Fe3+-siderophore translocation to proceed, significant conformational changes must occur; most notably the rearrangement of the N-terminal plug. Once inside the periplasm, the Fe3+-siderophore is bound by a periplasmic binding protein that can deliver it to a specific ATP-binding cassette (ABC) transporter, which catalytically utilizes ATP as a substrate to facilitate import into the cytoplasm.

Once the iron loaded siderophore is translocated into the cytoplasm the iron must be released for further use by the bacterial cell. This is achieved by one of two mechanisms: reduction of tightly bound Fe3+ to weakly bound Fe2+ by dedicated ferric- reductases or proteolytic cleavage of the siderophore peptide backbone (92, 99). The latter method results in destruction of the siderophore with some energetic cost in resynthesizing the siderophore. Alternatively, some pathogens, such as P. aeruginosa, can reduce the siderophore, in this case pyoverdine in the periplasm, allowing release of iron and efficient recycling of the siderophore for subsequent rounds of iron import (100).

1.5 Heme Acquisition Systems in Pathogenic Bacteria

1.5.1 Heme Uptake Across the Gram-Positive Bacterial Cell Wall

Heme serves as a primary reservoir of iron for invading pathogens and thus pathogens have evolved sophisticated mechanisms for the transport and utilization of heme. Gram-positive organisms have an extensive peptidoglycan cell wall that protects the cell from osmotic lysis and provides rigidity and support. This also provides a significant barrier that renders substrates impervious to the cell, including heme.

Therefore, heme is captured and transported in these organisms by a protein relay of cell wall anchored proteins termed the iron surface determinant (Isd) system (101). These systems have been identified in numerous Gram-positive pathogens but the most well characterized system is that from S. aureus (Figure 1.4). The isd locus encodes a sortase

(srtB), the cell wall anchored heme transport proteins (isdA, isdB, isdC, and isdH), a cytoplasmic membrane ABC transport system (isdD, isdE, and isdF), as well as heme degradation enzymes (isdG and isdI) (101-103). IsdH and IsdB are capable of extracting heme from hemoglobin (Hb) or Hb-haptoglobin complexes (104, 105). Heme loaded

IsdH or IsdB can then bi-directionally exchange heme between each other or unidirectionally transfer heme to IsdA (106). holo-IsdA can then in turn transport heme to

IsdC (107). It was additionally shown in vitro that IsdB could deliver heme to IsdC, however, this is an unlikely scenario in vivo due to the spatial localization of IsdC in the cell wall and most likely heme is transferred from IsdA (108). Only IsdC has been shown to deliver heme to IsdE, the cytoplasmic membrane lipid anchored heme acceptor that specifically delivers heme to the membrane permease IsdD/F (106, 108). There are

Figure 1.4 Heme uptake system of Gram-positive pathogens. Gram-positive pathogens have cell wall anchored proteins at specific locations within the cell wall to allow for efficient unidirectional transfer of heme into the cell. Heme is initially captured and stripped from hemoglobin by IsdB/H, which then transfers heme to IsdA. IsdA then transfers heme to IsdC, that passes it to IsdE before it is actively transported across the cytoplasmic membrane by the IsdDF transporter where it is subsequently degraded by the IsdG/I monooxygenases.

15 no genes encoding the catalytic ATP hydrolase subunits required for ABC transporters in the isd operon, however, studies have implicated the ATPase FhuC as fulfilling this requirement (109). Once internalized, heme is then degraded by the actions of IsdG and

IsdI to liberate iron for subsequent cellular utilization (110).

All cell wall anchored Isd proteins contain 1-3 near iron transport (NEAT) domains, an N-terminal signal peptide, and a C-terminal sortase anchoring signal (Figure

1.5A) (111). Structurally, NEAT domains share a common overall 8-stranded immunoglobin-like β-sandwich fold with a small yet flexible 310 α-helix between the β1 and β2 strands (Figure 1.5B) (112-115). The NEAT domains are separated by stretches of highly charged amino acids that are predicted to be highly disordered. Sequence analysis, spectroscopic, and structural studies revealed the presence of highly conserved residues important for forming contacts with and ligating the heme. For example, in

IsdA-N1 (IsdA NEAT domain 1) the conserved Tyr-166 on the loop connecting β7 and

β8 directly ligates the heme iron while a second invariant tyrosine four residues away

(Tyr-170) forms a hydrogen bond with the iron coordinating tyrosine and forms extensive

π-bonding interactions with one of the heme pyrrole rings. The IsdB-N2 heme binding

NEAT domain has an additional distal coordinating ligand to the heme iron in Met-362 that is crucial for transfer to IsdA-N1 (116). Additionally, many other conserved residues are present in IsdA-N1 such as Tyr-87 involved in π-stacking interactions with the tetrapyrrole skeleton, Trp-113 involved in interactions with one of the heme vinyls, and

Ser-82 that hydrogen bonds to one of the heme propionates (113). Curiously, His-83 in the distal pocket of IsdA-N1 makes close contacts with the heme and it has been shown when the iron is reduced, His-83 is capable of coordinating to the heme (117). This

Figure 1.5 Structural properties of the Isd proteins. A. Schematic of the Isd proteins NEAT domains. NEAT domains are numbered from their N-terminus. The N-terminal secretion signal is represented by white boxes. Dark grey boxes represent heme binding NEAT domains while the light grey boxes represent NEAT domains implicated in interactions with hemoglobin. The sortase signal sequences are indicated at the C- terminus of each Isd protein. B. Overall architecture of the Isd NEAT domains. NEAT domain 1 from IsdC is represented with heme ligated by the coordinating Tyr-132. Adapted from PDF file 2O6P.

17 histidine is absent in the heme binding domains of IsdH-N3 and IsdC-N1 and instead is substituted by hydrophobic residues Val-564 and Ile-48, respectively. This has been proposed to be a possible mechanistic explanation for unilateral transfer and release of heme from IsdH/B to IsdA. However, not all NEAT domains follow this similar architecture in the heme binding pocket. For example, IsdH-N1, IsdH-N2, and IsdB-N1 are not involved in heme binding and lack the conserved Tyr-87, Tyr-166, Tyr-170, Trp-

113, and Ser-82 residues. Instead, these NEAT domains, as in IsdH-N1, have a series of conserved but disordered aromatic residues Tyr-125, Tyr-126, His-127, Phe-128, and

Phe-129 in the second loop connecting β1- and β2-strands (112). These NEAT domains have been implicated in tightly binding and immobilizing host Hb or Hb-haptoglobin complexes on the cell surface and enhancing the capture of heme to the heme binding

NEAT domains (114). The cytoplasmic membrane anchored IsdE heme binding NEAT domain also has a unique heme binding pocket. The IsdE NEAT domain contains a six- coordinate heme through His-229 on the distal face and Met-78 on the proximal side

(118). Paramagnetic relaxation enhancement NMR studies have suggested that heme binding NEAT domains from holo-IsdA and apo-IsdC form a transient weak, but stereospecific “hand clasp” interaction allowing rapid transfer of heme (Figure 1.6) (119).

In this model, heme is contained within a hydrophobic binding cleft formed at the interface between the IsdA and IsdC heme binding NEAT domains. Contacts between residues in the loop connecting strands β7 and β8 in both IsdA and IsdC are located adjacent to the 310 α-helix of the opposite protein. Such an arrangement positions both of the conserved heme coordinating Tyr residues in a position to accept heme, however, does not explain how heme is released from IsdA to IsdC. Presumably, this

Figure 1.6 Proposed “hand clasp” model of heme transfer between IsdA and IsdC. A transient weak protein-protein interaction has been proposed to transfer heme between Isd proteins, where heme must translate 18 Å from the binding pocket in IsdA to IsdC. Adapted from PDB files 3QZO and 2O6P.

19 protein-protein interaction would have to weaken the Tyr bond to the heme in IsdA to favor transfer to IsdC. The 310 helix in IsdC makes contacts with the loop bearing the coordinating Tyr-166 of IsdA, possibly resulting in conformational changes capable of disrupting this bond. Additionally, the loop connecting strands β7 and β8 in IsdC make contacts with the 310 helix in IsdA which are responsible for forming stabilizing noncovalent interactions with the heme, which could further reduce the heme affinity for

IsdA. Although the same contacts are made with IsdC, the β7 and β8 strands extend further than that of IsdA, with the coordinating Tyr-132 positioned more centrally on strand β8, possibly explaining how IsdC can retain its affinity for heme upon complex formation.

In summary, the data supports a model where organizational arrangement of Isd proteins within the cell wall facilitate a unidirectional transfer of heme to the cytoplasmic membrane which is mediated by protein-protein interactions. IsdH and IsdB are located on the periphery of the cell wall, engaging and capturing host heme containing proteins.

The centrally located IsdA within the cell wall can accept heme from IsdH-N3 or IsdB-

N2 and subsequently pass it to IsdC, which is located furthest within the cell wall, presumably in close proximity to the cytoplasmic membrane to which IsdE is anchored.

ABC transport by the IsdDF/FhuC system, proposed to occur in a similar way as ABC transport through the cytoplasmic membranes of Gram-negative pathogens, and heme degradation by the IsdG/I proteins, will be discussed in greater detail in the following sections.

1.5.2 Active Transport of Heme Across the Outer Membranes of Gram-Negative

Bacteria

Gram-negative pathogens lack an extracellular cell wall and instead have a second asymmetrical OM lipid bilayer. Substrates must be actively transported across the OM and shuttled through the periplasmic space to reach the cytoplasmic membrane. As with the Fe3+-siderophore transporters, active heme transport across the OM of Gram-negative pathogens relies on the presence of specific TonB-dependent OM transporters (16). The heme receptors share the common overall fold of the TonB-dependent OM proteins, specifically a 22-stranded anti-parallel β-barrel forming the translocation channel, an N- terminal “plug” domain, and extracellular loops responsible for substrate recognition

(Figure 1.7). Additionally, the majority of heme transporters characterized thus far coordinate heme through two conserved histidines on the N-terminal plug and the

FRAP/PNPNL domain on one of the extracellular loops (120). Bacterial OM heme transporters fall into one of two classes, those that access heme through a specific secreted hemophore and those that bind directly to the host heme containing proteins.

Hemophores are low molecular weight proteins (< 20kDa) that are excreted into the environment where they scavenge heme from host heme containing proteins, such as hemoglobin, hemoglobin-haptoglobin, and hemopexin (121-124). The S. marcescens hemophore system has been the most extensively studied and characterized to date. Its hemophore, HasA, is excreted into the extracellular environment by the HasD/E secretion system and can bind heme with a reported picomolar affinity (125, 126). Heme loaded

HasA interacts with the OM receptor HasR for subsequent internalization of heme (127).

The OM receptor ShuA from S. dysenteriae is an example of a heme receptor that

Figure 1.7 TonB dependent outer membrane receptor architecture of apo-ShuA from S. dysenteriae. TonB dependent outer membrane receptors are composed of a 22-stranded anti-parallel β-barrel (blue) that insert into the lipid bilayer and an N-terminal “plug” (red) that occludes the translocation channel. Long extracellular loops extend into the extracellular space and are involved in substrate recognition. Conserved heme coordinating histidine residues (green sticks) are located on the apical face of the N- terminal plug and on the extracellular FRAP-loop. Adapted from PDB file 3FHH.

22 directly interacts with host heme binding proteins, notably oxidized hemoglobin (metHb) dimers (128). Binding of the hemeprotein substrate induces a conformational change in the receptor that results in the unwinding of the conserved N-terminal TonB-box on the periplasmic face of the receptor. TonB recognizes and binds this TonB-box motif forming an inter-protein β-sheet at the TonB-box and positioning of the TonB α-helix 1a in close proximity to the plug domain where electrostatic interactions drive a conformational change in the plug required for substrate translocation to the periplasm

(129, 130). A more in-depth review of OM transporters will be covered in Chapters 2 and 3.

1.5.3 ATP-Dependent Transport Across the Cytoplasmic Membranes

Translocation of heme across the cytoplasmic membrane (CM) is mediated by the action of an ATP binding cassette (ABC) transporter and its soluble periplasmic binding protein (PBP) (Figure 1.8). The PBP’s all share a common overall structure containing

N-and C-terminal lobes that form a substrate binding cleft connected by α-helices.

PBP’s are divided into three classes based on the number of α-helices connecting the two globular domains, with the heme PBP’s falling into the third class which have only one interconnecting α-helix (131, 132). The N- and C-terminal lobes of the PBP’s are structurally similar, consisting of a five-stranded β-sheet flanked by α-helices. In the

PBP ShuT of S. dysenteriae, heme is coordinated in the cleft between lobes by a conserved Tyr-94 with an apparent high affinity (132, 133). Minimal conformational changes are apparent upon heme binding to ShuT. The most noticeable difference between apo- and holo-ShuT is the rearrangement of Tyr-94 to a more inward facing

Figure 1.8 Model of the S. dysenteriae ShuUV ABC transporter in complex with its substrate binding protein ShuT. ShuU (blue) and ShuV (green) homodimers form an overall heterotetrameric complex. Conserved histidine residues (red) critical for transport have been highlighted in the translocation channel of ShuU. Holo-ShuT (purple) recognizes and interacts with the TMD of ShuU through surface exposed glutamate residues (yellow) from ShuT and arginine residues (orange) from each subunit of ShuU. ShuUV modeled from PDB file 4G1U. Holo-ShuT adapted from PDB file 2R7A.

24 conformation inducing a slight closure of the N- and C-terminal lobes around the heme in the binding cleft (132). This also orients surface exposed glutamate residues in a position conducive to binding to its specific ABC transporter ShuUV through corresponding conserved arginine residues (Fig 1.8).

ABC transporters represent one of the largest known superfamilies of proteins that are primarily responsible for the ATP-dependent transport of substrates across biological membranes (134, 135). The ABC transporters of Gram-negative bacteria are multi-subunit proteins consisting of two transmembrane domains (TMD), which form the translocation channel, and two cytoplasmic nucleotide binding domains (NBD), which are responsible for the binding and hydrolysis of ATP. The TMD can vary in chain length and sequence allowing for selectivity and/or diversity in transported substrates while the NBD is one of the most highly conserved proteins known (136). The NBD has a number of conserved structural features including the Walker A and the “signature”

LSGGQ motif in the Walker C region responsible for ATP-binding, the Walker B region responsible for binding a catalytic Mg2+ ion, and the Q-loop and switch region critical for

ATP hydrolysis (137, 138).

The recent structure of the Yersinia pestis ABC heme transporter, HmuUV, revealed a similar overall architecture to the well characterized E. coli vitamin B12 transporter BtuCD (139, 140). The overall architecture of the HmuUV transporter consists of a membrane spanning dimer of heterodimers (U2V2). Furthermore, alignment of all known heme ABC transporters revealed two conserved histidine residues in the

TMD that are critical for heme transport (141). In vitro spectroscopic and functional studies of BtuCD, HmuUV, and the heme transporter from S. dysenteriae, ShuUV,

25 suggest an overall mechanism of transport through an ATP-switching model (140-142).

In this model, the translocation channel is closed to the periplasm and upon protein- protein interaction between the cognate substrate loaded PBP and the TMD, a conformational change occurs that 1) triggers opening of the translocation channel to the

PBP and closure to the cytoplasm, 2) facilitates release of substrate from the PBP to the translocation channel, 3) substrate binding in the channel transduces a conformational switch to the cytoplasmic surface that positions the NBD within close proximity of each other inducting ATP-binding, and 4) hydrolysis of ATP causes a “power stroke” driving substrate release to the cytoplasm (143). A greater in-depth review on the finer mechanistic aspects of ABC transporters will be covered in Chapter 4.

Many Gram-negative bacteria also contain a cytoplasmic heme binding protein

(CBP) whose function has been the subject of some debate. The protein was initially annotated as a heme oxygenase, but more recently has been shown to be a regulator of the metabolic flux of heme into the cell and is required for efficient heme utilization (144,

145).

1.5.4 Heme Degradation and Utilization

On internalization of heme in the cytoplasm, the tetrapyrrole ring of heme must be cleaved to release the iron for further utilization by the bacteria. Heme degradation to release iron is achieved through the catalytic actions of heme monooxygenases (HO).

These enzymes fall into two classes: the biliverdin producing canonical HOs first discovered in mammals and higher organisms (146-148) and more recently Gram-

26 negative bacteria (149, 150), and the staphylobilin producing Isd-like proteins of S. aureus (110) and Mycobacterium tuberculosis (Figure 1.9) (151).

1.5.4.1 Canonical Enzymes of Heme Degradation

The canonical HO enzymes use heme as both a substrate and cofactor for the self- catalyzed degradation of heme to biliverdin, CO, and iron. Despite the lack of sequence identity (30-35%) the overall fold of the mammalian and bacterial enzymes are very similar. The structure comprises a six α-helical bundle that coordinates heme through a conserved histidine from the N-terminal helix (152, 153).

Furthermore, the reaction in both human and bacterial enzymes is similar and proceeds by binding of heme to the enzyme followed by the one electron reduction of ferric (Fe3+) heme to ferrous (Fe2+) heme with the reducing power being provided by

2+ NADPH (Figure 1.10). The Fe heme iron binds O2 and following a second electron transfer the active (Fe3+-OOH) hydroperoxy heme is formed. Electrophilic addition of the terminal O of the Fe3+-OOH oxidizing species at the α-meso carbon generates α- meso-hydroxyheme, which in the presence of oxygen rapidly converts to the second intermediate verdoheme, liberating CO in the process. A final four electron reduction

3+ and O2 binding step generates the reactive peroxy (Fe -OOH) verdoheme species that inserts a second O-atom to yield biliverdin and eventually iron release (146, 147, 154-

156). The overall reaction consumes 3 equivalents of O2 and 3 equivalents of NADPH, while liberating biliverdin, CO, and iron.

Figure 1.9 Structures of the heme degrading enzymes from Gram-negative and Gram- positive pathogens. A. Bundled helix of the canonical heme oxygenase HemO from the Gram-negative pathogen P. aeruginosa. Inset shows the planar conformation of heme in the active site. B. Structure of the non-canonical heme oxygenase IsdI from the Gram- positive pathogen S. aureus. Insets shows an extensive ruffled conformation of heme proposed to be important in the enzymatic mechanism for the generation of the alternative staphylobilins.

Figure 1.10 Heme degradation by the canonical heme oxygenases to BVIXα. The overall reaction converts heme into α-biliverdin consuming 3 equivalents of O2 and 7 electrons. The key intermediates during the catalytic cycle, α-meso-hydroxyheme and α- verdoheme are shown.

1.5.4.2 A Paradigm Shift in Heme Degradation and the Identification of the Isd-Like

Heme Monooxygenases

Many Gram-negative and Gram-positive pathogens have heme uptake systems for which a canonical HO has not been identified. However, it was recently discovered that

S. aureus and M. tuberculosis degrade heme through a similar reaction mechanism despite yielding different end products (Figure 1.11) (156). The IsdG/I proteins from S. aureus were originally classified as heme oxygenases (110), despite having no structural or sequence similarity to the canonical HOs as well as distinct end products in staphylobilin (157). It was later proposed that the extensive heme ruffling in these enzymes, as well as the altered electronic structure of the porphyrin ring, results in the alternate reaction products (158, 159). Mechanistic characterization of the M. tuberculosis enzyme MhuD showed that heme was converted to a similar bilin end product, mycobilin (160). Mechanistic studies confirmed there was no liberation of CO during the course of the reaction and the α-meso carbon was retained as an aldehyde with the adjacent meso-carbon at the β- or δ-position being oxidized to a carbonyl. The structural similarities and heme ruffling of MhuD is similar to the IsdG/I enzymes suggesting heme oxidation proceeds through the same reaction mechanism. Further investigation of the IsdG/I catalyzed reaction confirmed a similar reaction occurred, but with the α-meso carbon being released as formaldehyde (161). A detailed mechanism of how the IsdG-like proteins catalyze heme degradation reactions still remain to be determined, but recent studies ruled out verdoheme as an intermediate suggesting the enzymes have evolved to prevent the liberation of CO. In S. aureus, it has been shown that CO can have bactericidal effects (162, 163) while in M. tuberculosis CO acts as a

Figure 1.11 Heme degradation through HemO, IsdG/I, and MhuD. The mechanism of heme degradation is proposed to proceed through meso-hydroxyheme for all three enzymes at which point the mechanistic pathways diverge. HemO releases the meso- carbon as CO, IsdG/I releases the meso-carbon as formaldehyde, and MhuD retains the meso-carbon on the tetrapyrrole ring as an aldehyde. All pathways ultimately lead to the liberation of iron.

31 signal to activate a dormancy regulon through a heme-dependent two-component sensor kinase system (164). Either scenario would obviously lead to unfavorable outcomes for either pathogen when establishing an infection, therefore, they have evolved alternative mechanisms for liberating iron from heme.

1.5.5 Heme Utilization in Pseudomonas aeruginosa

P. aeruginosa is a Gram-negative bacteria commonly found in soil and water and is capable of infecting a variety of species, including humans, plants, and animals (165).

P. aeruginosa is an opportunistic pathogen responsible for hospital acquired nosocomial infections and is extremely prevalent in patients who are immune compromised, such as in AIDS, chemotherapeutic patients, burn victims, and especially in cystic fibrosis, often causing chronic and potentially life-threatening infections (166). P. aeruginosa is a difficult pathogen to treat due to its intrinsic resistance to multiple antibiotics, ability to establish biofilms upon colonization, and the expression of numerous virulence factors

(167). As for nearly all bacterial pathogens the availability and acquisition of iron plays a key role in the propensity of P. aeruginosa to establish and maintain an infection.

P. aeruginosa encodes at least two heme uptake systems, the Pseudomonas heme uptake (phu) and heme assimilation systems (has) (Figure 1.12) (168). The has system is comprised of the hemophore HasA and its cognate OM receptor HasR, as well as the

HasD/E export system required for HasA secretion. The phu system contains an OM receptor that interacts with host heme containing proteins as well the components necessary for complete import of heme into the cytoplasm. This includes PhuT, a soluble periplasmic binding protein that also serves as the soluble receptor for a specific ABC

Figure 1.12 Heme uptake system from the Gram-negative pathogen P. aeruginosa. Heme can be actively transported into the cell from host heme containing proteins, such as hemoglobin, or through a secreted hemophore utilizing TonB-dependent OM transporters PhuR or HasR, respectively. Heme is transported to the periplasm where PhuT shuttles it to a specific ABC transporter PhuUV which actively transports heme across the CM utilizing ATP as an energy source. Once internalized, PhuS chaperones heme to the iron- regulated heme oxygenase HemO that degrades heme to δ/β-biliverdin, liberating iron in the process.

33 transporter, PhuUV, and the soluble cytoplasmic heme chaperone PhuS. The has system lacks the cytoplasmic transport system, and is assumed to utilize the PhuT-UV periplasmic ABC transport system to translocate heme into the cytoplasm. P. aeruginosa also encodes two heme oxygenases, HemO and BphO (169, 170). The iron-regulated

HemO catalyzes the degradation of extracellular heme to δ/β-biliverdin (171).

Furthermore, PhuS has been shown to serve as a regulator for the flux of extracellular heme into the cell as a result of a specific protein-protein interaction with HemO (144,

145, 172). In contrast, the biliverdin α-regioselective BphO is not iron regulated and appears to be involved in maintaining the homeostasis of the intracellular biosynthetic heme pool (144). In addition, BphO is encoded in an operon with a histidine-sensor kinase, BphP, which is a red/far-red light sensor (170, 173, 174). The function of the two-component sensor kinase in heme metabolism is currently not well understood.

1.6 Conclusions

In the mammalian host, heme serves as a prime reservoir for invading pathogens to meet their iron requirements. This strict requirement for iron is evident from the array of sophisticated mechanisms pathogens have evolved to acquire and utilize this vital nutrient, while regulating its uptake and storage to curb the potentially toxic and damaging side effects. Many bacterial opportunistic pathogens have evolved multiple redundant pathways to increase their adaptability and survival within a variety of iron- restricted environmental niches.

The research presented herein aims to provide insight into the structure-function relationships of the heme uptake systems from the opportunistic pathogen P. aeruginosa.

In Chapter 2, the initial purification and characterization of both the hemophore- dependent HasR and PhuR OM receptors are presented. Through site-directed mutagenesis and spectroscopic analysis the heme coordination properties of the receptors have been characterized. Furthermore, the previously reported differential regulation of the OM receptors in response to heme and iron is investigated further. In Chapter 3, through a series of bacterial genetic and 13C-heme isotopic labeling experiments, the relative in vivo contributions each receptor to heme uptake is examined. In Chapter 4, focus is switched to heme transport across the cytoplasmic membrane. Optimization of a protein expression and purification system for a homolog of the PhuT-UV ABC transporter ShuT-UV from S. dysenteriae will be described. The structure-function of the system has been examined through site-directed mutagenesis in an in vitro proteoliposome system. Ultimately, these studies provide insight into the mechanisms by which pathogens acquire heme. The dependence on iron for survival and infection may serve as an Achilles’ heel in future drug development strategies. Advancing our understanding of how these systems work will aid in identifying potential therapeutic targets within the heme uptake systems.

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Chapter 2

Spectroscopic and Biochemical Characterization of the

Pseudomonas aeruginosa Outer Membrane Heme Receptors

2.1 Introduction

Bacterial pathogens have an absolute requirement for iron for survival and virulence. However, the transport of molecules across the outer membrane (OM) presents significant challenges for nutrient acquisition. The lipophilic nature of membranes prevents the diffusion of larger polar substrates. Additionally, the outer leaflet of Gram-negative bacteria consists of a lipopolysaccharide envelope that imparts a strong negatively charged surface that prevents the diffusion of hydrophobic molecules, such as heme. Furthermore, there is no conventional energy source available in the periplasm, such as nucleotide triphosphates, to support active transport of molecules.

Thus, Gram-negative pathogens utilize TonB-dependent OM receptors which are able to harness the energy generated by the proton motive force of the cytoplasmic membrane

(CM) via TonB to actively transport substrates into the cell (1, 2). There are two types of heme OM receptors; those that utilize a hemophore and those that directly interact with and strip heme from host heme containing proteins.

Hemophore systems have been identified in several Gram-negative pathogens such as S. marcescens (3), P. aeruginosa (4, 5), and Y. pestis (6), as well as many others.

The most well characterized system is that of S. marcescens where the biochemical and structural characterization of the hemophore-OM receptor pairing of HasA and HasR has led to much of what is known of heme transport across the OM (7-10). The hemophore,

HasA, is expressed under conditions of iron starvation where it is secreted to the extracellular space in its proteolytically processed active 17 kDa form by the action of the metalloprotease PrtSM (3, 11, 12). HasA scavenges heme from the extracellular environment mainly from host hemoglobin (Hb). HasA has an extremely high affinity

8 -1 for heme (Ka > 10 M ), yet is unable to form a stable complex with Hb, an observation that was also noted with the analogous hemophore from P. aeruginosa, HasAp (8, 13).

This was proposed to be due to a passive mode of heme acquisition as it spontaneously dissociates from Hb (13). Heme in HasA is coordinated axially by His-32 and Tyr-75, with His-83 forming a critical hydrogen bond to Tyr-75 (Figure 2.1) (14). Mutation of

His-83 to Ala resulted in a 256-fold decrease in heme affinity, demonstrating its importance in strengthening the Tyr-75 bond to heme (15). Upon heme binding, holo-

HasA delivers heme to the specific OM hemophore receptor HasR which, in conjunction with the energy transduced by TonB, is responsible for heme translocation to the periplasm (8). Early studies first demonstrated that an E. coli heme auxotroph recombinantly expressing HasR was able to grow sufficiently in iron restricted media when supplemented with heme or Hb concentrations at or above 10-4 M. However, the effectiveness at which this strain was able to utilize heme was increased 100-fold (10-6

M) when HasA was co-expressed or exogenously added to the culture media (16).

Figure 2.1 Structure of the hemophore holo-HasA from S. marcescens. Structure of holo- HasA showing key residues (yellow sticks) attributed to its high affinity for heme. Inset shows close up of the heme binding pocket, with the proximal His-32 and distal Tyr-75 heme coordinating residues, as well as His-83 involved in hydrogen bonding to Tyr-75. Adapted from PDB file 1DK0.

Perhaps somewhat surprisingly, both holo-HasA and apo-HasA were shown to form very tight complexes with HasR, both in vitro and in vivo (8, 15). Furthermore, it was shown that the transfer of heme from HasA to HasR occurs spontaneously and unidirectionally, even in the absence of the energy transducer TonB (10). This observation suggested that the thermodynamically unfavorable transfer of heme from the high affinity site of HasA to the lower affinity site of HasR must be driven by the highly exothermic and exergonic protein-protein interaction (10).

In contrast, recent spectroscopic and structural studies on the P. aeruginosa hemophore HasAp demonstrated that hydrophobic contacts between the heme and residues within the heme binding loops are primarily responsible for efficient heme capture, and that the heme coordinating residues Tyr-75 and His-32 have minimal impact on heme binding (13, 17, 18). Using time-resolved rapid freeze-quench resonance

Raman (RFQ-RR) it was shown HasAp binds heme in a two-step process where initial coordination to Tyr-75 occurs rapidly in the first binding step which then triggers closure of the loop bearing His-32 with subsequent heme coordination in a second slower step

(13). The H32A mutant only exhibited the first rapid binding phase, supporting initial coordination of heme to Tyr-75 (13). Interestingly, mutation of Tyr-75 or His-83 only minimally affected heme binding to HasAp and each exhibited biphasic binding kinetics as observed for the wild type protein, although with different intermediates (18).

Additionally, structural data revealed that Tyr-75 could still coordinate heme in the H83A mutant, an observation also seen in the RFQ-RR experiments, where instead a water molecule forms an H-bond with the phenolate oxygen of Tyr-75 (18). Taken together, the RFQ-RR and structural studies from HasAp suggest that hydrophobic interactions

60 govern efficient heme binding to HasAp and that the coordination of heme by Tyr-75 and

His-32 primarily play a role in slowing the dissociation of heme from HasAp and that these ligands may play an important role in controlled heme release from the hemophore to its cognate OM receptor.

Sequence alignments of the heme OM receptors, including the hemophore receptors, identified stretches of conserved sequence including two highly conserved histidine residues important for heme binding (19). In S. marcescens HasR, these correspond to His-189 on the apical face of N-terminal plug and His-602 on the conserved FRAP/PNPNL extracellular loop, and were confirmed as the heme binding ligands by site-directed mutagenesis and spectroscopic studies (10). The majority of heme OM receptors thus far characterized exhibit spectroscopic signatures characteristic of bis-His coordinated heme proteins, with a Soret band around 412 nm and distinct visible bands around 535 and 560 nm indicative of a 6-coordinate low-spin (6CLS) heme

(10, 20). Furthermore, the HasA-HasR-heme structure was solved and while answering some questions leaves much of the detailed mechanism of heme transfer unanswered (9).

The crystal structure revealed extracellular loops L6, L8 and L9 from HasR were all responsible for making contacts with and burying a large surface area of HasA, owing to the tight binding of the protein to the receptor. In this complex, heme was no longer bound to HasA and instead was axially coordinated by His-189 from the N-terminal plug and His-602 from the FRAP-loop in HasR, as previously predicted. A possible explanation for heme transfer was presented based on the structure of the HasA-HasR complex where extracellular loops L7 and L8 from HasR protrude into the HasA loop bearing the coordinating His-32, thereby weakening HasA affinity for heme.

Additionally, it was noted that Ile-671 from loop L8 is repositioned in the HasA pocket where heme would have been bound to HasA. This also in turn provides an explanation for heme displacement from HasA to HasR. The authors proposed a model whereby the conformational changes induced upon protein-protein interaction results in repositioning of extracellular loops L7 and L8, displacing the loop bearing the coordinating His-32 from HasA which displaces heme from HasA through a steric clash with the incoming

Ile-671 from L8. Heme is then captured by His-189 through a translocation channel capped by His-602 from the FRAP-loop, securing heme in the receptor and preventing back transfer to HasA (9). This theory was tested in the HasR I671G mutant, which retains the ability to bind heme but the heme remained bound to HasA, suggesting Ile-

671 is critical in displacing heme from HasA (Figure 2.2) (9).

Many heme receptors have been identified and characterized that directly interact and strip heme from host heme proteins. The receptor ShuA from S. dysenteriae has been shown to specifically bind host Hb and partially facilitate the transfer of heme to the receptor (20, 21). ShuA preferentially captures heme from oxidized Hb (metHb) dimers or tetramers, a physiologically relevant form of Hb upon erythrocyte lysis (20). It was further shown that ShuA was able to acquire heme from metHb on the order of magnitude of 3x103 faster than from oxyHb or free heme alone, and additionally this transfer was unidirectional (20). As with all previously characterized heme receptors, sequence alignments revealed key heme coordinating residues His-86 on the N-terminal plug and His-420 on the extracellular FRAP-loop. As expected, the crystal structure of apo-ShuA confirmed the heme coordinating ligands His-86 on the N-terminal plug and

His-420 on the extracellular FRAP-loop (22). The holo-ShuA protein exhibited the

Figure 2.2 Structure of the S. marcescens HasA-HasR complex. HasR (green) with N- terminal plug domain highlighted (red) in complex with HasA (purple). Inset shows close-up of protein interface, with heme (red sticks) coordinated by conserved His-189 on the N-terminal plug and His-602 on the extracellular FRAP-loop (cyan sticks). Ile-671 (orange sticks) is proposed to sterically clash with heme in the HasA heme binding pocket to facilitate heme transfer to HasR. Heme coordinating Tyr-75 (yellow sticks) is oriented towards the translocation channel of HasR, while the loop bearing His-32 is disordered and not resolved in the structure. Adapted from PDB file 3CSL.

63 spectroscopic signatures of a bis-His ligated heme with a Soret at 413 nm and visible bands at 535 and 565 nm (20). Mutation of either His-86 or His-420 to Ala resulted in reduced heme transfer rates from the metHb dimer and interestingly, complete loss of heme capture from metHb tetramer, whereas the double H86/H420A mutant completely loses the ability to bind heme. Additionally, in the single His-mutants, back transfer of heme from ShuA to metHb was observed, suggesting both histidine residues are important for securing heme within the translocation channel (20).

The OM receptors of Gram-negative bacteria must be energized in order to actively transport substrates such as heme across the OM. This energy is harnessed from the proton motive force of the cytoplasmic membrane (CM) and is transduced across the periplasm by the action of the TonB/ExbB/ExbD protein complex (23, 24). All TonB- dependent OM receptors contain a consensus N-terminal amino acid motif termed the

“TonB-box” which is critical for interactions with TonB (25, 26). Upon substrate binding this TonB motif becomes accessible to the periplasm to form an interprotein β- sheet with the C-terminal domain of the TonB protein (27, 28). This recognition is largely mediated through electrostatic interactions of oppositely charged residues between the C-terminus of TonB and the N-terminal TonB-box of the receptor. Once the protein contacts occur, structural and molecular dynamic simulation studies have suggested a “pulling” or “shearing” mechanical force that is applied perpendicular to the plug domain (24, 29). This mechanical force is thought to result in ejection of the intact plug into the periplasm or cause a partial unwinding of the N-terminal plug opening a permeation channel on one of the inner faces of the β-barrel translocation channel (29-

31). While no experimental evidence has been able to confirm either scenario, much of

64 the data so far more strongly supports the later model where a partial unwinding event occurs (29, 32). Once this permeation channel opens, it is proposed residues lining the inner face of the translocation channel can facilitate transfer of substrate into the periplasm where it is captured by a periplasmic binding protein (PBP). Furthermore, it has been proposed TonB may act as a protein scaffold for PBPs, positioning them near the channel opening to rapidly capture substrate for subsequent import by a specific

ABC-transporter (33). The details of the subsequent transport across the CM by the

ABC-transporter will be further elaborated upon in Chapter 4.

The opportunistic pathogen of interest in our laboratory, P. aeruginosa, encodes two systems; a hemophore-dependent and independent heme uptake system, each with their own designated OM receptor (4). Similar to that described for S. marcescens, the has system utilizes the hemophore HasAp, which specifically delivers scavenged heme to the HasR receptor. The phu system in contrast utilizes the PhuR receptor that directly interacts with host heme containing proteins. However, in PhuR, the conserved histidine on the FRAP-loop is absent, suggesting PhuR may utilize an alternate ligand set for heme coordination (Figure 2.3). To obtain a better understanding of how these pathogens acquire heme I undertook a biochemical and spectroscopic approach to examine the heme binding characteristics of both PhuR and HasR, so as to gain a better understanding of their individual contributions to heme acquisition.

Figure 2.3 Sequence alignment of the heme coordinating extracellular loop from known heme binding outer membrane receptors. Conserved heme coordinating histidine residues and FRAP/PNPNL domains in bold font. Potential heme coordinating tyrosine residues of PhuR are indicated in bold red font. Yp, Yersinia pestis; Ye, Y. entercolitica; Sd, Shigella dysenteriae; Vc, Vibrio cholerae; Pg, Porphyromonas gingivalis; Sm, Serratia marcescens; and Pa, Pseudomonas aeruginosa.

2.2 Materials and Methods

2.2.1 General Methods

Oligonucleotide primers for PCR and site-directed mutagenesis were obtained from Sigma-Aldrich. All detergents unless otherwise noted were purchased from

Anatrace. Deionized, doubly distilled water was used in all experiments. Heme was purchased from Frontier Scientific. Heme preparations were freshly made each day when required by dissolving in 0.1 N NaOH, diluting 5x in 100 mM Tris-HCl, pH 7.5, followed by filtration through a 0.20 μm polyethersulfone (PES) filter. Heme concentrations were determined by the pyridine hemochrome assay and concentrations were adjusted accordingly (34). All primary polyclonal antibodies for protein detection by Western blots were produced by Covance Custom Antibodies.

2.2.2 Bacterial Strains and Media

DNA manipulations were performed in Escherichia coli strain DH5α (F-, ara D

(lac-proAB) rpsL ϕ80dlacZ DM15 hasd R17) and E. coli strain BL21 (DE3) (B F- dcm ompT hsdS (rB-mB-) gal λ (DE3)) was utilized for protein expression. Luria-Bertani

(LB) media was routinely used for plasmid maintenance. Protein expression media varied as described below. Pseudomonas aeruginosa strain PAO1 was used for assessing the iron/heme regulation of PhuR and HasR. Pseudomonas isolation agar (PIA) was used for initial streaking and growth of PAO1 from -80°C glycerol stocks. LB media was used for liquid culture acclimation and M9 minimal media (Teknova) was used for culture maintenance under nutrient defined growth conditions.

2.2.3 Expression Vector Construction and Site Directed Mutagenesis

The hasR and phuR genes were PCR-amplified from P. aeruginosa PAO1 genomic DNA. The expression vector pET22b (Novagen) was chosen for expression of

HasR and PhuR. The first 22 amino acid residues corresponding to a P. aeruginosa signal peptide were omitted from the final gene constructs as predicted by the signal peptide program SignalP (35). The forward primer (PhuRF-5’-

TAATATGGCCATGGCCCTGGCGGGGAAC-3’) incorporated an MscI site preceding the codon for Ala-23 of phuR and allowed for incorporation of the E. coli PelB signal peptide from the plasmid backbone. The reverse primer (PhuRR-5’-

ACCATCTCGAGGATGTCCCAGACCAGGTTGACC-3’) was designed to remove the stop codon at the C-terminus of the phuR gene to allow the utilization of the His6 tag following the XhoI site in pET22b. Similarly primer HasRF-5’-

TATAGGATCCACAAGACGGGGCGGA-3’ was designed to omit the first 36 amino acids of the predicted native signal peptide of HasR and allow cloning at the BamHI restriction site preceding the E. coli PelB signal peptide. The reverse primer (HasRR-5’-

TAATGAGCTCGAACTGGTATTCGAGGGTGC-3’) removed the native stop codon to include the His6 tag utilizing the pET22b SacI restriction site. The resulting constructs, pHasR22b and pPhuR22b, incorporated the N-terminal pelB leader sequence of E. coli for targeting to the periplasm, and a His6 tag on the C-terminus for rapid purification of the proteins by metal affinity chromatography. Site directed mutants were generated by

PCR using the QuikChange mutagenesis kit (Agilent) and verified by DNA sequencing

(Eurofins MWG Operon).

2.2.4 Expression and Purification of HasAp

The expression construct of the P. aeruginosa truncated form of HasAp was provided by the Rivera lab (36). Freshly transformed E. coli BL21 (DE3) cells were used to inoculate 50 ml of LB media with added ampicillin (Amp) (100 μg/ml) and grown overnight at 37°C, 225 rpm. The following day, 10 ml of subculture was used to inoculate 1 L of LB media with added Amp (100 μg/ml). Cultures were grown at 37°C,

225 rpm until the OD600 reached ~0.8-0.9. The temperature was lowered to 25°C and cultures were induced with IPTG to a final concentration of 1 mM and allowed to grow an additional 3 h. Cells were harvested by centrifugation in a Beckman JLA8.1 rotor at

7,000 rpm and stored at -80°C until further purification.

Cells were resuspended in 4ml/g cell pellet 20 mM Tris-HCl (pH 7.6) containing

10 mM benzamidine, 2 protease inhibitor cocktail tablets (Roche Diagnostic), 1 mM

MgCl2, 20 μg/ml DNase, and 10 μg/ml lysozyme. Cells were stirred in the cold room for

1 h followed by sonication on ice to lyse the cells. Cellular debris was centrifuged

25,000 rpm for 1 h in a Beckman JA25.50 rotor and clarified supernatant was applied to a

Q-sepharose anion exchange column (2.5 cm x 10.0 cm) equilibrated in 20mM Tris-HCl

(pH 7.6). The column was washed with 10 column volumes (CV) of the same buffer and

HasAp was eluted with 0-600 mM NaCl gradient. Peak fractions were pooled and dialyzed against 4 L of 20 mM Tris-HCl (pH 7.6). Dialyzed protein was again subjected to another round of Q-sepharose anion exchange chromatography as before and peak fractions were collected and concentrated to ~ 5 ml. Concentrated HasAp was further purified by gel filtration using a Superdex S75 HR 10/300 column equilibrated in 20 mM

Tris-HCl (pH 7.6) containing 100 mM NaCl. Purified fractions were collected, pooled,

69 and concentrated to ~ 5 ml. To generate fully heme loaded HasAp, heme was prepared as described above and titrated to purified HasAp. Residual heme was removed by passing reconstituted HasAp over disposable desalting spin columns (Zeba, Thermo Scientific).

The UV-visible absorption spectra was checked to ensure proper heme loading of HasAp on an Agilent Cary 300 UV-visible spectrophotometer.

2.2.5 Expression of PhuR and HasR

Freshly transformed E. coli BL21 (DE3) cells with either pPhuR22b or pHasR22b were grown on LB agar plates with Amp (100 μg/ml) overnight at 37°C. The next day, a single colony was picked and used to inoculate 50 ml of the non-inducing media MDAG-

135 with added Amp (100 μg/ml) (37). Subcultures were allowed to grow at 37°C, 250 rpm, for 12 h. After the 12 h incubation period, 10 ml of subculture was used to inoculate 1 L of auto-inducing media ZYM-5052 or MDA-5052 (for obtaining primarily apo-protein) with added Amp (100 μg/ml) (37). Cultures were grown at 25°C, 200 rpm, for 12 h and then harvested by centrifugation at 7,000 rpm in a Beckman JLA8.1 rotor.

Cell pellets were weighed and stored at -80°C until further purification. To increase the fraction of heme loaded PhuR or HasR, heme or Hb was added to cultures during the last

1 h of incubation to a final concentration of 10 μM or 5 μM, respectively.

2.2.6 Outer Membrane Isolation from PhuR or HasR Expressing Cells

Cell pellets were thawed and resuspended in 4 ml/g of cell pellet of Buffer A (10 mM sodium phosphate (pH 7.5) containing 125 mM NaCl, 3 mM KCl, 10 mM benzamidine, 2 EDTA-free protease inhibitor cocktail tablets (Roche Diagnostic), 1 mM

MgCl2, 20 μg/ml DNase, and 10 μg/ml lysozyme. Cells were resuspended by stirring for

1 h at 4°C and lysed by passage through a French pressure cell at 18,000 psi (2 passages).

Cells were allowed to stir at room temperature for 30 min following which they were centrifuged at 12,000 rpm for 15 min in a Beckman JA17 rotor to pellet cellular debris.

Clarified lysate was retained and subsequently centrifuged again at 25,000 rpm for 1 h in a Beckman JA25.50 rotor to pellet total cellular membranes. Membrane pellets were resuspended 1 ml/g of original cell pellet in Buffer A with an added EDTA-free protease inhibitor cocktail tablet. Inner membrane proteins were selectively solubilized by addition of 2% (w/v) Triton X-100 (Sigma) and 0.5% (w/v) Sarkosyl (Teknova) and allowed to stir at room temperature for 1 h. Membranes were again pelleted at 25,000 rpm for 1 h. Triton X-100 treatment did not solubilize PhuR or HasR, at least to an extent that was detectable by visualization on SDS-PAGE and this step only served to further improve the purity of PhuR and HasR prior to ion exchange chromatography.

The resulting supernatant containing miscellaneous membrane proteins but not PhuR or

HasR was discarded. Pelleted outer membranes were resuspended in 30 ml Buffer A with an added EDTA-free mini-protease inhibitor cocktail table (Roche Diagnostic) and stirred at 4°C overnight.

2.2.7 Detergent Screening of PhuR and HasR

Total protein concentrations of resuspended OM pellets were determined using the RCDCTM protein assay (BioRad) and adjusted to 10 mg/ml. 500 μl aliquots were taken and centrifuged in a Sorvall MTX150 micro-ultracentrifuge at 50,000 rpm for 30 min in a S55A2-0187 rotor. Pellets were resuspended in 500 μl of Buffer A

71 supplemented with 2% (w/v) of 88 various detergents from a detergent screening kit

(Anatrace). Resuspended pellets were incubated on a rocking platform for 1 h at room temperature. Membranes were again pelleted and the presence of PhuR or HasR was assessed in both the solubilized fraction and membrane pellet and the extraction efficiency was visualized on 12% SDS-PAGE. The best 10 detergents were selected and further tested in secondary screens to determine extraction efficiency in the presence of varying salt, detergent concentrations, and solubilization temperatures.

2.2.8 Evaluation of Heme in the Presence of Detergent Micelles

Detergent solutions are able to give heme Sorets that can overlap with the natural

Sorets generated by heme-bound proteins. To ensure the Soret and visible bands in the

UV-visible spectrum obtained from solubilized PhuR or HasR was due to actual heme- protein interactions and not heme-detergent interactions, a suitable detergent had to be tested for and selected. Selected detergents from 10% (w/v) stocks were added to 20 mM

Tris-HCl (pH 7.5) and 100 mM NaCl to a final concentration of 3x their critical micelle concentration (CMC). A 500 μM stock of heme was prepared as described above and titrated into the detergent solution to a final concentration of 10 μM and the UV-visible absorption spectra were recorded.

2.2.9 Solubilization and Purification of PhuR and HasR

Resuspended total outer membrane protein concentrations were determined using the RCDCTM protein assay (BioRad) and total protein concentrations were adjusted to 10 mg/ml with Buffer A. Outer membranes were then solubilized by the addition of 2%

(w/v) Fos-Choline-15 (FC-15), allowed to stir at room temperature for 2 h, and then centrifuged in a Beckman JA25.50 rotor at 25,000 rpm for 1 h. FC-15 had the best efficiency at extracting PhuR and HasR from outer membranes which was determined by treating outer membrane pellets with 88 different detergents from a detergent screening kit purchased from Anatrace, as described above. Solubilized protein was then diluted 2x in Buffer B (50 mM Tris-HCl (pH 7.5) containing 250 mM NaCl, 5 mM imidazole, and

0.07% (w/v) lauryl-N,N-dimethylamine oxide (LDAO)) and loaded onto a Ni-NTA metal affinity column (2.5 x 3.0 cm) equilibrated in the same buffer. Flow through was collected and the column was washed again with 10 CV of Buffer B, followed by a second wash with 5 CV Buffer C (50 mM Tris-HCl (pH 7.5) containing 100 mM NaCl,

20 mM imidazole, and 0.07% (w/v) LDAO). Protein was eluted with Buffer E (50 mM

Tris-HCl (pH 7.5) containing 50 mM NaCl, 250 mM imidazole, and 0.07% (w/v) LDAO) and collected in 5 ml fractions. All fractions were analyzed for protein content by 10%

SDS-PAGE and peak fractions were pooled. Proteins were concentrated using Ultracel

YM-50 centrifugal filter units (Millipore) with a molecular weight cut off (MWCO) of 50 kDa. Concentrated protein was then dialyzed twice against 50 mM Tris-HCl (pH 7.5) containing 50 mM NaCl, and 0.07% LDAO using Spectra/Por CE dialysis membrane tubing (Spectrum Laboratories) with a MWCO of 50 kDa. Yields varied from protein preparation to preparation but generally averaged about 1 mg of purified protein per gram of cell pellet. Purified samples were stored at -80°C. Electronic absorption spectra were recorded on an Agilent Cary 300 UV-visible spectrophotometer after each subsequent purification column and again following dialysis to monitor the heme Soret on removal of imidazole.

2.2.10 Determination of Extinction Coefficients

The extinction coefficient of native PhuR at 280 nm was calculated to be 148.34

-1 -1 mM cm . The extinction coefficient for natively folded PhuR (εN280) was determined as follows. The absorbance at 280 nm for native PhuR (AN280) was measured. PhuR was then denatured in 6 M guanidinium hydrochloride, 0.1 M potassium phosphate (pH 7.6) and the absorbance of the denatured protein at 280 nm was measured (AD280). The concentration for denatured protein was determined using the theoretically calculated

-1 -1 extinction coefficient (εD280) of 134.08 mM cm (www.expasy.org). The extinction coefficient for the natively folded protein (εN280) was then determined according to equation 1.

εN280 = εD280 (AN280/AD280) (1)

Extinction coefficients for natively folded HasR could not be determined due to protein precipitation and aggregation. Instead, the theoretically calculated extinction coefficient at 280 nm of 126.06 mM-1cm-1 was used (www.expasy.org).

2.2.11 Circular Dichroism of PhuR and HasR

Purified PhuR and HasR were dialyzed overnight in 4 L of 20 mM sodium phosphate (pH 7.5) containing 50 mM NaCl, and 0.07% LDAO at 4°C. Proteins were diluted to 2 μM in the same buffer and circular dichroism (CD) spectra were recorded from 260 nm to 190 nm on a JASCO J-810 spectropolarimeter using a 1mm pathlength quartz cuvette. The spectra of three scans were recorded consecutively and averaged followed by subtraction from the spectrum of the buffer solution. The mean residue

74 ellipticity (deg cm2 dmol-1) was calculated after subtraction of the corresponding buffer using CDPRO software as recommended by JASCO.

2.2.12 Magnetic Circular Dichroism

Magnetic circular dichroism (MCD) spectra were recorded at 4oC and 1.41 T on a

JASCO J815 spectropolarimeter fitted with a MCD-1B electromagnet and interfaced with a Gateway PC through a JASCO IF-815-2 interface unit. JASCO software was used for data acquisition and manipulation as reported previously (38). UV-visible absorption spectra were recorded on a Cary 400 spectrometer, before and after each MCD measurement to track sample integrity. Heme concentrations were determined (based on the pyridine hemochromogen method) for the Soret absorption peaks of ferric wild type

-1 -1 -1 -1 PhuR (εSoret = 96.9 mM cm ), Y519A PhuR (εSoret = 114.0 mM cm ), Y519H PhuR

-1 -1 -1 -1 -1 - (εSoret = 100 mM cm ) H124A PhuR (εSoret = 105.8 mM cm ), and HasR (100 mM cm

1). The cyanide (CN-) adducts were prepared by adding aliquots of KCN stock solutions (1

M or diluted concentrations, pH adjusted to ~7) to the protein, respectively, until apparent saturation was reached.

2.2.13 Isolation and Characterization of Freshly Isolated Human Hemoglobin

10 ml of whole human blood was centrifuged in a Beckman JA-12 rotor at 2,000 g for 10 min. The supernatant was aspirated off and erythrocyte pellets were resuspended in

2 volumes of an isotonic buffer consisting of 20 mM Tris-HCl (pH 7.4) containing 5 mM

KCl, 2 mM CaCl2, 140 mM NaCl, and 1 mM MgSO4. Resuspended erythrocytes were pelleted again at 2,000 g for 10 min followed by aspiration of the supernatant. Three

75 additional wash cycles were performed and finally pelleted erythrocytes were resuspended in 10 volumes of a hypotonic buffer consisting of 5 mM Tris-HCl (pH 7.4) and 2 mM

EDTA and allowed to stir at 4°C for 20 min. Cells were further lysed on ice by sonication and then centrifuged in a Beckman JA25.50 rotor at 25,000 rpm for 1 h. The supernatant was collected and analyzed by 4-15% SDS-PAGE. Isolated Hb was then dialyzed overnight in 4 L of 50 mM Tris-HCl (pH 7.4), 100 mM NaCl or 50 mM Tris-HCl (pH 7.4),

500 mM NaCl. Stock solutions were kept above 600 μM to prevent spontaneous dissociation into αβ dimers and concomitant dissociation of heme upon rapid oxidation

(39).

The spectroscopic properties of the freshly isolated Hb were checked to ensure proper folding and functionality. Fresh oxyHb preps were generated by dilution of stock

Hb (≥ 600 μM) to 2 μM in 20 mM Tris-HCl (pH 7.4) containing 500 mM NaCl. deoxyHb preps were generated by addition of 1 mM sodium dithionite. metHb was generated by incubation with a 10-fold excess of potassium ferricyanide for 10 min at room temperature followed by removal of excess ferricyanide over a Zeba desalting spin column (Pierce).

All spectra were recorded on an Agilent Cary 300 UV-visible spectrophotometer.

The quaternary state of isolated human Hb was determined by FPLC analysis.

Proteins were loaded (100 μl) onto a Superdex S75 HR 10/300 gel filtration column equilibrated in 20 mM Tris-HCl (pH 7.4) containing 500 mM NaCl, at a flow rate of 0.35 ml/min. Hb dimer formation was induced by rapid dilution of stock solutions (≥ 600 μM) in 20 mM Tris-HCl (pH 7.4) containing 500 mM NaCl. metHb dimers could be formed upon incubation with 50 μM potassium ferricyanide at 37°C for 10 min. Tetrameric Hb was retained upon dilution to 20 μM in 20 mM Tris-HCl (pH 7.4) containing 100 mM

NaCl. Molecular mass markers were also run under the same conditions to generate a standard curve in the range of 200-12.5 kDa.

2.2.14 Hemin- and Hemoglobin-agarose Pulldowns of PhuR and HasR

Purified apo-PhuR or apo-HasR (10 μM) in 20 mM Tris-HCl (pH 7.5) containing

50 mM NaCl, and 0.07% LDAO in a final volume of 500 μl was incubated with hemin- or

Hb-agarose (20 μl). Samples were incubated with rotation at 25oC for 1 h. Hemin- and

Hb-agarose beads were briefly centrifuged and supernatant aspirated. Resin was washed four times in 50 mM Tris-HCl (pH 7.5) containing 50 mM NaCl and 0.07% (w/v)

LDAO. The resin was pelleted and resuspended in 50 μl 1X SDS-loading buffer, boiled, and analyzed on 10% SDS-PAGE

2.2.15 Heme Transfer from holo-HasA to apo-HasR and Hemoglobin to apo-PhuR

His-tagged apo-HasR or PhuR (5 μM) in 20 mM Tris-HCl (pH 7.5) containing 50 mM NaCl were incubated with either holo-HasA or Hb (4 μM) in a final volume of 1 ml, respectively. The proteins were incubated for 10 min at 37oC following which the receptors were immobilized on 200 μl Ni-NTA resin washed with 4 volumes of 20 mM Tris-HCl

(pH 7.5) containing 250 mM NaCl and 5 mM imidazole followed by 4 volumes 20 mM

Tris-HCl (pH 7.5) containing 50 mM NaCl and 20 mM imidazole. HasR and PhuR were eluted in the same buffer containing 250 μM imidazole. Imidazole was removed over desalting spin columns and the UV-visible spectrum for each fraction recorded to determine the extent of heme transfer from holo-HasA to HasR and metHb to PhuR. The fractions were further analyzed by SDS-PAGE for evidence of complex formation.

2.2.16 Chemical Cross-linking of PhuR and Hemoglobin or HasR and HasA

Chemical crosslinking of the HasR and PhuR receptors to their respective hemeprotein substrates was performed using two different conditions. The first involved direct crosslinking in a single step incubation period while the second involved a two step reaction where metHb or holo-HasA were first activated followed by a second reaction step with PhuR or HasR.

In the single step reaction, PhuR and HasR were dialyzed separately against 20 mM sodium phosphate (pH 7.4) containing 100 mM NaCl, 30 mM β-octylglucoside (BOG). metHb and holo-HasA were dialyzed separately against 20 mM sodium phosphate (pH 7.0) and 100 mM NaCl. PhuR and metHb or HasR and holo-HasA were mixed at 1 mg/ml each in a 500 μl total reaction volume. 10 mg of 1-ethyl-3-(3- dimethylaminopropyl)carbodiimide hydrochloride (EDC) was added directly to the reaction mixture. Samples were incubated at room temperature on a rotary platform and aliquots were taken at various time points and analyzed by SDS-PAGE.

In the two step reaction, PhuR and HasR were dialyzed separately as before. metHb and holo-HasA were dialyzed separately against 100 mM 2-(N-morpholino)ethane sulfonic acid (MES) (pH 5.0) and 100 mM NaCl. EDC and N-hydroxysulfosuccinimide (Sulfo-

NHS) were added directly to dialyzed metHb (600 μM) and holo-HasA (200 μM) to give a final concentration of 20 mM EDC and 50 mM Sulfo-NHS. Reactions were allowed to incubate at room temperature on a rotary platform for 15 min. 2 μl of 2-mercaptoethanol was added to inactivate EDC and the reaction mixtures were removed of excess EDC,

Sulfo-NHS, and 2-mercaptoethanol and buffer exchanged using Zeba desalting spin columns (Pierce) into 20 mM sodium phosphate (pH 7.4) and 100 mM NaCl. Activated

78 metHb or holo-HasA was added to PhuR (1 mg/ml) or HasR (1 mg/ml), respectively, to a final concentration of 1 mg/ml. Reactions were allowed to incubate at room temperature on a rotary platform and aliquots were taken at various time points and analyzed by SDS-

PAGE.

2.2.17 Iron and Heme Regulation of PhuR and HasR by Western Blot Analysis

Pelleted cells were resuspended in 150 μl Bugbuster HT (EMD Millipore) with an added protease inhibitor tablet (Roche Diagnostic) and incubated at room temperature for

30 min with gentle agitation. Total protein concentrations were determined by the

RCDCTM protein assay (BioRad). Samples were diluted 2x into SDS-loading buffer, boiled

15 min, and 10 μg total protein was loaded and separated by 4-15% (w/v) SDS-PAGE.

Following SDS-PAGE, proteins were electrophoretically transferred and immobilized on a PVDF membrane. The membranes were blocked overnight in a 5% blocking buffer (5%

(w/v) skim milk in Tris-HCl buffered saline with 0.2% (v/v) Tween-20 (TBST)).

Membranes were washed 3x with 25 ml of TBST and probed with a 1:2,500 dilution of primary anti-PhuR or anti-HasR polyclonal antibodies in TBST with 1% (w/v) skim milk.

Membranes were again washed 3x with 25 ml of TBST and probed again with a 1:10,000 dilution of goat anti-rabbit immunoglobin G conjugated to horseradish peroxidase (KPL,

Inc.) in TBST with 1% (w/v) skim milk. Proteins were visualized by enhanced chemiluminescence using the Super Signal Chemiluminescence kit (Pierce) using a

Fluorochem HD-2 imager (ProteinSimple).

2.3 Results

2.3.1 Expression and Purification of PhuR and HasR

The SignalP program predicted that the N-terminus of both PhuR and HasR encoded predicted signal peptides (Figure 2.4) (35). For recombinant expression the native signal peptides were removed and replaced with the E. coli signal peptide PelB following cloning into the expression vector pET22b. The PelB signal targets proteins to the periplasmic space for sorting and insertion into the OM. The pET22b receptor constructs on trial protein expression studies generated natively folded proteins that were successfully targeted to the OM. PhuR and HasR when analyzed on 12% SDS-PAGE ran at apparent molecular weights near their predicted masses of 83 kDa and 95 kDa, respectively. As previously reported for the purification of the S. marcescens hemophore receptor smHasR, apo- and holo-PhuR and HasR could be separated via anion exchange chromatography (10).

2.3.2 Detergent Screenings for PhuR and HasR Solubilization and Purification

Isolated OM preps of PhuR and HasR were subjected to solubilization screens to test for extraction efficiencies using a screening kit consisting of 88 different detergents.

From the screens, Fos-Choline 15 (FC-15) gave the best extraction efficiencies for both

PhuR and HasR. However, the expense of the detergent did not make it suitable to use throughout the remaining purification and heme binding studies. Instead, both proteins were found to be stable in a variety of detergents upon extraction, including lauryl-N,N- dimethylamine oxide (LDAO), n-dodecyl-β-D-maltoside (DDM), and at lower

Figure 2.4 Predicted signal peptide cleavage sites of PhuR and HasR. A. PhuR was predicted to cleave between residues Leu-22 and Ala-23. B. HasR was predicted to cleave between residues Ala-35 and Gln-37. Cleavage sites predicted by the program SignalP 4.1 (35).

81 concentrations, n-octyl-β-D-glucoside (BOG). However, careful consideration had to be taken in selecting the appropriate detergent, as detergent solutions with heme can yield a

Soret similar to that of known hemeproteins. DDM displays a detergent-heme Soret at

404 nm, which almost exactly overlaps with the heme-protein Soret of PhuR. Therefore various detergents utilized in the purification of membrane proteins were tested in solutions with heme to find a suitable detergent that did not exhibit these overlapping

Sorets as well as detergents that kept both membrane proteins stable once solubilized from outer membranes (Figure 2.5). In summary, while BOG exhibited desirable spectroscopic profile with heme, the high CMC value and relative instability of proteins when concentrated made this detergent unfavorable. As stated above, the cost of FC-15 does not make it a suitable choice either. Zwittergent 3-14 (ZW3-14), while used in a variety of other purification and heme binding studies with membrane proteins, began to eventually form crystalline needle precipitates at 4°C, and thus the soluble concentration cannot be assumed to remain at desired levels. LDAO exhibited the best properties to proceed with protein purification and subsequent spectroscopic studies.

2.3.3 Spectroscopic Characterization of Wild Type PhuR and HasR

As was expected, HasR showed spectroscopic signatures characteristic of the previously characterized OM receptors that contain a bis-His coordination to the heme iron (10, 19, 20, 40). Upon purification the protein had a visible pinkish-red color and

UV-visible spectroscopy revealed the Soret was centered at 411 nm with distinct visible bands at 535 and 554 nm (Figure 2.6A). Conversely, purified PhuR had a dark brownish- red color, suggesting the heme coordination may be distinct from that of HasR. Indeed,

Figure 2.5 Absorption spectra of heme in various detergents. Freshly prepared heme was titrated into various detergents at 3x their CMC value to a final concentration of 10 μM. Detergent solutions are listed in the figure legend. Values in parenthesis represent the absorbance maxima (nm) of heme in each detergent.

83 the UV-visible spectrum of PhuR revealed a blue shifted Soret to 405 nm and visible bands at 530 and 609 nm, with a slight shoulder at 564 nm (Figure 2.6B). The band at

609 nm is indicative of a HS heme species, however, the slight shoulder at 564 nm suggests there is also some LS species. This spectroscopic signature is similar to that of the well characterized hemophore HasA, which has an axial Tyr-His coordination (14,

17). Sequence alignment of all known OM heme receptors reveals conserved histidine residues on the N-terminal plug as well as in the conserved FRAP/PNPNL domain on an extracellular loop responsible for heme coordination (Figure 2.3). In HasR, these residues correspond to His-189 and His-624, whereas in PhuR the N-terminal plug His-

124 is conserved, but the conserved histidine in the FRAP-loop is absent. Other potential coordinating ligands are present, including Tyr-519 and Tyr-529, which given the similarity of the holo-PhuR spectrum to that of the Tyr coordinated hemes in the hemophore HasA and the periplasmic binding protein PhuT would give rise to the spectrum observed for holo-PhuR.

Additionally, it was noted the heme content of HasR and PhuR could be increased upon addition of exogenous heme or Hb to expression culture media during the last hour of incubation. However, only a fraction of the receptor could be isolated with heme bound and achieving 100% heme loading was not possible. Typically, the heme content of HasR or PhuR would vary from prep to prep but would on average yield around 40-

50% holo-protein when the media was supplemented with exogenous heme or Hb

(Figures 2.6C and D, respectively). The inability to obtain fully loaded receptor is perhaps not unexpected considering the proteins function to actively transport heme into the cell.

Figure 2.6 Purification of HasR and PhuR from cultures supplemented with heme or hemoglobin. A. Electronic absorption spectrum of partially heme loaded Fe(III) holo- HasR. Inset shows SDS-PAGE of purified HasR (5 μM). Lane 1, molecular weight markers; Lane 2, 10 μg purified HasR. B. Electronic absorption spectrum of partially heme loaded Fe(III) holo-PhuR (20 μM). Inset shows SDS-PAGE of purified PhuR. Lane 1, molecular weight markers; Lane 2, 10 μg purified PhuR. C. HasR purified from unsupplemented cultures ( ) or cultures supplemented with 10 μM heme ( ). D. PhuR purified from unsupplemented cultures (- - -), cultures supplemented with 10 μM heme ( ), or cultures supplemented with 10 μM metHb ( ).

The HasR and PhuR receptors were further characterized by magnetic circular dichroism (MCD). MCD spectroscopic signatures for heme proteins correlate with axial ligands and ligand field strength. Therefore, such signatures can be valuable in determining the heme coordination in unknown heme environments through comparison with structurally determined heme proteins, or porphyrin model complexes. The MCD spectrum of HasR compared to the bis-His coordinated cytochrome b5 shows similar features, although some differences in intensity are evident (Figure 2.7A). On the basis of the absorption and MCD spectrum, we conclude that holo-HasR is six coordinate low- spin (6CLS). This is consistent with the previously characterized S. marcescens hemophore-dependent HasR receptor with which the P. aeruginosa receptor has significant homology (10).

Given the potential Tyr-ligation in PhuR, the MCD spectrum was compared to phenol-bound Fe(III)-leghemoglobin A (Figure 2.7B). The absorption and MCD spectra of Fe(III) holo-PhuR at ~570 and ~420 nm suggest that PhuR is significantly more LS than leghemoglobin A. Taken together, the absorption and MCD spectra suggest that in contrast to HasR, the heme in PhuR is coordinated by histidine and an anionic O-bound ligand.

2.3.4 Identification of Tyr-519 as the Heme Ligand on the Extracellular FRAP-Loop

To determine if either Tyr-519 or Tyr-529 provide the anionic O-ligand to the heme, a series of Tyr-mutants were constructed. The absorption spectrum of the Y529A

PhuR mutant remained unchanged following purification. In contrast, the PhuR Y519A mutant upon metal affinity purification gave a red-shifted Soret at 412 nm with visible

Figure 2.7 Absorption and MCD spectra of the Fe(III) holo-HasR and PhuR receptors. A. Absorption (lower panel) and MCD (upper panel) of Fe(III) holo-HasR ( ) recorded at 4oC in 50 mM Tris-HCl (pH 7.5), 50 mM NaCl, and 0.07% LDAO and Fe(III) cytochrome b5 ( ) in 50 mM potassium phosphate (pH 7.0). The cytochrome b5 spectrum was re-plotted from reference (41). B. Conditions as in A for Fe(III) holo- PhuR ( ) and phenol-bound leghemoglobin A ( ).

87 bands at 534 and 553 nm (Figure 2.8A). However, upon dialysis and removal of exogenous imidazole, the Soret maximum shifted to 406 nm with the reappearance of the

HS marker band at 612 nm, features similar to the spectrum of the wild type PhuR

(Figure 2.8A). The coordination of an exogenous imidazole within the heme binding site of the Y519A mutant suggests that in the absence of Tyr-519, an exogenous ligand, in this case imidazole, can coordinate to the heme.

The absorption spectrum of the Y519A PhuR mutant upon dialysis resembles that of the wild type PhuR, suggesting a water ligand may coordinate in place of the Tyr-519.

However, further analysis of the Y519A PhuR mutant by MCD showed distinct differences as evident from the decreased intensity of the troughs at 420 nm and 570 nm, indicating that Y519A PhuR is more HS than the wild type protein (Figure 2.8B).

Similarly, while the Y519A PhuR mutant showed some broad similarities to the Fe(III)

6CHS met-aqua myoglobin (Mb), distinct differences are observed (Figure 2.8B). The heme environment of the Fe(III) heme in Y519A PhuR in the absence of a protein- donated ligand from the extracellular loop is likely to be highly dynamic, which may account for the unusual spectroscopic signatures observed. Consistent with the dynamic nature of the heme environment, titration of Y519A with cyanide (CN-) did not result in a completely low spin system. Titration with CN- shifts the Soret band from 407 nm to 418 nm with a LS visible peak at 540 nm (Figure 2.8C). Similarly, the MCD spectra showed a 2-fold increase in the Soret peak to trough intensities, consistent with the increase in LS character. However, the HS charge transfer band around 610 nm in the absorption spectrum, and the 630 nm trough in the MCD spectra are not completely lost even in the presence of excess CN- (Figure 2.8C).

Figure 2.8 Absorption and MCD spectra of the Fe(III) PhuR Y519A mutant. A. Absorption spectra of the imidazole coordinated PhuR Y519A (8.5 μM) ( ) and following dialysis ( ) at 4oC in 50 mM Tris-HCl (pH 7.5), 50 mM NaCl, and 0.07% LDAO. B. MCD spectra of the PhuR Y519A mutant ( ) compared with wild type PhuR ( ) as in A, and Fe(III) Mb ( ) in 50 mM Tris (pH 7.5), 50 mM NaCl. C. Absorption (lower panel) and MCD (upper panel) of PhuR Y519A (___) and Y519A-CN (aliquots of 1 mM KCN until apparent saturation) (___) at 4oC in 50 mM Tris-HCl (pH 7.5), 50 mM NaCl, and 0.07% LDAO.

In contrast to the Y519A mutant, the Y519H variant displays a Soret at 411 nm, with visible bands at 533 and 558 nm, and a complete disappearance of the HS marker feature at 610 nm. The spectrum of the Y519H protein was unchanged following dialysis, suggesting that the heme is coordinated through endogenous His-ligands, namely His-124 on the N-terminal plug and His-519 on the extracellular loop (Figure

2.9A). Consistent with the bis-His ligation, the MCD spectrum of the Fe(III) Y519H

PhuR overlays well with that of cytochrome b5 and the bis-imidazole coordinated H93G

Mb (Figure 2.9A). Furthermore, the absorption and MCD spectra are similar to those of

HasR reported herein (Figure 2.7A), and the structurally characterized OM heme receptors that contain heme coordinated by two histidines (10, 20, 40).

2.3.5 Characterization of the N-Terminal Plug Mutant H124A

To further confirm His-124 as the N-terminal plug heme ligand, the H124A mutant was constructed and characterized. The purified protein following dialysis has a blue shifted Soret peak at 402 nm and a HS marker at 607 nm (Figure 2.9B). The spectrum is reminiscent of the tyrosine ligated proteins which typically exhibit characteristic Soret maxima around 400 nm with bands in the visible region close to

~500, ~520, and ~615 nm. Absorption and MCD comparison of the H124A mutant to that of the structurally characterized 5CHS Tyr-ligated holo-ShuT showed close similarities with the exception of a decrease in the trough at ~540 nm (Figure 2.9B).

Figure 2.9 Absorption and MCD spectra of the Fe(III) PhuR Y519H and H124A mutants. A. Absorption (lower panel) and MCD (upper panel) spectra of PhuR Y519H (_ __), ______Fe(III) cytochrome b5 ( ), and Fe(III) Mb H93G-bis-Im complex (1 mM Im) ( ) at 4oC in 50 mM Tris-HCl (pH 7.5), 50 mM NaCl and 0.07% LDAO in case of the PhuR Y519H receptor. The Fe(III) Mb H93G-bis-Im complex spectrum was re-plotted from reference (42). B. Absorption (lower panel) and MCD (upper panel) of PhuR H124A (____) and holo-ShuT (____) at 4oC in 50 mM Tris-HCl (pH 7.5), 50 mM NaCl and 0.07% LDAO in the case of the PhuR H124A receptor.

2.3.6 Heme Binding and Transfer to PhuR and HasR

Attempts at loading either receptor in detergent micelles in vitro with heme were unsuccessful due to nonspecific binding and eventual precipitation of protein. This observation was consistent with previous studies done on the S. dysenteriae ShuA (20) and N. meningitidis HmbR (43) receptors. Most likely, addition of free heme to detergent solubilized OM receptors results in non-specific binding. However, both receptors as expressed were functional and able to bind heme when it was added to expression cultures during the last hour of incubation. Additionally, both HasR and PhuR in detergent micelles could bind to hemin-agarose but not to Hb-agarose resins (Figures

2.10A and 2.11A, respectively). As free heme is not likely what is encountered in the host, attempts were made to load apo-HasR with purified holo-HasA and apo-PhuR with freshly isolated metHb. Incubating apo-HasR with holo-HasA did not result in a shift in

Soret from 406 nm to 411 nm, which would be indicative of heme transfer from HasA to

HasR (Figure 2.10B). Additionally, incubation of His-tagged apo-HasR with holo-HasA did not result in a stable HasR-HasA complex that co-eluted off of Ni-NTA resin (Figure

2.10C). This is in stark contrast to what has previously been reported for the S. marcescens HasR receptor. The HasR receptor was reported to have low nanomolar affinity for its hemophore HasA in S. marcescens, both in the apo- and holo-form (8, 15).

Additionally, it was reported that holo-HasA could transfer heme to HasR in detergent micelles in a 1:1 stochiometry (10). The lack of an interaction between the P. aeruginosa

HasR and HasA proteins was not due to an effect of detergent solubilization, as no protein complex was detected when using only OM preps of HasR.

Figure 2.10 Heme binding properties of detergent solubilized HasR. A. apo-HasR following elution (50 ul) from hemin- and Hb-agarose in SDS-loading buffer. HasR (10 ul) was assessed by 10% SDS-PAGE. Lane 1, molecular weight markers; Lane 2, elution from hemin-agarose; Lane 3, elution from Hb-agarose. B. Spectra of 5 uM apo-HasR ( ), 4 uM holo-HasA ( ), and holo-HasA following incubation with apo-HasR ( ). Inset shows 5x magnification of visible regions 500-800 nm. C. Elution of His-tagged HasR (5 uM) from Ni-NTA resin following incubation with holo-HasA (4 uM). Lane 1, molecular weight markers; Lane 2, flow through; Lane 3, wash in 20 mM Tris-HCl (pH 7.5), 250 mM NaCl, 5 mM Im, and 0.07% LDAO; Lane 4, wash in 20 mM Tris-HCl (pH 7.5), 100 mM NaCl, 20 mM Im, and 0.07% LDAO; Lanes 5-7, elutions in 20 mM Tris- HCl (pH 7.5) containing 50 mM NaCl, 250 mM Im, and 0.07% LDAO.

Similarly, incubating His-tagged apo-PhuR with metHb did not result in a stable protein complex that co-eluted off Ni-NTA resin (Figure 2.11B). Due to the overlap of the heme Soret of metHb (406 nm) with that of holo-PhuR (405 nm), analysis of the heme content following elution of His-tagged apo-PhuR from metHb over Ni-NTA agarose was checked. From this experiment, there was no transfer of heme from metHb to PhuR evident (Figure 2.11C). It should be noted that the presence of detergent does have an effect on the heme Soret of metHb, possibly altering protein structural elements rendering a form that is not viable to transfer heme to PhuR. Different detergents were tested, yet all perturbed the spectrum of metHb (Figure 2.11D). This same detergent effect with holo-HasA was not evident as the electronic absorption spectrum remained unchanged.

The fact that no stable complex between apo-PhuR and Hb or apo-HasR and holo-

HasA was detected could mean that a transient complex could still exist. To test this, chemical crosslinking experiments were conducted. Under both circumstances, however, no detectable complex was formed. Additionally, to confirm the inability of PhuR and

HasR to extract heme from their respective substrates was not due to a compromise of the structural integrity of the detergent purified receptors, a CD spectrum was obtained for both PhuR and HasR (Figure 2.12). In keeping with the predicted 22-stranded β-barrel structure of the OM receptors, the CD spectra are consistent with secondary structures dominated by β-sheets. The CD spectrum of HasR does appear to have a higher α-helical content. This is consistent with the elongated extracellular loops in the hemophore receptors which have a higher degree of α-helical content and possibly due to the extended N-terminal domain that is involved in cell surface signaling.

Figure 2.11 Heme binding properties of detergent solubilized PhuR. A. apo-PhuR following elution (50 ul) from hemin- and Hb-agarose in SDS-loading buffer. PhuR (10 ul) was assessed by 10% SDS-PAGE. Lane 1, molecular weight markers; Lane 2, elution from hemin-agarose; Lane 3, elution from Hb-agarose. B. Elution of His-tagged PhuR (5 uM) from Ni-NTA resin following incubation with metHb (4 uM). Lane 1, molecular weight markers; Lane 2, flow through; Lanes 3-4, wash in 20 mM Tris-HCl (pH 7.5), 250 mM NaCl, 5 mM Im and 0.07% LDAO; Lane 5, wash in 20 mM Tris-HCl (pH 7.5), 100 mM NaCl, 20 mM Im, and 0.07% LDAO; Lanes 6-8, elutions in 20 mM Tris-HCl (pH 7.5), 50 mM NaCl, 250 mM Im, and 0.07% LDAO. C. Spectra of metHb with apo-PhuR in 20 mM Tris-HCl (pH 7.5), 250 mM NaCl, 5 mM Im, and 0.07% LDAO (), wash fraction in 20 mM Tris-HCl (pH 7.5), 250 mM NaCl, 5 mM Im, and 0.07% LDAO (), and elution fraction in 20 mM Tris-HCl (pH 7.5), 50 mM NaCl, 250 mM Im, and 0.07% LDAO (). D. Absorption spectra of metHb (2.5 μM) in various detergents at 3x CMC values, as indicated.

Figure 2.12 Evaluation of secondary structural elements of HasR and PhuR by circular dichroism. Spectra of HasR (2 μM) ( ) and PhuR (2 μM) ( ) in 20 mM sodium phosphate (pH 7.5), 50 mM NaCl, and 0.07% LDAO were recorded at 25°C. Spectra are averages of three consecutive scans.

2.3.7 Iron/Heme Regulation of PhuR and HasR

In S. marcescens the HasR receptor was proposed to have higher affinity than the non-hemophore receptor HemR, but required greater iron restriction for expression (44).

However, these studies were based solely on growth profiles of the hasR and hemR deletion strains in the presence of varying concentrations of heme and the iron chelator dipyrridyl. The hemR deletion strain, similar to wild type, could access Hb via the HasR receptor at 10-fold lower concentrations than the HemR dependent hasR mutant, but required greater iron restriction for expression. In contrast the hasR deletion strain could only access Hb at the higher heme concentration even under extreme iron restriction. It is not clear from these earlier studies if the differences in the ability to acquire heme by

HasR or HemR are due to protein abundance or intrinsic heme binding affinities of the respective receptors. To evaluate the effect of iron restriction on the expression levels of the PhuR and HasR receptors of P. aeruginosa PAO1, cells were grown in defined minimal media and supplemented with varying concentrations of iron and heme. Cells were grown to mid-log phase and protein expression levels were determined by Western blot. The protein levels of HasR and PhuR are similarly regulated by iron in the absence or presence of heme, with no evidence that the expression of HasR requires greater iron restriction than PhuR (Figures 2.13A and B). Interestingly, the PhuR and HasR receptors, as well as the cytoplasmic heme binding protein PhuS are constitutively expressed at a low level, even in the presence of 100 μg/ml iron. In contrast, the expression of heme oxygenase (HemO) is more tightly regulated by iron (Figure 2.13C).

This would suggest that on encountering heme in the environment the bacteria are

Figure 2.13 Western blot analysis of the heme uptake proteins. PAO1 cells were grown in M9 minimal media supplemented with indicated amounts of iron and/or heme. Detection of PhuR (A) and HasR (B); Lanes 1,7, and 13, molecular weight markers; Lanes 2-6, cells supplemented with 0, 1, 5, 10, and 100 μg/ml FeCl3; Lanes 8-12, cells supplemented with 0, 1, 5, 10, and 100 μg/ml FeCl3 + 0.5 μM heme; Lane 14, purified PhuR (A) or HasR (B) (10 ng). C. Detection of PhuS (upper panel) or HemO (lower panel); Lanes 1 and 9, molecular weight markers; Lanes 2-8, 0, 1, 5, 10, 25, 50, and 100 μg/ml FeCl3 + 0.5 μM heme; Lane 10, purified PhuS (upper) or HemO (lower) (30 ng).

“primed” for heme uptake. The data is consistent with previous metabolism studies from our laboratory that showed heme uptake, despite the expression of the heme uptake proteins, is dependent on the presence of a catalytically active HemO (45).

2.4 Discussion

The ability of bacterial pathogens to access and acquire iron is critical to their virulence and survival within the host. Central to this adaption is the ability to utilize heme, an abundant iron source within the host. P. aeruginosa encodes two outer membrane receptors in order to maximize its ability to acquire heme across a wide range of physiological conditions. In an effort to understand the contributions of PhuR and the hemophore-dependent HasR to P. aeruginosa heme acquisition I report here on the initial purification and characterization of the P. aeruginosa HasR and PhuR OM receptors.

Site-directed mutagenesis and spectroscopic characterization confirmed PhuR coordinates heme through His-124 and Tyr-519. This is in contrast to all previously characterized OM receptors including the structurally characterized S. marcescens HasR that have a bis-His coordination (9). Interestingly, the PhuR heme coordination is identical to that of the secreted S. marcescens and P. aeruginosa HasA hemophores (14,

17). Furthermore, Tyr ligation has emerged as a common motif across a spectrum of high affinity heme acquisition proteins and systems. These include the lipid anchored surface exposed Isd heme binding proteins of Gram-positive pathogens (46-48), and the soluble periplasmic binding proteins in Gram-negatives (49, 50). Recently, Rivera and co-workers in a series of structural and spectroscopic studies of the P. aeruginosa hemophore HasAp have proposed a model whereby heme binding is driven largely by

99 hydrophobic and π-π stacking interactions, with the heme affinity being attributed to a slow off rate. The axial Tyr ligation is stabilized by hydrogen bond interactions with either a His as in the HasA hemophores (14, 17, 51), a Tyr in the Isd heme receptors (46-

48), or an Arg in the case of the periplasmic binding proteins (50). Thus, interaction of the protein with its cognate receptor drives conformational rearrangement destabilizing the hydrogen bond interaction with the Tyr ligand and releasing the heme to its receptor

(18, 51). The fact that the S. marcescens HasR in the absence of HasA is less efficient in acquiring heme would suggest the emergence of the His-Tyr motif in surface exposed heme receptors provides as advantage in the scavenging and acquisition of heme over the bis-His coordinated OM receptors. Furthermore, if in P. aeruginosa HasR and PhuR are not differentially regulated in the presence of heme or iron, one might hypothesize that the His-Tyr coordination in PhuR provides a high affinity system that does not require the expression and secretion of a high affinity hemophore.

However, it is difficult to directly attribute heme uptake efficiency by PhuR or

HasR by simply characterizing the thermodynamic and kinetic parameters of heme binding to the isolated receptors. Heme binding and reconstitution of the purified receptors has proven to be remarkably challenging. Furthermore, in vitro heme binding affinities that have been reported for surface exposed receptors from both Gram-negative and Gram-positive bacteria are in the μM range (10, 52), a concentration well above physiological accessible concentrations of heme in the host. Our lab has previously shown the S. dysenteriae ShuA receptor following reconstitution in lipid bicelles could be partially heme loaded from metHb but not free hemin (20). However, low heme affinity is likely attributed to the absence of additional contributions from interaction with

100 specific heme protein substrates, as well as the energy transducing TonB system.

Perhaps not surprisingly, given their role in active transport the PhuR and HasR receptors are purified only partially heme loaded, even following addition of Hb to the media. In keeping with the sub-stoichiometric heme levels reported herein, similar results have been reported for other heterologous expressed OM heme receptors (10, 43).

To date, much of the information on the heme OM receptors come from studies conducted on the hemophore system from S. marcescens. However, there are several discrepancies reported in the literature to these findings. Initial studies using recombinantly expressed HasR in a heme auxotroph strain of E. coli was able to show that HasA was able to bind HasR in vivo. However, using a tonB deletion strain and 3H- heme bound holo-HasA, the authors showed that in the absence of a functional TonB system, 100% of heme remained with the hemophore HasA (8). From this they concluded that HasA binding to HasR occurred subsequent to steps involved with the

TonB energy transducer and that perhaps TonB was required for heme transfer from

HasA to HasR. In contrast to these findings, a few years later the same authors reported a

1:1 stochiometric transfer of heme from holo-HasA to apo-HasR in detergent micelles

(10). This contradiction to their previous reports would suggest that possibly the heme transfer event in vitro was an artifact of detergent solubilization and could possibly be non-specific heme transfer. The initial in vivo studies suggesting that TonB was required for heme transfer fits with our data above from the HasR system in P. aeruginosa, where heme transfer from HasA to HasR appears to be dependent upon a functional TonB. The same observation is true for that of PhuR and Hb. With the reported extremely high affinities of heme with HasA, this would seem a plausible assumption that an energy

101 source must be required for stripping heme from these heme proteins. In addition, the tight binding complex of HasR with HasA was not observed. While a tight complex between holo-HasA and HasR could be plausible, it does not seem likely that apo-HasA would have such a high affinity for HasR (reported even tighter than holo-HasA), given that the HasR system also functions as a regulator by transducing a signal through an

ECF-σ signaling cascade (53, 54). If so, the HasR signaling cascade would be constitutively activated under such a scenario, both in the presence or absence of heme.

For the reasons stated above, the data presented within is more in line with what is happening physiologically in vivo than that of the previous reports from the S. marcescens systems.

Despite the difficulties in translating such in vitro observations to the relative roles of PhuR and HasR in heme uptake, several in vivo lines of evidence support PhuR being the primary heme receptor in infection. Firstly, phuR is located in an operon encoding the necessary components required for complete translocation of heme into the cell whereas the hemophore-dependent HasR requires the phu encoded periplasmic ABC- transporter PhuT-UV. Furthermore, P. aeruginosa clinical isolates are more efficient at metabolizing heme as an iron source through up regulation of the Phu system (55).

Finally, mutations in the promoter of PhuR leading to increased expression also coincide with the loss of pyoverdine biosynthesis and an increased dependence on heme (56). The current characterization of the PhuR His-Tyr coordination along with recent in vivo studies suggests the PhuR receptor is the primary mechanism by which P. aeruginosa acquires heme, with the HasA-HasR system playing a supplementary role. To further address the contributions of PhuR and HasR to heme uptake, I have undertaken a

102 bacterial genetic and 13C-heme isotopic labeling approach in P. aeruginosa hasR and phuR deletion mutants. These studies are described in Chapter 3 and provide significant insight into the specific contributions of the PhuR and HasR receptors and their distinct heme coordination properties to heme utilization in P. aeruginosa.

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110

Chapter 3

Examining the Individual Contributions of PhuR and HasR to

Heme Acquisition in Pseudomonas aeruginosa

3.1 Introduction

Several pathogens including Y. pestis (1), S. marcescens (2), N. meningitidis (3),

V. cholerae (4), and P. aeruginosa (5), among others, encode multiple heme uptake systems. Genetic and bioinformatics approaches have made the identification of these receptors significantly easier, however, much information is still lacking on the mechanistic aspects of transport and to what extent each receptor has towards heme uptake. P. aeruginosa encodes both the Pseudomonas heme uptake (phu) and heme assimilation systems (has) (5). Each system contains its own OM receptor, the hemophore dependent HasR receptor, and the non-hemophore dependent PhuR, which can directly interact with host heme containing proteins. The P. aeruginosa HasR receptor shows high sequence homology with the S. marcescens HasR receptor (47% identities, 61% similarities) (6, 7). The S. marcescens HasR receptor has been reported to mediate signal transduction through a signaling domain located in an extra stretch of residues on the N-terminal region of the transporter (7, 8). Signal transduction is

111 relayed through the extracytoplasmic function sigma (ECF-σ) and anti-sigma factors HasI and HasS, respectively (7, 8). Binding of the hemophore holo-HasA to HasR induces a signal to the periplasm which inactivates the anti-sigma factor HasS which activates and releases of the sigma factor HasI in the cytoplasm to allow transcription of the hasRA operon (7). It was further shown that the transcription of hasS was autoregulated by HasI in a Fur-independent manner (8). This allows for accumulation of inactivated HasS, which can then rapidly respond to extracellular levels of heme to fine-tune the fluctuating extracellular heme concentrations. Bioinformatics and sequence analysis revealed P. aeruginosa HasR contains an N-terminal signaling domain similar to that of the S. marcescens HasR as well as genes upstream of HasR corresponding to a transmembrane sensor anti-sigma factor and ECF-σ factor.

P. aeruginosa is unique in that it also encodes two heme oxygenases responsible for degrading heme to biliverdin (BVIX), liberating CO and iron in the process (9, 10).

The iron-regulated heme oxygenase, HemO, is upregulated in response to iron deplete conditions due to alleviation of Fur-mediated repression. HemO degrades heme into

BVIXβ and BVIXδ isomers, distinct from the “classical” characterized heme oxygenases from other bacterial pathogens as well as from humans, which degrade heme to BVIXα.

The regioselective degradation to BVIXβ and BVIXδ by HemO is due a 110° rotation of heme that positions the δ-meso carbon in the catalytic active site. Both BVIXδ or BVIXβ isomers are possible due to rotation about the α/γ heme axis by 180° and it has been shown the δ-isomer is the predominant product with a 70:30 ratio of δ:β isomers produced in catalytic turnover studies (11). The crystal structure of the N. meningitidis

HemO revealed that key surface residues, Lys-16 and Tyr-112, were responsible for

112 hydrogen bonding to the heme propionates which positioned heme in the active site for α- regioselective degradation (12). These residues were instead replaced by Asn-19 and

Phe-117 in P. aeruginosa, and the N19K/F117Y mutant resulted in an enzyme that produced predominantly BVIXα (55%) yet still retained some BVIXδ (35%) and BVIXβ

(10%) heme oxygenase activity (11). Instead, the crystal structure of HemO from P. aeruginosa revealed Lys-132 was responsible for forming hydrogen bonds with the heme propionates which resulted in positioning and stabilization of the β- or δ-meso carbon in the active site for catalytic attack by oxygen (13). The importance of this stabilization was demonstrated upon the K132A mutation which yielded BVIXα (12%), BVIXβ

(26%), and BVIXδ (62%) (14). Additionally, it was shown that Phe-189 was critical in influencing the β/δ ratio produced by the enzyme (14).

The second heme oxygenase, BphO, is non-iron regulated and is located in an operon with the bacteriophytochrome BphP. BphO degrades heme to BVIXα which serves as a red/far-red light sensing chromophore for BphP, although it was also shown the BVIXδ was also capable of binding to BphP, albeit with a much lower affinity (10).

This is in contrast to many plant and cyanobacterial phytochromes incorporate bilins as the chromophore (15). BVIXα binds BphP through covalent attachment of the endo- vinyl group from the A-ring to Cys-12 in BphP and photoconversion between the active and inactive forms is mediated by 15Z/15E isomerization of BVIXα (16). These bacteriophytochromes functions as histidine sensor kinases that activate a response regulator for transcriptional control of specific gene targets. However, a response regulator is not located in the bphOP operon nor has it yet been identified. The conversion of heme to BVIXα by BphO relies on reducing equivalents of electrons

113 supplied from ferrodoxins or ascorbate in vitro (10). Interestingly, BVIXα is co-purified with BphO and product release was dependent on the presence of BphP or by titration with heme in vitro (10). However, while both BphO and HemO function to degrade heme, it is proposed both serve separate cellular functions in vivo. It has been previously shown in our lab that PhuS specifically delivers heme to HemO, but not BphO (17).

Thus, it is apparent that HemO is responsible for utilization of heme as an iron source.

The in vivo function of the BphO/BphP proteins and their role in sensing and regulation remains to be determined.

The BVIX isomers generated by catalytic degradation of extracellular heme are excreted into the extracellular environment either by passive diffusion or more likely an as yet unidentified exporter. Additionally, it is unknown as to what function the secreted

BVIX isomers may serve, if any at all. However, this does provide a unique opportunity to assess the efficiency at which P. aeruginosa is able to utilize extracellularly derived heme by monitoring the accumulation of BVIX in the extracellular media. In order for this to be a feasible approach, one has to be able to distinguish BVIX isomers that were derived from heme actively transported into the cell versus that which was derived from their own intracellular biosynthesized heme. P. aeruginosa does have the biosynthetic necessary necessary for heme biosynthesis and metabolizes intracellular hemein the process of amintinaing the intracellular heme and iron omeostasis. Therefore, cultures supplemented with isotopically labeled 13C-heme as a sole source of iron provides a means to distinguish between the two pathways. It has previosuy been demonstrated by our lab that when provided with 13C-heme as the sole iron source P. aeruginosa can efficiently utilize it as a source of iron (18).

114

In the current chapter, I will focus on determining the in vivo contributions of the individual OM heme receptors, PhuR and HasR, in extracellular heme uptake. We propose based on the evolution of the His-Tyr motif found in high affinity heme uptake systems studies that PhuR is the primary “high capacity” heme uptake receptor while the

HasA-HasR system may serve a supplementary role in heme uptake but a primary role as a sensor of extracellular heme status. This hypothesis is further supported by several recent reports in the literature. First, heme uptake by HasR requires is dependent on the cytoplasmic PhuT-UV transporter as the has system does not encode the proteins necessary for translocation of heme into the cytoplasm (5). Furthermore, it has been shown in a longitudinal P. aeruginosa cystic fibrosis isolates that heme is the preferred source of iron and these isolates have adapted to become extremely efficient at utilizing heme at the expense of siderophore uptake (19). Lastly, a separate independent study evaluated several different strains of P. aeruginosa from cystic fibrosis patients and determined that loss of pyoverdine mediated iron acquisition correlated with the positive selection of strains harboring mutations in the promoter region of the phu operon. These mutations resulted in significant up regulation of the PhuR receptor suggesting within the host the cells adapt to utilize heme (20). Furthermore, mutations within the phu promoter coincided with loss of the pyoverdine biosynthesis genes.

To test the hypothesis that the OM heme receptors have non-redundant roles in heme utilization we have employed bacterial genetics of P. aeruginosa PAO1 to generate individual and multiple receptor mutants to be evaluated in combination with 13C-heme isotopic labeling studies. 13C-heme isotopic labeling and liquid chromatography tandem mass spectrometry (LC-MS/MS) methodology allows for indirect analysis of

115 extracellular heme uptake efficiency in these mutants by assaying the extracellular BVIX isomer patterns. In addition qRT-PCR and Western blot analysis allowed us to evaluate the expression levels of the heme uptake proteins. This multifaceted approach allowed for the evaluation of heme uptake efficiency as well as regulation of the individual receptors and their relative contributions in maintaining intracellular iron homeostasis.

3.2 Materials and Methods

3.2.1 General Methods

Unlabeled (12C) δ-aminolevulinic acid (ALA) was purchased from Sigma.

Labeled (13C) ALA was purchased from Cambridge Isotope Laboratories. Heme was purchased from Frontier Scientific. Heme preparations were made by resuspending in

0.1 N NaOH then diluted 5x with 100 mM Tris-HCl (pH 7.5), followed by filtration through a 0.20 μm polyethersulfone (PES) filter. Heme concentrations were measured by the pyridine hemochrome method (21). All solvents used during mass spectrometry were purchased from Fisher Scientific and were LC/MS grade. All primary polyclonal antibodies for protein detection by Western blot were produced by Covance Custom

Antibodies.

3.2.2 Bacterial Strains and Culture Conditions

P. aeruginosa cultures were freshly streaked onto Pseudomonas Isolation Agar

(PIA) from -80°C glycerol freezer stocks. Colonies from freshly streaked plates were used to inoculate Luria-Bertani (LB) media and cultured overnight at 37°C, 225 rpm.

Aliquots of culture were used to inoculate fresh LB media to an OD600 of 0.1 and

116 incubated 12 h at 37°C, 225 rpm. These cultures were then used to inoculate M9 minimal media (Teknova) to an OD600 of 0.1 and grown overnight at 37°C, 225 rpm, to allow for adjustment to iron restricted conditions. An aliquot of the overnight culture was then transferred to fresh M9 minimal media to an OD600 of 0.08 and supplemented with various concentrations of iron or heme as indicated for the following growth, HPLC-

MS/MS metabolomic, qRT-PCR, and Western blot studies unless otherwise stated.

Gentamycin (75 μg/ml), tetracycline (150 μg/ml), and ampicillin (200 μg/ml) were utilized where appropriate for maintenance of the marked P. aeruginosa receptor mutants during the generation of unmarked deletions.

E. coli strain DH5α (F’, ara D (lac-proAB) rpsL ϕ80dlacZ DM15 hasd R17) was used for maintenance of plasmid pFLP2. E. coli strain S17-1 (pro thi hsdR_ Tpr Smr; chromosome::RP4-2 Tc::Mu-Km::Tn7/λpir) was used for diparental matings with P. aeruginosa mutants. E. coli strain BL21 (DE3) (B F- dcm ompT hsdS (rB-mB-) gal λ

(DE3)) was utilized for protein expression. Fresh E. coli transformants were maintained on LB agar and liquid cultures in LB. Ampicillin (100 μg/ml) was used as the selection antibiotic in E. coli for plasmid maintenance. A list of bacterial strains and plasmids used in these studies are listed in Table 3.1.

3.2.3 Generation of PAO1 Receptor Mutants

The PAO1 ΔphuR::gmFRT, ΔhasR::tcFRT, ΔhxuC, ΔphuR::gm ΔhasR::tc, and

ΔphuR::gm ΔhasR::tcΔhxuC mutants were created and generously provided by the

Amanda Oglesby-Sherrouse, PhD and Michael Vasil, PhD labs. To make the unmarked

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Strain Description Source or reference E. coli

DH5α F’, ara D (lac-proAB) rpsL φ80dlacZ DM15 Stratagene hasd R17 S17-1 pro thi hsdR_ Tpr Smr; chromosome::RP4-2 (22) Tc::Mu-Km::Tn7/λpir

P. aeruginosa

PAO1 Wild type (23)

ΔphuR::gmFRT PAO1 with 0.4 kb of phuR replaced by GmR Oglesby-Sherrouse cassette harboring flanking flip recombinase (Unpublished) sites ΔhasR::gmFRT PAO1 with 1.5 kb of hasR replaced by TcR Oglesby-Sherrouse cassette harboring flanking flip recombinase (Unpublished) sites ΔphuR::gmΔhasR::tc PAO1 with 1.1 kb of phuR replaced by GmR (5) cassette and 1.5 kb of hasR replaced by TcR cassette ΔphuR::gmΔhasR::tc PAO1 with 1.1 kb of phuR replaced by GmR Oglesby-Sherrouse ΔhxuC cassette, 1.5 kb of hasR replaced by TcR (Unpublished) cassette, and 1 kb hxuC removed by flip recombinase ΔphuR PAO1 with 1.1 kb of phuR removed by flip This study recombinase ΔhasR PAO1 with 1.5 kb of hasR removed by flip This study recombinase ΔhxuC PAO1 with 1 kb of hxuC removed by flip Oglesby-Sherrouse recombinase (Unpublished)

Plasmids pFLP2 AmpR; source of FLP recombinase (24)

MRL2 AmpR; pET-11a derivate harboring the rat liver (25) outer mitochondrial membrane cytochrome b5 gene encoding a water-soluble domain of the cytochrome b5

Table 3.1 Bacterial strains and plasmids used in the current studies.

118 mutants, the flippase (FLP) recombinase encoded from the pFLP2 plasmid used. Briefly,

E. coli S17 cells were transformed with pFLP2 and used to mate with marked mutants of

PAO1 harboring the FRT sites flanking the antibiotic cassette. After matings, PAO1 cells were streaked onto PIA plates and then challenged on LB gentamycin+ (for

ΔphuR::gmFRT) or LB tetracycline+ (for ΔhasR::tcFRT) agar plates. Sensitive colonies were selected and further streaked on LB 5% sucrose to remove the pFLP2 plasmid. All strains were verified for loss of the cassette by PCR DNA sequencing (Eurofins MWG

Operon).

3.2.4 Preparation and Isolation of 13C-Heme

δ-ALA or [4-13C]-δ-ALA was used as a biosynthetic precursor for the production

12 13 of C- or C-heme, respectively. Expression of cytochrome b5 in the presence of δ-

ALA and iron induces heme biosynthesis in E. coli where the produced cytochrome b5 captures and acts as a reservoir for the synthesized heme. 13C-heme was prepared as previously reported with minor modifications (25). A soluble form of the mitochondrial rat cytochrome b5 was expressed in E. coli BL21 (DE3) cells in M9 minimal media with the following supplements: 2 mM MgSO4, 100 µM CaCl2, 150 nM (NH4)6Mo7O24, 40

µM FeSO4 (acidified with 1 N HCl), 17 µM EDTA, 3 µM CuSO4, 2 µM Co(NO3)2, 7.6

µM ZnSO4, 9.4 µM Na2B4O7·10 H2O, and 1 mM NTA. Cells were allowed to grow at

37°C to an OD600 of ~0.9 where cytochrome b5 expression was induced by addition of 1 mM IPTG. After 10 min of incubation, heme biosynthesis was induced by addition of

13 0.1 mM [4- C]-δ-ALA and 100 mg/L FeSO4 and was further incubated for 3 h at 25°C.

Cells were harvested and lysed by sonication and cellular debris was removed by

119 centrifugation. The cellular lysate was loaded onto a Q-sepharose anion exchange column (2.5 x 10 cm) equilibrated in 50 mM Tris-HCl (pH 7.4), 5 mM EDTA, and 50 mM NaCl. The column was washed with 10 column volumes (CV) of the same buffer followed by an additional wash with 5 CV of 50 mM Tris-HCl (pH 7.4), 5 mM EDTA, and 125 mM NaCl. Cytochrome b5 was eluted in 50 mM Tris-HCl (pH 7.4), 5 mM

EDTA, and 250 mM NaCl and dialyzed overnight in 50 mM Tris-HCl (pH 7.4), 5 mM

EDTA, and 50 mM NaCl. Heme extraction from cytochrome b5 was performed by the acid-butanone method as previously described (11). The pyridine hemochrome method was used on an aliquot of extracted heme to determine final yield (21). The labeling pattern of biosynthesized heme using [4-13C]-δ-ALA, along with the BVIX fragmentation patterns and corresponding m/z are shown in Figure 3.1.

3.2.5 Extraction of Biliverdin Isomers from P. aeruginosa Supernatants

Analysis of the BVIX products were performed as previously reported with minor modifications (18). PAO1 strains were grown as described above and supplemented with either 0.5 µM or 5 µM 13C-labeled heme. 50 ml cultures were allowed to grow for 8 h at

37˚C in 250 ml baffled flasks with shaking at 225 rpm and harvested by centrifugation.

Supernatants were acidified to pH ~3 with 10% trifluoroacetic acid (TFA) and loaded over a C18 SepPak column. The column was washed with 4 ml of 0.1% TFA, 4 ml of acetonitrile:0.1% TFA (20:80), 4 ml of methanol:0.1% TFA (50:50), 450 μl 100% methanol, and then BVIX isomers were eluted with 650 μl 100% methanol. Purified

BVIX isomers were air dried and stored for up to a week at -80˚C prior to liquid chromatography tandem mass spectrometry (LC-MS/MS) analysis.

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Figure 3.1 13C-heme labeling and BVIX isomer fragmentation patterns. Incorporation of 13C carbons indicated by bold dots on heme or BVIX structures. BphO catalytically degrades heme to BVIXα. HemO catalytically degrades heme to a mixture of either BVIXβ or BVIXδ. MS/MS fragmentation site of BVIX isomers indicated by squiggly line and respective m/z of 12C- and 13C-BVIX isomers are listed.

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3.2.6 Mass Spectrometry Analysis of Extracted BVIX Isomers

BVIX isomers were separated on an ACQUITY UPLC (Waters) using a reverse phase Phenomenex Ultracarb 5 μm ODS analytical column (4.6 x 250 mm) by resuspending dried samples in 10 μl DMSO and mixing with 50 μl mobile phase (20 mM formic acid:acetone (50:50, v/v)). Particulate matter was removed by centrifugation and filtration through a 0.45 μm PTFE syringe and 10 μl of sample was injected per run at a flow rate of 0.6 ml/min. BVIX isomers were detected by monitoring the UV-visible absorption at 377 nm. Isomer masses were detected using multiple reaction monitoring

(MRM) on an ACQUITY TQD (Waters) for parent ions of m/z = 583.1 (12C-BVIX) and m/z = 591.1 (13C-BVIX). The source temperature set to 150°C, the capillary voltage at

3.30 kV, and a cone voltage set to 72 V. Collision energies were set to 34 V for BVIXα,

36 V for BVIXβ, and 30 V for BVIXδ. Fragment ions were detected at m/z = 297.1 (12C) and 301.1 (13C) for BVIXα, m/z = 343.1 (12C) and 347.1 (13C) for BVIXβ, and m/z =

402.1 (12C) and 408.1 (13C) for BVIXδ.

3.2.7 PAO1 RNA Extraction and Purification

PAO1 WT and mutant strains were grown as described above in M9 minimal media and or supplemented with or without 0.5 µM heme, 5 µM heme, or 100 µM FeCl3 as a positive control for Fur mediated iron regulation. All conditions were performed in triplicate. 1 ml aliquots were taken from cultures grown at the 8 h time point and cells were harvested by centrifugation at 7,000 rpm for 10 min. Total RNA was extracted from the cell pellets using the RNeasy mini kit (Qiagen). Residual DNA bound to the column was removed by treatment with RNase-free DNaseI (Qiagen) for 30 min and

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RNA was eluted in 40 µl RNase-free water. Eluted RNA was further treated for DNA contamination with RNase-free DNaseI (New England Biolabs) for 2 h at room temperature. RNA samples were precipitated overnight at -20˚C by the addition of 5 µl 3

M sodium acetate (pH 5.5) and 150 µl 100% ethanol. Precipitated RNA was centrifuged

13,000 rpm for 30 min at 4˚C, washed with ice cold 100% ethanol, and resuspended in 50

µl RNase-free water. RNA concentrations were determined by measuring the absorbance at 260 nm on a NanoDrop 2000c spectrophotometer (Thermo Scientific) and adjusted to

50 ng/µl.

3.2.8 Generation of cDNA and qRT-PCR Analysis

cDNA was generated from 50 ng total RNA using the ImPromII Reverse

Transcription System (Promega). Briefly, 1 µl RNA (50 ng/µl) was added to 1 µl

Random Primer (0.5 µg/µl) and adjusted to 5 µl with RNase-free water. Samples were incubated 65˚C for 10 min and cooled on ice. Master mix (4 µl 5X Buffer, 4 µl MgCl2, 1

µl dNTP’s, 1 µl Reverse Transcriptase, 5 µl RNase-free water) was added to the priming reaction and incubated at 25˚C for 5 min, 42˚C for 1 h, and 70˚C for 15 min. qPCR reactions were performed using a Light Cycler 480 (Roche) using the TaqMan Gene

Expression Master Mix (Life Technologies). CT values were calculated and relative amounts of cDNA were normalized by dividing the expression values by the relative amounts of the housekeeping gene omlA. All primers and probes used for qRT-PCR studies are listed in Table 3.2.

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Primers and Probes Sequence omlA Probe 5'-/56-FAM/CCC GAC GCC AAG TGC GGT TT/3BHQ_1/-3' omlA F Primer 5'-ACG CAG GAC ATG ATA GAC CAG TT-3' omlA R Primer 5'-TCG TTG AAG AAC AGG CTG ACG-3' phuR Probe 5'-/56-FAM/CTA CGC GCA GAC CCA CCG CAA C/3BHQ_1/-3' phuR F Primer 5'-TGA CCA ACG ACT TCT TCA GC-3' phuR R Primer 5'-CTT TAC GAT GTC CGG ATC GAC-3' hasR Probe 5'-/FAM/CTG GCC TAC GGG CAG CTC TCC TA/3BHQ_1/-3' hasR F Primer 5'-CGT GGC GTC GAG TAC CAG-3' hasR R Primer 5'-GGT CTT CGA ACA GAA GTC GTT G-3' phuS Probe 5'-/56-FAM/CTT TCG GCC GCC GCT TCG A/3BHQ_1/-3' phuS F Primer 5'-TGC CGA CGA ACA CCA TGA-3' phuS R Primer 5'-TGG CGA CCT GGC GAA A-3' hemO Probe 5'-/56-FAM/TTC GTC GCC/ZEN/GCC CAG TAC CTC TTC CAG CAT /3IABkFQ/-3' hemO F Primer 5'-TGG TGA AGA GCA AGG AAC CCT TC-3' hemO R Primer 5'-TTC GTT GCG ATA AAG AGG CTC CA-3' hasA Probe 5'-/56-FAM/TCG ACC CGA GCC TGT/3BHQ_1/-3' hasA F Primer 5'-ATC GAC GCG CTG CTG AA-3' hasA R Primer 5'-TGG TCG AAG GTG GAG TTG ATC-3'

Table 3.2 qPCR primers and probes used in the current studies.

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3.2.9 SDS-PAGE and Western Blot Analysis of Wild Type and Mutant Strains of

PAO1

1 ml aliquots were harvested at the 8 h time point as described for the isolation of total mRNA above. Cell pellets were resuspended in 100 ul per OD600 of 1.0 in

Bugbuster (Novagen). Cells were incubated at room temperature for 30 min with occasional agitation to ensure complete cell lysis and then boiled for 10 min. Total protein concentration was then determined using by BioRad RCDC Assay. 20 μl of sample was then diluted with 20 μl 2X SDS-loading dye and 30 μg total protein was loaded and run on a 4-15% SDS-PAGE. Proteins were electrophoretically transferred to a PVDF membrane (Millipore) as previously described (26). Membranes were blocked with blocking buffer (5% (w/v) skim milk in Tris-buffered saline (TBS) with 0.2% (v/v)

Tween 20), washed, and probed with a 1:2,000 dilution of anti-PhuR or anti-HasR primary antibody and a 1:2,500 dilution of anti-PhuS or anti-HemO primary antibody in

1% (w/v) skim milk in TBS with 0.2% (v/v) Tween 20. The membrane was then washed and probed with goat anti-rabbit immunoglobin G conjugated to horseradish peroxidase

(KPL) at a dilution of 1:10,000 in 1% (w/v) skim milk in TBS with 0.2% (v/v) Tween 20.

Proteins were visualized and enhanced by chemiluminescence detection using the Super- signal chemiluminescence kit (Pierce) on a Fluorochem HD2 imager (ProteinSimple).

PhuS and HemO primary polyclonal antibodies were produced from previous studies.

PhuR and HasR antibodies were generated from a 1 mg/ml stock of purified PhuR or

HasR protein in rabbits. All antibodies were produced by Covance Custom Antibodies.

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3.3 Results

3.3.1 Generation of Unmarked PAO1 Receptor Mutants

The marked antibiotic cassette single receptor mutants with flanking flippase recognition target sequences (ΔphuR::gmFRT and ΔhasR::tcFRT) of P. aeruginosa

PAO1 were created and generously provided by the labs of Dr. Amanda Oglesby-

Sherrouse and Dr. Mike Vasil, and subsequent generation of the unmarked mutants was performed utilizing flippase site-directed recombination. Unmarked mutants were verified by challenging with respective antibiotics as well as PCR sequencing of the phuR and hasR genes. The resulting mutants were determined from the sequencing results to have had a 1.1 kb DNA excision in phuR and 1.5 kb DNA excision in hasR. The marked double receptor mutant was also generously provided (ΔphuR::gmΔhasR::tc) and was also used for further studies. The deletion of these genes was also confirmed by examining the lack of protein expression in these mutants by Western blotting as discussed in the studies below. These mutants were generated and used for establishing individual contributions of each receptor to heme uptake in vivo.

3.3.2 Growth Phenotypes of PAO1 Wild Type and Receptor Mutants

The growth phenotypes of PAO1 WT and receptor mutants were analyzed under conditions of iron restriction in M9 minimal media supplemented with or without heme or iron. All strains grew similarly in low iron (Figure 3.2A) or supplemented with 100

μM FeCl3 (Figure 3.2D). When cells were supplemented with 0.5 μM heme all strains grew similarly, however, the ΔhasR strain consistently grew at a slightly faster rate

(Figure 3.2B). When supplemented with 5 μM heme, the WT and single receptor

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Figure 3.2 PAO1 wild type and receptor mutant growth curves. A. Growth analysis of PAO1 wild type (●), ΔphuR (○), ΔhasR (▼), and ΔphuR::gmΔhasR::tc (□) in M9 minimal media. B. Same as A supplemented with 0.5 μM heme. C. Same as A supplemented with 5 μM heme. D. Same as A supplemented with 100 μM FeCl3. All growth curves were performed in triplicate and standard deviations represented as error bars.

127 mutants all grew at similar rates (Figure 3.2C). Not surprisingly, when grown at 5 μM heme the ΔphuR::gmΔhasR::tc strain which lacks both OM receptors displayed a slow growth phenotype (Figure 3.2C). However, it was surprising that the

ΔphuR::gmΔhasR::tc strain was able to grow when supplemented with 0.5 μM heme, suggesting this strain was still able to obtain trace amounts of extracellular heme in the absence of both receptors.

3.3.3 Heme Metabolism Profiles of PAO1 Wild Type and Receptor Mutants

Deletion of either one or both of the receptors should have an impact on the ability of P. aeruginosa to utilize extracellular heme as an iron source. To analyze the impact this on intracellular iron homeostasis, we looked at the efficiency of heme uptake indirectly by monitoring the production of BVIX generated from the catalytic action of heme oxygenase. Previous LC-MS studies have utilized isotopic 13C-heme labeling to determine BVIX derived from the degradation of extracellular versus intracellular heme

(18, 27). Additionally, since BVIX gets excreted to the media, a simple extraction of the media could isolate BVIX isomers for analysis by LC-MS/MS methods. Furthermore, tandem ESI-MS/MS analysis can distinguish between BVIXδ and β by the action of

HemO versus BVIXα from cleavage by BphO (Figure 3.1). By monitoring the 13C-heme metabolite profile of the WT and mutant strains in combination with measurements of the steady state mRNA and protein levels of the Phu and Has systems, we were able to evaluate the individual contributions of each receptor to extracellular heme utilization.

PAO1 wild type cultures when supplemented with either 0.5 or 5 μM 13C-heme as the sole source of iron preferentially utilize extracellular heme as an iron source, as

128 evidenced from the LC-MS/MS profile where essentially all BVIX isomers detected were

13C-labeled (Figures 3.3A and 3.4A, respectively). The majority of the heme was degraded through HemO to 13C-BVIXβ and δ as judged by peaks corresponding to fragment ions at m/z 347.1 and 408.1, respectively. There was a smaller fraction of 13C-

BVIXα (m/z 301.1) present as a result of extracellular heme being “shunted” through

BphO. However, the appearance of 12C-BVIX was essentially undetectable when cultures were supplemented at both 0.5 and 5 μM heme (Figures 3.3A and 3.4A, respectively).

In contrast, when either the single ΔhasR or ΔphuR mutants were supplemented with 0.5 μM 13C-heme as the sole iron source, a shift in the ratio of intracellular to extracellular heme metabolism (Figures 3.3B and C, respectively). In these cultures the majority of BVIX produced was still 13C-labeled and catalyzed by HemO, indicating the cells were still able to uptake extracellular heme, however, were much less efficient as evidenced by the increase in 12C-BVIX. This observation was most pronounced in the

ΔphuR mutant, where the ratio of 12C-BVIX to 13C-BVIX isomers was essentially equal, suggesting a greater impact on heme acquisition efficiency (Figure 3.3C). Additionally, when compared to the WT strain, the single receptor mutants generate about three times the amount of BVIX by comparing signal intensities. Also interesting was the apparent change in isomer ratio between BVIXβ and δ in the ΔphuR mutant (Figure 3.3C). It was previously determined that the BVIXβ and δ ratio is determined by a rotation of the heme around in the α/γ axis in the HemO active site (11, 13). Therefore, this ratio is determined by how heme is “handed off” to HemO and suggests heme taken up via PhuR

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Figure 3.3 BVIX isomer fragmentation patterns for PAO1 wild type and mutant strains supplemented with 0.5 μM heme. A. LC-MS/MS fragmentation pattern of the eluting isomers in PAO1 wild type. B. As in A for the ΔhasR strain. C. As in A for the ΔphuR strain. D. As in A for the ΔphuR::gmΔhasR::tc strain. LC-MS/MS was performed as described in Materials and Methods with multiple reaction monitoring.

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Figure 3.4 BVIX isomer fragmentation patterns for PAO1 wild type and mutant strains supplemented with 5 μM heme. A. LC-MS/MS fragmentation pattern of the eluting isomers in PAO1 wild type. B. As in A for the ΔhasR strain. C. As in A for the ΔphuR strain. D. As in A for the ΔphuR::gmΔhasR::tc strain. LC-MS/MS was performed as described in Materials and Methods with multiple reaction monitoring.

131 or HasR may be delivered to PhuS in alternate orientations. However, the shift in 12C- heme metabolism and switch in BVIX isomer ratios could be suppressed upon supplementation of cultures with the higher levels of heme at 5 μM (Figures 3.4B and C).

The ΔhasR and ΔphuR mutants at 5 μM heme were also able to generate higher levels of

BVIX by about ten and five times when compared to the WT strain, respectively (Figures

3.4B and C, respectively). The reason for the apparent increase in heme metabolism when supplemented with both low and high heme was not evident until mRNA profiles and protein expression studies were examined, as will be discussed in the next section.

Not surprisingly, the ΔphuR::gmΔhasR::tc mutant was inefficient at utilizing extracellular heme, as evidenced by the predominant generation of 12C-BVIX in the LC-

MS/MS at both 0.5 and 5 μM heme (Figures 3.3D and 3.4D, respectively). The majority of intracellular heme was shunted through HemO to 12C-BVIXβ and δ as judged by the fragment ions at m/z 343.1 and 402.1, respectively. At the higher heme concentration of

5 μM, increased background levels of 13C-BVIX were detected, presumably as a result of heme diffusing across the membrane (Figure 3.4D).

3.3.4 qPCR and Western Blot Analysis of the Heme Uptake Genes in the Receptor

Mutant Strains

From the LC-MS/MS metabolite studies above, P. aeruginosa is still able to grow in the presence of heme as a sole source of iron in the absence of PhuR or HasR.

However, the heme metabolism studies suggest there is a disruption in intracellular iron homeostasis due to inefficient extracellular heme uptake, leading to subsequent degradation of intracellular heme in these mutants. To assess the effect of deletion of

132 either receptor, quantitative real time PCR (qRT-PCR) was used to evaluate the steady state mRNA transcript and protein levels of the heme uptake and utilization proteins in the mutant strains compared to PAO1 wild type.

The ΔhasR strain shows a modest increase in the levels of phuR mRNA by about

2.5- and 2-fold when compared to PAO1 wild type mRNA levels at 0.5 and 5 μM heme, respectively (Figure 3.5A). In keeping with the increase in phuR mRNA, increased levels of phuS and hemO were also observed at both heme concentrations (Figure 3.5D and E, respectively). This increase in mRNA levels directly translated to increased levels of

PhuS and HemO protein expression levels when compared to wild type PAO1 (Figure

3.6B).

Similarly, the ΔphuR strain also displayed an increase in the mRNA transcript levels of hasR, although to a much greater extent. The steady state mRNA levels of hasR are up regulated about 6- and 30-fold when compared to PAO1 wild type mRNA levels at

0.5 and 5 μM heme, respectively (Figure 3.5B). As HasR utilizes the hemophore HasA as a substrate, the steady state mRNA levels of hasA were also examined to rule out that inefficient heme utilization was a consequence of a lack of concurrent hasA transcription with hasR. qPCR analysis revealed that hasA levels were also up regulated in the ΔphuR strain when compared to PAO1 wild type by about 6- and 4-fold in 0.5 and 5 μM heme, respectively (Figure 3.5C). In keeping with the up regulation of the receptors, levels of phuS and hemO mRNA were similarly elevated (Figures 3.5D and E, respectively). This also translated to an increase in the PhuS and HemO protein levels (Figure 3.6B). The significant up regulation of HasR and the intracellular heme utilization proteins PhuS and

HemO, coupled with the increased metabolic flux of intracellular 12C-heme through

133

Figure 3.5 Effect of heme and iron on mRNA transcription levels of the heme utilization systems. RNA isolated from the indicated strains at the 8 h time point supplemented with either heme or iron was quantified by qRT-PCR as described in the Materials and Methods. mRNA levels of phuR (A), hasR (B), hasA (C), phuS (D) and hemO (E). The indicated p-values were normalized to mRNA levels of the PAO1 under the same conditions where *p < 0.05, ** p < 0.005, or ***p < 0.001.

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Figure 3.6 Effect of heme and iron on the heme uptake protein expression levels in PAO1 WT and receptor mutant strains. A. Western blot analysis of PhuR (upper panel) and HasR (lower panel) at the 8 h time point in P. aeruginosa strains as indicated. B. Western blot analysis of PhuS (upper panel) and HemO (lower panel) at the 8 h time point in P. aeruginosa strains as indicated.

135

HemO as evidenced in the metabolic studies, suggests HasR is less efficient than PhuR at acquiring heme.

Interestingly, Fur-mediated repression of PhuS appears to have been disrupted in the ΔphuR mutant in the presence of 100 μM FeCl3, even though sequence analysis revealed the Fur box in the promoter region remained intact (Figure 3.5D). This was also noted in a ΔphuS mutant where Fur-mediated repression of PhuR was similarly effected

(unpublished data). PhuR and PhuS are divergently transcribed from the same promoter region, however, the fact that the Fur box remained intact in both mutants suggests there may be additional regulatory elements upstream of the currently defined promoter (5).

Also of interest was the OM receptors PhuR and HasR (Figure 3.6A), as well as the cytoplasmic chaperone PhuS (Figure 3.6B), were expressed at low basal levels even in the presence of 100 μM FeCl3. In contrast, HemO expression is more tightly regulated by iron (Figure 3.6B). This would suggest that upon encountering heme in the environment the bacteria are “primed” for heme uptake. Furthermore, the data is consistent with previous metabolism studies showing extracellular heme uptake is dependent on the presence of a catalytically active HemO (27).

3.4 Discussion

Due to the complications in extrapolating heme binding affinities of the detergent isolated receptors in vitro, the current work presented in this chapter was aimed at evaluating the relative individual contributions of PhuR and HasR towards heme acquisition in vivo. Previous studies in S. marcescens suggested that the hemophore dependent HasR represented the high affinity receptor system while HemR represented

136 the lower affinity system (2). However, the data presented within suggests that at least in

P. aeruginosa the HasA-HasR system has a lower capacity for heme uptake and instead may serve primarily as a sensor of extracellular heme. Activation of the HasA-HasR signaling cascade may lead to up regulation of the high capacity Phu system, the major facilitator of extracellular heme acquisition, as heme uptake through the Has system must utilize the Phu system proteins for complete import to the cytoplasm. Therefore, as heme acquired through HasR is dependent upon the periplasmic and cytoplasmic PhuT-UV transporter, we hypothesize some “cross-talk” exists between the hemophore signaling cascade and the phuSTUV genes, which are divergently transcribed yet under the same promoter as phuR (Figure 3.7).

The data presented above would suggest that both systems are optimally required for heme uptake, as deletion of either receptor results an imbalance in iron homeostasis as evidenced by the catalytic turnover of biosynthesized 12C-heme by HemO. The deletion of the phuR gene, however, appeared to have a more drastic perturbation on intracellular iron homeostasis, as evidenced by an increase in the ratio of intracellularly derived 12C-

BVIX to extracellularly derived 13C-BVIX. This was also apparent from the qPCR and western data, where both HasR and HasA are significantly up regulated to counteract the loss of PhuR. Therefore, the PhuR receptor and the emerging His-Tyr motif in high affinity heme uptake systems indicate that PhuR is the primary mechanism for heme transport, whereas the HasR receptors primary function is in sensing extracellular heme and activating heme uptake.

137

Figure 3.7 Proposed model for the Pseudomonas aeruginosa heme cell surface signaling system. A. In the absence of heme HasS binds the HasI σ-factor inhibiting its activity. B. In the presence of heme HasA binds to HasR initiating TonB-dependent import of heme into the cell and signaling displacement of the anti-σ factor freeing HasI for increased expression of the heme uptake systems. Positive regulation of hasS allows inactive HasS to accumulate so that as extracellular heme levels decrease HasI can be readily inactivated, rapidly down regulating the heme uptake systems.

138

Further support for PhuR being the primary receptor in heme acquisition comes from recent studies utilizing clinical strains of P. aeruginosa in chronic infection.

Natural mutations within the phuR promoter of these clinical isolates resulted in increased expression levels of PhuR, conferring a growth advantage in the presence of hemoglobin. Furthermore, this evolution within the host towards increased reliance on heme coincided with a loss of the alternative iron-scavenging siderophore systems (20).

While the HasA-HasR system is important for efficient heme acquisition, it may play a more significant role in acute infection, while the evolution towards utilizing the higher capacity Phu system may provide an advantage during chronic infection.

It is interesting that the double receptor mutant was able to grow in the presence of heme and that when looking at the heme metabolite profiles that 13C-BVIX was still being produced. This suggests that P. aeruginosa is able to still uptake extracellular heme in the absence of both PhuR and HasR. There may possibly be an as yet unidentified third receptor responsible for heme acquisition. Another possibility would be these cells become stressed and their membrane integrity becomes compromised, allowing for passive diffusion of heme. The homologous HxuC protein from H. influenzae was proposed to be a heme OM receptor based on the observation it was able to bind heme in vitro (28). However, the relevance of this is uncertain, as previous attempts at loading the OM receptors PhuR and HasR with heme resulted in non-specific binding and protein aggregation, an observation also reported with ShuA from Shigella dysenteriae (29) and HmbR from Neisseria meningitidis (30). The BVIX isomer patterns using a single mutant ΔhxuC strain and triple mutant ΔphuR::gmΔhasR::tcΔhxuC strain were analyzed and compared, yet did not deviate from that of PAO1 wild type and the

139

ΔphuR::gmΔhasR::tc strains, respectively. It is clear that deletion of hxuC had no effect whatsoever on heme uptake, discrediting HxuC as a heme transporter in P. aeruginosa.

Furthermore, the alleviation of Fur-mediated iron suppression of PhuS in the

ΔphuR mutant is quite intriguing. The same effect was seen in our lab for PhuR in a

ΔphuS mutant (unpublished data). These two genes are located in the phu operon and are under negative control by the same Fur-box, but are divergently transcribed from each other (5). It is interesting to note that this genetic arrangement is only seen in the pathogenic strains of P. aeruginosa, indicating this particular genetic arrangement may instill some advantage toward heme uptake. Sequence analysis of the promoter region, specifically the Fur-box, revealed no disruption in the genetic sequence resulting from the genetic manipulations in these studies that could foreseeably result in disruption of Fur binding to its target DNA sequence. It is entirely possible that there are other genetic elements encoded in the coding region of phuR or phuS that are responsible for co- regulation with Fur.

Although we cannot say with certainty PhuR represents a high affinity receptor, the heme coordinating ligands His-124 on the N-terminal plug and Tyr-519 on the extracellular FRAP-loop, as detailed in Chapter 2, represent a common motif among high affinity heme binding proteins. The His-Tyr heme binding motif is found in the high affinity hemophores from Gram-negative pathogens (31, 32) and the lipid anchored, surface exposed receptors of Gram-positive pathogens (33-35). In conclusion, we propose P. aeruginosa utilizes both systems to rapidly respond to and utilize heme across a range of physiological conditions within the host, with PhuR evolving the high affinity

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His-Tyr motif to serve as the major facilitator of heme acquisition with HasR acting supplementary to PhuR as well as functioning as an extracellular heme sensor.

3.5 References

1. Rossi MS, et al. (2001) Identification and characterization of the hemophore-

dependent heme acquisition system of Yersinia pestis. Infect Immun 69(11):6707-

6717.

2. Benevides-Matos N & Biville F (2010) The Hem and Has haem uptake systems in

Serratia marcescens. Microbiology 156(Pt 6):1749-1757.

3. Perkins-Balding D, Baer MT, & Stojiljkovic I (2003) Identification of

functionally important regions of a haemoglobin receptor from Neisseria

meningitidis. Microbiology 149(Pt 12):3423-3435.

4. Wyckoff EE, Schmitt M, Wilks A, & Payne SM (2004) HutZ is required for

efficient heme utilization in Vibrio cholerae. J Bacteriol 186(13):4142-4151.

5. Ochsner UA, Johnson Z, & Vasil ML (2000) Genetics and regulation of two

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Chapter 4

Heme Transport Across the Cytoplasmic Membrane:

Reconstitution of the Shigella dysenteriae ShuUV ABC Transporter

in a Proteoliposome System

4.1 Introduction

Upon import of extracellular heme into the periplasm via dedicated TonB- dependent outer membrane (OM) receptors, heme must traverse the periplasmic space, and be translocated across the cytoplasmic membrane (CM) into the cytoplasm.

However, this must be a carefully orchestrated process to prevent the potentially toxic and damaging side effects of free heme in the cell. Thus, pathogenic bacteria utilize high-affinity periplasmic transport systems that sequester and translocate heme in a coordinated fashion facilitated by specific protein-protein interactions. Shigella dysenteriae like most other pathogenic bacteria encode a periplasmic transport system comprising a soluble periplasmic heme binding protein (PBP) ShuT and a CM ATP- binding cassette (ABC) transporter ShuUV, which imports heme into the cytoplasm where it is captured by a soluble cytoplasmic heme binding protein (CBP) ShuS (Chapter

146

1 Figure 1.8) (1). The ligand driven conformational changes coupled to specific protein- protein driven ligand release is an effective method to control the unidirectional transport of heme into the cell while abrogating potentially toxic side effects of free heme release.

S. dysenteriae is able to capture and transport heme from the host heme containing proteins such as hemoglobin through the TonB-dependent OM receptor ShuA

(2). Upon active transport into the periplasm heme is sequestered by the PBP ShuT with very high affinity through a conserved Tyr residue (3). These proteins form an extensive class of substrate specific binding proteins that are responsible for binding and shuttling diverse substrates across the periplasmic space. They all have in common an overall protein architecture that is composed of N- and C-terminal lobes with a substrate binding cleft between the two domains. In ShuT, each of these domains consist of a central 4 stranded β-sheet arrangement flanked by 5 α-helices that are interconnected by a single long α-helical linker or “hinge” domain (4). A narrow but deep substrate binding cleft is sandwiched between the two domains that nearly completely buries the heme molecule within the protein, attributing to this protein’s extremely high affinity for heme. These

PBPs can be structurally grouped according to the number of interconnecting linkers between the two domains (5, 6). The PBPs involved in binding heme, such as ShuT (4),

2+ vitamin B12, BtuF (7), iron-siderophores, FhuD (8), and Zn ions, TroA (9), all contain a single interconnecting α-helical linker and are classified as type III PBPs. The type I

PBPs, such as MalE which binds maltose (10), and type II PBPs, such as GlnBP which binds glutamine (11), contain three and two linkers interconnecting the two domains, respectively.

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It was shown in the type I PBPs, such as the maltose binding protein MalE, that large conformational changes occurred upon substrate binding that act as a “Venus flytrap” by clamping the N- and C-terminal lobes around the binding cleft (10, 12, 13).

These large conformational reorientations were proposed to be important for both ABC transporter recognition as well as release of substrate from the PBP to the transporter. In contrast, the crystal structure of apo- and holo-ShuT revealed minimal conformation changes upon substrate binding, indicating the type III PBPs may be more rigid (Figure

4.1) (4). This was also noted in FhuD and to a lesser extent in BtuF (14). Molecular dynamic simulation studies have supported the finding that the type III PBPs are more rigid, however, may be more flexible than previously thought from interpretation of the crystal structures alone (15-17). Nonetheless, it is the “closed” holo-protein that is then functionally capable of recognizing and interacting with its cognate ABC transporter.

Once heme is transported through the CM by the ABC transporter it must be sequestered on entering the cytoplasm. The cytoplasmic binding proteins (CBP) such as those encoded within the heme uptake operons were hypothesized to play this role.

However, the function of the cytoplasmic heme binding proteins has been a topic of debate in the literature, where the E. coli cytoplasmic heme binding protein ChuS was proposed to function as a heme oxygenase, despite sharing no sequence or structural similarities to previously identified heme oxygenases (18-21). These in vitro experiments demonstrated the protein’s ability to degrade heme when provided with reducing equivalents, however, the studies did not take into consideration the heme degradation reaction was due to non-catalytic coupled oxidation by H2O2 (22, 23). Contrary to these reports, the cytoplasmic binding proteins PhuS from P. aeruginonsa and ShuS from S.

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Figure 4.1 Structure of the periplasmic heme binding protein ShuT. A. apo-ShuT and B. holo-ShuT showing slight conformational rearrangements associated with heme binding. Heme coordinating Tyr-94 (green) adopts an outward facing conformation in the apo- structure and adopts an inward facing conformation upon binding heme, securing heme within the binding cleft. Adapted from PDB files 2R7A and 2RG7.

149 dysenteriae were shown to be important for heme utilization, but did not function as heme degrading enzymes (24-26). Furthermore, PhuS has been shown to be a heme chaperone to the iron regulated heme oxygenase HemO (26-28). From these reports, it is clear the function of the heme binding CBPs is to sequester heme as it enters the cytoplasm and, at least in the case of PhuS, shuttle it to the heme degrading enzymes.

The heme binding CBPs share a common overall fold that has been described as a

“β-propeller” from similarities to the heme binding protein hemopexin (19, 20, 29, 30).

The overall structure is composed of two structurally similar N- and C-terminal domains each composed of a core of 9 β-sheets flanked by α-helices (Figure 4.2). Heme is bound by a conserved histidine residue in the C-terminal domain, corresponding to His-193 in

ShuS. From the homologous protein PhuS it was determined the proximal histidine residue only minimally had an impact on heme binding and instead, a deep and predominantly hydrophobic cleft was responsible for heme binding affinities associated with this protein (28, 31, 32). Conformational changes induced in PhuS upon binding heme drive the interaction with its heme delivery partner, the iron regulated heme oxygenase HemO (28). While no heme oxygenase has yet been identified in S. dysenteriae, ShuS is still proposed to function as a heme chaperone. Furthermore, it was shown apo-ShuS can interact with and bind DNA and this effect was reversible upon titration with heme, suggesting ShuS may be involved in other cellular functions in addition to serving as a heme chaperone (24, 33).

ABC transporters are a ubiquitous class of importers/exporters involved in a wide variety of cellular processes in eukaryotes, prokaryotes, and archaea, and represent one of the largest known superfamily of proteins (34, 35). They are responsible for the

150

Figure 4.2 Structure of holo-ChuS from E. coli. Structure of the cytoplasmic heme binding proteins displaying symmetrical N- and C-terminal domains. Core consists of a β-propeller (purple) flanked by α-helices (green). Heme coordinating histidine is displayed (red sticks). Adapted from PDB file 4CDP.

151 unidirectional transport of substrates across biological membranes, by coupling substrate translocation with hydrolysis of ATP. These transporters are generally comprised of two transmembrane domains (TMD) forming the translocation channel and two cytosolic nucleotide binding domains (NBD) responsible for binding and hydrolysis of ATP. The

TMD’s, owing to their abilities to transport chemically diverse substrates, are highly variable in both polypeptide chain length and sequence. In contrast the NBD’s, have among the most conserved phylogenetic sequences of any protein family, indicating a conserved mechanism for coupling the hydrolysis of ATP to substrate translocation (36).

These highly conserved regions include the Walker A and the “signature” LSGGQ motif in the Walker C regions responsible for binding ATP, the Walker B region responsible for binding Mg2+, and the invariant Gln on the Q-loop and His in the “switch” region critical for catalyzing ATP-hydrolysis (37). This high degree of conservation among catalytically essential groups initially lead to the proposal that all ABC transporters would function by a common mechanism, however, recent studies in ABC transporter structure and function have revealed many differences in this large superfamily of proteins.

In S. dysenteriae, holo-ShuT in its “closed” conformation is able to recognize and interact with its heme specific ABC transporter ShuUV. This interaction is primarily mediated by electrostatic interactions through two conserved acidic glutamate resides on each lobe of ShuT and conserved basic arginine resides extending from the surface of a periplasmic loop from each ShuU subunit that form salt bridges (38). As with most ABC transporters, ShuUV is comprised of two TMD subunits of ShuU and two NBD subunits of ShuV, to form an overall ShuU2V2 heterotetramer, based on homology modeling with

152 the E. coli vitamin B12 transporter BtuCD and the Y. pestis heme transporter HmuUV (39,

40). The two ShuU subunits are each composed of 10 α-helices that span the CM, forming a heme translocation pathway consisting of a total of 20 α-helices. The two soluble ShuV subunits are responsible for binding and hydrolysis of ATP that energizes the transporter for import of heme into the cytoplasm.

ABC transporters can be grouped into exporters or importers, which can further be subgrouped based on their overall membrane domain architecture and topology (41,

42). Recently, it was proposed these subgroups could be further divided by their mechanism of transport (43). ShuUV is a type II ABC transporter, which also includes the vitamin B12 transporter BtuCD of E. coli and the metal chelate transporter Hi1470/71 of H. influenzae. BtuCD has been extensively studied both structurally and functionally, and has contributed to most of how we understand these transporters to function. The crystal structure of BtuCD revealed a TMD architecture consisting of 20 membrane spanning α-helices that form the translocation channel, typical of type II transporters

(39). In this structure, an outward facing conformation of the TMD’s in the absence of bound nucleotides in the NBD’swas observed, in contrast to that seen in the type I transporters such as MalFGK or the other type II transporter Hi1470/71, which adopt an inward facing conformation (44, 45). This structure was somewhat controversial and it was proposed the outward facing conformation of BtuCD in the absence of nucleotide was artificial due to detergent selection in the crystallization process. However, numerous studies have since suggested that this conformation is physiologically relevant and that BtuCD may indeed have a distinct mechanism for substrate translocation (46-

49). Additionally, the recently solved crystal structure of the heme ABC transporter

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HmuUV from Y. pestis further supported these findings (40). The cumulative data supports a proposed “peristaltic pumping” mechanism for substrate import (Figure 4.3).

In this model, the transporter adopts an outward facing conformation, ready to accept substrate from its cognate periplasmic binding protein, while sealed to the cytoplasmic side through gate I formed by transmembrane helix (TM) 5 (39). Upon substrate binding and release to the translocation channel, the transporter adopts an intermediate conformation by opening of TM5, however, is still sealed to the cytoplasm by repositioning of TM2 and TM3 which form gate II, creating a sealed cavity within the receptor (50). A separate study on the type II molybdate transporter MolBCA also suggested a centrally closed cavity prior to substrate release to the cytoplasm (51).

Subsequent hydrolysis of ATP results in substrate release to the cytoplasm followed by the release of ADP which resets the receptor for subsequent rounds of import.

While the data supporting a mechanism of a “peristaltic pump” has been proposed, it is evident that the type II transporters may have subtle differences depending on the substrate being transported (43). While the crystal structure of Y. pestis HmuUV heme transporter implies a similar mechanism to BtuCD due to overall transporter architecture and the observation of an outward facing conformation in the resting state, slight variations in the TMDs proposed to be involved in accommodating heme as a substrate may separate the mechanistic properties between the two (40). This was noted when the crystal structure of HmuUV was compared to that of the BtuCD structure, where a narrower substrate binding channel in the transmembrane domains more likely accommodates the less bulky and smaller heme group than that of the substrate cobalamin for the BtuCD transporter, which had a much larger substrate channel (40).

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Figure 4.3 Proposed model of transport for heme through the periplasmic ABC transport system. A. In the resting state of the transporter, the TMDs are in a closed conformation to the periplasmic side and the NBDs are separate. Heme bound PBP recognizes and interacts with its specific ABC transporter in this conformation. B. The holo-PBP binds to its cognate receptor, inducing conformational changes in the TMDs resulting in opening of the translocation channel and opening of the PBP to facilitate transfer of heme to the channel. This in turn facilitates dimerization of the NBDs, enhancing their affinity for ATP. Subsequent hydrolysis of ATP to ADP and inorganic phosphate by the NBDs generates a “power stroke” that energizes the transporter for heme translocation. C. The resultant “power stroke” upon transport of heme resets the transporter and releases the PBP for subsequent rounds of transport. Upon the completion of heme translocation, heme is captured by the CBP in the cytoplasm.

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Furthermore, the observation that the S. dysenteriae cytoplasmic heme binding protein

ShuS was additionally required for ShuUV function also possibly hints at additional mechanistic steps involved in heme transport and release to the cytoplasm (38). We have previously reconstituted the ShuUV transporter in proteoliposomes and shown transport from holo-ShuT encapsulated in the lumen of the liposome to ShuS in the external medium (38). In order to dissect the mechanism of heme translocation across the cytoplasmic membrane a combination of site-directed mutagenesis and spectroscopic studies was proposed to trap intermediates in the translocation channel. As a first step toward this goal I undertook a redesign and optimization of the ShuUV expression system to generate protein levels required for such studies. In this chapter, constructs of

S. dysenteriae ShuUV were generated and tested to optimize protein expression and reconstitution efficiency for the future structural and functional studies that will address the mechanism of heme translocation and possible mechanistic differences between type

II ABC transporters.

4.2 Materials and Methods

4.2.1 General Methods

Deionized, doubly distilled water was used in all experiments. Oligonucleotide primers for PCR were purchased from Sigma-Aldrich. Sequence verification of plasmid constructs was performed by Eurofins MWG Operon. All DNA modifying enzymes and restriction enzymes were purchased from New England Biolabs. Solvents used for peptide extractions were all LC-MS grade and purchased from Fisher Scientific.

Detergents used throughout purification of ShuUV were purchased from Anatrace unless

156 otherwise stated. Lipids for in vitro reconstitution of ShuUV proteoliposomes were purchased from Avanti Polar Lipids. Heme was purchased from Frontier Scientific.

Heme preparations were freshly made each day when required by dissolving in 0.1 N

NaOH, diluting 5x in 100 mM Tris-HCl (pH 7.5), followed by filtration through a 0.20

μm polyethersulfone (PES) filter. Heme concentrations were determined by the pyridine hemochrome assay and concentrations were adjusted accordingly (52). All electronic absorption spectra were recorded on an Agilent Cary 300 UV-visible spectrophotometer.

4.2.2 Bacterial Strains and Culture Conditions

Plasmid DNA expression and maintenance was performed in E. coli strain DH5α

(F’, ara D (lac-proAB) rpsL ϕ80dlacZ DM15 hasd R17). E. coli strain BL21 (DE3) (B F- dcm ompT hsdS (rB-mB-) gal λ (DE3)) was utilized for protein expression of both

- - - ShuUV and ShuS and E. coli strain BL21 (DE3) pLysS (F ompT hsdSB (rB mB ) gal dcm

(DE3)) was used for expression of ShuT. Luria-Bertani (LB) media was routinely used for plasmid maintenance. Protein expression media varied as described below.

4.2.3 Construction of Over-expression Vectors for Optimal ShuUV Production

shuUV was PCR amplified from plasmid pHTL106 (53). chuUV was PCR amplified from E. coli O157:H7 str. EDL933 genomic DNA. The first encoded α-helix of btuC was PCR amplified from E. coli O157:H7 str. EDL933 genomic DNA. Various pET (Novagen) expression vectors were used for testing protein expression. All restriction enzymes were purchased from New England Biolabs. A list of constructs along with their descriptions are listed in Table 4.1.

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4.2.4 Test Expressions of Optimized Constructs

A single colony from freshly transformed cells was used to inoculate 50 ml of LB with ampicillin (Amp) (100 μg/ml) and incubated overnight at 37°C, 225 rpm. 5 μl of overnight subcultures were used to inoculate 5 ml of various media with Carb (100

μg/ml) to test expression profiles: M9 minimal media, LB, Terrific Broth (TB), and

ZYM-5052 auto-inducing media (54). Cultures were set up in duplicates and grown at either 25°C or 37°C, 200 rpm. Duplicate culture sets were also set up in triplicate to be induced with either 0, 0.3, or 1 mM IPTG once the OD600 reached ~0.7. Aliquots of cells were taken pre-induction and 3, 6, 12, 24, and 36 h post-induction and analyzed by 12%

SDS-PAGE in comparison with empty control vector for over expression of ShuUV.

4.2.5 Construction of the Chimeric Over-expression Construct pCh1SdUV

The sequence corresponding to about 350 bp upstream of the ATG start codon of the btuC gene to the end of sequence encoding the first α-helix was cloned from E. coli

O157:H7 str. EDL933 genomic DNA utilizing the forward primer BtuCh1F (5’-

TCGAAGATCTGCTTACAAAAGCTGAAATGTCAG-3’) which incorporated the BglII restriction site, and the reverse primer BtuCh1R (5’-

GGTATCTAGAAAGGCTTAAGAGAAGCGCC-3’) which substituted the 6 bp XbaI restriction site in place of the codons for Cys-32 and Ala-33. The PCR product was subcloned into pET16b resulting in construct pBtuCh1.

The sequence starting at the C-terminus of the first predicted α-helix of shuU

(Gly-30) through the stop codon of shuV was cloned from plasmid pHTL106 (53) utilizing the forward primer ShuUΔh1F (5’-

158

GGTCTCTAGAGGAGCGTTAAAGATCAGTTTTG-3’) which incorporated the XbaI restriction site in place of codons for Ser-28 and Gln-29 of shuU, and the reverse primer

HmuVR3 (5’-TTCAGGATCCTTAATGATCGAGGACGATCAG-3’) which incorporated the BamHI restriction site following the native stop codon of shuV. The

PCR product was subcloned into pBtuCh1 resulting in construct pShuUVΔh1, which contained the 5’ untranslated region of btuC and the chimeric shuUV gene harboring the first encoded α-helix of btuC.

The full sequence containing the chimeric gene of shuUV was cloned from pShuUVΔh1 using the forward primer BtuCF (5’-ATTGCATATGCTGACACTTGCCG-

3’) which incorporated the NdeI restriction site at the start of the btuC gene, and the reverse primer HmuV3 which incorporated the BamHI restriction site following the stop codon for shuV. The PCR product was cloned into pET16b resulting in construct pCh1SdUV which incorporated an N-terminal deca-histidine tag on the chimeric protein.

4.2.6 In-gel Tryptic Digestion and Extraction of Expressed Proteins

Proteins from cell extracts were separated by SDS-PAGE and visualized by

Coomassie blue staining. Protein bands of interest were excised from the gel and cut into small pieces (1 x 1 mm) and placed in 1.5 ml Eppendorf tubes pre-rinsed with acetonitrile. Gel pieces were rinsed twice with 50% methanol for 10 min each. Solvent was removed and gel pieces were stored at -20°C until ready for peptide extraction.

To prepare for in-gel tryptic digestion, gel pieces were swollen in 50 μl 20 mM

NH4HCO3 for 10 min. Excess buffer was removed and the gel pieces were dehydrated in

50 μl acetonitrile for 10 min. Following removal of solvent the gel pieces were reswollen

159 in 100 mM NH4HCO3 with 10 mM DTT for reduction of excised protein. Nitrogen was blown into the tubes, sealed, and incubated for 1 h at 56°C. Samples were cooled back to

o 25 C and the reducing buffer was discarded. 50 μl of 100 mM NH4HCO3 with 55 mM iodoacetamide was added to alkylate reduced cysteines of excised protein. Nitrogen was blown into the tubes, sealed, and incubated for 45 min in the dark at room temperature.

Alkylating buffer was discarded and gel pieces were washed in 50 μl of 20 mM

NH4HCO3 for 10 min. Buffer was discarded and gel pieces were dehydrated in 100 μl acetonitrile for 10 min. Solvent was discarded and gel pieces were reswollen again in 20 mM NH4HCO3 for 10 min. Buffer was discarded and gel pieces were again dehydrated in 100 μl acetonitrile for 10 min. Solvent was removed and gel pieces were placed in a rotary speed vacuum for 15 min until completely dried. Gel pieces were then reswollen with 25 μl of 20 mM NH4HCO3 containing 10 ng/μl of sequencing grade trypsin

(Promega). Nitrogen was blown into the tubes, sealed, and incubated for 1 h on ice.

Excess trypsin digest buffer was removed and gel pieces were covered in 25 μl of 20 mM

NH4HCO3 and nitrogen was blown into the tubes and sealed. The remaining digestion was allowed to incubate overnight at 37°C.

For peptide extraction, the digestion reaction was briefly centrifuged and the supernatant was extracted and transferred to 1.5 ml Eppendorf tubes pre-rinsed with acetonitrile. Peptides were then extracted once with 25 μl of 20 mM NH4HCO3 and then

3x with 50 μl of 50% acetonitrile/5% formic acid. Each extraction step was performed for 20 min in a sonicating water bath and each extraction step was pooled with the first extracted supernatant. Extracted peptides were placed in a rotary speed vacuum until completely dry.

160

4.2.7 Peptide Mass Fingerprinting of Extracted Peptides by MALDI-MS

Dried peptide extracts were resuspended in 5 μl of a saturated solution of α- cyano-4-hydroxycinnamic acid (4-HCCA) in 50% acetonitrile and 0.1% trifluoroacetic acid. 2 μl of resuspended peptides were spotted on the MALDI target plate and air dried.

MALDI-MS data were collected on a Bruker Autoflex Speed MALDI-TOF/TOF mass spectrometer in reflectron positive mode. At least ten different shots were taken per spotted sample, with 1,000 pulses per shot, collected at a frequency of 200 Hz, and a laser power at 75%. Mass suppression was set to 450 Da and a mass range of 600-3,000 Da were collected. Calibration was performed using a ProteoMass Peptide MALDI-MS

Calibration Kit (Sigma). Collected data was analyzed using the Bruker Daltonics

FlexAnalysis 3.4 software package. Analyzed data was compared to predicted tryptic digest patterns based off of primary amino acid sequences that were generated by the program PeptideMass (http://www.expasy.org) (55). Parameters were set to allow for 3 maximum missed cleavages, setting all cysteine residues as carbamidomethyl-cysteine, and only selecting for monoisotopic masses ([M+H]+ ions).

4.2.8 Expression and Purification of ShuUV using the pCh1SdUV Construct

A single colony from freshly transformed E. coli BL21 (DE3) cells was used to inoculate 100 ml of TB media with added Amp (100 μg/ml). Cultures were incubated overnight at 37°C, 225 rpm. The following day, 10 ml of overnight subculture was used to inoculate 1 L of TB media (8 L total) with added Carb (100 μg/ml) and cultures were allowed incubate at 25°C, 180 rpm, for 36 h. Cultures were harvested at 6,000 rpm in a

Beckman JLA8.1 rotor.

161

Harvested cells were resuspended in 5 ml/g cell pellet in 50 mM Tris-HCl (pH

7.3) with 100 mM NaCl, 15% (w/v) glycerol, 10 mM benzamidine, and 2 EDTA-free protease inhibitor tablets (Roche Diagnostic). Lysozyme (20 μg/ml) and DNase (10

μg/ml) were added and cells were stirred at 4°C until completely resuspended, followed by cell lysis by French pressure treatment at 18,000 p.s.i. (2 passages). Cellular debris was pelleted at 12,000 rpm for 15 min in a Beckman JA-17 rotor and the supernatant was collected and further centrifuged 25,000 rpm for 1 h in a Beckman JA25.50 rotor to pellet cellular membranes. Membrane pellets were resuspended in 1 ml/g cell pellet of the above buffer and membranes were solubilized by addition of 4% (w/v) Triton X-100 and stirred at 4°C overnight. Membranes were pelleted again at 25,000 rpm for 1 h in a

Beckman JA25.50 rotor and the supernatant was saved. Membrane pellets were resuspended in 30 ml of the same buffer and solubilized a second time with 1% (w/v) lauryl-N,N-dimethylamine oxide (LDAO). Membranes were stirred at room temperature for 1 h and again centrifuged at 25,000 rpm for 1 h in a Beckman JA25.50 rotor. The supernatant was combined with the first solubilization step and loaded onto a Ni-NTA metal affinity column (2.5 x 3.0 cm) equilibrated in 50 mM Tris-HCl (pH 7.3), 100mM

NaCl, 5mM imidazole, 15% (w/v) glycerol, and 0.07% LDAO. The column was then washed with 10 column volumes (CV) of the same buffer followed by 3 CV of the same buffer containing 20 mM imidazole. ShuUV was eluted in ~5 ml fractions of the same buffer containing 250 mM imidazole. All fractions were analyzed by 14% SDS-PAGE.

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4.2.9 Expression and Purification of ShuT

ShuT was expressed from the plasmid pETShuT as previously described with minor modifications (3). Briefly, a single colony of freshly transformed BL21 (DE3) pLysS cells was used to inoculate a 50 ml subculture of LB with Amp (100 μg/mL) and grown overnight at 37°C, 225 rpm. 10 ml of overnight subculture was used to inoculate

1 L fresh cultures of LB with Amp (100 μg/ml) and grown to an OD600 of ~0.7 at 37°C,

225 rpm. Cultures were then induced with 1 mM IPTG and allowed to grow an additional 3 h at 25°C, 225 rpm. Cultures were harvested by centrifugation at 6,000 rpm in a Beckman JLA8.1 rotor.

Harvested cells were resuspended in 4 ml/g cell pellet in 50 mM Tris-HCl (pH

8.0) with 10 mM benzamidine, 1 mM MgCl2, and an added EDTA-free protease inhibitor tablet (Roche Diagnostic). Lysozyme (20 μg/ml) and DNase (10 μg/ml) were added and cells were stirred at 4°C until completely resuspended and lysed by sonication. Lysed cells were centrifuged at 20,000 rpm for 45 min in a Beckman JA25.50 rotor. The supernatant containing soluble periplasmic fractions was loaded onto a Ni-NTA agarose column (1 x 10 cm) equilibrated in 50 mM Tris-HCl (pH 7.8), 300 mM NaCl, and 5 mM imidazole. The column was washed with 10 column volumes (CV) of the same buffer.

The column was then washed with 10 CV of the same buffer containing 20 mM imidazole. ShuT was eluted in 50 mM Tris-HCl (pH 7.8) and 250mM imidazole. Peak fractions were determined by SDS-PAGE and dialyzed against 20 mM Tris-HCl (pH

7.8). The protein was further purified over a Q-sepharose anion exchange column (1 x 10 cm) equilibrated in 20 mM Tris-HCl (pH 7.8). The column was washed with 10 CV of the same buffer and ShuT was eluted using a linear salt gradient from 0-200 mM NaCl.

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Purified protein was concentrated to 10 mg/mL and stored at -80°C until further use.

Protein concentrations were determined using the extinction coefficient value of 30.71

-1 -1 mM cm at 280 nm (ε280) as previously determined (3).

4.2.10 Heme Reconstitution of ShuT

Heme stock solutions were freshly prepared as described above. Fully reconstituted holo-ShuT was prepared by titration of heme stocks to purified apo-ShuT until the heme:protein ratio reached 2:1. Excess heme was removed by passage over a

Sephacryl S-100 column (1 x 100 cm) equilibrated in 20 mM Tris-HCl (pH 7.5). Heme loading of ShuT was checked by UV-visible absorption spectroscopy. Heme concentration was determined using the extinction coefficient value of 102.70 mM-1cm-1 at 400 nm (ε400) for holo-ShuT as previously determined (3).

4.2.11 Expression and Purification of ShuS

ShuS was expressed from plasmid pEShuS as previously described with minor modifications (24). Briefly, a single colony of freshly transformed BL21 (DE3) cells was used to inoculate a 50 ml subculture of LB with Amp (100 μg/ml) and grown overnight at

37°C, 225 rpm. 10 ml of overnight subculture was used to inoculate 1 L fresh cultures of

LB with Amp (100 μg/ml) and grown to an OD600 of ~0.7 at 37°C, 225 rpm. Cultures were then induced with 1 mM IPTG and allowed to grow an additional 3 h at 25°C, 225 rpm. Cultures were harvested by centrifugation at 6,000 rpm in a Beckman JLA8.1 rotor.

Cells were resuspended in 4 ml/g cell pellet 50 mM Tris-HCl (pH 8.0), 10 mM benzamidine, 1 mM EDTA, 1 mM MgCl2, and an added protease inhibitor tablet (Roche

164

Diagnostic). Lysozyme (20 μg/ml) and DNase (10 μg/ml) were added and cells were stirred at 4°C until completely resuspended and lysed by sonication. Lysed cells were centrifuged 20,000 rpm for 45 min in a Beckman JA25.50 rotor. The supernatant containing soluble cytoplasmic proteins was loaded onto a Q-sepharose anion exchange column (3 x 15 cm) equilibrated in 20 mM Tris-HCl (pH 8.0) and washed with 10 CV of the same buffer containing 50 mM NaCl. ShuS was eluted using a linear salt gradient of

50-400 mM NaCl. Peak fractions of ShuS as assessed by SDS-PAGE were applied to a

Bio-Gel HTP column (1.5 x 10 cm) equilibrated in 10 mM potassium phosphate (pH 7.4), and washed with 10 CV of the same buffer. ShuS was eluted with a linear gradient of 10-

50 mM potassium phosphate, pH 7.4. Purified ShuS was concentrated to 10 mg/ml and stored at -80°C until further use. Protein concentrations were determined using the

-1 -1 predicted extinction coefficient value of 53.19 mM cm at 280 nm (ε280) determined from the program ProtParam (http://www.expasy.org) (56).

4.2.12 Preparation of ShuUV Reconstituted in Proteoliposomes

100 mg of E. coli total lipid extracts and 33 mg of L-α-lysophosphatidylcholine was weighed and mixed to 20 mg/ml in 50 mM Tris-HCl (pH 7.3) in a glass vial.

Nitrogen was blown into the vial and then sealed and allowed to incubate at 42°C for 3 h with occasional mixing to hydrate the lipid mixture. To ensure complete resuspension, lipids were briefly sonicated on ice. Liposomes were then subjected to 3 freeze-thaw cycles in a dry ice/ethanol bath and homogenous lipid micelles were produced by extruding the resuspended lipids using a Mini Extruder (Avanti Polar Lipids) through a

400 nm polycarbonate membrane. Purified ShuUV was added to the liposome mixture at

165 a ratio of 50:1 (lipid:protein) and diluted with 50 mM Tris-HCl (pH 7.3) and 0.14% (w/v)

Triton X-100 to a final lipid concentration of 4 mg/ml. The mixture was mixed on a rotary platform shaker and gently rocked at room temperature for 30 min. 40 mg/ml (wet weight) of BioBeads SM2 (BioRad) were added to the mixture to remove detergent and allowed to rock at room temperature for 30 min. Similar aliquots were added 4 more times and incubated at 4°C for 30 min each, with the last addition being incubated overnight. BioBeads SM2 were removed by filtration through a disposable spin column.

The solution was diluted ~5x with 50 mM Tris-HCl (pH 7.5), and centrifuged 25,000 rpm for 1 h in a Beckman JA25.50 rotor. Proteoliposomes were washed in the same buffer and pelleted again as before. Proteoliposomes were then resuspended to 20 mg/ml lipids in 50 mM Tris-HCl (pH 7.5) and 150 mM NaCl, flash frozen, and stored at -80°C.

4.2.13 Reconstitution of ShuT into Proteoliposomes

Proteoliposomes containing reconstituted ShuUV were thawed on ice. To reconstitute ShuT in the lumen of proteoliposomes, lipids (1.7 mg/ml) and ShuT (50 μM) were mixed in reaction buffer (50 mM Tris-HCl (pH 7.5) and 150 mM NaCl). The mixture was subjected to 3 rounds of freeze-thaw cycles in a dry ice/ethanol bath followed by extrusion through a 400 nm polycarbonate membrane. Proteoliposomes were pelleted at 25,000 rpm for 1 h in a Beckman JA25.50 rotor. The supernatant was removed and pelleted proteoliposomes were washed again with reaction buffer followed by resuspension of lipids to 1.7 mg/ml.

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4.2.14 ATPase and Heme Transport Assays

To initiate in vitro functional transport assays, 50 μM apo-ShuS was added to 500

μl aliquots of the resuspended proteoliposomes and incubated at 37°C for 5 min prior to initiation of the transport cycle. ATP hydrolysis and heme transport was initiated by the addition of 2 mM ATP and 10 mM MgCl2 to fully reconstituted liposomes and 50 μl aliquots were taken at several time points and added directly to 50 μl of a 12% (w/v) sodium dodecyl sulfate (SDS) stop solution. To determine the ATPase activity of ShuUV under various conditions of transport, the levels of inorganic phosphate generated from the ATPase reaction was determined using a NaH2PO4 calibration curve generated by the modified molybdate method (57). Each measurement was done in triplicate and averaged. Heme transport was monitored by centrifugation of proteoliposome pellets and analysis of the supernatant containing heme bound ShuS by UV-visible electronic absorption spectroscopy. Heme concentrations were quantified by the pyridine hemochrome assay (52).

4.3 Results

4.3.1 Plasmid Construction and Test Expressions of ShuUV Constructs

The expression and purification of ShuUV has been previously reported by our laboratory, however, the expression levels were extremely low and not sufficient to conduct spectroscopic experiments (38). Therefore, design and generation of alternate expression constructs was performed to improve ShuUV protein expression levels. A summary of generated constructs can be found in Table 4.1 along with a description of the observed expression profiles. The original construct of ShuUV incorporated an N-

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Plasmid N-terminus Description pET22bShuUV PelB-MGLSLLLL Original construct (38) pShuUV MGLSLLLL PelB leader removed from original construct. 6x His-tag C-term ShuV. ShuV expression only. pSdUV MLKDSFSS Incorporation of extra 12 N-terminal amino acids. 6x His-tag C-term ShuV. ShuV expression only pSdUVPelB PelB-MLKDSFSS Incorporation of PelB to pSdUV. No expression. pShuU MLKDSFSS Clone of only shuU from pSdUV. 6x His-tag C-term ShuU. No expression. pShuV N/A Clone of only shuV from pSdUV. ShuV expression. pShuUShuV MLKDSFSS ShuU and ShuV expressed from separate promoters on same pETDuet plasmid. No expression. pChuUV MLKDSFSS Complete clone of chuUV from E. coli. 6x His- tag C-term HmuV. HmuV expression only. pCh1SdUV BtuC α-helix 1 Chimeric ShuU with first α-helix swapped with BtuC. 10x His-tag on N-term ShuU. ShuUV expression.

Table 4.1 List of generated ShuUV expression constructs and description of observed expression profiles.

168 terminal PelB leader sequence for targeting of proteins to the periplasm in E. coli. It was assumed that keeping a PelB leader sequence for targeting to the periplasm would result in the transporter being targeted to the cytoplasmic membrane. This seemed unnecessary as the native proteins do not require nor contain any sorting signal for specific membrane targeting. Removal of the PelB leader sequence was the first step in optimizing the expression construct as a means of increasing the yields of ShuUV. However, examination of the expression profiles resulted in no protein expression. Since E. coli and S. dysenteriae have a highly conserved genetic code, the sequence of ShuUV deposited in the protein database was compared with that of E. coli ChuUV to optimize the sequence for expression in E. coli. Not surprisingly, the sequence identities between

ShuUV and ChuUV were >99%, with only two amino acid substitutions between the two proteins. However, the E. coli ChuU sequence contained an extra 12 amino acids at the

N-terminus not found in the original predicted ShuU sequence (Figure 4.4A). Further examination of the upstream sequence of the shuUV genes revealed this sequence was also conserved in ShuU along with a strong ribosomal binding site (RBS) located directly upstream at the -10 position of the alternate start codon, suggesting that the start codon was erroneously assigned for shuU in the database (Figure 4.4B). Primers were made to re-clone the shuUV genes from genomic DNA to incorporate this extra stretch of 12 amino acids. Expression analysis of this construct revealed excellent over-expression of a single band at ~30 kDa, but did not reveal any indication that both subunits were being expressed (Figure 4.4C). As both ShuU (35.4 kDa) and ShuV (29.9 kDa) are close in relative size it was difficult to determine which protein was over-expressed based on

169

Figure 4.4 Sequence analysis and expression profile of predicted ShuUV. A. Amino acid sequence alignment of the N-terminal residues of ChuU (E. coli) and ShuU (S. dysenteria) revealed an extra 12 amino acids not encoded for ShuU (top). Examination of regions upstream of predicted start site revealed amino acids were present and conserved (bottom). B. DNA and amino acid sequences upstream of the shuU predicted translation initiation site. Predicted translation initiation site for shuU (green) and chuU (blue). Potential -10 ribosomal binding sites for each translation initiation site in red. C. 14% SDS-PAGE of ShuUV test expression from expression construct pSdUV (right) which incorporated the extra 12 amino acid residues predicted in the ChuU sequence compared with empty control vector pET22b (left). M, molecular weight markers; 1, wash; 2, elution from Ni-NTA resin.

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SDS-PAGE alone. In-gel tryptic digests of the protein followed by MALDI-MS analysis revealed a mass fingerprint corresponding to ShuV (Table 4.2). The masses identified covered 82.3% of the predicted 81.4% total sequence coverage available (Figure 4.5).

All constructs tested failed to yield over-expression of both ShuU and ShuV.

Since ShuU and ShuV are expressed from a single transcript and are genetically linked by an overlapping stop (ShuU) and start (ShuV) codon, the appearance of ShuV in expression screens indicated that the complete mRNA transcript was being transcribed.

However, the absence of ShuU suggested instability in the 5’ end of the mRNA transcript that prevented translation initiation or alternatively, instability of translated ShuU protein that was subsequently being targeted for proteolysis. Separate constructs of ShuU and

ShuV were constructed in an attempt to express the proteins separately. ShuV expressed and folded in the absence of ShuU, whereas the ShuU construct did not show any significant protein levels. A similar observation was reported with the E. coli vitamin B12

ABC transporter BtuCD, in which co-translation of both proteins was required for BtuC expression, implying cooperative folding between subunits (58). This was also confirmed when trying to express ShuU and ShuV from separate promoters on a single plasmid.

It was recently reported that the 5’ untranslated regions of mRNA transcripts encoding iron-regulated genes, including the OM heme receptor shuA, contained a post- transcriptional thermoregulator (59). At 25°C, the mRNA transcript formed a stable stem-loop structure that occludes the Shine-Dalgarno (SD) sequence, effectively prohibiting translation initiation. However, at 37°C the stem-loop structure destabilizes, allowing access of RNA polymerase to the SD sequence. To test the possibility that

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Expected m/z Observed m/z #MC Position Peptide Sequence 2603.31 2603.35 0 158-178 WLFLDEPTSALDIHHQQHLFR 2596.41 2596.36 1 15-40 RLTDNVSLTFPGGEIVAILGP NGAGK 2594.14 2594.38 0 105-127 TGNQDNETAQIMALCDCQAL ANR 2206.05 2206.03 0 82-100 QNSHMAFPFSVQEVIQMGR 1929.94 1929.79 0 188-203 QFNVCCVLHDLNLAAR 1903.03 1903.86 0 60-75 LFNKPLNEWSITELAK 1623.74 1623.87 0 46-59 QLTGLQPDSGECR 1579.83 1579.93 0 1-14 MISAQNLVYSLQGR 1507.85 1507.96 0 145-157 LLVQLWEPTPSPK 1002.49 1002.48 0 131-139 QLSGGEQQR 781.43 781.48 0 182-187 QLVHER

Table 4.2 Peptides identified by in-gel tryptic digestion and mass fingerprint analysis by MALDI-MS. For peptide mass predictions, the primary ShuV amino acid sequence was input into the program ProteoMass (55). Predicted peptides were generated using trypsin, allowing for up to 3 missed cleavages (#MC), all cysteine forming carbamidomethyl- cysteine, and selecting for monoisotopic masses only [M+H]+. Mass filter from 500- 3,000 Da was selected.

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Figure 4.5 Sequence coverage of ShuV from MALDI-MS mass fingerprinting analysis of tryptic digests. A. Primary sequence coverage identified by MALDI-MS finger printing. Underlined capital letters indicate residues predicted to be identified by MALDI-MS using the constraints detailed in Methods and Materials. Lower case letters indicate residues not predicted to be identified. Highlighted (cyan) residues indicate actual sequence coverage observed by MALDI-MS analysis. B. ShuV model highlighting peptide sequence coverage observed (cyan) and missing (orange) from the predicted sequence coverage. Residues not predicted or observed (grey) are indicated as well. ShuV modeled from PDB file 2QI9.

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ShuUV was also under similar thermoregulation expression cultures were grown at a constant 37°C, however, production of both protein subunits was still unsuccessful.

The E. coli vitamin B12 transporter BtuCD has been successfully expressed and purified to sufficient levels for both structural and functional characterization (39, 47).

As a way to overcome the issues with the possible 5’ instability of mRNA or N-terminal instability of the protein, a chimeric construct of ShuUV was designed. This construct removed the first predicted N-terminal α-helix of ShuU and replaced it with the first α- helix of BtuC. The BtuCD protein was also designed with an N-terminal deca-histidine tag on BtuC, a strategy also incorporated into the chimeric ShuUV construct. Subsequent expression studies and isolation of membranes for protein analysis revealed the presence of a bright pink color that had not been previously seen, possibly indicating residual heme was associated with the membrane fractions. Detergent extraction and purification over Ni-NTA metal affinity column revealed the presence of two unique bands that were not present in empty vector controls, as assessed by SDS-PAGE (Figure 4.6A). Further, absorption spectroscopy of the partially purified protein following dialysis revealed a distinct spectrum with a heme Soret at 414 nm and visible bands at 532 and 561 nm

(Figure 4.6B). This spectrum is reminiscent of the bis-His coordinated hemes from the

TonB dependent OM transporters HasR and the Y519H PhuR mutant previously reported in Chapter 2. Additionally, it was shown two conserved histidines (His-252 and His-262) within the translocation channel were critical for heme transport, although no direct evidence has been reported yet as to whether they coordinate heme during the transport cycle (38). Reconstitution into liposomes and SDS-PAGE analysis revealed the protein could be carried over into proteoliposome preparations (Figure 4.6C).

174

Figure 4.6 Analysis of the expression construct pCh1SdUV. A. 14% SDS-PAGE of test expressions of pCh1SdUV in BL21 (DE3) cells grown for 36 h at 25°C in TB media. M, molecular weight markers; 1, control vector pET22b; 2, pCh1SdUV. B. Absorption spectra of chimeric ShuUV after Ni-NTA purification. C. 14% SDS-PAGE of chimeric ShuUV after reconstitution into liposomes. M, molecular weight marker; 1, 10 μg load of total protein; 2, 20 μg load of total protein.

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4.3.2 Functional Assay of Chimeric ShuUV

The reconstituted chimeric ShuUV protein into proteoliposomes was then assessed for its ability to couple the hydrolysis of ATP to substrate translocation. In this proteoliposome system, ShuUV is incorporated into the lipid micelle bilayer that has holo-ShuT inside the lumen, while apo-ShuS is added to the external medium after complete reconstitution to accept heme as it is transported outside of the lumen (Figure

4.7A). Addition of ATP to the resuspended proteoliposomes could then initiate the heme transport cycle. A stimulation in ATP hydrolysis could then be monitored by the appearance of inorganic phosphate and measured by a modified molybdate assay while heme translocation could be monitored by the pyridine hemochrome assay. As seen with other reconstituted ABC transporters, there was a low basal level of ATP hydrolysis with

ShuUV alone in lipids (40, 43, 47). However, upon incorporation of holo-ShuT inside the lumen, a sustained increase in ATP-hydrolysis accompanying heme translocation was observed (Figure 4.7B). Addition of apo-ShuS to the buffer only slightly increased the rate of ATP hydrolysis, suggesting ShuS is not required to stimulate heme transport.

4.4 Discussion

Many cytoplasmic membrane proteins contain a series of positively charged residues at the N-terminus. The presence of these charged residues are thought to serve as an anchor to the negatively charged polar head groups of the phospholipids in the cytoplasmic membrane. Interestingly, the BtuC TMD contains such a series of charged residues but this sequence is absent in ShuU, possibly providing an explanation why

ShuU was not expressed in previous expression trials or constructs lacking this sequence.

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Figure 4.7 Functional assay for in vitro heme transport in proteoliposomes. A. ShuUV (blue and green) is incorporated into liposomes with holo-ShuT (purple) in the lumen. Addition of ATP, Mg2+, and/or ShuS to the extracellular buffer begins the ATP dependent transport of heme across the lipid bilayer. B. In vitro functional assay showing sustained increase in ATP hydrolysis upon addition of ATP and Mg2+ in the presence or absence of ShuS (red and purple traces, respectively). Basal levels of ATP hydrolysis are elevated with ShuUV in proteoliposomes alone (green trace) or when ShuS is additionally added to the extracellular buffer (blue trace) compared to empty liposomes (black trace).

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The absence of these positive residues to anchor ShuU in the membrane could mean that insertion of ShuU in native S. dysenteriae membranes may require an additional chaperone protein that it must associate with in the membrane. By inserting an N- terminal deca-histidine tag, as well as utilizing the first α-helix of BtuC which has a sequence of 8 positively charged residues (RQQQRQNIR) resulted in the creation of a functionally stable protein targeted to the cytoplasmic membrane. Prior to the construction of the chimeric protein the possibility of a thermoregulated mRNA transcript was examined. However, this possibility seemed unlikely during recombinant expression, as the promoter and RBS are derived from an expression plasmid. As expected, induction of protein expression was not dependent upon the temperature.

However, as stated previously, shuA was identified as an iron-regulated gene involving the 5’ untranslated region as being under thermoregulatory control (59). A string of 4 T nucleotides were located 8 base pairs upstream of the SD region that could form a potential stem loop structure. shuA is in the same operon as the other genes encoding heme uptake (shuTWXYUV) but is divergently transcribed. Examination of the upstream region of the identified SD region in shuU revealed a string of 4 T nucleotides about 8 base pairs away, indicating in vivo, under its native promoter, protein expression from these transcripts may also be thermally regulated (Figure 4.3B).

Reconstitution of the complete chimeric ShuUV proteoliposomes revealed that replacement of the first α-helix did not affect its function. ShuUV exhibited a basal level of ATP-hydrolysis that could be stimulated in the presence of holo-ShuT. Surprisingly, this was the minimal requirement for sustained increase in ATPase activity, contrary to the previous report that a cytoplasmic heme binding protein was also required (38). This

178 could possibly be explained by the replacement of the first α-helix, which is slightly longer in the BtuCD crystal structure and is responsible for making contacts with the

NBDs, although minimal, which could possibly alter ATP hydrolysis and transport of substrate (39). Indeed in the homologous Y. pestis HmuUV transporter, the native helix is slightly shorter and does not appear to make contact with the NBD, and is therefore unable to directly influence interaction with the cytoplasmic binding protein (Figure 4.8)

(40). In keeping with our current data the HmuUV transporter was fully functional in the absence of the cytoplasmic heme binding protein (40).

In the current studies I have optimized the expression and purification of a fully functional ABC transport system from S. dysenteriae. This system will provide the basis for future site-directed mutagenesis and resonance Raman spectroscopic studies aimed at probing the mechanistic details of heme translocation from ShuT, through ShuUV, and finally to ShuS. Through a series of site-directed mutants of ShuUV in conjunction with functional assays utilizing the in vitro proteoliposome system, the identification of residues critical for heme transport and function will be identified. These mutants will be further utilized in future spectroscopic studies aimed at capturing intermediates along the heme translocation pathway that will be paramount in further elucidating the mechanistic details of heme transport across biological membranes.

179

Figure 4.8 Comparison of α-helix 1 from the ABC transporters involved in heme and vitamin B12 uptake. Both models show one HmuUV (left) and one BtuCD (right) heterodimer subunit. The first α-helix from E. coli BtuC (orange) extends further into the cytoplasm and forms minimal contacts with the ATPase subunit BtuD. This is in contrast to the first α-helix of Y. pestis HmuU (dark blue), providing a possible explanation for N- terminal instability seen in the ABC transporters involved in heme uptake and possible mechanistic differences seen between the two transporters. Adapted from PDB files 4G1U and 2QI9.

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188

Chapter 5

Summary and Future Directions

5.1 Summary

Pathogens are highly dependent upon their ability to acquire iron for survival and virulence. Upon infection, the host’s innate immune response deploys active countermeasures that diminish and restrict accessible iron to the pathogen. In response, pathogenic bacteria have evolved sophisticated mechanisms to acquire this scarce micronutrient, including TonB-dependent receptors capable of acquiring iron in the form of heme from host heme containing proteins. Despite recent advances in the identification and characterization of these systems from genetic and structural studies, much of the mechanistic details on heme transport through biological membranes remains to be elucidated. The focus of the current research presented herein initially focused on developing and optimizing expression and purification systems for the outer membrane and cytoplasmic membrane transporters, as a prerequisite for spectroscopic and structural characterization. In addition to the initial biochemical characterization of the systems, we have further developed bacterial genetic and isotopic labeling methodologies to dissect in vivo the mechanism of heme uptake across the OM, and the

189 contributions of the hemophore (Has) and non-hemophore (Phu) dependent systems of

Pseudomonas aeruginosa. Additionally, a periplasmic ABC transport system from

Shigella dysenteriae was optimized for utilization in future studies that will be directed at elucidating mechanistic steps involved in heme transport across the cytoplasmic membrane.

In Chapter 2, I conducted the initial expression and purification studies of the hemophore dependent OM receptor HasR and the host heme protein receptor PhuR of P. aeruginosa. Initial sequence analysis and spectroscopic characterization of the HasR receptor identified conserved His-189 on the N-terminal plug domain and His-624 on the

FRAP-loop are responsible for heme coordination. As isolated HasR displayed an absorption spectra typical of the previously characterized bis-His receptors (1, 2), with a heme Soret at 411nm and bands in the visible region at 535 and 561 nm. In contrast, sequence analysis of PhuR revealed the conserved histidine residue in the FRAP-loop was absent. PhuR on purification and spectroscopic analysis gave a spectrum distinct from the typical bis-His coordinated receptors. Instead, the spectra more resembled that of the high affinity hemophores, with a heme Soret at 405 nm and visible bands at 530 and 609 nm.

Site-directed mutagenesis combined with absorption and magnetic circular dichroism spectroscopy confirmed PhuR coordinates heme through His-124 on the N-terminal plug and Tyr-519 on the extracellular FRAP-loop, a ligand set not seen in any OM receptors to date. However, a recent report by Mokry et al, identified a five-coordinate Tyr complex in the isolated HmbR receptor of N. meningitidis (3). Interestingly, Tyr ligation has emerged as a common motif across a spectrum of high affinity heme acquisition proteins and systems. These include the lipid anchored surface exposed Isd heme binding proteins of

190

Gram-positive pathogens (4-6), and the soluble periplasmic binding proteins in Gram- negatives (7, 8). The implications of the emergence of these alternate heme coordination profiles may reflect differences in heme affinity or capacity of the receptor that provides an advantage within the ecological niche occupied by these pathogens.

Neither HasR nor PhuR were competent in vitro to extract heme from HasA or metHb, respectively. Therefore, I could not directly compare the thermodynamic and kinetic parameters of heme binding to the isolated receptors PhuR and HasR as a measure of their potential contribution to heme acquisition. I confirmed the inability to extract heme from HasA or metHb was not due to diminished structural integrity of the receptors as judged by the CD spectra. Additionally, incubation of HasR with HasA did not result in the formation of a stable protein-protein complex as reported for the S. marcescens HasR receptor (2). I cannot reconcile these results with those of the previously characterized S. marcescens HasA-HasR receptor system but the present data is consistent with previous reports on the OM heme receptors ShuA and HmbR from S. dysenteriae and N. meningitides, respectively (1, 3). My current data is further complicated by the fact that previous in vivo reports on the S. marcescens system indicated heme release from HasA to

HasR required interaction with the TonB-dependent energy transducing system (9). My studies are consistent with the latter observation and the recent characterization of the purified N. meningtidis HmbR receptor (3), that the OM receptors require the TonB- dependent energy transducing system for heme extraction.

However, as I was unable to determine meaningful thermodynamic and kinetic parameters for heme binding to the detergent solubilized receptors in vitro, I addressed the contributions of the PhuR and HasR receptors to heme acquisition by in vivo methods

191 as outlined in Chapter 3. Deletion of hasR resulted in a less than 2-fold up regulation of

PhuR and did not affect the cells ability to acquire extracellular heme. The hasR mutant was unable to down regulate the flux of heme into the cell with the increase in steady state levels of the intracellular heme utilization proteins accounting for the “shunting” of intracellular biosynthesized heme through HemO. The inability to regulate PhuS and

HemO is not Fur-dependent as in the presence of iron the steady state mRNA levels of phuR, phuS and hemO were down regulated to the same degree as the wild type strain.

The HasR receptor in P. aeruginosa is similar to that previously described for S. marcescens functions as a signal transduction receptor through the extra cytoplasmic function (ECF) sigma and anti-sigma factors HasI and HasS, respectively (10). In S. marcescens, binding of the holo-HasA to HasR inactivates the anti-sigma factor HasS and activates the sigma factor HasI allowing transcription of the hasAR and hasSI operons.

Therefore, accumulation of inactive HasS can rapidly decrease HasR expression in response to the fluctuating extra cellular heme concentrations. Therefore, as heme acquired through HasR requires the periplasmic and cytoplasmic components of the Phu uptake system, “cross-talk” between the hemophore signaling cascade and the phuSTUV genes may account for the elevated levels on loss of HasR.

Deletion of phuR resulted in a significant up regulation of HasR to compensate for loss of PhuR, however, despite the up regulation and increase in steady state levels of hasR a significant increase in intracellular 12C-heme metabolism was observed. The increased steady state levels of the PhuS and HemO proteins coupled with the decreased efficiency of the HasR receptor accounts for this decreased efficiency in extracellular heme uptake.

The increase in PhuS in contrast to that observed in the ΔhasR deletion was due to

192 disruption in Fur-regulated repression of phuS in the presence of iron. Our laboratory has observed a similar disruption in Fur-regulation of phuR on deletion of phuS as a result of upstream regulatory factors that are currently being determined (Wilks et al, unpublished data). I also observed that increased PhuS protein levels in both the ΔhasR and ΔphuR strains appeared to be coupled to increased steady state HemO levels. This regulatory link between phuS and hemO is heme dependent as in presence of iron hemO transcription was suppressed to the same degree as in the wild type strain. We are currently investigating the heme dependent regulatory link between the phu and hemO operons.

Taken together the present data confirms that both the Has and Phu systems are required for the regulation and efficient utilization of heme in P. aeruginosa. Based on the present data I propose a model where the HasA-HasR system has a lower capacity for heme uptake but acts as a sensor of extracellular heme (Figure 3.7). Activation of the HasA-

HasR signaling cascade leads to up regulation of the high capacity Phu system, the major facilitator of extracellular heme acquisition. The signaling cascade also positively regulates the levels of the anti-sigma factor HasS, which accumulates in an inactive state.

As extracellular heme levels decrease, HasS inactivates the sigma-factor HasI and down regulates the heme acquisition systems. This fine-tuning allows the cell to rapidly respond to external heme levels in a Fur-independent manner. Furthermore, support for PhuR being the primary receptor for heme acquisition in chronic infection comes from recent studies where mutations within the phuR promoter in P. aeruginosa clinical isolates leading to increased PhuR expression conferred a growth advantage in the presence of hemoglobin.

Additionally, this evolution within the host toward increased reliance on heme via up regulation of PhuR coincides with loss of alternate iron-scavenging systems (11). In acute

193 infection, the HasA-HasR system may play a more significant role, however, on chronic infection the bacteria adapt to utilize heme as the primary iron source through the high capacity Phu system.

Finally, in Chapter 4 the expression and purification of the ABC transporter ShuUV was optimized and enhanced to conduct future experiments aimed at elucidating a detailed mechanism of heme transport into the cytoplasm. Through testing a series of ShuUV expression constructs, it became evident the N-terminal stability of ShuU upon translation initiation greatly dictated the success of generating a stably folded protein. By replacing the N-terminal α-helix of ShuU with the first α-helix of BtuC, a chimeric ShuUV protein was stably expressed and successfully incorporated into bacterial membranes. Purification of the transporter and inspection by absorption spectroscopy revealed the presence of residual heme bound to the receptor. Reconstitution of ShuUV in proteoliposomes with holo-ShuT resulted in a functional ABC transport system upon the addition of ATP as an energy source, as evidenced by a sustained increase in the evolution of inorganic phosphate. Contrary to the previous ShuUV report (12), the chimeric construct generated in these studies only minimally required holo-ShuT and the ABC-transporter ShuUV to stimulate ATPase activity in proteoliposomes. Addition of ShuS had negligible impact on the overall rate of ATP hydrolysis when holo-ShuT was also incorporated inside the lumen, however, it remains to be seen if heme release from the transporter is prevented in the absence of ShuS.

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5.2 Future Directions

The 13C-heme metabolism studies represent a powerful tool for evaluation of the in vivo efficiency of heme uptake in light of the difficulties associated with examining heme uptake in vitro, as evidenced in Chapter 2. Reincorporation of non-functional mutant variants of PhuR or HasR into the chromosome of the PAO1 ΔphuRΔhasR deletion strain will further elucidate the relative contributions of the receptors towards heme uptake and will provide insight into the relative affinities of the His-Tyr motif in

PhuR. In PhuR, complementation with the H124A, Y519A, and Y519H variants and monitoring heme utilization by the 13C-heme metabolism studies will allow for direct comparison between the His-Tyr and bis-His ligand motif in heme acquisition.

These studies also allow for elucidation of the cell surface signaling system of

HasR in sensing and regulating extracellular heme uptake. Generation of the H189A and

H624A HasR variant could result in a receptor non-functional for heme transport, yet may still be able to bind its substrate holo-HasA, inducing the signaling cascade.

Subsequent metabolism studies in conjunction with qRT-PCR analysis would confirm

HasR’s role primarily as an extracellular heme sensor. Additionally, the contribution of

HasR to heme uptake in the absence of PhuR will be further tested by supplementing cells with 13C-heme loaded holo-HasA. This may also give insight into the observed switch in BVIXβ:BVIXδ ratios when heme is solely funneled through HasR. HasR mutants targeting the N-terminal signaling domain can also be constructed to evaluate heme uptake efficiency in the absence of the cell surface signal cascade. The above studies will provide further insight into cell surface signaling, regulation, and, contributions of the P. aeruginosa Phu and Has systems.

195

Finally, in Chapter 4, optimization of the ShuUV system from S. dysenteriae was paramount for developing future assays aimed at probing the mechanistic details of heme transport across the cytoplasmic membrane. The transporter was able to be expressed and purified at levels required for functional reconstitution in a proteoliposome system.

The current over-expression construct is a chimeric protein containing partial sequence of the vitamin B12 transporter BtuC which has an extended N-terminus. While studies will continue utilizing this construct, further optimization of the expression construct can be performed to obtain a more native-like ShuUV transporter. Specifically, cloning of the shuUV gene encoding the extra N-terminal 12 amino acids and incorporation of the N- terminal 10x His-tag on ShuU may enhance protein expression with native ShuUV. This is currently being tested in the laboratory.

While many functional studies have been performed with the vitamin B12 transporter BtuCD and heme transporters ShuUV and HmuUV, there is still a gap in the information regarding the mechanistic details of how heme is transported through the membrane (12-14). By generation of a series of site-directed mutants and utilizing the in vitro proteoliposome system, residues critical for steps along the translocation of heme into the cytoplasm can be identified. These include residues in the Walker A loop involved in ATP binding, the Walker B motif involved in binding a catalytic Mg2+ ion, and the invariant glutamine critical for ATP hydrolysis. Additionally, using non- hydrolyzable ATP analogues, such as sodium orthovanadate, we can selectively trap intermediates in the steps up to binding of ATP and before ATP hydrolysis. The previously identified histidine residues (His-152 and His-162) critical for heme transport will also be utilized in attempts of trapping stable intermediates (12). By generating a

196 series of mutants from identified residues critical for heme transport and ATPase activity, we will be able to trap heme intermediates along the translocation pathway that can be further investigated by spectroscopic methods. In this way we can begin to develop a detailed mechanism of heme transport through the periplasmic ABC transport system.

5.3 Conclusions

The rise in cases of multi-drug resistant bacterial infections has become a global health concern, placing severe economical burdens on the current healthcare systems

(15). This problem is compounded with the alarmingly fast rate of emerging cases of antimicrobial resistance and the lack of new therapeutic targets and drugs reaching the market. The need for novel therapeutic treatments is more evident than it has ever been.

Recently, a shift in the conventional wisdom of developing antibiotics that target functions essential for the pathogens survival, to functions essential for virulence, provides a means to combat bacterial infection, while reducing the propensity for antibiotic resistance by placing less selective pressure on the bacteria to mutate.

Targeting systems essential for virulence would then provide the host’s immune system the opportunity to effectively reduce bacterial load and perhaps clear the infection. As iron is a central metabolic requirement and is also linked to virulence, alteration in the metabolic competence of the bacteria represents a novel approach. Heme represents one of the most abundant and readily available sources of iron, and has been shown to be the optimal source in P. aeruginosa. Therefore, targeting heme uptake and utilization may provide a means to curb virulence associated with these pathogens and reduce their propensity to establish an infection. Advancing our knowledge on the mechanistic details

197 involved in heme transport and utilization are thus fundamental for the development of novel antibiotics targeted to these systems.

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