interactions of l. monocytogenes with host cellular defenses

Interactions of L. monocytogenes with Host Cellular

Defenses

by

Grace Ying-Tung Lam

A thesis submitted in conformity with the requirements

for the degree of Doctor of Philosophy

Institute of Medical Sciences

University of Toronto

© Copyright by Grace Lam 2012

ii

Interactions of L. monocytogenes with Host Cellular

Defenses

Grace Lam

Doctor of Philosophy

Institute of Medical Sciences

University of Toronto

2012

Abstract

Listeria monocytogenes is an intracellular bacterium that utilizes two phospholipases C (PLCs)

and a pore-forming cytolysin (listeriolysin O, LLO) to escape the phagosome. However, prior to

escape, the bacterium must overcome a number of phagosomal defenses, including autophagy

and NOX2 NADPH oxidase production of reactive oxygen species (ROS).

Autophagy, the cellular process of self-digestion, is a key component of innate immunity.

Previously, it has been shown that L. monocytogenes is targeted by autophagy (LC3+) at 1 h post

infection (p.i.) but the mechanism remains elusive. Here, I show that at 1 h p.i., diacylglycerol

(DAG) and ROS production are required for autophagy targeting to the bacteria, which are

predominantly in phagosomes. It has been shown that autophagy targeting of cytosolic L.

monocytogenes is mediated via protein ubiquitination. However, protein ubiquitination is not

associated with LC3+ bacteria at 1 h p.i.. Thus, my data suggest that distinct signals mediate

autophagy targeting of L. monocytogenes depending on the location within host cells.

iii

Given that ROS mediate autophagy targeting to L. monocytogenes and that previous studies have

demonstrated that ROS production limits bacterial escape, I investigated how L. monocytogenes

overcomes ROS production prior to phagosomal escape. I found that LLO inhibits ROS

production by preventing NOX2 NADPH oxidase localization to L. monocytogenes-containing

phagosomes. LLO-deficient bacteria can be complemented by perfringolysin O, a related

cytolysin, suggesting that other pathogens may also use pore-forming cytolysins to inhibit ROS

production. While PLCs can activate ROS production, this effect is alleviated by LLO pore-

formation. Therefore, the combined activities of PLCs and LLO allow L. monocytogenes to

efficiently escape the phagosome while avoiding microbicidal ROS.

Together, this thesis provides a clearer understanding of the balance between host defense versus

bacterial evasion. Greater insight into host-bacterial interaction may lead to better therapeutics

that can “tip the balance” in the host’s favour.

iv

Acknowledgments

I must first express my deepest gratitude for my supervisor, Dr. John H. Brumell, for his

guidance, support and patience throughout the course of my graduate studies. I am particularly

thankful for the scientific freedom I was given to pursue my own theories and hypotheses. I am

also grateful to the members of my supervisory committee, Drs. Scott Gray-Owen, Nicola Jones

and Philip Sherman, for their support, criticism and suggestions, pushing me beyond the

boundaries of what I thought I can achieve.

This thesis would not have been possible without the input and support of all past and current

Brumell Lab members. I am particularly grateful to Dr. Malina Bakowski for showing me the

ropes of the lab when I first arrived, Dr. Cheryl Birmingham for her hard work on the L.

monocytogenes project I was to inherit, and Dr. Ju Huang for providing both emotional and

scientific advice, insight and support. I am thankful for Veronica Canadien who ensures the

smooth operation of the lab. I would also like to thank Michelle Ang and Dr. Ramzi Fattouh, my

lab husband, for their help with the FACS experiments and Dr. Darren Higgins from Harvard

Medical School for his scientific advice and tutelage on performing in vivo L. monocytogenes

infections in mice. Furthermore, I wish to express my gratitude to the staff at the SickKids

imaging facility, the Mt. Sinai Hospital TEM facility as well as SIDNET at SickKids. I would

also like to thank the members of the Grinstein, Jones, Klip, Robinson and Trimble laboratories

for the daily laughs, reagents, and equipment usage.

Finally, I would like to thank my parents, Raymond and Elaine, for their enduring love, endless

patience and unquestioning (monetary and emotional) support during my lengthy educational

journey. Lastly, I must thank Dr. Mathew Estey for his insightful scientific advice (especially

while on Greyhound buses), patience, emotional support, and unconditional love. You are the

North star to my wandering bark; the light during the good times and the bad.

v

Contributions

The majority the work presented in this thesis was conducted by me. Experiments were

conceived by Dr. John Brumell and myself with advice from my supervisory committee, which

comprises of Drs. Scott Gray-Owen, Nicola Jones and Phil Sherman. Further advice on the

direction and development of my projects was given by Drs. Sergio Grinstein, Aleixo M. Muise

and Darren E. Higgins. Dr. Ramzi Fattouh helped perform the flow cytometric assays in Chapter

20 with technical assistance from Michelle Ang and David Rizutti. Mike Woodside and Paul

Paroutis provided technical support for confocal microscopy. Post fixation processing of samples

for transmission microscopy (TEM) in Chapter 20 was conducted by Bob Tempkin at the Mt.

Sinai TEM facility.

I am supported by a M.D/Ph.D Studentship and Canadian Graduate Scholarship Doctoral

Research Award from the Canadian Institutes of Health Research. This work was supported by

an operating grant from the Canadian Institutes of Health Research (MOP#97756).

vi

Table of Contents

ACKNOWLEDGMENTS..................................................................................................................................................4

CONTRIBUTIONS.............................................................................................................................................................5

TABLE OF CONTENTS ...................................................................................................................................................6

LIST OF ABBREVIATIONS..........................................................................................................................................11

LIST OF TABLES ............................................................................................................................................................15

LIST OF FIGURES ..........................................................................................................................................................16

CHAPTER 1 BACKGROUND .....................................................................................................................................1

1 INTRODUCTION TO PHAGOCYTOSIS ..............................................................................................................1

1.1 MACROPHAGES.........................................................................................................................................................2

1.2 PHAGOSOMAL MATURATION .............................................................................................................................3

2 INNATE HOST DEFENSES IN THE PHAGOSOME..........................................................................................5

2.1 REACTIVE OXYGEN SPECIES (ROS) ......................................................................................................................7 2.1.1 Intracellular Sources of ROS ........................................................................................................................7 2.1.2 NOX2 NADPH oxidase – Main source of ROS in Immunity.......................................................................9 2.1.2.1 Components of the NOX2 NADPH oxidase ..................................................................................................9 2.1.2.2 Activating signals for NOX2 NADPH oxidase assembly ...........................................................................13 2.1.2.3 Loss of NOX2 NADPH oxidase function results in Chronic Granulomatous Disease (CGD)................14 2.1.3 Types of ROS ................................................................................................................................................15 2.1.4 Functions of ROS .........................................................................................................................................17 2.1.4 Common techniques of ROS quantification................................................................................................20

2.2 AUTOPHAGY..........................................................................................................................................................28 2.3 AUTOPHAGY CORE MACHINERY...........................................................................................................................28 2.3.1 INITIATION .............................................................................................................................................................29 2.3.2 ELONGATION..........................................................................................................................................................29

2.3.1 Maturation: Non-selective vs selective autophagy ....................................................................................32 2.4 AUTOPHAGY AND INTRACELLULAR PATHOGENS................................................................................................33

2.3.3 Common techniques used to study autophagy............................................................................................46 2.3.3.1 Techniques used to quantify changes in autophagy ...................................................................................46 2.3.3.2 Techniques used to assess flux .....................................................................................................................47

2.4 OTHER DEFENSES .................................................................................................................................................50

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3 BACTERIAL EVASION STRATEGIES IN RESPONSE TO HOST DEFENSES........................................50

4 L. MONOCYTOGENES .............................................................................................................................................52

4.1 GENETIC DIVERSITY OF L. MONOCYTOGENES ......................................................................................................53 4.2 EPIDEMIOLOGY OF CLINICAL LISTERIOSIS ..........................................................................................................53

4.2.1 Clinical Presentation and Treatment..........................................................................................................53 4.2.2 Clinical Isolates ...........................................................................................................................................54

4.3 L. MONOCYTOGENES INFECTION AND INTRACELLULAR LIFESTYLE.....................................................................54 4.3.1 Entry into host cells .....................................................................................................................................55 4.3.2 Escape from the phagosome........................................................................................................................57 4.3.3 Cytosolic Replication...................................................................................................................................61 4.3.4 Secondary Spread ........................................................................................................................................61 4.3.5 PrfA Regulon ................................................................................................................................................62

4.4 PHAGOSOMAL IMMUNE DEFENSES AND L. MONOCYTOGENES.............................................................................63 4.4.1 Autophagy and L. monocytogenes ..............................................................................................................63 4.4.2 ROS AND L. monocytogenes.......................................................................................................................67

CHAPTER 2 AIMS AND HYPOTHESES ...............................................................................................................72

5 AIM ..............................................................................................................................................................................72

5.1 HYPOTHESIS #1:....................................................................................................................................................73 5.1.1 Aim ................................................................................................................................................................73

5.2 HYPOTHESIS #2:....................................................................................................................................................74 5.2.1 Aim ................................................................................................................................................................74

CHAPTER 3 METHODS ............................................................................................................................................77

6 BACTERIAL STRAINS AND CULTURE CONDITIONS................................................................................77

7 MACROPHAGE GENERATION AND CULTURE CONDITIONS ...............................................................77

8 PLASMIDS AND TRANSFECTIONS ...................................................................................................................78

9 NEUTROPHIL PURIFICATION AND RECOMBINANT LLO (RLLO) TREATMENT ..........................79

10 IN VIVO INFECTION ............................................................................................................................................79

11 MACROPHAGE REPLICATION ASSAY AND INFECTION ......................................................................79

12 REAGENTS AND ANTIBODIES.........................................................................................................................80

13 IMMUNOSTAINING AND IMMUNOFLUORESCENCE .............................................................................80

14 DYNAMIC LUMINOL BASED ASSAYS OF ROS PRODUCTION .............................................................81

15 NITROBLUE TETRAZOLIUM (NBT) QUANTIFICATION OF SUPEROXIDE PRODUCTION........81

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16 FLOW CYTOMETRIC QUANTIFICATION OF ROS ...................................................................................82

17 CERIUM PERHYDROXIDE PRECIPITATE QUANTIFICATION OF PHAGOSOMAL ROS ............82

18 STATISTICAL ANALYSIS...................................................................................................................................83

CHAPTER 4 RESULTS...............................................................................................................................................84

19 HOST AND BACTERIA FACTORS REGULATE AUTOPHAGIC TARGETING OF L.

MONOCYTOGENES DURING EARLY STAGES OF INFECTION IN MACROPHAGES ..............................84

19.1 AUTOPHAGY TARGETS WILD TYPE L. MONOCYTOGENES WITHIN MACROPHAGE PHAGOSOMES PRIOR TO

PHAGOSOMAL ESCAPE.....................................................................................................................................................84 19.2 HOST AND BACTERIAL FACTORS PROMOTE DAG ACCUMULATION ON PHAGOSOMES CONTAINING L.

MONOCYTOGENES.............................................................................................................................................................89 19.3 DAG PRODUCTION IS REQUIRED FOR AUTOPHAGIC TARGETING OF L. MONOCYTOGENES AT EARLY STAGES OF

INFECTION........................................................................................................................................................................95 19.4 ROS PRODUCTION BY THE NOX2 NADPH OXIDASE IS REQUIRED FOR AUTOPHAGIC TARGETING OF L.

MONOCYTOGENES AT EARLY STAGES OF INFECTION. .....................................................................................................97 19.5 LC3 TARGETING IS DEPENDENT ON LLO PORE-FORMATION..........................................................................100 19.6 ROS PRODUCTION BY THE NOX2 NADPH OXIDASE IS REQUIRED FOR SLAP FORMATION AT LATER STAGES

OF INFECTION. ...............................................................................................................................................................102 19.7 SUMMARY .........................................................................................................................................................105

20 LISTERIOLYSIN O SUPPRESSES PLC-MEDIATED ACTIVATION OF THE MICROBICIDAL

NADPH OXIDASE TO PROMOTE LISTERIA MONOCYTOGENES INFECTION.........................................106

20.1 LLO INHIBITS ROS PRODUCTION BY THE NOX2 NADPH OXIDASE ............................................................106 20.2 LLO CAN INHIBIT ROS PRODUCTION BY OTHER CELL TYPES ........................................................................112 20.3 LLO INHIBITS PHAGOSOMAL PRODUCTION OF ROS .......................................................................................113 20.4 L. MONOCYTOGENES PLCS CONTRIBUTE TO THE INDUCTION OF ROS PRODUCTION BY THE NOX2 NADPH

OXIDASE.........................................................................................................................................................................113 20.5 LLO DEFICIENT BACTERIA SURVIVE IN NOX2 NADPH OXIDASE DEFICIENT MACROPHAGES ....................117 20.6 LLO DEFICIENT BACTERIA SURVIVE IN NOX2 NADPH OXIDASE DEFICIENT MICE .....................................117 20.7 LLO INHIBITS NOX2 NADPH OXIDASE ASSEMBLY AT THE PHAGOSOME....................................................121 20.8 PERFRINGOLYSIN O (PFO) EXPRESSION CAN COMPLEMENT NOX2 NADPH OXIDASE INHIBITION BY LLO

DEFICIENT BACTERIA ....................................................................................................................................................121 20.9 LLO INHIBITION OF NOX2 NADPH OXIDASE ASSEMBLY IS NOT MEDIATED THROUGH ALTERING KEY

SIGNALING EVENTS THAT MEDIATE NOX2 NADPH OXIDASE ASSEMBLY ................................................................125 20.10 SUMMARY .......................................................................................................................................................128

CHAPTER 5 DISCUSSION ......................................................................................................................................129

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21 DECIPHERING THE COMPLEXITY OF L. MONOCYTOGENES AND AUTOPHAGY

INTERACTION ..............................................................................................................................................................132

21.1 DIFFERENT DOWNSTREAM SIGNALING CONSEQUENCES OF DAG PRODUCTION BY

BACTERIA OR HOST ..................................................................................................................................................132

21.2 INDUCTION OF AUTOPHAGY – ADVANTAGE TO THE CELL OR BACTERIA?............................133

21.3 HOW IS LC3 TARGETED TO L. MONOCYTOGENES CONTAINING PHAGOSOMES? ...................134

21.4 L. MONOCYTOGENES APPEARS TO POSSESS THE ABILITY TO LIMIT AUTOPHAGIC

TARGETING...................................................................................................................................................................135

21.4.1 FUTURE DIRECTIONS ....................................................................................................................................137

21.5 MULTIPLE SIGNALS MAY BE ACTING IN THE SAME PATHWAY TO MEDIATE AUTOPHAGY

TARGETING...................................................................................................................................................................138


22 WALKING A TIGHT ROPE: THE IMPORTANCE OF L. MONOCYTOGENES VIRULENCE

FACTOR ANTAGONISM IN PRESENCE OF NOX2 NADPH OXIDASE ACTIVITY ..................................139

22.1 MULTIPLE PEAKS OF ROS PRODUCTION MAY BE DUE TO DIFFERENT ACTIVATION

SIGNALS..........................................................................................................................................................................139


22.2 RELATIONSHIP BETWEEN ROS AND LLO PROVIDES INSIGHT INTO L. MONOCYTOGENES

PATHOGENESIS IN DIFFERENT CELL TYPES..................................................................................................141


22.3 UNDERSTANDING THE MECHANISM OF LLO MISLOCALIZATION OF NOX2 NADPH

OXIDASE .........................................................................................................................................................................143


22.4 MULTIPLE MECHANISMS MAY BE INVOLVED IN NOX2 NADPH OXIDASE INHIBITION.......147

22.5 OPPOSING CONSEQUENCES OF LLO AND PLCS ACTIVITY ALLOWS FOR ROS PRODUCTION

IN THE PHAGOSOMAL COMPARTMENT ...........................................................................................................148

22.6 NOX2 INHIBITION BY PORE FORMING CYTOLYSINS AND VIRULENCE FACTOR

ANTAGONISM MAY BE COMMON STRATEGIES FOR INTRACELLULAR SURVIVAL ......................151

22.7 UNDERSTANDING LLO PORE-FORMATION AND ROS IN THE TARGETING OF AUTOPHAGY

............................................................................................................................................................................................152


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23 CONCLUDING REMARKS................................................................................................................................156

REFERENCES ................................................................................................................................................................157

COPYRIGHT ACKNOWLEDGEMENTS ................................................................................................................197

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List of Abbreviations

AMP Adenosine monophosphate

AMPK AMP-activated protein kinase

APF 2-[6-(4’-amino)phenoxy-3H-

xanthen-3-on-9-yl]benzoic

acid

Atg Autophagy-related genes

BHI Brain-Heart Infusion

BKGD Background

BMDM Bone marrow derived

macrophages

BMDN Bone marrow derived

neutrophils

cAMP Cycic AMP

CBD-YFP Cell wall binding domain –

yellow fluorescent protein

CD Crohn’s Disease

CDC Cholesterol dependent

cytolysins

CFU Colony forming units

CGD Chronic granulomatous

disease

CLEM Correlative light microscopy-

electron microscopy

CM-

H2DCFDA

5-(and-6)-chloromethyl-2-7-

dichlorodihydrofluorescein

diacetate

CMA Chaperone-mediated

autophagy

CPS Counts per second

CREB cAMP response element-

binding

DAG Diacylglycerol

DCF 2’-7’-dichlorofluorescein

DHC Dihydrocalcein

DHE Dihydroethidium

DHR Dihydrorhodamine 123

DMEM Dulbecco’s modified Eagle’s

medium

DMSO Dimethyl sulfoxide

xii

DN Dominant negative

DNA Deoxyribonucleic acid

DPI Diphenyliodonium

EEA-1 Early endosomal autoantigen-

1

Ena/VASP Enabled/vasodilator-

stimulated phosphoprotein

ER Endoplasmic reticulum

ERK1/2 Extracellular regulated kinase

1/2

FBS Fetal bovine serum

FcγR Fc-gamma-receptor

GFP Green fluorescent protein

GILT γ-interferon inducible

lysosomal thiolreductase

GTP Guanosine triphosphate

HPF 2-[6-(4’-hydroxy) phenoxy-

3H-xanthen-3-on-9-yl]

benzoic acid

HRP Horseradish peroxidase

IBD Inflammatory Bowel Disease

IBMX 3-isobutyl-1-methylxanthine

IFN Interferon

Inl Internalin

IP3 Inositol 3,4, 5 triphosphate

IPTG Isopropyl b-D-11-

thiogalactopyranoside

LAMP Lysosomal-associated

membrane proteins

LC3 Microtubule-associated protein

light chain 3

LD50 50% lethal dose

LLO Listeriolysin O

LLR Leucine-rich repeats

LPS Lipopolysaccharide

MAPK Mitogen-activated protein

kinase

MEFs Mouse embryonic fibroblasts

MnSOD Manganese superoxide

dismutase

MOI Multiplicity of infection

MPO Myeloperoxidase

xiii

NADPH Nicotinamide adenine

dinucleotide phosphate

NBR1 Neighbor of BRCA1 gene 1

NBT Nitroblue tetrazolium chloride

Ndk Nucleoside diphosphate kinase

NDP52 Nuclear dot protein 52 kDa

NETs Neutrophil extracellular traps

NF-κB Nuclear factor-kappa-B

NLR Nod-like receptor

NOD Nucleotide-binding

oligomerization domain

NOS Nitric oxide synthase

p.i. Post infection

Pak p21-activated kinase

PAMP Pathogen associated molecular

pattern

PAS Phagophore assembly site

PBMC Peripheral blood mononuclear

cells

PBS Phosphate buffered saline

PC-PLC Phosphatidylcholine specific

phospholipase C

PAP Phosphatidic acid phosphatase

PBD Pak-binding domain

PDE Phosphodiesterase

PE Phosphatidylethanolamine

PFA Paraformaldehyde

PFO Perfringolysin O

Phox Phagocytic oxidase

PI Phosphatidylinositide

PI3K Phosphatidylinositol 3-kinase

PI3P Phosphatidylinositol 3-

phosphate

PI-PLC Phosphatidylinositol specific

phospholipase C

PKA Protein kinase A

PKC Protein kinase C

PLD Phospholipase D

PMA Phorbol 12-myristate 13-

acetate

RPMI Roswell Park Memorial

Institute medium

xiv

PR Proline rich

PrfA Positive regulation factor A

PRR Pathogen pattern recognition

receptor

Px PhoX domain

RFP Red fluorescent protein

RNS Reactive nitrogen species

ROS Reactive oxygen species

SEM Standard error of the mean

SH3 SRC-homology 3

SLAPs Spacious Listeria containing

phagosomes

SOD Superoxide dismutase

SPI-2 Salmonella pathogenicity

island 2

ssRNA Single stranded ribonucleic

acid

TEM Transmission electron

microscopy

TIGAR TP53-induced glycolysis and

apoptosis regulator

TLR Toll-like receptor

TNF-α Tumor necrosis factor-alpha

Ub Ubiquitin

UPEC Uropathogenic E. coli

Vcc Vibrio cholera cytolysin

VPS34 Vacuolar protein sorting 34

WASP Wiskott Aldrich syndrome

protein

Yop Yersinia outer protein

xv

List of Tables

Table 2.3 Summary of known interactions between pathogens and autophagy

Table 4.4. List of pathogens that have been reported to actively modulate NOX activity.

Table 20.1. Mean and SEM of luminol and isoluminol readings from BMDM infected with WT

or ΔLLO bacteria.

xvi

List of Figures

Figure 1. Various Stages of Phagosomal Maturation.

Figure 2.1. Key host phagosomal defenses against invading pathogens.

Figure 2.2. Intracellular sources of ROS.

Figure 2.3. A Schematic of NOX2 assembly and activation.

Figure 2.4. The generation of common ROS products in phagocytes.

Figure 2.5. Steps involved in the induction of autophagy.

Figure 2.6. Regulation of autophagy by immune signals in response to invading pathogens.

Figure 2.7. The known pathways that are dependent on ROS induction of autophagy.

Figure 2.8. Examples of methods to assess autophagy.

Figure 4. Intracellular lifecycle of L. monocytogenes with associated electron micrographs.

Figure 19.1.1. Autophagy targets wild type L. monocytogenes during macrophage infection.

Figure 19.1.2. Autophagy targets wild type L. monocytogenes within macrophage phagosomes

during early stages of infection.

Figure 19.1.3. LC3+ L. monocytogenes containing phagosomes are not targeted by Ub at 1 h.

Figure 19.2.1. Host and bacterial factors promote DAG accumulation on phagosomes containing

L. monocytogenes.

Figure 19.2.2. Inhibition of host DAG producers can further decrease DAG localization to ΔPI-

PLCΔPC-PLC L. monocytogenes.

Figure 19.3. DAG production is required for autophagic targeting of L. monocytogenes during

early stages of infection.

xvii

Figure 19.4. ROS production by the NOX2 NADPH oxidase is required for autophagic targeting

of L. monocytogenes during early stages of infection.

Figure 19.5. LC3 targeting is dependent on LLO pore-forming ability secreted from live L.

monocytogenes

Figure 19.6 ROS production by the NOX2 NADPH oxidase is required for the generation of

SLAPs.

Figure 20.1.1. ΔLLO mutants induce greater ROS production compared to WT or ΔLLOΔPI-

PLCΔPC-PLC L. monocytogenes during infection in macrophages.

Figure 20.1.2. LLO inhibits PLC-mediated ROS production by the NOX2 NADPH oxidase.

Figure 20.1.3. LLO inhibits ROS production by NOX2 NADPH oxidase.

Figure 20.2. LLO is capable of inhibiting ROS production by PMA activated human neutrophils.

Figure 20.3. LLO inhibits ROS production in phagosomes.

Figure 20.4. Both PLCs are required to induce ROS production by macrophages during

infection.

Figure 20.5. ΔLLO bacteria survive in NOX2 NADPH oxidase-deficient macrophages in vitro.

Figure 20.6. ΔLLO bacteria survive in NOX2 NADPH oxidase-deficient macrophages in vitro

and in vivo.

Figure 20.7.1. LLO inhibits NOX2 NADPH oxidase assembly at the phagosome.

Figure 20.7.2. Wild type L. monocytogenes inhibit ROS production through LLO to mislocalize

p47phox and p67phox to the phagosome.

Figure 20.8. LLO inhibition of ROS production can be complemented by perfringolysin O

(PFO).

xviii

Figure 20.9. LLO mediated inhibition of ROS production is not dependent on altering the

phosphorylation states of key signaling molecules responsible of NOX2 NADPH oxidase

activation.

Figure 21.1. Autophagy targeting to L. monocytogenes at early and late stages of infection are

mediated via different mechanisms.

Figure 21.2. The “Push and Pull” dynamics of host and bacteria interaction.

Figure 21.3. Proposed model of autophagic targeting of latex beads or bacteria.

Figure 22.5. Methods of ROS inhibition employed by L. monocytogenes.

Figure 22.7. Putative Model of L. monocytogenes interaction with early host defenses against

phagocytosed pathogens.

1

Chapter 1

Background

1 Introduction to Phagocytosis

Host-bacterial interaction is a dynamic process where opposing measures are taken by the host

and pathogen with the goal of ensuring survival. The host immune system provides defenses

against invading microbial pathogen, whereas pathogens have an arsenal of virulence factors to

counteract the microbicidal effects of host immunity. This interaction applies a selective pressure

on both parties to evolve diverse defensive or evasion strategies. In recent years, much work has

focused on the early stages of infection as it has become increasingly apparent that the course of

infection can be significantly influenced by events early in an infection. As such, a number of

reviews have focused solely on this critical early stage of infection (Flannagan et al., 2009;

(Kirkegaard et al., 2004; Kumar and Valdivia, 2009). Depending on the invading microbe, the

host can mount different responses against the pathogen, be it bacterial, fungal, parasitic or viral

in nature. Given the focus of this thesis, host responses to bacterial infection will be

predominantly discussed.

Upon entry in the body, bacteria first encounter the early branch of the immune system: the

innate immune response. The central players of this response are phagocytes, which are typically

the first responders to an infection. These cells are so-termed for their ability to ingest debris,

dead cells, or invading pathogens in a process called phagocytosis, “cell-eating”. These cells

either reside in peripheral tissues or in the blood stream until they are activated and recruited to

the site of infection.

While a class of “non-professional” phagocytes exists with the limited ability to phagocytose

bacteria, a range of “professional” phagocytes, including neutrophils, monocytes, macrophages,

dendritic cells and mast cells, are efficient at phagocytosis. Of these, neutrophils and

macrophages are the most efficient at phagocytosis and thus are often the key cell types for

studying bacterial-host interactions. Given the focus of this thesis, discussion of phagocytosis

2

will be restricted to that of macrophages.

1.1 Macrophages

Macrophages differentiate from hematopoetic stem cells first as the immature form, monocytes.

Monocytes mainly circulate in the blood stream and exhibit limited ability to divide and

phagocytose particles. Upon activation, monocytes migrate from the blood to the site of infection

and depending on the type of activation signal, they can differentiate into macrophages that are

either highly proliferative or efficient at phagocytosis and killing.

Phagocytosis by macrophages is initiated upon the detection of extracellular bacteria. The

presence of pathogens can be identified via the activation of Toll-like receptors (TLRs), various

types of opsonin receptors, or scavenger receptors. Toll-like receptors (TLRs) on phagocytes

interact with bacterial cell wall components named pathogen associated molecular patterns

(PAMPs). There is a diverse family of PAMPs, the best characterized of which include

lipopolysaccharides (LPS), peptidoglycans (PGs), flagellin and lipoteichoic acids (LTAs).

Opsonin receptors, including both Fc-gamma receptors (FcγRs) and complement receptors, can

also trigger phagocytosis. Soluble factors, such as antibodies and complement, can often bind

directly to extracellular bacteria. In doing so, antibodies can neutralize the invasiveness of the

pathogen by physically blocking interaction between bacterial and host proteins while

complement components can act in a concerted fashion to directly form holes on the bacterial

surface, resulting in bacterial lysis. In addition, antibodies and complement on the surface of the

bacteria can interact with FcγRs and complement receptors on macrophages to increase the

efficiency of phagocytosis in a process called opsonization. Finally, scavenger receptors act in a

manner similar to TLRs, binding to bacterial cell wall proteins. While the activation of these host

receptors can mediate phagocytosis, it also results in a series of downstream signaling cascades

that mediate the activation of a number of important cellular innate immune defenses. Many

these mechanisms are involved in activating other bactericidal mechanisms, thus allowing for

more efficient bacterial killing inside the phagosome.

Upon activation, macrophages mediate the rapid clearance of bacteria. This can be achieved via

3

either the extracellular secretion of bactericidal compounds or the intracellular degradation of

ingested or phagocytosed pathogens. Extracellular killing of bacteria can occur following the

secretion of reactive compounds, such as reactive nitrogen species (RNS) or reactive oxygen

species (ROS), or bactericidal enzymes, such as lysozymes. Intracellular killing of bacteria

involves phagocytosis of pathogens followed by digestion in the “stomach” of the cell, the

lysosome. Not only is phagocytosis critical for bacterial clearance but also promotes antigen

presentation, directing the establishment of the late branch of the immune system: the adaptive

immune response.

1.2 Phagosomal Maturation

To degrade phagocytosed pathogens, phagocytes have a number of mechanisms to breakdown

the internalized bacteria. One key degradative mechanism is the delivery of the bacteria to the

lysosome for digestion. This is a multi-step process that begins when the bacteria is

phagocytosed, creating a bacteria containing compartment called a phagosome (Figure 1).

Typically, phagosomes mature through the acquisition and concerted loss of various vesicular

and signaling proteins that mediate transition through different stages of maturation and

ultimately fuse with the lysosome. Markers of early phagosome include early endosomal

autoantigen (EEA-1) and the small GTPase, Rab5 (Bucci et al., 1992). These markers are

recruited by the changes in lipid composition of the phagosomal membrane. VPS34, a class III

phosphatidylinositol-3-kinase, generates phosphatidlinositol-3-phosphate (PI(3)P), which allows

the binding of these early markers (Vieira et al., 2001). Next, the phagosome progresses to the

late phagosome via the acquisition of V-ATPase, a proton-pump, creating an acidic luminal

environment (Desjardins et al., 1994). Furthermore, lysosomal-associated membrane proteins

(LAMPs) and small GTPase Rab7 are recruited (Bucci et al., 1992). Degradative enzymes, such

as cathepsins and hydrolases are delivered to the lumen of the phagosome. Thus, changes to the

membrane of the phagosome are associated with changes in the lumen, transitioning to a more

acidic, oxidative and degradative environment as the phagosome matures. Finally, late

phagosome fusion with the lysosome, which has an average pH of 4.5, results in the formation of

the phagolysosome in which microbial degradation occurs (Desjardins et al., 1994) (Figure 1).

4

Figure 1. Various Stages of Phagosomal Maturation. Interaction of the bacteria containing

phagosome with various subcompartments of the endocytic pathway. This mediates the transition

of the phagosome from (A) early, (B) intermediate, (C) late phagosome to ultimately the

phagolysosome where degradation occurs. EEA1, early endosome antigen 1; LAMP, lysosomal-

associated membrane protein; PI(3)P phosphatidylinositol-3-phosphate. Adapted with permission

from Macmillian Publishers Ltd: Nat. Rev. Microbiol. Flannagan et al. Antimicrobial

mechanisms of phagocytes and bacterial evasion strategies. 7:5 (355-66), 2009.

5

2 Innate Host Defenses in the Phagosome

Prior to delivery of phagocytosed bacteria to the lysosome, there are a number of host

mechanisms that are activated in the phagosome that either directly or indirectly targets the

engulfed bacteria. Such strategies include the production of antimicrobial peptides, such as

defensins, lactoferrins (and other metal chelators), proteases, cathepsins, RNS and ROS (Figure

2.1) (reviewed in (Flannagan et al., 2009)). Antimicrobial peptides are charged peptides that

directly form pores in the bacterial membrane, resulting in lysis. Metal chelators in the

phagosomal lumen can bind essential nutrients, such as iron, thus restricting bacterial growth by

limiting bacterial access to free metals. Furthermore, as assortment of peptide specific proteases

and cathepsins, as well as hydrolases against carbohydrates and lipids, are also present inside the

phagosome, thus aiding in bacterial degradation. Finally, reactive compounds, such as RNS and

ROS can interfere with bacterial survival by modifying bacterial proteins, lipids and DNA to

prevent proper function or folding. It must be noted that while there are a number of antibacterial

mechanisms inside the phagosome, they are activate during different stages of phagosomal

maturation. For example, the optimal pH for the activity of hydrolytic enzymes is often pH < 4.0.

As such, these enzymes are most active during the late phagosome stage. Conversely, NOX2

NADPH oxidase- mediated production of ROS is among the earliest and most robust defenses

employed by phagocytes to combat microbial infection. The recruitment and assembly of the

NOX2 NADPH oxidase complex begin as early as the formation of the nascent phagocytic cup,

long before the phagosome has been fully sealed.

6

Figure 2.1. Key host phagosomal defenses against invading pathogens. Inside the

phagosome, the bacteria encounters a series of bacteriocidal measures, including reactive oxygen

species (ROS) generated by the NOX2 NADPH oxidase and reactive nitrogen species (RNS)

generated by the inducible nitric oxide synthase (iNOS). Furthermore, antimicrobial peptides,

such as defensins, and proteases all contribute to direct bacterial disruption and degradation.

SOD, superoxide dismutase, MPO, myeloperoxidase. Adapted with permission from Macmillian

Publishers Ltd: Nat. Rev. Microbiol. Flannagan et al. Antimicrobial mechanisms of phagocytes

and bacterial evasion strategies. 7:5 (355-66), 2009.

7

2.1 Reactive Oxygen Species (ROS)

ROS are a group of highly reactive oxygen based free radicals that can be toxic to both the host

and pathogens (Rada et al., 2008). ROS can interact with proteins, lipids and DNA, resulting in

damage or structural changes to the molecule that can interfere with proper function. However, a

number of important host enzymes produce ROS as a byproduct. Furthermore, ROS also have a

number of important direct and indirect functions in immunity where ROS production by

phagocytes can directly kill pathogens and signal the activation of further immune responses to

effectively clear the infection. Thus, given the “double-edged sword” nature of ROS, there are a

number of host mechanisms that regulate the level of ROS both spatially and temporally.

2.1.1 Intracellular Sources of ROS

There are a number of cellular sources that generate ROS including NOXs, the mitochondrial

electron transport chain (ETC), xanthine oxidoreductase, peroxisomes, and the endoplasmic

reticulum (ER) (Antonenkov et al.; Harrison, 2002; Murphy, 2009; Nauseef, 2008; Santos et al.,

2009). In cells that do not express NOX2 NADPH oxidase, the main source of ROS is derived

from the mitochondrial ETC. The ETC removes and transports electrons from the donor

compound nicotinamide adenine dinucleotide (NADH) to generate an electrical gradient that

ultimately drives the production of adenosine triphosphate (ATP) (Raha and Robinson, 2000).

Two components of the ETC, NADH dehydrogenase (Complex I) and Coenzyme Q -

cytochrome C reductase (Complex III), are prone to electron leakage out of the ETC. When

combined with oxygen, this results in the production of superoxides (Raha and Robinson, 2000).

Besides the mitochondria, there are other organelles that also harbor enzymes capable of

producing ROS. Peroxisomes are organelles that contain a number of oxidases that function to

catabolize long and very long fatty acid chains. Many of these enzymes, such as acyl-CoA

oxidase, urate oxidase or sarcosine oxidase, produce ROS as a byproduct (Antonenkov et al.). In

the ER, ER oxidoreductin (Ero1) is responsible for ROS production. Proper folding and disulfide

bond formation of oxidative proteins require the maintenance of a suitable redox environment.

Ero1 is a flavin-containing enzyme that produces ROS to create an oxidizing environment in the

ER (Travers et al., 2000; Tu et al., 2000). In absence of Ero1, reduced and misfolded proteins

8

Figure 2.2. Intracellular sources of ROS. Superoxide anions (O2–•) can be generated by

xanthine oxidase in catalyzing the oxidation of hypoxanthine to xanthine; by NADPH oxidase

(e.g. Nox2 on phagosomal membrane in phagocytic cells) in converting NADPH to NADP+; and

by mitochondrial electron transport chain reaction, in which electrons (e) are transferred from the

mitochondria complex I (I) to CoQ via the complex II (II), then to cytochrome c via the complex

III (III) and finally to O2 via the complex IV (IV) and H2O is produced. Electrons can leak from

the electron transport chain and O2–• is generated from the complex I and III. Other sources of

ROS include ER and peroxisome. Adapted with permission from Mary Ann Liebert, Inc:

Antioxidants Redox Signaling, Huang J et al. Autophagy Signaling Through Reactive Oxygen

Species. 14(11): 2215-31.

9

accumulate in the ER (Frand and Kaiser, 1998; Pollard et al., 1998). In addition to organelle-

associated proteins, cytosolic enzymes can also make ROS. Xanthine oxidoreductase, a flavin-

containing enzyme in the liver that catalyzes the conversion of the purine derivative,

hypoxanthine, to xanthine and ROS (Harrison, 2002). Lastly, the membrane bound protein

complex, NOX, also produces ROS.

2.1.2 NOX2 NADPH oxidase – Main source of ROS in Immunity

The NOX family of protein complexes, comprised of 7 members (NOX1-5 and DUOX1-2), is

characterized by their ability to produce ROS by transferring an electron from NADPH to

oxygen (Nauseef, 2004). However, the members of this family of complexes greatly differ in

their activation signals, the kinetic of ROS production as well as the cell type in which they are

expressed. Macrophages predominantly express NOX2 NADPH oxidase as other cell types

express a low or negligible level of this protein complex. Compared to other NOX family

members, NOX2 NADPH oxidase is the most rapid and efficient producer of ROS, making it the

main source of ROS during infection.

2.1.2.1 Components of the NOX2 NADPH oxidase

NOX2 NADPH oxidase is comprised of functional transmembrane heterodimers, gp91phox and

p22phox (also known collectively as the cytochrome b558), and 4 regulatory cytosolic subunits –

p40phox, p47phox, p67phox and the small GTPase, Rac2 (Figure 2.3). In the inactive state,

cytochrome b558 resides in either Rab5+ early endosomes (Casbon et al., 2009; Li et al., 2006)

or Rab11+ recycling vesicles (Casbon et al., 2009), p40phox, p47phox and p67phox are in a

dephosphorylated trimeric state in the cytosol (Lapouge et al., 2002), while cytosolic Rac2

remains inactive in the GDP bound state via interaction with RhoGDI (Figure 2.3 A) (Ando et

al., 1992; Mizuno et al., 1992). Upon stimulation or the initiation of phagocytosis, rapid transit of

cytochrome b558 from vesicles to the nascent phagocytic cup occurs (Figure 2.3 B) (Diebold and

Bokoch, 2001). Concurrently, p47phox is phosphorylated, and undergoes a conformational change

in the C-terminal “autoinhibitory domain” that now exposes two SRC-homology 3 (SH3) regions

10

to interact with the proline rich (PR) motif on p22phox (Dusi et al., 1993). Furthermore, Phox

homology (PX) domains on p47phox bind to phosphatidic acid (PA) and phosphatidylinositol 3, 4-

bisphosphate PI(3,4,)P2, transient phosphoinositides that are generated only at the plasma

membrane upon phagocytosis (Bravo et al., 2001; Karathanassis et al., 2002). This further

mediates p47phox localization to cytochrome b558 (Nauseef, 2004). Since p47phox, p67phox and

p40phox are trimerized in the cytosol, the translocation of p47phox brings the other two regulatory

subunits to the membrane as well (Figure 2.3 B) (Lapouge et al., 2002; Park et al., 1994).

However, there are specific regulatory mechanisms in place that control the activation state of

both p67phox and p40phox, which are also required for the proper functioning of the NOX2

complex. Phosphorylated p67phox interacts with cytochrome b558, inducing a conformational

change in the functional subunit that is necessary for ROS production (Zhao et al., 2003). The

role that p40phox plays is less clear but studies suggest that phosphorylated p40phox may undergo a

conformation change exposing its PB1 domain, which mediates p67phox translocation, and its PX

domain, which allows the interaction with PI(3)P (Ellson et al., 2001a; Ellson et al., 2001b;

Kanai et al., 2001). In the inactive state, p40phox PX and PB1 domains are bound in an

intramolecular interaction, preventing interaction with p67phox or PI(3)P (Honbou et al., 2007).

As a result, NOX2 NADPH oxidase activation may also be modulated by a p40phox

conformational change (Bissonnette et al., 2008). Finally, GDP-Rac2 is converted to GTP-Rac2

through the activity of a Rac guanine nucleotide exchange factor (Figure 2.3 B). Active Rac2 can

now translocation to the phagosomal membrane to bind p67phox, thereby completing the assembly

of a functional NOX2 NADPH oxidase complex (Figure 2.3 C).

11

A B

C

12

Figure 2.3 A Schematic of NOX2 assembly and activation. A. In the resting stage, cytochrome

b558 (gp91 and p22phox) resides in vesicles. Rac2, in the inactive GDP bound form, remains in

the cytosol. The regulatory subunits, p47phox, p67phox and p40phox, are in a trimeric complex in the

cytosol. B. Upon receiving signals for activation, cytochrome b558 is recruited to the membrane.

Simultaneously, protein kinase C (PKC) phosphorylates p47phox, resulting in a conformational

change thus allowing p47phox to interact with p22phox and the lipids, PA and PI(3,4)P2. RhoGDI

inhibition of Rac2 is now released to allow GTP binding. ROS production commences at the

nascent phagocytic cup prior to phagosome sealing. C. As the lipid composition changes during

the late stages of phagosome sealing, the membrane becomes enriched with PI(3)P. p40phox

undergoes a conformational change to allow interaction with PI(3)P and p67phox, thereby

maintaining NOX2 NADPH oxidase function and localization to the phagosome. Adapted with

permission from Mary Ann Liebert, Inc: Antioxidants Redox Signaling, Leto T et al. Targeting

and Regulation of Reactive Oxygen Species Generation by NOX Family NADPH Oxidases.

2009. 11(10): 2607-19.

13

2.1.2.2 Activating signals for NOX2 NADPH oxidase assembly

There have been many reports about the identity of the kinases responsible for p47phox, p67phox

and p40phox phosphorylation, and include several PKC isoforms (PKCα (Bengis-Garber and

Gruener, 1996; Fontayne et al., 2002), PKCδ (Dang et al., 2001a; Fontayne et al., 2002), PKCβ

(Dekker et al., 2000; Fontayne et al., 2002), PKCγ (Bey et al., 2004) and PKCζ (Dang et al.,

2001b; Fontayne et al., 2002)), PKA (Kramer et al., 1988), p21 activated kinase (PAK) (Martyn

et al., 2005), ERK1/2 (Dang et al., 2003; Dewas et al., 2000), AKT (Chen et al., 2003), PI3K

(Lehmann et al., 2009; Yamamori et al., 2004), and possibly others. The complexity of NOX2

regulation by such a large number of kinases not only suggests that perhaps a high threshold of

activating signals may be required for NOX2 activity but also that there may be differing

amounts of ROS produced by each individual NOX2 complex, depending on the type of local

activating signal it receives. In fact, a recent study finds that NOX2 assembly and activity is

highly heterogeneous - where only 50% of phagosomes formed upon Fc-γ receptor (FcγR)

mediated phagocytosis have proper p40phox localization and NOX2 function (Tian et al., 2008).

After assembly and activation, NOX2 then produces ROS in a reaction on the cytoplasmic region

of the gp91phox subunit, converting NADPH to NADP+ in a process that liberates two electrons

and one H+. Only the two electrons are transported to the lumen of the phagosome where they

react with two oxygen molecules to form two superoxide ions.

While NOX2 is the main producer of ROS upon phagocytosis (Lambeth, 2004), it must be noted

that other NOX family members, including NOX1, NOX3 and NOX4 are also expressed in

macrophages and can be activated upon stimulation with specific signals. For example, in the

field of ROS and atherosclerosis, oxidation of low-density lipoproteins (oxLDL) leads to

macrophage to foam cell conversion, one of the key steps in plaque formation (Carnevale et al.,

2007). It was recently found that lipopolysaccharide (LPS), a toll-like receptor (TLR)-4 agonist,

can lead to increased NOX1 expression, thereby expediting macrophage conversion to foam cells

(Lee et al., 2009; Park et al., 2009). Other groups have found that NOX4 also plays a critical role

in the formation of oxLDL, leading to macrophage death (Lee et al.). Additionally, NOX3

expression, albeit low, has been reported in RAW macrophages (Sasaki et al., 2009). Thus, while

NOX2 is the predominant form of NADPH oxidase expressed in macrophages and other

phagocytes, the expression and contributions of other NOX members during an infection

14

warrants further investigation. However, it must be noted that other NOX family member are

unlikely to be able to compensate for NOX2 NADPH oxidase activity in the event of NOX2

dysfunction for several reasons. First, NOXes have different activation signals. NOX5, DUOX1

and DUOX2 are activated by increases in intracellular Ca2+ levels while NOX4 activity is not

regulated by the same trimeric phox complex (Bedard and Krause, 2007). Second, other NOXes

have slower kinetics of ROS production when compared to NOX2. Upon macrophage activation,

NOX1 require ~90 min before ROS is produced (Lee et al., 2009) while NOX2 can produce

ROS within minutes (Tian et al., 2008). Finally, other NOXes produce much lower levels of

ROS when compared to NOX2. Under conditions of NOX2 transcriptional downregulation and

NOX1 and NOX3 upregulation, the level of ROS production by stimulated macrophages is

roughly 30x less than when NOX2 transcription is normal (Sasaki et al., 2009). Thus, while other

NOXes can be upregulated to compensate for ROS production in the event of NOX2 NADPH

oxidase dysfunction (Sasaki et al., 2009), they cannot, in a timely fashion, compensate for the

burst of ROS required to effectively clear certain infections. As such, patients bearing mutations

in NOX2 NADPH oxidase genes exhibit an inability to clear infections by organisms that have

antioxidant defenses.

2.1.2.3 Loss of NOX2 NADPH oxidase function results in Chronic

Granulomatous Disease (CGD)

The critical role that NOX2 NADPH oxidase-derived ROS play in host immunity was clearly

demonstrated by genetic studies done in late 1980s that showed patients with mutations in NOX2

NADPH oxidase component genes develop chronic granulomatous disease (CGD) (reviewed by

(Segal, 1996)). CGD is characterized by recurrent and severe infections by a particular spectrum

of bacteria (such as Staphylococcus aureus, Salmonella and Klebsiella species) and fungi (such

as Aspergillus and Candida species) as a result of the host’s inability to mount an effective innate

immune response despite chronic sterile inflammation observed in the body (Winkelstein et al.,

2000). On histological examination, infiltration of large amounts of immune cells at the site of

infection (termed granulomas) can be characteristically observed. However, these granulomas

persist as the immune cells present lack the ability to produce ROS and thus are unable to resolve

the infection. Given that the NOX2 NADPH oxidase is comprised of a number of components,

15

the loss of any one can result in the inability to make ROS. As such, CGD refers to a group of

hereditary diseases that result from a mutation in any one of the NOX2 NADPH oxidase

components, characterized by a defect in ROS production. As such, CGD patients are more

susceptible to catalase expressing pathogens. Since that time, much work has been done to

elucidate the contribution of ROS to innate immunity and the defense against invading

pathogens.

2.1.3 Types of ROS

Due to its highly reactive nature, superoxide can readily form a number of other compounds

(reviewed in (Bartosz, 2009; Lam et al., 2010; Lambeth, 2004)) (Figure 2.4). Superoxide can

react with nitric oxide to produce peroxynitrite, an even stronger oxidizing agent (Ramos et al.,

1995). The combination of two superoxide ions, catalyzed by superoxide dismutase (SOD),

results in the production of hydrogen peroxide (H2O2). H2O2 acts mainly upon thiol groups in

cysteine residues, leading to either oxidation (Cho et al., 2004; Rhee et al., 2000) or disulfide

bond formation (Biswas et al., 2006; Georgiou, 2002). H2O2 can also produce hypochloric acid

(HOCl) when combined with Cl- in a reaction catalyzed by myeloperoxidase. Both H2O2 and

HOCl have been shown to be present in sufficient concentrations in the phagosome to kill

microbes (Jiang et al., 1997; Nauseef, 2001). H2O2 can also interact with transition metal ions,

such as ferrous and ferric ions, to produce hydroxyl radicals (OH•), or with superoxides to

generate singlet oxygen (1O•). OH• in particular, while short lived, is the most highly oxidizing

member of the ROS family, reacting rapidly and non-discriminately with DNA, lipids and

proteins.

16

Figure 2.4. The generation of common ROS products in phagocytes. Reprinted with

permission from Springer Science + Business Media: Seminars in Immunopathology, Lam G et

al. The many roles of NOX2 NADPH oxidase-derived ROS in immunity, 32(4): 415-30

17

2.1.4 Functions of ROS

2.1.3.1 Microbicidal Activity of ROS

While superoxide is the main product of NOX2 NADPH oxidase, it remains controversial

whether it is the main antimicrobial compound. Some researchers have noted that superoxide is

relatively unreactive when compared to the rest of the ROS family and thus may not be sufficient

enough of a defensive strategy on its own (Reeves et al., 2003). Conversely, others have argued

that in the low pH environment of the phagosome, the majority of superoxide is protonated

(HO2•), becoming a much more reactive compound (Klebanoff, 2005).

While the direct antimicrobial effect of superoxides remains contentious, the indirect

antimicrobial effect of superoxides via its reactive products is certain. Due to its highly unstable

nature, superoxide can readily form a number of other compounds (reviewed in (Bartosz, 2009;

Lambeth, 2004)). Superoxide can react with nitric oxide to produce peroxynitrite, an even

stronger reducing agent (Ramos et al., 1995). The combination of two molecules of superoxide

ions, catalyzed by the activity of the enzyme, superoxide dismutase (SOD), results in the

production of hydrogen peroxide (H2O2). H2O2 acts mainly upon thiol groups in cysteine

residues, leading to either oxidation (Cho et al., 2004; Rhee et al., 2000) or disulfide bond

formation (Biswas et al., 2006; Georgiou, 2002). H2O2 can in turn produce hypochloric acid

(HOCl) when combined with Cl- in a reaction catalyzed by myeloperoxidase. Both H2O2 and

HOCl have been shown to be present in sufficient concentrations in the phagosome to kill

microbes (Jiang et al., 1997; Nauseef, 2001). H2O2 can also interact with transition metal ions,

such as ferrous and ferric ions, to produce hydroxyl radicals (OH•), or with superoxides to

generate singlet oxygen (1O•). OH• in particular, while short lived, is the most highly oxidizing

member of the ROS family, reacting rapidly and non-discriminatorily with DNA, lipids and

proteins.

2.1.3.2 ROS modulation of bacterial virulence factors

In addition to its direct microbicidal effects on pathogens, ROS can also arrest pathogen survival

and growth either via the inactivation of critical bacterial products or the modulation of the

phagosomal or extracellular environment. For example, leukotoxin, a key virulence factor of

18

Actinobacillus actinomycetemcomitans, has been shown to induce apoptosis in human

macrophages and neutrophils (Simpson et al., 1988; Tsai and Taichman, 1986).

Myeloperoxidase, through the production of HOCl, has been implicated in the oxidation and

subsequent inactivation of leukotoxin (Clark et al., 1986; Korostoff et al., 1998; Yamaguchi et

al., 2001). Pneumolysin, a vital virulence factor secreted by Streptococcus pneumoniae that is

required for survival in human neutrophils, has also been shown to be likewise oxidized by

HOCl. Pneumolysin is a pore-forming cytolysin whose function is regulated by the ability to

form disulfide bonds, without which proper oligomerization for pore formation cannot occur

(Geoffroy et al., 1981; Hotze et al., 2001). Myeloperoxidase was found to be effective in

inhibiting pneumolysin activity, presumably via the prevention of disulfide bond formation

(Clark, 1986). Another pore forming cytolysin, listeriolysin O (LLO), has been suggested to be

inhibited in much the same fashion (Myers et al., 2003; Singh et al., 2008). LLO is one of the

key virulence factors produced by Listeria monocytogenes that mediates bacterial survival and

growth in both human and murine macrophages (Portnoy et al., 1988; Schnupf and Portnoy,

2007). It has long been known that LLO requires a reducing environment for its activity

(Geoffroy et al., 1987; Portnoy et al., 1992). Recent work has further identified the particular

host protein - γ-interferon inducible lysosomal thiolreductase (GILT) - that is responsible for the

oxidation of LLO to mediate disulfide bond formation (Singh et al., 2008). It is therefore likely

that the creation of an oxidative environment by ROS can impede disulfide bond formation in

LLO. However, virulence factors are not the only reported microbial target of ROS as quorum

sensing by Staphylococcus aureus both in vitro and in vivo has also been shown to be affected

(Rothfork et al., 2004). Quorum sensing is the process by which bacteria modulate their behavior

once they “sense” their population has expanded above a certain threshold number. Rothfork and

colleagues demonstrated that HOCl and peroxynitrite inhibited the autoinducer peptide activity, a

critical player in quorum sensing, leading to an inability to upregulate virulence factor expression

once the bacteria has reached sufficient numbers.

19

2.1.3.3 Cellular Signaling Effects

ROS, in particular H2O2, have a wide range of reported signaling effects that will only be briefly

summarized for completeness (this topic is extensively reviewed by (Bedard and Krause, 2007;

Oakley et al., 2009)).

One mechanism of ROS effect on signaling is mediated through the modification of the cysteine

residue via one of two ways. First, the oxidation of the sulfur molecule to form sulfenic (Cys-

SOH), sulfinic (Cys-SO2H), or sulfonic (Cys-SO3H) acid can occur (Bindoli et al., 2008).

Second, the oxidation of cysteines can result in reactive thiols, leading to the formation of

disulfide bonds (Biswas et al., 2006; Georgiou, 2002). The significance of the oxidation of

cysteines is that ROS can influence cellular signaling via the inhibition of tyrosine phosphatases,

G proteins and certain ion channels (Cho et al., 2004; Rhee et al., 2000), as well as the activation

of kinases, such as mitogen-activated protein (MAP) kinases (Djordjevic et al., 2005). Another

major effect of ROS on signaling is the modulation of Ca2+ signaling. Oxidation of the ryanodine

receptor, which contains ROS sensitive cysteine residues (Liu and Pessah, 1994), results in the

release of intracellular Ca2+ stores (Favero et al., 1995; Kawakami and Okabe, 1998; Suzuki et

al., 1998). Similarly, ROS can also activate the inositol trisphosphate (IP3) receptor family Ca2+

release channels (Germano et al., 2004; Hu et al., 2002). An additional consequence of ROS

signaling is the activation of transcription factors. Stimulation of TLR results in the increase of

ROS signaling, leading to enhanced expression of NF-κB (Bubici et al., 2006). ROS signaling

also results in the expression of a number of genes involved in antioxidative and tumor

suppressive action, such as Nrf2 (Copple et al., 2008), forkhead box containing transcription

factor O (FoxO) (Tothova et al., 2007), and p53 (Liu et al., 2008).

2.1.3.4 Other functions of ROS

In addition to the roles mentioned, ROS have a plethora of functions in immunity. ROS have

been reported to participate in the formation of neutrophil extracellular traps (NETs), fibrous

meshworks comprised of DNA and protein expelled into the extracellular space for the purpose

of entrapping and killing pathogens (Brinkmann et al., 2004; Urban et al., 2006). ROS have also

been implicated in the modulation of the immune system to help create an environment that

20

allows for an efficient and effective immune response. As such, there has been a wealth of

research focused on the interaction between NOX2 NADPH oxidase-derived ROS and various

key members of both the innate and adaptive immune system. The results of these efforts paint a

complex picture of ROS function as ROS have been reported to participate in both pro-

inflammatory and anti-inflammatory signaling. Finally, recent work has demonstrated a role for

ROS in the induction of autophagy. The details of this ROS function are discussed later in the

section 2.1.3.3.1: ROS-mediated induction of autophagy.

2.1.4 Common techniques of ROS quantification

Given the transient nature and diversity of ROS, attempts at ROS quantification or visualization

can be challenging. A range of spectroscopic, chemiluminescent, electrophysiologic and other

imaging-based methods have been developed to address this issue (Eruslanov and Kusmartsev,

2010; Tarpey et al., 2004; Yeung et al., 2005). Of these, only a select few are useful for

quantification of intracellular ROS production. Here, I will highlight some of these commonly

employed techniques (summarized in Table 2.1).

2.1.4.1 Oxidizable fluorescent probes

In general, intracellular ROS production can be assessed using cell permeant probes that

fluoresce upon oxidization. This fluorescence can then be monitored via spectroscopy,

microscopy or flow cytometry. Fluorescein and calcein based probes are the most commonly

used indicators of ROS given their cell permeant nature.

Dihydroethidium (DHE), dihydrorhodamine 123 (DHR) and 2’-7’-dichlorodihydrofluorescein

(H2DCF) are among the most commonly employed fluorescein based probes for ROS

visualization and quantification. DHE is a blue, cell permeant probe that is specific for the

detection of superoxides (Tollefson et al., 2003). Upon oxidation by ROS, DHE is converted to

ethidium bromide, a red fluorescent compound, which then intercalates into cellular DNA

(Tollefson et al., 2003). However, DHE produces high background fluorescence even in resting

cells, rendering this probe relatively insensitive to small changes in ROS production. Thus, while

21

Name of compound Type of ROS detected Location of

ROS detected

Oxidizable fluorescent dyes

Dihydroethidium (DHE) Superoxide Intracellular

Dihydrorhodamine 123 (DHR) Superoxide, H2O2, HOCl and

peroxynitrite anions

Intracellular(Nakagawa

et al., 2004)

2’-7’-dichlorodihydrofluorescein (H2DCF) Superoxide, H2O2, peroxyl

radicals and peroxynitrite

anions

Intracellular

2-[6-(4’-hydroxy)phenoxy-3H-xanthen-3-on-

9-yl]benzoic acid (HPF)

Hydroxyl radicals and

peroxynitrite anions

Intracellular

2-[6-(4’-amino)phenoxy-3H-xanthen-3-on-9-

yl]benzoic acid (APF)

Hydroxyl radicals,

peroxynitrite anions and

HOCL

Intracellular

Dihydrocalcein (DHC) H2O2 Intracellular

Other oxidizable chemical compounds

Luminol (5-amino-2,3,-dihydro-1,4-

phthalazinedione)

Superoxide, H2O2, HOCL,

peroxyl radicals, and

peroxynitrite anion

Intracellular and

extracellular

Isoluminol (4-aminophthalhydrazide) Superoxide, H2O2, HOCL,

peroxyl radicals, and

peroxynitrite anion

Extracellular

Lucigenin Superoxide Extracellular

22

Nitroblue tetrazolium chloride (NBT) Superoxide Intracellular

Cerium Chloride (+ TEM) Superoxide, H2O2 Phagosomal

Table 2.1. Table of common compounds used for ROS detection.

23

DHE is highly specific for superoxides, it lacks the ability to distinguish the specific cellular

location of ROS generation and is not sensitive enough to detect low levels of ROS production.

In contrast, DHR can be used to detect a range of different types of ROS, including H2O2, HOCl

and peroxynitrite anions (Crow, 1997). DHR is a non-fluorescent probe that is converted into a

red fluorescent product called rhodamine 123 upon oxidation. Since both DHR and its coloured

product are freely diffuseable across membranes, this technique is likewise unable to distinguish

the location of ROS generation.

The limitations of DHE and DHR gave rise to a third class of fluorescent probes, H2DCF.

Similar to DHR, colourless H2DCF is converted to the green fluorescent product, 2’-7’-

dichlorofluorescein (DCF) only upon oxidation. The key advantage of DCF over rhodamine 123

is its limited permeability, thus allowing the product to be trapped inside the host cell upon

generation. The limitation of this technique, however, is that H2DCF is also poorly permeant. A

number of commercially available H2DCF probes have been generated, which bear chemical

modifications that render them more specific to intracellular ROS with improved rates of cellular

retention. Among the most widely used H2DCF derivatives is H2DCFDA, the chemically

reduced and acetylated version of the probe. Chemical reduction improves the probe’s sensitivity

to be oxidized, while acetylation ensures oxidation can only occur inside the host cell since the

probe remains non-fluorescent until the acetate groups have been removed by intracellular

esterases. Furthermore, carboxylation of H2DCFDA (carboxy-H2DCFDA) creates an additional

negative charge, improving retention of the probe. Addition of a thiol-reactive chloromethyl

group (CM- H2DCFDA) or an amine-reactive succinimidyl ester group (H2DCFDA, SE) allows

the probe to be covalently linked to intracellular structures, giving maximal cellular retention.

Depending on the modifications made, each H2DCFDA derivative has different sensitivities to

specific types of ROS. Carboxy-H2DCFDA is sensitive to H2O2, peroxyl radicals and

peroxynitrite anions, while CM-H2DCFDA is sensitive to all these ROS as well as hydroxyl

radicals. H2DCFDA, SE is specific for superoxides (Myhre et al., 2003).

Recently, new generations of fluorescein-based probes have been developed for the detection of

specific ROS. 2-[6-(4’-hydroxy)phenoxy-3H-xanthen-3-on-9-yl]benzoic acid (HPF) is specific

for the detection of hydroxyl radicals and peroxynitrite anions, while 2-[6-(4’-amino)phenoxy-

3H-xanthen-3-on-9-yl]benzoic acid (APF) is specific for both these ROS as well as hypochlorous

24

acid. The fluorescence of both probes is quenched. However, upon oxidation, quenching is

relieved and fluorescence can then be observed.

Of the calcein-based agents, dihydrocalcein (DHC) is one of the more popular dyes used in the

field. DHC is a non-fluorescent compound that becomes green-fluorescent calcein upon

oxidation. This probe is specific for H2O2 and has good cell permeability and retention

properties. However, one major drawback of this probe is that the fluorescence can be quenched

by metals (including iron, cobalt, nickel, and copper) at neutral pH (Keller et al., 2004).

2.1.4.1.1 Assessing fluorescence

Upon oxidization, all of the above-mentioned probes become fluorophores, compounds that have

the potential to fluoresce (or emit light) upon excitation by the proper wavelength. To assess

fluorescence, a light source is required to excite the fluorophore with light of the correct

wavelength, and the resulting emission of light from the excited fluorophore is detected with a

detector. Quantification of the amount of light generated by each cell gives an indication of the

concentration of fluorophore present, which gives an indication of the concentration of ROS

present to generate these fluorophores. However, it must be noted that one important limitation

of all these probes is that they are photo-oxidizable. In other words, the very process to assess

fluorescence will generate more fluorophore that can now fluoresce. Thus, repeated light-based

imaging will activate the probe, giving increased fluorescence simply due to the process of

imaging. This drawback may be limited by using low light conditions during fluorescence

microscopy; however, given the set-up time required for microscopic imaging, the effect of

photo-oxidation will be difficult to eliminate using this mode of detection. Certainly, exposure of

a stained sample to light results in a noticeable increase in fluorescence within seconds.

One viable alternative to fluorescence microscopy is flow cytometric analysis of fluorescence. In

flow cytometry, cells of increased fluorescence are defined or “gated” using H2DCF unstained

and stained controls. By comparing the histograms of fluorescence between the unstained and

stained controls, the population of cells that exhibit increased fluorescence in the stained control

is thus gated as ROS+. This gate can then be applied to subsequent samples in the same

experiment in order to give a percentage of cells that are ROS+ (out of a constant number of total

25

cells examined) that have increased fluorescence as defined using the stained control. Thus, each

single cell will be exposed to the excitation light once. Furthermore, the advantage of flow

cytometric analysis of fluorescence allows a population based assessment of fluorescence, giving

insight into the proportion of cells in a population that has increased ROS production.

Additionally, if there are different populations of cells that produce high versus low levels of

ROS, flow cytometric analysis will be capable of distinguishing them. While flow cytometry of

H2DCFDA stained cells allows for the detection of intracellular ROS, one key limitation of this

technique is the inability to indicate the specific cellular location of ROS generation.

2.1.4.2 Other oxidizable chemical compounds

Given the problems of photo-oxidation, a number of compounds have been developed to assess

ROS production via non-fluorescence-based methods. Among the most commonly employed

reagents are luminol, lucigenin, nitroblue tetrazolium chloride, and cerium chloride.

Luminol (5-amino-2,3,-dihydro-1,4-phthalazinedione) is a light coloured compound that

becomes chemiluminescent in the presence of ROS (Allen, 1986). The interaction of luminol

with ROS results in the formation of luminol dianion, a reaction that is catalyzed by the enzyme,

horseradish peroxidase (HRP) (Cormier and Prichard, 1968; Prichard and Cormier, 1968).

Reaction of luminol dianion with O2 produces unstable aminophthalate. The electrons in this

unstable compound undergo decay from an excited state to the ground state, emitting a photon of

light in the process. Thus, due to the 1:1 ratio of luminol and ROS to generate a luminol dianion,

quantification of light generated gives an indication of the concentration of ROS in the system.

Since luminol is cell permeable, both intracellular and extracellular ROS can be detected using

this reagent (Briheim et al., 1984; Dahlgren et al., 1989; Lock and Dahlgren, 1988). Luminol is

one of the most broad-spectrum ROS sensitive reagent, as it can detect the majority of ROS

including H2O2, hypochlorous acid, peroxyl radicals, peroxynitrite anion, and superoxides.

Derivatives of luminol, such as isoluminol, lack the ability to enter cells. As such, isoluminol is

specific for the detection of extracellular ROS generation (Lundqvist and Dahlgren, 1996). While

the generation of intracellular ROS can be extrapolated by comparing the results of the luminol

(intra- and extracellular ROS) and isoluminol (extracellular ROS) chemiluminescence assays,

direct detection of intracellular ROS production can be conducted by introducing superoxide

26

dismutase (SOD) to the luminol chemiluminescence assay (Chou et al., 2004; Lundqvist and

Dahlgren, 1996). SOD is cell impermeable and can eliminate extracellular superoxide

production. Given that both luminol and isoluminol assays are based on chemiluminescence, the

quantification of ROS production is assessed on a whole population basis using a luminometer.

Specific assessment of superoxide production can be conducted in a number of ways. Lucigenin

is a chemiluminescent compound commonly used to assess superoxide production. However,

since lucigenin is cell impermeable, it is useful in the quantification of extracellular superoxide

production (Liochev and Fridovich, 1998; Tarpey and Fridovich, 2001). To assess intracellular

production of superoxide, the nitroblue tetrazolium chloride (NBT) quantification assay can be

employed. NBT is a yellow, water-soluble compound comprised of two tetrazole moieties. Upon

interaction with superoxides, NBT is converted to a dark blue precipitate called formazan

(Beauchamp and Fridovich, 1971). NBT can freely diffuse into ROS-producing cells and

insoluble formazan precipitants are formed at the site of superoxide production. Differential

interference contrast imaging of fixed NBT treated cells allows for visualization of where

superoxide is being produced. Alternatively, cells can be treated with DMSO and KOH to allow

solublization of formazan precipitants. Subsequent spectrophotometric assessment at 570nm then

allows for the quantification of formazan precipitants, which indicates the concentration of total

superoxide in the system. Prior treatment of NBT treated cells with methanol will remove all

extracellular formazan precipitants allowing the detection and quantification of intracellular

formazan precipitants (protocol described in (Keith et al., 2009)).

2.1.4.3 Assessing phagosomal ROS by TEM

With the exception of NBT and the H2DCFDA probe, the techniques discussed above cannot

specifically distinguish phagosomal ROS from ROS generated elsewhere inside or outside of the

cell. The H2DCFDA,SE probe can be pre-conjugated to latex beads, zymosan particles or

bacteria via covalent insertion into amino groups. Labeled particles or microorganisms can then

be fed to macrophages. This technique has been successfully performed in the context of

studying superoxide production in phagosomes of neutrophils (Ryan et al., 1990). However, the

limitation of this technique is that the fluorescent probe is photo-oxidizable and therefore it is

technically challenging to limit oxidization during the labeling procedure.

27

Perhaps the most direct method of visualization of phagosomal ROS is via cerium chloride

treatment coupled with transmission electron microscopy. Much like NBT, cerium chloride also

forms an insoluble precipitant, cerium perhydroxide, in presence of ROS. Cerium perhydroxide

is a black compound that is deposited where it is formed, and is also easily visualized using

TEM. Thus, phagosomes containing bacteria or other particles can be assessed for ROS

production in the phagosomal compartment using cerium chloride. A number of studies have

successfully performed this technique to visualize ROS production in phagosomes containing

Salmonella enterica serovar Typhimurium or Burkholderia cenocepacia (Keith et al., 2009;

Vazquez-Torres et al., 2000). While this method is not ideal for quantifying the absolute amount

of intracellular ROS, it is a good method of assessing the percentage of bacteria containing

phagosomes that are producing ROS in the lumen of the phagosome.

2.1.4.4 Concluding remarks on ROS quantification or visualization

techniques

The development of ROS sensitive probes and chemical reagents that are sensitive to only one or

two ROS members have allowed for greater insight into the physiological roles of different ROS

members. It must be noted that one key limitation of all ROS sensing techniques is that they are

sensitive to oxidation by air. Thus, all dyes and reagents must be kept in air tight environments,

be it tightly lidded containers with moisture absorbing chips or nitrogen/argon rich

environments, to reduce spontaneous oxidation. Despite efforts of controlling oxidation, ROS

reagents are typically only be usable within months of opening. Thus, given the reactivity of

ROS to air, diversity of ROS and the specificities of each probe, multiple assays should be done

to gain a more complete and accurate picture of intracellular ROS production.

28

2.2 Autophagy

Autophagy is the controlled process of cellular self-digestion where cellular contents are

delivered to the lysosome for degradation in order to maintain cellular homeostasis (Levine and

Deretic, 2007). This process can be subdivided into three types – chaperone mediated autophagy

(CMA), microautophagy and macroautophagy (reviewed in (Mizushima et al., 2008)). With

respect to immunity, there has been a wealth of research demonstrating that macroautophagy,

hereafter referred to as autophagy, is a key component of the innate immune defense against

many pathogenic microorganisms, by removing them from the cytosol, limiting their escape

from phagosomes and promoting phagosome maturation (reviewed in (Deretic and Levine, 2009;

Hussey et al., 2009)). Autophagy is characterized by the presence of autophagosomes, double

membranous lamellar vesicles bearing the autophagy marker, microtubule-associated protein

light chain 3 (LC3) (reviewed in (Hussey et al., 2009)). LC3, or Atg8 in yeast, is a member of a

family of over thirty autophagy-related (Atg) proteins that act sequentially to mediate the

formation of autophagosomes. While the initial steps of autophagosome formation are currently

poorly understood, autophagy proteins are thought to recruit a crescent-shaped isolation

membrane (the source of which is currently unclear) that elongates around and captures the

cytoplasmic cargo, such as damaged organelles, cytosol, macromolecules or pathogens.

2.3 Autophagy core machinery

Autophagy is a process that is highly conserved from yeast to drosophila to man. Autophagy is

characterized by the formation of intracellular double/multi-membranous vesicles, called

autophagosomes. It is typically maintained at a low basal level under non-stress condition.

However, upon stress or stimulation, autophagic activity can be greatly induced (Huang and

Klionsky, 2007). Over thirty key components of the autophagy machinery encoded by

autophagy-related genes (ATG) function at different steps of this process (Klionsky et al., 2003),

which can be grouped into several specific stages: initiation (Figure 2.5 A), elongation (Figure

2.5 B), and maturation (Figure 2.5 C), which includes autophagosome trafficking, cargo

degradation and nutrient recycling.

29

2.3.1 Initiation

The Atg1 (homolog of mammalian ULK1) kinase complex (Atg1-Atg13-Atg17-Atg29-Atg31 in

yeast, and ULK1/2-Atg13-FIP200-Atg101 in mammals) is critical for autophagy induction

(Mizushima, 2010). The class III phosphatidylinositol 3-kinase (PI3K) complex comprising of

VPS34 (the PI3K), Vps15, Atg14 and Beclin 1/Atg6, functions at the stage of phagophore (the

autophagosome formation site or preautophagosomal structure (PAS) in yeast) initiation and

assembly (Funderburk et al., 2010) (Figure 2.5 A).

2.3.2 Elongation

Phagophore elongation involves two ubiquitin-like conjugation complexes that are essential for

expansion and finally autophagosome formation (Figure 2.5 B). One is the Atg12-Atg5-Atg16

complex, in which Atg12 is catalyzed by the E1-like enzyme Atg7 and E2-like enzyme Atg10,

and then conjugated to Atg5 (Mizushima et al., 1998). The resultant Atg12-Atg5 conjugate

further interacts with Atg16 and forms a complex that associates with autophagosomes

(Mizushima et al., 2003). The other ubiquitin-like system is the Atg8 (homolog of mammalian

LC3)-phosphatidylethonolamine (PE) conjugate (Ichimura et al., 2000). Under the activity of

Atg7, Atg3 (another E2-like enzyme) and Atg12-Atg5 complex (acts as an E3), Atg8/LC3 is

attached to the phospholipid PE and remains on the autophagosomal membrane throughout the

autophagy process (Hanada et al., 2007; Ichimura et al., 2000; Kabeya et al., 2000). Before its

conjugation, Atg8/LC3 needs to be processed by Atg4, a cysteine protease, to expose a glycine

residue that is prerequisite for PE conjugation (Kirisako et al., 2000). Atg4 also regulates the

deconjugation of Atg8/LC3-PE to recycle free Atg8/LC3 after the autophagosome is completely

formed (Kirisako et al., 2000).

31

Figure 2.5. Steps involved in the induction of autophagy. Upon starvation or other types of

stress, VPS34 and beclin 1 interaction as well as mTOR inhibition results in activation of

autophagy. This sets off a chain of events that includes (A) initiation, (B) elongation, and (C)

maturation of autophagy. (B) The elongation step of autophagy is regulated by two

ubiquitylation systems, Atg12 and LC3II pathways. Atg 12 is first conjugated to Atg7, and then

is transferred to Atg10. This allows for Atg12 conjugation to an Atg5 lysine residue. The

resulting Atg12-Atg5 complex is stabilized by Atg16. This trimeric complex enhances the

conversion of LC3I to LC3II. Atg4 mediates in the recycling of LC3II to LC3I. (C) The

autophagosome then fuses with the lysosome, resulting in the degradation of the autophagosome

and its contents. Reprint by permission from Macmillian Publishers Ltd: Nat. Rev. Immunol.

Levine et al. Unveiling the roles of autophagy in innate and adaptive immunity. 7:5 (767-77),

2007.

32

2.3.1 Maturation: Non-selective vs selective autophagy

Autophagy can be classified into non-selective bulk autophagy and selective autophagy that

targets specific cargoes. Though starvation-induced bulk autophagy has long been thought to be

nonspecific, recent proteomic studies suggested a highly regulated and ordered manner of

organelle degradation by autophagy under amino acid starvation condition (Kristensen et al.,

2008). It was shown that at early time points after starvation, cytosolic proteins and proteasomes

were degraded by autophagy, and ribosomes were degraded in lysosomes at later time points

(about 12 hours post-starvation). Organelle degradation started even later (about 24 hours after

starvation) (Kristensen et al., 2008).

Selective autophagy can be subdivided into many different types, depending on the nature of the

cargo being degraded. Selective targeting and degradation of peroxisome (Dunn et al., 2005),

mitochondria (Kim et al., 2007) and endoplasmic reticulum (ER) (Bernales et al., 2006) by

autophagy are called pexophagy, mitophagy and reticulophagy, respectively. Furthermore,

misfolded protein aggregates, such as Huntingtin protein aggregates, can be selectively degraded

by autophagy (named aggrephagy) (Rubinsztein, 2006). Finally, autophagy is able to target

invaded viruses, bacteria and parasites by a process called xenophagy (Orvedahl and Levine,

2009).

The selective cargo recognition depends on cargo receptor proteins that can be recognized by the

autophagy machinery. Mitochondria-localized protein Nix (also known as Bcl2/E1B 19 kDa-

interacting protein 3 (BNIP3)-like protein (BNIP3L)) was recently shown to be a receptor on

mitochondria, mediating mitophagy through interaction with LC3 via the N-terminal LC3

interacting region (Novak et al., 2010). Yeast Pichia pastoris (P. pastoris), a model organism for

pexophagy studies, expresses a receptor protein PpAtg30 on peroxisomal membrane and

interacts with PpAtg11 for autophagy recognition (Farre et al., 2008). Moreover, several recent

studies have shown that the ubiquitin (both mono- and poly-ubiquitin)-binding protein

p62/SQSTM1 is a cargo receptor that links autophagy machinery to different cargo targets,

including polyubiquitinated protein aggregates (Pankiv et al., 2007), monoubiquitin-tagged

peroxisomes (Kim et al., 2008) and bacterial pathogens, such as Salmonella enterica serovar

Typhimurium (S. Typhimurium) (Zheng et al., 2009), Shigella flexneri (S.flexneri) (Dupont et

al., 2009) and Listeria monocytogenes (L. monocytogenes) (Yoshikawa et al., 2009).

33

2.4 Autophagy and Intracellular Pathogens

Autophagy has been extensively studied for its role in immunity. It is now believed that

autophagy is an important innate immune defense mechanism, and plays multiple roles in

adaptive immunity (extensively reviewed in (Deretic, 2009; Orvedahl and Levine, 2009; Virgin

and Levine, 2009)). Autophagy can limit the intracellular growth of invading pathogens by

degrading them through autophagosome-lysosome pathway. The mechanism of LC3 targeting in

a wide range of intracellular pathogens involves, at least in part, protein ubiquitination and

p62/SQSTM1 adaptor protein recruitment. P62/SQSTM1 can interact with both LC3 and

ubiquitinated cargo through an LC3 interacting region and a C-terminal ubiquitin-associated

(UBA) domain, respectively (Ichimura et al., 2008; Pankiv et al., 2007). Thus, p62/SQSTMI acts

in an adapter capacity to bridge the interaction between the ubiquitinated bacteria and LC3.

Recently, additional players involved in LC3 targeting of pathogens have surfaced. Kirkin and

colleagues reported that another ubiquitin-binding protein NBR1 (Neighbor of BRCA1 gene 1),

which shares similar domain organizations to p62/SQSTM1 and directly interacts with

p62/SQSTM1, also acts as a receptor for selective autophagy targeting of ubiquitinated protein

aggregates (Kirkin et al., 2009a). However NBR1 does not play a major role in autophagy of S.

Typhimurium (Zheng et al., 2009), suggesting that NBR1 and p62 may have different cargo

preferences. Another adaptor protein NDP52 (nuclear dot protein 52 kDa), has also been recently

shown to be involved in autophagy targeting of S. Typhimurium by recognizing the ubiquitin

coat on the bacteria and bringing them to the autophagosomes via binding to LC3 (Thurston et

al., 2009). A focused analysis of the two adaptor proteins during S. Typhimurium infection

revealed that p62/SQSTM1 and NDP52 bind to distinct microdomains on the bacteria and that

the loss of either adaptors results in a decrease in autophagy targeting to the bacteria (Cemma et

al., 2010). These observations suggest that while two adaptor proteins appears to be involved in

autophagy targeting of S. Typhimurium, they may play different roles in the autophagy pathway

that is currently unappreciated. Furthermore, examination of LC3-interacting proteins may reveal

additional receptors that are responsible for selective autophagy cargo recognition.

While a wide range of bacteria, viruses and parasites have been shown to be targets of

autophagy, some have evolved strategies to either evade autophagy recognition or utilize

autophagy for their survival and replication (Hussey et al., 2009; Orvedahl and Levine, 2009)

34

(listed in Table 2.3). Autophagy also allows the detection of certain single-stranded RNA

(ssRNA) viruses through the transportation of viral replication intermediates into acidic

endosomes for recognition by host pathogen pattern recognition receptor (PRR), Toll-like

receptor 7 (TLR7) (Lee et al., 2007). Moreover, autophagy is involved in pro-inflammatory

cytokine production via either positive or negative effects in different cell types. In the above

example TLR7 activation, autophagy is required for viral infection-initiated type I interferon

(IFN) production in plasmacytoid dendritic cells (Lee et al., 2007). However other studies using

ATG5 knockout mouse embryonic fibroblasts (MEFs) for ssRNA virus infection found enhanced

production of type I IFN in these autophagy-deficient cells, suggesting a suppression of cytokine

release by autophagy (Jounai et al., 2007; Tal et al., 2009). In addition, studies of mice

expressing inefficient level or a truncated Atg16L1 showed upregulated transcription of

adipocytokine genes, or in the latter study elevated production of interleukin (IL)-1β and IL-18

when mice were stimulated with bacterial endotoxin, lipopolysaccharide (LPS) (Cadwell et al.,

2008; Saitoh et al., 2008). Therefore autophagy has paradoxical roles in both pro- and anti-

inflammatory responses.

Autophagy targeting mechanism Bacterial evasion strategy Reference

Bacteria

Group A Streptococcus

Cytosolic Streptococci is targeted by

LC3; lack of Atg5 results in delayed

bacterial clearance

(Nakagaw

a et al.,

2004)

L. monocytogenes

PGRP-LE recognition of bacterial

peptidoglycan induces autophagy;

bacteria targeted by LC3 at 8 h p.i.

when treated with chloramphenicol;

ActA prevents ubiquitination

and LC3 targeting of

bacteria; PI-PLC and PC-

PLC may play an unknown

(Birmingham et

al., 2007a; Py et

al., 2007; Rich

et al., 2003;

35

peak of LC3 targeting at 1 h p.i.;

autophagy limits bacterial replication as

L. monocytogenes infection of mice

with ATG5-/- macrophages results in

higher bacterial load over control

role in autophagy evasion as

bacteria deficient in the

PLCs exhibit increased

growth in ATG5-/- MEFs

Yano et al.,

2008;

Yoshikawa et

al., 2009; Zhao

et al., 2008)

Shigella flexneri

Atg5 binds to VirG Bacterial IcsB masks VirG to

prevent recognition and

LC3 targeting

(Ogawa et

al., 2005)

Salmonella enterica

SPI-1 TTSS-damaged Salmonella-

containing vacuoles are targeted by

autophagy; lack of Atg5 results in

increased bacterial survival

(Birmingham et

al., 2006;

Kageyama et

al., 2011)

Francisella tulerensis

Cytosolic bacteria targeted by autophagy (Checroun et

al., 2006)

Helicobacter pylori

Autophagy limits the stability and toxic

effect of VacA; autophagy inhibition

increases bacterial replication;

autophagy induction decreases

replication

(Terebiznik et

al., 2009)

Vibrio cholerae

Autophagy targets bacterial VCC (Gutierrez et

al., 2004)

36

Burkholderia pseudomallei

Cytosolic bacteria targeted by autophagy;

autophagy limits bacterial replication

Bacterial BopA reduce LC3

targeting

(Cullinane et

al., 2008)

Mycobacterium tuberculosis

LC3 targeting to mycobacteria containing

phagosomes, limiting survival (Gutierrez et al.,

2004)

Parasite

Leishmania major

Differentiation into infective

form requires autophgy

induction

(Besteiro et

al., 2006)

Toxoplasma gondii

LC3 targeting to Toxoplasma containing

compartments to limit survival

(Andrade et

al., 2006)

Trypanosoma cruzi

Encode orthologs of Atg8

conjugation system;

autophagy induction

needed for survival

(Alvarez et

al., 2008)

Viruses

Poliovirus

Viral replication inside

LC3+ double membrane

(Taylor and

Kirkegaard,

37

vesicles; replication

dependent on autophagy

2007)

Human immunodeficiency virus (HIV)

Autophagy inhibited but

autophagy components

needed for replication

(Zhou and

Spector,

2008)

Hepatitis C (HCV)

Autophagy induction and

Atg5 interaction with viral

NS5B factor allows viral

replication

(Ait-

Goughoulte

et al., 2008;

Guevin et al.,

2010)

Sindbis virus

Autophagy restricts viral replication (Liang et al.,

2006;

Orvedahl et

al., 2010)

Herpesvirus (HSV-1)

Viral ICP34.5 factor inhibits

autophagy by targeting

PKR and beclin1

(Orvedahl et

al., 2007;

Talloczy et

al., 2006)

Table 2.3. Summary of known interactions between pathogens and autophagy

38

2.3.2.1 Signals for Autophagy Induction

While autophagy has been identified as a host innate immune defense, the mechanisms of

autophagy signaling remain largely unknown. Autophagy regulation in immunity is summarized

in Figure 2.6. Pathogen detection receptors, including TLRs and NOD-like receptors (NLRs), are

crucial host immune factors that sense extra- or intracellular microbes and initiate downstream

signaling pathways to activate antimicrobial immune responses (Delbridge and O'Riordan,

2007). A number of TLRs have been reported to stimulate autophagy in murine and human

phagocytes (Delgado et al., 2008; Xu et al., 2007). Furthermore, agonist-stimulated receptor

(such as TLRs and FcγR) signaling during phagocytosis is shown to induce autophagy targeting

of the phagosome and promote phagosomal maturation (Huang et al., 2009; Sanjuan et al.,

2007).

Besides TLRs, NLR members, such as Naip5, Ipaf, PGRP-LE and NOD2, can regulate

autophagy as well. Naip5 is involved in promoting efficient autophagy of Legionella

pneumophila (L. pneumophila) (Amer and Swanson, 2005). Ipaf has been shown to negatively

regulate autophagy of S. flexneri in macrophages (Suzuki et al., 2007). PGRP-LE is crucial for

autophagy induction and limiting bacterial growth during L. monocytogenes infection of

Drosophila (Yano et al., 2008). NOD2 mediates autophagy induction and targeting of S. flexneri

(Travassos et al., 2010). Autophagy can also be upregulated by IFN-γ and its downstream

effector, Irgm1 (or human IRGM), to eliminate M. tuberculosis and Toxoplasma gondii

(Orvedahl and Levine, 2009). Conversely, autophagy can also be negatively regulated by NF-κB

during tumor necrosis factor (TNF)-α-stimulation (Djavaheri-Mergny et al., 2006). Viral

infection can induce autophagy via the cytosolic double-stranded viral RNA-sensing kinase

PKR, activating the transcription factor eIF2α (Talloczy et al., 2002). In addition, viruses can

inhibit autophagy by modulating autophagy proteins. An α-herpesvirus, HSV-1, produces the

neurovirulence factor, ICP34.5, to directly target Beclin 1 and block autophagy (Orvedahl et al.,

2007).

39

Figure 2.6. Regulation of autophagy by immune signals in response to invading pathogens.

Induction of autophagy is observed upon activation by Naip5 by L. pneumophila, Ipaf or

Atg16L1 by S. flexneri, PGRP-LE by L. monocytogenes. Some viruses can induce autophagy via

elF2α but others can inhibit autophagy via inhibition of Beclin1. Host immune modulators, such

as IFN-γ, TNF-α and ROS can also induce autophagy. Reprinted by permission from Mary Ann

Liebert, Inc: Antioxidants Redox Signaling, Huang J et al. Autophagy Signaling Through

Reactive Oxygen Species. 14(11): 2215-31.

40

2.3.2.1.1 ROS-mediated induction of Autophagy

Mounting oxidative stress can result in damaged organelles, proteins, and DNA and the

clearance of which is one of the main functions of autophagy in a number of cell types (Kaushik

and Cuervo, 2006; Lemasters, 2005; Xiong et al., 2007) (reviewed in (Kiffin et al., 2006)). If

proper clearance cannot take place, increasing ROS levels would then lead to autophagic cell

death (Chen et al., 2008), as seen in the context of TNF-α induced autophagic cell death in

Ewing sarcoma cells (Chen et al., 2008) or LPS induced autophagic macrophage cell death (Xu

et al., 2006). If autophagy cannot be properly activated, cellular dysregulation in diseases of age,

such as Alzheimer’s (Cataldo et al., 1996), or uncontrolled inflammation, such as Diabetes

mellitus (Sooparb et al., 2004), would result. However, the mechanism by which ROS

production leads to activation of autophagy is not clear.

The first direct link between ROS and autophagy was provided in context of starvation-induced

autophagy and mitochondrial-derived ROS (Scherz-Shouval and Elazar, 2007; Scherz-Shouval et

al., 2007a; Scherz-Shouval et al., 2007b). In response to nutrient and serum starvation, it was

found that a number of cell lines – HeLa, HEK293, CHO and MEFs – greatly induced both

superoxide and H2O2 production in a class III PI3K-dependant manner when compared to

control. Furthermore, there was more ROS production in wild type MEFs over MEFs without

Atg5 (Scherz-Shouval et al., 2007b). These two observations combined indicate that PI3K is

somehow upstream of ROS production, leading to autophagy. ROS production was proposed to

inhibit the redox sensitive cysteine protease, Atg4 - specifically the cysteine residue residing in

the active site of the enzyme. Atg4 is one of the autophagy proteins involved in the regulation of

LC3 phosphatidylethanolamine (PE) lipidation (LC3-II) and delipidation (LC3-I) (Nakatogawa

et al., 2007), a post-translational event essential for autophagy to occur. When Atg4 encounters

H2O2, Cys81 becomes oxidized in a reversible reaction, rendering the enzymatic activity null

(Scherz-Shouval et al., 2007b). The lack of Atg4 activity is thought to allow LC3 to remain

lipidated and localized to the autophagosomal membrane, leading to autophagy.

The link between mitochondrial ROS and starvation-induced autophagy was further confirmed

by Chen and colleagues (Chen et al., 2009). Superoxides but not H2O2 were found to be the

critical ROS member for the induction of autophagy in HEK293, human glioma cell line U87

and HeLa cells. This study found that pharmacological inhibition or siRNA silencing of SOD,

41

which converts superoxide to H2O2, results in increased autophagy when compared to WT

controls. Conversely, the overexpression of SOD resulted in a decrease in superoxide levels and

increase in H2O2, leading to decreased autophagosome formation. Superoxides were also found

to be important for the induction of autophagy in the context of muscular atrophy. It was found

that transgenic mice with a SOD deletion exhibit increased numbers of autophagosomes

(Dobrowolny et al., 2008). Since the superoxides are quickly converted to H2O2 and other

reactive products, there are limitations in ROS assays that make it hard to distinguish one

particular type from another. Furthermore, ROS products are linked together in a web of

reactions (Figure 2.4), thus it becomes difficult to take away any one enzyme without affecting

any other products in a compensatory manner. For example, by knocking out SOD, not only is

the level of H2O2 affected, but also the levels of HOCl, peroxynitrite and OH•. The limitation of

detection techniques available, coupled with the transient nature of ROS members makes it

challenging to pinpoint specific triggers of autophagy. Regardless of the type of ROS, however,

it is clear that mitochondrial ROS are indeed involved in the activation of starvation-induced

autophagy.

Further evidence has surfaced relating ROS to starvation-induced autophagy. One of the

mechanisms of glycolysis downregulation is mediated through TIGAR, TP53-induced glycolysis

and apoptosis regulator. An indirect consequence of TIGAR activity on glucose metabolism is

the decrease in global intracellular ROS levels (Bensaad et al., 2006). TIGAR activity was found

to block autophagosome formation in the human osteosarcoma U2OS cells, even under

starvation conditions (Bensaad et al., 2009). This inhibition of autophagy was correlated with

lowered ROS levels. Adding TIGAR exogenously results in an even greater decrease in ROS and

autophagy. Thus, TIGAR is potentially expressed only under nutrient rich conditions to inhibit

autophagy activation via the inhibition of ROS.

Unlike starvation-induced autophagy, the mechanism by which antimicrobial autophagy is

activated is less clear. The first detection of invading pathogens occurs via the binding of pattern

recognition receptors, such as TLRs, to its pathogen associated molecular patterns (PAMPs),

resulting in the activation of autophagy in both murine RAW 264.7 macrophage cell line and

primary bone marrow derived macrophages (Delgado et al., 2008; Sanjuan et al., 2009; Xu et al.,

2008) (reviewed by (Delgado and Deretic, 2009)). Nucleotide-binding oligomerization domain

(NOD) and NOD-like receptor (NLR) signaling has also been shown to regulate autophagy

42

(Suzuki and Nunez, 2008; Travassos et al., 2010). Upon the activation of TLR, FcγR,

complement receptors or others, phagocytosis of microbes is initiated and autophagy is triggered

(Park, 2003; Sanjuan et al., 2007). Seminal work done by Sanjuan and colleagues showed that

the activation of TLR upon phagocytosis results in the recruitment of LC3 to the phagosome

(Sanjuan et al., 2007) forming a single membrane LC3+ phagosome. The induction of autophagy

leads to phagosomal maturation and subsequent degradation of the phagosomal content in the

autophagolysosome. TLR signaling has also been shown to be upstream of NOX2 NADPH

oxidase assembly and activation (Laroux et al., 2005; Quinn and Gauss, 2004). Given that TLR

activation leads to both NOX2 NADPH oxidase activation and autophagy, we hypothesized that

ROS might also play a role in the activation of antimicrobial autophagy. To this end, we first

examined if TLR or FcγR activated autophagy is mediated through ROS production. Using bone

marrow derived neutrophils (BMDN), we challenged cells with IgG-coated latex beads or dead

yeast cells, zymosan, and found that LC3 was recruited to the phagosome (Huang et al., 2009).

This recruitment was further examined using either pharmacological agents to block/neutralize

ROS or comparing wild type with NOX2 NADPH oxidase deficient BMDN with respect to LC3

recruitment upon stimulation. We found that in absence of NOX2 NADPH oxidase activity, LC3

recruitment to the phagosome was impaired. An analysis of the purified phagosomes revealed

enhanced localization of Atg12 (an essential autophagy protein) and LC3-II on IgG-coated latex

bead containing phagosome and this recruitment was inhibited by the addition of the NOX2

NADPH oxidase inhibitor, DPI. Thus, ROS was found to be critical for LC3 targeting to latex

bead containing phagosomes (Figure 2.7). We further established that other cell types (epithelial

cells) also employ member(s) of the NOX family for antimicrobial autophagy, suggesting that

the NOX family may be a general mechanism of antimicrobial defense in a range of cell types.

Using p22phox siRNA silencing in the human embryonic epithelial cells, Henle-407, we observed

that the autophagy of Salmonella enterica serovar Typhimurium decreased. Recently, another

independent study confirmed some of our observations in human neutrophils, suggesting that

NOX2 NADPH oxidase-derived ROS induction of autophagy is not a murine only phenomenon

(Mitroulis et al., 2010).

44

Figure 2.7. The known pathways that are dependent on ROS induction of autophagy. (A)

Phagocytosis of microbes results in the recruitment of NOX2 NADPH oxidase complex to the

phagosome. The presence of ROS signals the induction of autophagy through an unknown

mechanism, resulting in the recruitment of autophagy protein, LC3, to the phagosome, where

LC3 is then conjugated to PE. Subsequent fusion with the lysosome allows for microbial

degradation in the autophagolysosome. (B) Upon nutrient starvation, ROS are generated in the

mitochondria as a byproduct of electron transport chain activity. Loss of mitochondrial integrity

results in the release of ROS into the cytosol, triggering autophagy. The damaged mitochondria

and partial cytoplasm are then contained in an autophagosome that is likewise marked for

lysosomal fusion and degradation. Reprinted by permission from Springer Science + Business

Media: Seminars in Immunopathology, Lam G et al. The many roles of NOX2 NADPH oxidase-

derived ROS in immunity, 32(4): 415-30

45

2.3.2.1.2 Diacylglycerol-mediated induction of Autophagy

Diacylglycerol (DAG) was also found to be a key signal for mediating autophagic targeting to

Salmonella enterica serovar Typhimurium containing phagosomes at 1 h post infection (p.i.)

(Shahnazari et al. 2010). Inhibition of DAG production by host DAG-producing enzymes,

phospholipase D and phosphatidic acid phosphatase, results in a reduction in LC3 targeting to S.

Typhimurium. DAG mediated autophagy targeting to the bacteria was further found to be

dependent on the activity of the downstream signaling kinase, protein kinase C-delta (PKCδ)

(Shahnazari et al. 2010). How PKCδ mediates autophagic targeting remains to be understood.

2.3.2.1.3 Ubiquitinated protein-mediated induction of Autophagy

Finally, ubiquitinated proteins can also be a signal for autophagy and has been extensively

reviewed (Dupont et al., 2010; Kirkin et al., 2009b; Randow, 2011). Ubiquitin (Ub) is a small 76

amino acid protein that can be conjugated to proteins as single ubiquitin subunits or poly

ubiquitin chains. Ubiquitinated S. Typhimurium is targeted by autophagy (Birmingham et al.,

2006) via the recruitment of adaptor proteins, p62 (Cemma et al., 2010; Zheng et al., 2009) and

NDP52 (Cemma et al., 2010; von Muhlinen et al., 2010). L. monocytogenes mutants that lack the

ability to recruit actin, ΔActA, (see section on L. monocytogenes below) can also be

ubiquitinated in the cytosol and subsequently targeted by autophagy (Yoshikawa et al., 2009).

Finally, the remnants of the phagosome left behind after Shigella flexneri has escaped into the

cytosol can also be ubiquitinated and targeted for autophagic clearance (Dupont et al., 2009).

It remains controversial how LC3 is targeted to pathogens. Autophagic targeting may be the

result of either the fusion of LC3-positive (LC3+) vesicles, which elongates to form a double

membrane compartment or direct LC3 conjugation to a single membrane compartment (Huang

and Brumell, 2009; Sanjuan et al., 2007). Work done by Rich and colleagues indicates that

cytosolic ΔActA L. monocytogenes mutants were found in LC3+, endoplasmic reticulum (ER)

markers bearing, double membrane compartments in presence of the bacteristatic agent,

chloramphenicol. This observation suggests that cytosolic autophagic targeting of mutant L.

46

monocytogenes is mediated by the fusion of small LC3+ vesicles from the ER (Rich et al., 2003).

Similarly, a recent study using correlative light microscopy-electron microscopy (CLEM)

imaging indicates that autophagy targeted Salmonella containing vacuoles are double-membrane

structures (Kageyama et al., 2011). One possible explanation for these different observations

may be that the mechanism of LC3 recruitment is different depending on where the pathogen is

located. LC3+ vesicles may target cytosolic bacteria, while LC3 may be directly conjugated to

phagosomes containing bacteria. Another possible explanation is that the mechanism of LC3

recruitment is pathogen specific (ie: Gram-positive vs. Gram-negative). Thus, many questions

remain surrounding the mechanism of autophagic targeting to intracellular pathogens.

2.3.3 Common techniques used to study autophagy

Given the diversity of the field of autophagy, there is a vast range of techniques that are used to

study the different types of autophagy. However, there are some key methods that are universal

in the field to examine autophagic flux. Here, I will discuss these cornerstone techniques.

As mentioned, the concept of autophagy flux has evolved as the field of study matured. An

observed increase in LC3-I to LC3-II conversion is no longer enough to make the argument that

autophagy is increased. This is due to the fact that an accumulation of LC3-II may be due to a

block in autophagosome clearance (Tanida et al., 2005), as well as an increase in the rate of

autophagy. Thus, a number of other methods have been described to assess mammalian

autophagy and autophagic flux (extensively reviewed in (Klionsky et al., 2008; Klionsky et al.,

2007; Mizushima, 2004; Mizushima et al., 2010)).

2.3.3.1 Techniques used to quantify changes in autophagy

Currently, changes in LC3-II is most commonly assessed using western blot analysis, flow

cytometry, LC3 colocalization to particle containing phagosomes, or increase in LC3+ puncta.

LC3-I runs at 19KDa while LC3-II runs at 17KDa. As such, higher percentage SDS-PAGE gels

will have the power to resolve these two distinct bands (Figure 2.8 A). However, it must be noted

that LC3-I is often difficult to resolve for reasons that are currently unknown. Thus, LC3-II

47

remains the reliable indicator of autophagy. LC3-II levels have been described to increase upon

autophagic induction but decrease over time (> 2h) as a result of increased turnover (Mizushima

and Yoshimori, 2007). Changes in LC3 expression can also be assessed using flow cytometry.

Endogenous LC3 can be stained using immunofluorescent antibodies and increased fluorescence

can be assessed via flow cytometry on a population basis. Alternatively, increased cellular levels

of LC3 can also be visualized by immunofluorescence imaging or transmission electron

microscopy. Increases in autophagy result in increases in LC3+ punta (Figure 2.8 B; white

arrowheads). Thus, quantification of the changes in numbers of LC3+ punta per cell can give an

indication of changes in autophagy. Lastly, in the field of selective autophagy, LC3 targeting to

phagosomes containing foreign particles or pathogens can likewise be used to assess changes in

autophagic targeting. Quantification of the percentage of LC3+ particle containing phagosomes

(Figure 2.8 C; white arrowheads) over total number of particle containing phagosomes allows an

assessment of autophagic targeting to a selective target.

Autophagosomes are defined strictly as LC3+ double membrane compartments. However, LC3+

single membrane phagosomes have also been reported. Sanjuan and colleagues have previously

demonstrated that TLR or FCγR - mediated phagocytosis of IgG-coated latex beads or zymosan

particles results in formation of LC3+ single membrane particle containing phagosomes (Sanjuan

et al., 2007). As such, these structures, deemed LC3-associated phagosomes, must be

distinguished from autophagosomes. To determine if a LC3+ structure of interest is double or

single membraned, CLEM is currently the gold standard for this assessment. CLEM is a process

where LC3+ structures identified in immunofluorescence microscopy is matched with the

corresponding electron micrograph to determine the number of membranes that are on the LC3+

structures. Other techniques, such as electron microscopy of LC3 immunogold labeling, can also

be used.

2.3.3.2 Techniques used to assess flux

In tandem with studies of autophagy, techniques for assessing autophagic flux are often

employed to determine if increases in autophagy is due to increased rate of autophagy or if the

turnover is blocked, resulting in an accumulation of LC3-II. The rate of protein turnover can be

48

assessed by examining levels of p62 by western blot analysis. p62 (also known as

SQSTM1/sequestome 1) is an adaptor protein that is recruited to autophagosomes through

binding to LC3 and is degraded by autophagy (Bjorkoy et al., 2005). An increase in LC3-II that

is accompanied with a decrease in p62 correlates well with an increase in the rate of autophagy.

Conversely, an increase in both LC3-II and p62 correlates with a block in autophagic flux or

clearance (Mizushima et al., 2010). However, it must be noted that p62 is not solely degraded in

the autophagic pathway and thus changes in p62 levels may not solely reflect changes in

autophagy (Bjorkoy et al., 2005). To address this limitation, other assays have recently been

developed to assess autophagic flux.

An alternative method to measure autophagic flux is that of the RFP-GFP-LC3 tandem probe

(Kimura et al., 2007). This probe was designed based on the concept of differences in lysosomal

quenching of GFP versus RFP fluorescence. In the acidic environment of the lysosome, GFP

fluorescence is quenched while RFP fluorescence is not. Thus, under conditions of proper

autophagosome delivery to the lysosome, cells expressing the RFP-GFP-LC3 will display

predominantly red puncta since GFP, but not RFP fluorescence is quenched in the lysosome.

However, if a blockage in autophagosome delivery to the lysosome, cells would display

predominantly yellow puncta since both GFP and RFP fluorescence will be observed in the

autophagosomes prior to delivery to the lysosome. Thus, autophagic flux can be assessed by

quantifying the ratio of red to yellow puncta. Cells with increased rate of autophagy will display

a higher red to yellow puncta ratio while cells experiencing a block in autophagic flux will

display a lower red to yellow puncta ratio. One limitation is that this technique is based on the

assumption that cells retain normal rates of acidification and lysosome enzyme activity. If these

parameters are altered, LC3 puntae may remain yellow for reasons other than a block in

autophagic flux. As such, this assay does not give an indication of when degradation in the

lysosome is occurring. One technique that can address this limitation is the GFP-LC3 cleavage

assay. Upon delivery to the lysosome, the GFP-LC3 fusion protein will be partially degraded,

librating GFP (Gao et al., 2008). Thus, following free GFP accumulation via immunoblotting can

give an indication of lysosome activity upon fusion with autophagosomes. While this technique

has been commonly used in the study of yeast autophagy, its use in mammalian cells remains

limited.

49

Figure 2.8. Examples of methods to assess autophagy. (A) Western blot analysis of lysates

generated from RAW 264.7 macrophage infected with wild type L. monocytogenes for the

indicated time points p.i. and probed for LC3. Bands roughly at 19 and 17 KDa, correponding to

LC3-I or LC3-II, can be resolved. (B) Henle 207 cells expressing GFP-LC3 with or without S.

typhimurium infection. White arrowheads indicate LC3 puncta. (C) Raw 264.7 macrophages

expressing GFP-LC3 infected with L. monocytogenes. White arrowheads indicate LC3 targeting

to the intracellular bacteria.

WT

60’

WT

15’

WT

120’

Uni

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50

2.4 Other Defenses

In addition to ROS and autophagy, there are a number of other key host defenses that mediate the

microbicidal activity of the phagosome, including low phagosomal pH, reactive nitrogen species

(RNS), antimicrobial peptides and defensins. These processes were reviewed by Flannagan and

colleagues (Flannagan et al., 2009) and will only be summarized here for completeness.

Phagosomal acidification by the vacuolar ATPase is a vital step in phagosomal maturation. Not

only can a low pH environment (pH 4-5) be directly bacteriocidal, but can also allow the activity

of a number of pH regulated degradative enzymes. These enzymes include cathepsins,

hydrolyases and other proteases. RNS are produced in macrophages by the inducible nitric oxide

synthase (NOS) in response to inflammation (Fang, 2004). The main NOS product is nitric

oxide. Like superoxide, nitric oxide can be converted into a number of other RNS, all of which

are directly bacteriocidal. While RNS are produced in the cytosol, they can diffuse across

membranes to the sites of infection (Webb et al., 2001). Together, ROS and RNS can act

synergistically to damage proteins, lipids and microbial DNA (Fang, 2004). Lastly, a wide range

of soluble factors such as antimicrobial peptides, bacteriocidal granules, defensins, lysozymes,

lipases, proteases and peptidases target specific bacterial components and contribute to direct

bacterial killing in the phagosome (Flannagan et al., 2009).

3 Bacterial Evasion Strategies in Response to Host

Defenses

A number of pathogens have evolved mechanisms to evade degradation by phagocytes.

Phagocytosis has been described as a two-step event requiring pathogen uptake and clearance

and several pathogens have devised mechanisms to resist both these steps (Flannagan et al.,

2009; Kumar and Valdivia, 2009; Steinberg and Grinstein, 2008; Woolard and Frelinger, 2008).

Pathogens can hinder their uptake by phagocytes in a number of ways. Some bacteria secrete

polysaccharides, which form a slippery capsule or coating around the bacteria, making

phagocytosis difficult. Other pathogens, such as Pseudomonas aeruginosa, Escherichia coli,

Staphylococcus aureus, express factors that cleave or inactivate host proteins that enhance

51

phagocytosis (such as antibodies or complement), thus decreasing the efficiency of uptake (Hong

and Ghebrehiwet, 1992; Prasadarao et al., 2002; Rooijakkers et al., 2005). Finally, P.

aeruginosa, and uropathogenic E. coli (UPEC) secrete virulence factors that disrupt actin

dynamics that are required for uptake (Davis et al., 2005; Garrity-Ryan et al., 2000). Yersinia

enterocolitica also secretes virulence factors that impair uptake; however, the mechanism for this

inhibition remains elusive (Grosdent et al., 2002).

Intracellular pathogens that do not resist uptake have been shown to employ other strategies to

either inhibit or alter the process of phagosomal maturation to avoid lysosomal clearance. Some

pathogens, such as Mycobacterium tuberculosis, Coxiella burnetti, Legionella pneumonphila and

Salmonella enterica serovar Typhimurium, can withstand the low pH of the lysosomal

compartment either by expressing pH insensitive proteins or enzymes that neutralize the pH in

their immediate vicinity (Heinzen et al., 1996; Park et al., 1996; Sauer et al., 2005; Sturgill-

Koszycki and Swanson, 2000; Vandal et al., 2008). Similarly, some pathogens encode bacterial

virulence factors that can inhibit the activity or production of antibacterial peptides or small

molecules (McCaffrey and Allen, 2006; Peschel et al., 2001; Schmidtchen et al., 2002; Vazquez-

Torres et al., 2000). However, these survival mechanisms are inadequate if the bacteria is to

colonize and flourish in the host cell. Thus, more elaborate and sophisticated mechanisms are

needed in order to avoid phagosomal fusion with the lysosome.

Mechanisms of preventing phagosomal fusion with the lysosome can be classified into three

categories: 1) inhibition of phagosomal maturation, 2) alteration of phagosomal maturation, and

3) escape from the phagosome. Inhibition of maturation is exemplified by M. tuberculosis as this

bacterium prevents the recruitment of surface proteins (EEA1 and VPS34) (Fratti et al., 2001)

and the generation of specific phosphoinositides (PI3P) (Fratti et al., 2001; Fratti et al., 2003),

without which fusion with the lysosome cannot occur (Fratti et al., 2001; Pethe et al., 2004).

Similarly, maturation of the phagosome can be altered by recruiting surface proteins that mediate

phagosomal fusion with organelles other than the lysosome. For example, L. pneumophila

expresses effectors that slow maturation and allow recruitment of host factors, Rab1

(Ingmundson et al., 2007; Murata et al., 2006) and ARF1 (Nagai et al., 2002), which mediate

fusion with the endoplasmic reticulum instead of the lysosome (Robinson and Roy, 2006).

Finally, some pathogens, such as Listeria monocytogenes, encode virulence factors that mediate

escape from the phagosome (Portnoy et al., 2002).

52

The process of phagosomal escape is highly complex and intricate. It is one of the most efficient

methods of colonization as bacterial replication occurs rapidly in the nutrient-rich cytosol. In

addition to L. monocytogenes, a number of other pathogens, including S. Typhimurium and

Shigella flexneri, utilize this method for lysosome avoidance. Consequently, the use of L.

monocytogenes as a model pathogen to study the battle between host and bacteria will shed light

on the pathogenesis of a number of other pathogens. Thus, the focus of this thesis is to gain

insight into the early host defenses that are activated upon L. monocytogenes infection and the

bacterial attempts to avoid clearance.

4 L. monocytogenes

Listeria monocytogenes is a Gram-positive bacterium that is well adapted to survive both in the

environment (Fenlon et al., 1995) and inside mammalian cells (Dussurget et al., 2004). This rod-

shaped organism was first described in 1926 by E.G.D Murray following an epizootic outbreak

in rabbits and guinea pigs in an animal care facility in England (Murray et al., 1926). The

potential for this bacterium to infect humans was not realized until 1952 (Potel, 1952).

L. monocytogenes is uniquely resistant to conventional food sterilizing techniques, such as

refrigeration, low pH, high salt concentrations and other such environmental extremes (Schnupf

and Portnoy, 2007). Furthermore, this organism also produces biofilms, an extracellular polymer

matrix material that aids in prolonged survival (Djordjevic et al., 2002). Thus, given the

ubiquitous nature of this organism, the potential for food borne outbreaks is relatively high. The

first reported case of food borne L. monocytogenes outbreak occurred in Nova Scotia in 1981,

and resulted in 18 deaths (Schlech et al., 1983). Since then, there have been numerous cases of

outbreaks or product recalls due to L. monocytogenes contamination found on consumer goods

including ready-to-eat meats, soft unpasteurized cheeses, and raw produce.

53

4.1 Genetic Diversity of L. monocytogenes

L. monocytogenes belongs to the genus, Listeria, which is also comprised of L. innocua, L.

ivanovii, L. seeligeri, L. welshimeri and L. grayi (Schmid et al., 2005). Of these, only L.

monocytogenes and L. ivanovii are pathogenic (Hitchins, 1998). However, L. invanovii is only

pathogenic in animals. Understanding of L. monocytogenes pathogenesis has been improved

since early 2000 when the first L. monocytogenes genome was sequenced (Glaser et al., 2001).

To date, 22 different L. monocytogenes clinical isolates have been sequenced, with each genome

comprising roughly 3 Mbps, and an average of 2900 predicted protein-coding genes and an

average GC content of 38% (Glaser et al., 2001; Nelson et al., 2004).

4.2 Epidemiology of Clinical Listeriosis

4.2.1 Clinical Presentation and Treatment

L. monocytogenes infection in humans can result in listeriosis, a gastroenteritis that is self-

limiting in healthy individuals but may become severe and systemic for immunocompromised

individuals, the elderly and pregnant women (Rocourt and Bille, 1997). In susceptible

individuals, bacterial infection can progress to septicemia, meningoencephalitis and

maternofetal/neonatal listeriosis, culminating in a 20-30% mortality rate despite intervention

(Swaminathan and Gerner-Smidt, 2007). Listeriosis may also present as local infections

following hematogenous spread, commonly in the peritoneum, endocardium, joints or eyes

(Swaminathan and Gerner-Smidt, 2007). Furthermore, these bacteria can persist

asymptomatically even after the acute illness has subsided as L. monocytogenes can be detected

in the stool of 1-10% of the general population (Ramaswamy et al., 2007). The incidence rate of

listeriosis ranges between 0.1 to 11.3 per million individuals in different countries (WHO, 2004).

In Canada, there are on average 100 to 140 reported cases of listeriosis per year. However, it

must be noted that investigation of the epidemiology of L. monocytogenes infections can often be

complicated by the variability in incubation period, which can range from 3 to 60 days.

L. monocytogenes infection is commonly identified via blood and urine cultures. Treatment for

symptomatic listeriosis in the high-risk population involves supportive therapy along with high

54

doses of penicillin or ampicillin in combination with an aminoglycoside. While reports of

antibacterial resistance in clinical isolates remain rare (Hansen et al., 2005), more cases of

resistance in animal isolates have surfaced (Srinivasan et al., 2005).

4.2.2 Clinical Isolates

Clinical isolates of L. monocytogenes can be classified into 13 serovars based on gene content

and virulence. While serotypes 1/2a, 1/2b and 1/2c are the most abundant in the environment,

serotypes 1/2a, 1/2b and 4b are responsible for 95% of all human infections (Swaminathan and

Gerner-Smidt, 2007). Specifically, serotype 4b is responsible for the majority of L.

monocytogenes outbreaks and is associated with more severe symptoms and greater mortality in

infected individuals (Gerner-Smidt et al., 2005; Srinivasan et al., 2005). These clinical

observations suggest that different serotypes may confer different levels of virulence. While

there have been attempts to perform a comparative analysis between the different serotypes

(Jacquet et al., 2004), a systematic attempt has yet to be made. Thus, it remains currently unclear

why serotype 4b is more virulent than others.

Most notably, one serotype 1/2a strain, termed 10403S, is the most widely used strain in L.

monocytogenes research worldwide. 10403 was originally isolated from a human skin lesion in

the USA and 10403S is a streptomycin resistant mutant of the parent 10403 strain. Other

common laboratory strains include LO28, a serotype 1/2c strain that is not a clinical isolate, and

EGDe, a sertoype 1/2a strain that was originally isolated from rabbit tissue in England.

4.3 L. monocytogenes infection and intracellular lifestyle

Upon food borne entry into the human host, L. monocytogenes has the ability to cross the

intestinal, blood-brain and maternal-fetal barriers (Lecuit and Cossart, 2002). This organism is

also a facultative intracellular pathogen that can survive and replicate inside epithelial cells or

macrophages (Dussurget et al., 2004). As previously described, the cytosol of host cells is an

ideal environment for pathogens since it is nutrient rich and sheltered from a number of

55

extracellular immune defenses, such as direct killing by complement or antimicrobial peptides

and antibody neutralization and opsonization. The intracellular lifecycle of L. monocytogenes

can be divided into four distinct stages: 1) entry into the host cell, 2) phagosomal escape, 3)

cytosolic replication, and 4) secondary spread. L. monocytogenes expresses two proteins called

Internalin A and B (InlA and B), which mediate entry into the host cell by induced endocytosis

or phagocytosis.

Upon entry, L. monocytogenes expresses a number of virulence factors that subvert host defenses

in order to avoid death in the lysosome. These include listeriolysin O (LLO) and two

phospholipases (PLCs), whose activity mediates escape from the primary phagosomal vacuole.

Upon escape, L. monocytogenes can then replicate rapidly in the nutrient rich cytosol. Another

virulence factor, ActA, then mediates the nucleation of an actin tail on one end of the bacteria.

Polymerization of the actin tail allows the bacteria to “rocket” into neighbouring cells entering in

a double-membraned secondary vacuole. This therefore allows for secondary spread (Figure 4).

4.3.1 Entry into host cells

InlA and InlB mediate the uptake of L. monocytogenes into epithelial cells. They belong to the

family of internalin proteins that are characterized by the presence of leucine-rich repeats (LLRs)

(Gaillard et al., 1991). Specifically, InlA and B posses the LPXTG cell wall-sorting motif, which

allows linkage of these proteins to the bacterial surface (Dussurget et al., 2004). InlA and B also

have the capacity to bind to specific proteins in the host cell plasma membrane. InlA binds to

host E-cadherin, an interaction that is dependent on proline 16 in E-cadherin (Mengaud et al.,

1996). Mouse E-cadherin has a glutamic acid at this residue, which prevents InlA mediated entry

into mouse epithelial cells (Lecuit et al., 1999). Thus, oral administration of L. monocytogenes is

an inefficient method of establishing an in vivo infection in wild type mice. InlB binds to the

hepatocyte growth factor/scatter factor, or Met, receptor (Shen et al., 2000) and the globular part

of the complement component C1q receptor (C1qR) (Bierne and Cossart, 2002). A number of

studies have found that InlB alone is sufficient to mediate uptake of latex beads or other

noninvasive organisms (Braun et al., 1997; Braun et al., 1999; Braun et al., 1998). Interaction

between InlB and the Met receptor triggers phosphorylation and activation of this tyrosine kinase

receptor (Shen et al., 2000). As a result, downstream effectors are recruited (Ireton et al., 1999)

56

Figure 4. Intracellular lifecycle of L. monocytogenes with associated electron micrographs.

Intracellular survival of L. monocytogenes is mediated at different stages by the activity of

different virulence factors. 1) Internalin A and B (InlA and InlB) mediate bacterial entry. 2)

Bacterial escape from the phagosome is mediated by listeriolysin O (LLO) and PI-PLC. 3) Rapid

bacterial replication occurs in the cytosol. 4) Actin rocket tail formation is mediated by ActA. 5

& 6) Secondary spread into the neighbouring cell is mediated by the actin rocket tail. 7 & 8)

Escape from the double-membraned secondary phagosome is mediated by PC-PLC. Here, the

cycle of L. monocytogenes infection repeated. Reprinted from L. monocytogenes, a unique model

in infection biology: an overview. Pascale Cossart, Alejandro Toledo-Arana. Microbes and

Infection. 10: 1041-50 (2008), with permission from Elsevier.

57

to mediate actin rearrangement to allow for bacterial uptake via a zipper-like mechanism (Bierne

and Cossart, 2002). It must be noted that there are other factors that have been found to be

involved in bacterial mediated uptake. However, since the focus of this thesis is on L.

monocytogenes infection in macrophages and the predominant mode of bacterial uptake in this

cell type is mediated via phagocytosis, further details on bacterial mediated uptake mechanisms

will not be further discussed.

4.3.2 Escape from the phagosome

Upon entry, L. monocytogenes resides inside a phagosome that is destined for degradation in the

lysosome. To overcome this fate, the bacteria employ three virulence factors, listeriolysin O

(LLO) and two phospholipase Cs (PLCs) to escape from the phagosome. While the PLCs

enhance the efficiency of escape, LLO is necessary and sufficient for phagosomal escape

(Beauregard et al., 1997; Berche et al., 1987; Henry et al., 2006; Kuhn et al., 1988; Portnoy et al.,

1988). Thus, LLO is the critical virulence factor for L. monocytogenes survival in macrophages.

L. monocytogenes mutants that lack hly, the gene that encodes LLO, are avirulent in in vivo

mouse models. While it is not clear exactly how LLO mediates escape, it has been shown that

LLO inhibits fusion with the lysosome by generating small pores in the phagosome that block

acidification and the accumulation of Ca2+ in this compartment through uncoupling of the H+

pump (Shaughnessy et al., 2006). This provides a “window of opportunity” for the PLCs to lyse

the phagosomal membrane (Shaughnessy and Swanson, 2007). LLO and the PLCs have are also

necessary for escape from double membrane vacuole formed after secondary cell-to-cell spread

(Gedde et al., 2000).

4.3.2.1 Listeriolysin O (LLO)

LLO is a cholesterol dependent cytolysin (CDC) belonging to a family of pore-forming

cytolysins that are produced by a number of Gram-positive bacteria (Alouf, 2000). A diverse

range of pathogens secrete pore-forming cytolysins, including Clostridium perfringens (encodes

perfringolysin O), Streptococcus pyogenes (encodes streptolysin O), S. pneumoniae (encodes

58

pneumolysin), Aeromonas hydrophila (encodes aerolysin) and Bacillus anthracis (encodes

anthrolysin) (Tweten et al., 2001). Unlike other pore-forming cytolysins, LLO exhibits pH

sensitivity, with its optimal activity at pH <6 (Schuerch et al., 2005). Under acidic conditions,

LLO monomers bind to cholesterol rich regions on the membrane and undergo a conformational

change, which allows for the formation of hydrogen bonds with other monomers (Gekara et al.,

2005). This oligomerization results in the formation of the pre-pore complex. Via a cholesterol

independent mechanism (Gekara et al., 2005; Jacobs et al., 1998), two α-helical bundles on each

monomer extend to form transmembrane β-hairpins that insert into the membrane (Shatursky et

al., 1999; Shepard et al., 2000). This generates a functional pore of up to 350Å in diameter that is

comprised of roughly 33-50 monomers (Gilbert et al., 2000). Nascent LLO pores are thought to

only allow the passage of small molecules (< 550 Da), thus rendering the phagosome permeable

to Ca2+ and H+ (Shaughnessy et al., 2006). Changes in calcium concentration and pH can delay

phagosomal maturation, thus allowing time for the PLCs to enlarge the pores, which ultimately

dissolves the phagosomal membrane (Luzio et al., 2003; Shaughnessy and Swanson, 2007).

Since pore-forming cytolysins have the potential to form pores on any cholesterol containing

membrane, unregulated activity may result in host cell death. Indeed, it has been demonstrated

that genetically modified L. monocytogenes strains that produce perfringolysin O (PFO) in place

of LLO cause increased cellular toxicity compared to wild type L. monocytogenes (Glomski et

al., 2002; Jones and Portnoy, 1994). This work suggested that L. monocytogenes has evolved

“failsafe” mechanisms which ensure that LLO is only active at the appropriate time and cellular

location (reviewed in (Schnupf and Portnoy, 2007)). Subsequent studies have shown that a

number of unique factors ensure optimal LLO activity specifically in the phagosome.

Acidification of the phagosome allows structural reconfigurations that render LLO stable and

active. At neutral pH, LLO undergoes a rapid and irreversible alteration of its structure,

rendering it unable to form pores despite being oligomerized (Schuerch et al., 2005). However,

under cholesterol rich conditions, low levels of LLO activity can be observed even at neutral pH

(Bavdek et al., 2007). Thus, while optimal LLO function is limited to the late acidified

phagosome, LLO may still function to a lower extent at neutral pH. Furthermore, the reduction

of disulfide bonds in LLO is needed for efficient formation of pores (Geoffroy et al., 1987;

Portnoy et al., 1992). This is mediated by a host phagosomal protein called gamma-interferon

inducible lysosomal thiolreductase (GILT) (Singh et al., 2008). LLO activity is further

59

modulated by other host factors in the phagosome, such as cathepsin D (Carrasco-Marin et al.,

2009) and defensins (Arnett et al., 2011).

In addition to mechanisms that ensure optimal LLO activity is restricted to the phagosome, there

are also mechanisms that limit LLO activity in the cytosol once the bacteria has escaped the

phagosome. LLO encodes a PEST-like region (a sequence that marks proteins for proteasome

degradation) which functions to limit the levels of cytoplasmic LLO to prevent cytolysis

(Schnupf et al., 2006b; Schnupf et al., 2007). Cytosolic LLO levels are further controlled by a

negative feedback loop between LLO concentration and LLO translation (Schnupf et al., 2006a).

Finally, cytoplasmic LLO can aggregate and form protein complexes that are ubiquitinated

presumably by the host and are therefore targeted for clearance by the proteasome (Viala et al.,

2008). This is dependent, in part, on the N-terminal region of LLO which contains an

ubiquitination site. Infection with a L. monocytogenes mutant that lacks this region results in

higher levels of LLO and increased cytotoxicity compared to infection with wild type bacteria

(Decatur and Portnoy, 2000; Glomski et al., 2003).

Different signaling events have been reported to occur in different cells upon LLO stimulation.

LLO results in pro-inflammatory cytokine production in macrophages (Nishibori et al., 1996;

Yoshikawa et al., 1993), apoptosis in dendritic cells (Guzman et al., 1996), and degranulation

and leukotriene production in neutrophils (Sibelius et al., 1996b; Sibelius et al., 1999). In

epithelial HeLa cells, LLO induces mitogen-activated protein kinase (MAPK) (Tang et al., 1996)

and extracellular regulated kinase 1/2 (ERK1/2) activation (Tang et al., 1998). Studies in

embryonic kidney and goblet cells all reported changes in signaling or cell function upon LLO

stimulation, such as increased expression of cell adhesion molecules (Krull et al., 1997), mucin

production (Coconnier et al., 1998; Coconnier et al., 2000) and NF-κB activation (Kayal et al.,

2002).

In general, LLO mediated signaling can be divided into pore-dependent and pore-independent

events. Pore-dependent signaling may occur as a result of a breach in membrane integrity, the

formation of Ca2+ permeable pores in the endoplasmic reticulum, and phosphoinositide (PI)

metabolism (see the Phospholipase C section for details). Pore-independent signaling may result

from LLO oligomerization in lipid rafts or in the cytosolic recognition of specific LLO domains.

Furthermore, pore-forming cytolysins have been suggested to act as novel Toll-like receptor

60

(TLR) agonists. TLR activation is responsible for the induction of a proinflammatory response in

immune cells. LLO, like its pore-forming cytolysin family members, pneumolysin, anthrolysin,

PFO, and streptolysin O, can bind TLR4 and mediate NF-κB activation and TNF-α production

(Park et al., 2004; Srivastava et al., 2005). Furthermore, TLR2-/- mice are unresponsive to LLO-

deficient (ΔLLO) bacteria, displaying a deficiency in neutrophil recruitment to the site of

infection (Gekara et al., 2009). Thus, it is clear that LLO is a multi-functional virulence factor

that may have a number of other unappreciated functions.

4.3.2.2 Phosphatidylinositol – Phospholipase C (PI-PLC) and

phosphatidylcholine – Phospholipase C (PC-PLC)

L. monocytogenes produces two soluble phospholipase Cs (PLCs): phosphatidylinositol (PI)

specific PI-PLC and broad-spectrum phosphatidylcholine (PC)-PLC. PC-PLC is produced as a

proenzyme, requiring activation by the bacterial metalloprotease after it is exported from the

bacteria (Domann et al., 1991; Mengaud et al., 1991). The importance of these two PLCs to L.

monocytogenes pathogenesis can be seen in murine infection models. While infection with single

PLC deletion mutants results in a modest increase in 50% lethal dose (LD50), infection with

double PLC deletion mutants result in a dramatic 500-fold increase in LD50 (Smith et al., 1995).

There are a number of consequences that result from PLC activity. First, PLCs can mediate

membrane lysis (Goldfine et al., 1993; Smith et al., 1995), allowing for bacterial escape from the

phagosome (Grundling et al., 2003; Marquis et al., 1995). Studies indicate a role for PI-PLC in

mediating lysis of the single membrane primary vacuole (Camilli et al., 1993) while PC-PLC

mediates the lysis of the double membrane secondary vacuole (Vazquez-Boland et al., 1992).

However, both PLCs can act synergistically to allow for optimal lysis of either single or double

membrane vacuole (Camilli et al., 1993; Gedde et al., 2000). Intriguingly, the requirement for

PLC activity during phagosomal escape is cell-type dependent. In epithelial cells, PLCs are

necessary to allow for phagosomal escape (Grundling et al., 2003), while LLO alone is necessary

for escape and survival in macrophages (Gedde et al., 2000). However, it must be noted that

efficient escape from the primary vacuole in macrophages requires both LLO and PI-PLC

(Poussin et al., 2009; Wadsworth and Goldfine, 2002).

61

The second consequence of PLC activation is the activation of specific downstream signals. The

generation of DAG and IP3 by PLC mediates many changes in the cell. Most notably, DAG

production results in the recruitment and activation of protein kinase Cs (PKCs). Specifically in

phagocytes, DAG production requires both LLO and PI-PLC (Sibelius et al., 1999; Wadsworth

and Goldfine, 2002). Generation of DAG recruits host PKCβ and PKCδ. PKCδ activation

recruits host phospholipase D (PLDs) (Goldfine et al., 2000). In addition, IP3 mediates the

release of Ca2+ from the endoplasmic reticulum, resulting in an influx of intracellular Ca2+.

Changes in calcium concentration results in the activation of host PLCs (Titball, 1993). These

studies together suggest a model of escape in which LLO pore formation allows the passage of

PI-PLC from inside the phagosome to reach the cytosolic leaflet of the phagosomal membrane

(Sibelius et al., 1999). Here, PI-PLC will generate DAG and increase Ca2+ concentration, which

then recruits host PKC and activates host PLCs. Activation of PKCs results in recruitment and

activation of host PLDs, which together with bacterial PLCs can result in lysis of the phagosomal

membrane, thus allowing bacterial escape. Therefore, both bacterial and host enzymes contribute

to efficient bacterial escape (Poussin et al., 2009; Wadsworth and Goldfine, 2002).

4.3.3 Cytosolic Replication

Upon escape into the cytosol, L. monocytogenes undergoes a phase of rapid growth. In primary

bone marrow derived macrophages, this bacterium replicates in the nutrient rich cytosol with a

doubling time of roughly 40 min (de Chastellier and Berche, 1994). Quantification by

transmission electron microscopy reveals that phagosomal escape begins as early as 30 min p.i..

At 1 h p.i., 20% of intracellular L. monocytogenes has escaped while at 4 h p.i., 50% of

intracellular L. monocytogenes has escaped (de Chastellier and Berche, 1994).

4.3.4 Secondary Spread

After escape into the cytosol, another virulence factor called ActA induces polymerization of

actin filaments to promote cytosolic movement and cell-to-cell spread (Domann et al., 1992;

Kocks et al., 1992). ActA is anchored covalently to one pole of the bacterial surface via the C-

62

terminal transmembrane motif. Actin polymerization is mediated through an interaction between

the central four proline-rich repeat domains in ActA and members of the enabled/vasodilator-

stimulated phosphoprotein (Ena/VASP) family of actin nucleators (Lasa et al., 1995). This

interaction modulates bacterial speed and directionality (Auerbuch et al., 2003; Geese et al.,

2002; Laurent et al., 1999). The N-terminal region of ActA mimics host Wiskott Aldrich

syndrome protein (WASP) family proteins by interacting with Arp2/3, a seven-protein complex

that initiates the formation of branched actin filaments (Boujemaa-Paterski et al., 2001; Cossart,

2000; Skoble et al., 2000). This interaction modulates bacterial movement via the polymerization

of actin filaments (termed “actin-comet tails”). Bacteria driven by actin-comet tails can form

protrusions from one cell into the next, allowing for secondary spread of L. monocytogenes.

Upon entry into neighbouring cells, the invading bacteria insides in a double membrane vacuole

which is comprised of the plasma membrane from the previous and current host cell. Here, PC-

PLC is critical for mediating escape from the secondary vacuole (Grundling et al., 2003).

4.3.5 PrfA Regulon

The expression of the canonical L. monocytogenes virulence factors described above are encoded

in the prfA regulon, regulated by a transcription factor called the positive regulation factor A,

PrfA (extensively reviewed in (Freitag, 2009; Freitag et al., 2009; Gray et al., 2006)). PrfA

expression is regulated by a number of environmental factors that are unique to the host

environment. One such factor is temperature. PrfA transcription is directed in part by a

thermosensitive promoter (Leimeister-Wachter et al., 1992) that undergoes structural

rearrangements at temperatures above 30oC (Johansson et al., 2002). Independent of temperature,

phagosome specific conditions have also been observed to induce expression of the prfA regulon.

Inside the phagosome, L. monocytogenes is subjected to oxidative stress, reduced nutrient and

metals availability and low pH. These factors have all been observed to cause increased

expression of the prfA regulon (Becker et al., 1998; Ferreira et al., 2001; Ferreira et al., 2003;

Makino et al., 2005). Consistent with the need to rapidly modulate gene expression upon changes

in the environment, it must be noted that there is a pool of prfA transcripts inside the bacteria that

allow for rapid translation of the prfA regulon and its virulence factors immediately upon

introduction into the host (Leimeister-Wachter et al., 1992).

63

4.4 Phagosomal Immune Defenses and L. monocytogenes

A number of phagosomal defenses play an important role in limiting L. monocytogenes escape

into the cytosol. For example, antimicrobial peptides, specifically defensins, have recently been

shown to play an important role in limiting L. monocytogenes escape from the phagosome in

macrophages (Arnett et al., 2011). Intriguingly, L. monocytogenes has evolved a mechanism to

D-alanylate cell wall components, resulting in a more positively charged bacterial surface, thus

effectively repelling cationic antimicrobial peptides (Thedieck et al., 2006). In addition to

antimicrobial peptides, two other critical host defenses, ROS and autophagy, have likewise been

shown to be important phagosomal defenses against L. monocytogenes phagosomal escape.

4.4.1 Autophagy and L. monocytogenes

The importance of autophagy in limiting L. monocytogenes replication is seen in a number of in

vivo studies. Mice with Atg5 deficient macrophages (Atg5flox/flox-Lyz-Cre) exhibit a 50% drop in

survival 21 d p.i. with L. monocytogenes when compared to wild type mice (Zhao et al., 2008).

In particular, significantly greater bacterial load was observed in the livers at day 3 p.i. of the

Atg5flox/flox-Lyz-Cre mice when compared to control. Furthermore, drosophila mutants deficient

in Atg5 or the pattern-recognition receptor, peptidoglycan recognition protein (PGRP)-LE, fail to

induce autophagy in response to L. monocytogenes infection (Yano et al., 2008). PGRP-LE

recognition of the L. monocytogenes cell wall component, diaminopimelic acid-type

peptidoglycan, results in autophagy targeting of L. monocytogenes, as assessed by increased

LC3+ double-membrane L. monocytogenes containing compartments (Yano et al., 2008).

Drosophila lacking in PGRP-LE or expressing a mutant PGRP-LE are unable to induce

autophagy, resulting in increased susceptibility to L. monocytogenes infection. This results in a

four-fold decrease in the number of surviving drosophila mutants 8 d p.i. over the wild type

(Yano et al., 2008). Thus, these in vivo studies provide strong evidence that autophagic induction

during L. monocytogenes infection is a critical host defense against the bacteria.

64

In vitro studies reveal an extremely complex picture of how autophagy targets L. monocytogenes.

Depending on the stage of infection or where the bacteria is located, autophagy targeting of L.

monocytogenes may be mediated via different mechanisms. Roughly 30-40% of L.

monocytogenes is targeted by autophagy with the peak of LC3 targeting occurring in the early

stages of infection at 1 h p.i. (Birmingham et al., 2007a; Meyer-Morse et al., 2010; Py et al.,

2007). Autophagic targeting was found to be dependent on LLO as LLO deficient L.

monocytogenes does not become significantly LC3+ at any point during infection (Birmingham

et al., 2007a; Meyer-Morse et al., 2010; Py et al., 2007). However, during later stages of

infection at 8 h p.i., when most bacteria have escaped the phagosome, only 10% of wild type and

ActA deficient (ΔActA) L. monocytogenes are LC3+. Interestingly, while ΔActA mutants treated

with the bacteristatic agent, chloramphenicol, become 30% LC3+, wild type L. monocytogenes

treated with chloramphenicol remain 10% LC3+ (Birmingham et al., 2007a). This data suggests

that cytosolic autophagy targeting of L. monocytogenes may be evaded by ActA (Birmingham et

al., 2007a; Rich et al., 2003). It is known that the ΔActA mutant colocalizes with ubiquitinated

proteins (Ub) at 8 h p.i. (Perrin et al., 2004). While it remains unclear how L. monocytogenes is

ubiquitinated, a number of studies have provided evidence for LLO ubiquitination. LLO in the

cytosol has been described to form Ub+ and p62+ aggregates (Viala et al., 2008), and is thought

to be directly ubiquitinated via its N-terminal lysine residues (Schnupf et al., 2007). Recent work

by Sasakawa and colleagues suggests that this protein ubiquitination event mediates recruitment

of adaptor proteins such as p62/SQSTM1, which leads to autophagic targeting of the ΔActA

mutant (Yoshikawa et al., 2009). Thus, these observations combined suggest that ActA may play

a critical role in evading LC3 targeting by preventing ubiquitination of L. monocytogenes in the

cytosol. It remains unclear if ubiquitination also plays a role in LC3 targeting during the early

stage of infection.

While it is clear from in vivo observations that autophagy is capable of limiting bacterial

infection, in vitro studies of L. monocytogenes replication in autophagy deficient environments

have yielded less clear results. Mild increases have been reported when comparing the number of

wild type L. monocytogenes recovered from ATG5 deficient over wild type MEFs at various

time points p.i. (Birmingham et al., 2007a; Py et al., 2007; Yoshikawa et al., 2009). However, L.

monocytogenes mutants deficient in PI-PLC exhibit markedly greater bacterial replication in

ATG5 deficient when compared to wild type MEFs (Birmingham et al., 2007a). This observation

65

suggests that PI-PLC, or perhaps both PLCs, also play a role in the evasion of autophagy that is

currently unappreciated.

Despite these studies, many questions about autophagic targeting of L. monocytogenes still

remain. Since the bacteria can rapidly escape from the phagosome, it is unclear whether L.

monocytogenes is within phagosomes or in the cytosol at the peak of autophagy targeting at 1 h

p.i.. While the available data in the field suggests that ubiquitination is a process that occurs once

the bacteria has entered the cytosol, there lacks direct conclusive evidence to support this claim.

Given that damaged vacuoles have been shown to be a target of ubiquitination in S. flexneri

infection (Dupont et al., 2009), phagosomes containing L. monocytogenes may be damaged by

LLO pores and thus mediate ubiquitination. Furthermore, it is also unknown which host and/or

bacterial factor(s) mediate autophagy targeting of L. monocytogenes at 1 h p.i.. Thus, much less

is known about the mechanism of early autophagy targeting of L. monocytogenes (Figure 4.4 B)

and how it might differ from that of later targeting of L. monocytogenes (Figure 4.4 A).

66

Figure 4.4. Current model of autophagy targeting of L. monocytogenes. (A) In presence of

chloramphenicol, L. monocytogenes ΔactA mutant (but not wild type) is ubiquitinated and

targeted by autophagy at the later stages of infection. (B) The mechanism of autophagy targeting

of L. monocytogenes at 1 h remains unclear. It is unknown which host or bacterial factors recruits

LC3 at this early stage of infection. Prime candidates include ROS, DAG, and Ub.

67

4.4.2 ROS AND L. monocytogenes

Given that ROS are a potent bactericidal defense in the phagosome and they can act as a signal

for autophagy targeting during S. Typhimurium infection, how does L. monocytogenes evade

ROS in order to escape the phagosome? It is known that ROS are an important host defense

against L. monocytogenes replication. In vivo infection of gp91phox-/- mice with wild type L.

monocytogenes results in a higher bacterial load at 48 h post injection when compared to

infection of wild type mice (Dinauer et al., 1997; LaCourse et al., 2002; Shiloh et al., 1999).

ROS were likewise found to be important for in vitro macrophage clearance of L. monocytogenes

(Ohya et al., 1998a; Ohya et al., 1998b). Furthermore, L. monocytogenes exhibits greater escape

from the phagosome upon infection of gp91phox-/- bone marrow derived macrophages (BMDM)

compared to wild type BMDM (Myers et al., 2003). Taken together, the observed increase in L.

monocytogenes load recovered from gp91phox-/- infected mice may be attributed to increased

bacterial escape from the phagosome in the absence of ROS production. Thus, it is clear that

ROS is an important host defense that must be overcome in order for bacterial escape into the

cytosol.

Since ROS are an effective and important host defense against infection by intracellular

pathogens, it is not surprising that many intracellular pathogens have evolved mechanisms to

overcome ROS production by NOX2 NADPH oxidase (Table 4.4). The Salmonella

pathogenicity island 2 (SPI-2) is critical for the evasion of NOX2 NADPH oxidase targeting in

Salmonella enterica serovar Typhimurium infection in murine macrophages (Vazquez-Torres et

al., 2000). A similar inhibition of NOX2 NADPH oxidase targeting has likewise been reported in

other infections, including Burkholderia cenocepia (Keith et al., 2009), Chlamydia trachomatis

(Boncompain et al., 2010), Coxiella burnetti (Siemsen et al., 2009), and Francisella tulerensis

(McCaffrey and Allen, 2006) among others. The most common effect of these bacteria on NOX2

NADPH oxidase is the mislocalization of one or multiple members of the NOX2 NADPH

oxidase complex. However, it is not known how these bacteria induce mislocalization of the

NOX2 NADPH oxidase.

In addition, some pathogens have evolved different means of inhibiting NOX2 NADPH oxidase

activity. The inhibition of ROS by Anaplasma phagocytophilum has been attributed to the

virulence factor, AnkA, which decreases transcription of gp91phox (Garcia-Garcia et al., 2009).

68

More recently, F. tulerensis production of an acid phosphatase has been shown to inhibit ROS

production, presumably via dephosphorylation of the regulatory subunits of the NOX2 NADPH

complex (Mohapatra et al., 2010). Likewise, the parasite Cryptosporidium parvum also encodes

an acid phosphatase that inhibits ROS production (Aguirre-Garcia and Okhuysen, 2007). Finally,

the YopD pore-forming cytolysin secreted by Yersinia enterocolitica inhibits ROS production;

however, the mechanism by which this occurs remains unknown (Hartland et al., 1994). It should

be noted that other pathogens also inhibit NOX2 NADPH oxidase activity, but the bacterial

factors and mechanisms involved are not understood. Table 4.4 provides a summary of the

pathogens with reported abilities and known mechanisms to inhibit ROS production.

Little work has been done to examine whether L. monocytogenes also combats ROS production.

A previous study found that the LLO deficient L. monocytogenes induces higher levels of ROS

than wild type at 10 min post infection in neutrophils (Sibelius et al., 1999), suggesting that LLO

might modulate ROS production. While NOX2 NADPH oxidase localizes to phagosomes for

efficient L. monocytogenes killing, LLO has been shown to alter the localization of the early

phagosomal protein, Rab5a, thereby inhibiting phagosome maturation (Alvarez-Dominguez et

al., 1997; Shaughnessy et al., 2006). Furthermore, work done by Makino and colleagues revealed

that oxidative stress induces expression of the LLO encoding gene, hly (Makino et al., 2005).

Specifically, exogenous addition of ROS induces hly expression, which is reduced upon addition

of ROS neutralizing enzymes, such as superoxide dismutase or catalase (Makino et al., 2005).

Together, these observations suggest that L. monocytogenes may overcome ROS production in

an LLO-dependent manner in order to escape the phagosome.

69

Organism Mechanism of NOX

interaction

Cell Type Reference

Bacteria

Anaplasma

phagocytophilum

AnkA nuclear effector protein interacts with the

transcriptional regulatory regions the gp91

gene

HL-60 (Garcia-Garcia et

al., 2009)

Bacillus anthracis Edema toxin inhibits NOX1 production of ROS

via increase in cAMP and PKA activation

HT-29 (Kim and

Bokoch,

2009)

Burkholderia cenocepia p22phox and p40phox translocation delayed Raw macrophage (Keith et

al., 2009)

Chlamydia trachomatis Mislocalization of Rac2 HeLa (Boncompain et

al.)

Coxiella burnetii p47phox and p67phox translocation to NOX

inhibited

Human PBMC (Siemsen et al.,

2009)

Francisella tulerensis gp91, p22phox, p47phox and p67phox translocation

to LVS inhibited

Human PBMC (McCaffrey and

Allen, 2006;

Schulert et al.,

2009)

migR and fevR mutants unable to prevent ROS

production

Human PBMC (Buchan et al.,

2009)

Acid phosphatases inactivate NOX Human PBMC (Mohapatra et al.)

Helicobacter pylori Early cessation of p47phox and p67phox

translocation to NOX

Human PBMC (Allen and

McCaffrey,

2007)

Legionella pneumophila ROS inhibition via p47phox translocation

inhibition

U937 (Harada et al.,

2007)

70

Salmonella typhimurium SPI-2 dependent exclusion of p22phox and

p47phox

Mouse macrophage (Vazquez-Torres

and Fang, 2001;

Vazquez-Torres et

al., 2001;

Vazquez-Torres et

al., 2000)

Vibrio vulnificus RtxA1 induce expression of Rac2 to induce

ROS

Mouse macrophage

in vitro & in vivo

model

(Chung et

al.)

Yersinia enterocolitica YopD prevent ROS Production BMDM (Hartland

et al.,

1994)

Yersinia pestis Type three secretion system dependent

inhibition of ROS production

Human neutrophils (Spinner et

al., 2008)

Yersinia

pseudotuberculosis

YopE prevent ROS Production via inactivation

of Rac2

In vivo mouse model (Songsungt

hong et al.,

2010)

Parasite

Cryptosporidium

parvum

Acid phosphatase prevent ROS production Human PBMC (Aguirre-Garcia

and Okhuysen, 2007)

Leishmania donovani Lipophosphoglycan inhibition of PKC

phosphorylation leads to exclusion of p47phox

and p67phox

Mouse and RAW

macrophage

(Descoteaux et al.,

1992; Lodge et al.,

2006)

Virus

Epstein Barr Virus

(EBV)

EBNA-1 induces ROS production to increase

host genetic instability

DG75 and BJAB

(human B cell

lines)

(Gruhne et al.,

2009)

Hepatitis C Virus

(HCV)

NS3 superoxide production in macrophages

and neutrophils, via recruitment of p47phox

and p67phox

Human PBMC (Bureau et al.,

2001; Thoren et

al., 2004)

71

HIV Nef directly binds to p22phox increasing ROS

production

Human PBMC (Salmen et al.;

Vilhardt et al.,

2002)

Japanese encephalitis

virus

Virus induces ROS production via

mechanism unknown

Murine

neuroblastomas

(Raung et al., 2001)

Respiratory syncitial

virus (RSV)

Induction of ROS production via

mechanism unknown

Human PBMC (Kaul et al., 2000)

Table 4.4. List of pathogens that have been reported to actively modulate NOX activity.

Reprinted by permission from Springer Science + Business Media: Seminars in

Immunopathology, Lam G et al. The many roles of NOX2 NADPH oxidase-derived ROS in

immunity, 32(4): 415-30

72

Chapter 2

Aims and Hypotheses

5 Aim

The cytosol of a host cell is a nutrient rich environment that is capable of supporting rapid

microbial growth. As such, the host has a number of defensive strategies aimed at clearing

invading pathogens and preventing colonization. Reactive oxygen species (ROS) production by

NOX2 NADPH oxidase and autophagy are among the best-characterized host defenses against

invading pathogens. Not only does ROS confer direct antimicrobial activity, but also can activate

other antimicrobial defenses. Similarly, autophagy is a key player of the antimicrobial response.

Autophagy is the process of controlled self-digestion upregulated in times of cellular stress. In

the context of intracellular pathogens, autophagy can be activated to selectively target pathogens

and mediate bacterial clearance in the lysosome.

L. monocytogenes is a cytosol-adapted pathogen that colonizes host cells, including

macrophages, during infection (Portnoy et al., 2002). Upon invasion, a population of bacteria are

able to escape from the phagosome and colonize the nutrient-rich cytosol. Phagosomal escape is

mediated by the bacterial pore forming toxin Listeriolysin O (LLO) and two phospholipase C

enzymes (PLCs). PLCs are thought to not only degrade the phagosome to promote escape, but

can also activate signal transduction cascades within the host cell as a result of their catalytic

activity (Goldfine and Wadsworth, 2002; Portnoy et al., 2002). Upon entry to the cytosol, the

bacteria use a cell surface protein, ActA, to drive actin-based motility in the host cytosol, and

eventual cell-cell transfer (Portnoy et al., 2002). However, prior to escape, L. monocytogenes

will encounter a number of phagosomal host defenses that the bacteria must overcome. Among

these defenses are ROS and autophagy.

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5.1 Hypothesis #1:

Here, I examined the mechanisms that regulate autophagic targeting of L. monocytogenes in

macrophages at early stages of infection. I hypothesize that LC3 targets L. monocytogenes within

phagosomes at 1 h p.i. in a manner requiring both DAG production and ROS generated by the

NOX2 NADPH oxidase but not protein ubiquitination.

5.1.1 Aim

Autophagy is the controlled process of self-digestion through which cellular contents are

delivered to the lysosome (Levine and Deretic, 2007). There are many types of autophagy, all of

which play a vital role in maintaining cellular homeostasis. Autophagy has been shown to be a

key component of the innate immune defense against many pathogenic microorganisms (Levine

et al.). Autophagy can promote phagosomal maturation and also limit bacterial escape from these

compartments (Birmingham et al., 2006; Gutierrez et al., 2004; Sanjuan et al., 2007). Autophagy

can also target cytosolic bacteria, mediating their delivery to lysosomes (Nakagawa et al., 2004;

Rich et al., 2003). Despite the importance of autophagy, the mechanisms that regulate targeting

of pathogens to the autophagy pathway remain unclear. Recent studies of Salmonella enterica

serovar Typhimurium (S. Typhimurium) infection demonstrated the involvement of ubiquitinated

proteins (Ub) and ubiquitin-binding adaptors (Birmingham et al., 2006; Cemma et al., 2010;

Randow and Lehner, 2009; Zheng et al., 2009), reactive oxygen species (ROS) (Huang et al.,

2009), and diacylglycerol (DAG) (Shahnazari et al., 2010) as key signals for the targeting of

these bacteria to the autophagy pathway. Whether these signals contribute to the regulation of

autophagy during infection by other bacterial pathogens is largely unclear.

Previous studies have shown that L. monocytogenes is targeted by autophagy in host cells, as

evidenced by colocalization of bacteria with the autophagy marker microtubule-associated

protein light chain 3 (LC3). The peak of LC3 colocalization with bacteria occurs at 1 hour post

infection (p.i.) (Birmingham et al., 2007a; Meyer-Morse et al., 2010; Py et al., 2007). However,

many questions about this process still remain. Since L. monocytogenes can rapidly escape from

the phagosome, it is unclear whether bacteria are targeted by autophagy within phagosomes or in

the cytosol at this early stage of infection. Furthermore, it is also unknown which host and/or

74

bacterial factor(s) regulate autophagy of L. monocytogenes. Work from the Brumell laboratory

previously demonstrated that a non-motile (ΔactA) mutant of L. monocytogenes colocalizes with

ubiquitinated proteins in the cytosol (Perrin et al., 2004). Recent work by Sasakawa and

colleagues suggests that this protein ubiquitination event mediates recruitment of adaptor

proteins such as p62/SQSTM1 that lead to autophagic targeting of the ΔactA mutant of L.

monocytogenes inside the cytosol, a process initiated after 2 hours p.i. (Rich et al., 2003;

Yoshikawa et al., 2009). However, it is not known if protein ubiquitination plays a role in

autophagic targeting of wild type L. monocytogenes at early stages of infection, the time of

maximal autophagy of these bacteria.

Thus, Aim 1.1 will be to determine if LC3+ L. monocytogenes at 1 h p.i. is targeted within a

phagosome. Given that a previously published study indicates that the majority of L.

monocytogenes reside in phagosomes at 1 h p.i. (de Chastellier and Berche, 1994), I predict that

LC3+ L. monocytogenes at 1 h p.i. are inside phagosomes. Next, Aim 1.2 will be focused on

understanding how LC3 is targeted to L. monocytogenes at 1 h p.i.. I will examine if LC3+ L.

monocytogenes also colocalize with DAG or ubiquitin. Finally, I will modulate the levels of

DAG and ROS to determine if altering the levels of DAG and ROS will influence LC3 targeting

to L. monocytogenes at 1 h p.i..

5.2 Hypothesis #2:

Since L. monocytogenes escape from the phagosome is a slow process that begins 30 min p.i.,

this prompted me to examine ROS production in the L. monocytogenes containing phagosome

prior to escape. I hypothesize that L. monocytogenes, via the activity of LLO, can inhibit

phagosomal production of ROS by mislocalizing NOX2 NADPH oxidase components.

5.2.1 Aim

The NOX2 nicotinamide adenine dinucleotide phosphate (NADPH) oxidase (also referred to as

gp91phox; phagocyte oxidase) plays a key role in immune responses via the production of reactive

75

oxygen species (ROS) (Lam et al., 2010). The loss of NADPH oxidase activity in mice lacking

the gp91phox subunit (gp91phox-/-) results in increased replication of L. monocytogenes during the

first 24 h of infection compared to wild type animals (Dinauer et al., 1997; Shiloh et al., 1999).

In vitro studies suggest that the NOX2 NADPH oxidase limits escape of L. monocytogenes from

the phagosome in macrophages (Myers et al., 2003).

To ensure survival in the phagosome, a number of pathogens have evolved mechanisms to

overcome ROS production by NOX2 NADPH oxidase. Such pathogens include Chlamydia

trachomatis (Boncompain et al., 2010), Coxiella burnetti (Siemsen et al., 2009), Francisella

tulerensis (McCaffrey and Allen, 2006) among others (Lam et al., 2010). Specifically,

Salmonella enterica serovar Typhimurium and Burkholderia cenocepia inhibit ROS production

by preventing NOX2 NADPH oxidase targeting to the bacteria containing phagosomes

(Vazquez-Torres et al., 2000) (Keith et al., 2009). However, the mechanism by which pathogens

inhibits NOX2 NADPH oxidase activity remains unknown.

Previously, it has been shown that L. monocytogenes infected neutrophils generate ROS 10 min

post infection (p.i.) (Sibelius et al., 1999). However, dynamic studies of where ROS is produced

have never been performed. Furthermore, phagosomal analysis of ROS production during L.

monocytogenes infection has likewise never been studied. It is known that LLO can inhibit

phagosome maturation by preventing fusion with the lysosome (Alvarez-Dominguez et al., 1997;

Shaughnessy et al., 2006). Furthermore, work done by Makino and colleagues revealed that

oxidative stress induces expression of the LLO encoding gene, hly (Makino et al., 2005).

Specifically, exogenous addition of ROS induces hly expression, which is reduced upon addition

of ROS neutralizing enzymes, such as superoxide dismutase or catalase (Makino et al., 2005).

Thus, Aim 2.1 is to first compare ROS production by macrophages infected with either wild type

or ΔLLO L. monocytogenes. Production of ROS over the course of infection will be assessed to

determine the temporal changes in ROS levels. Furthermore, intracellular, extracellular, and

phagosomal production of ROS will be studied separately to determine the site of ROS

production. Given my hypothesis that macrophages infected with wild type L. monocytogenes

will produce less ROS than macrophages infected with ΔLLO L. monocytogenes, Aim 2.2 will

be to examine how LLO can inhibit ROS production. Specifically, the effect of LLO on

signaling events upstream of NOX2 NADPH oxidase activation will be examined. Furthermore,

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proper assembly of the NOX2 NADPH oxidase components will also be assessed in

macrophages infected with either wild type or ΔLLO L. monocytogenes. Lastly, Aim 2.3 will be

to assess if other LLO-related pore-forming cytolysins can also inhibit ROS production during

infection.

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Chapter 3

Methods

6 Bacterial strains and culture conditions

Bacterial strains used in this study were as follows: L. monocytogenes 10403S (Bishop and

Hinrichs, 1987), ΔLLO (DP-L2161) (Jones and Portnoy, 1994), ΔLLO + LLO (DP-L4818)

(Lauer et al., 2002), iLLO (DH-L1239) (Alberti-Segui et al., 2007), ΔPI-PLCΔPC-PLC (DP-

L1936) (Smith et al., 1995), ΔLLOΔPI-PLCΔPC-PLC (DP-L2391) (O'Riordan et al., 2002) (gifts

from Dr. D. Portnoy, UC Berkley), iPFO (DH-L1745), 10403S expressing sGFP (DH-L1039)

(Shen and Higgins, 2005) and ΔLLO expressing sGFP (DH-L1137) (Agaisse et al., 2005). Both

GFP expressing strains were expressed under the control of the same Hyper-SPO1 promoter

fused to the 5’ UTR of the LLO gene, hly. LLO was purified from Escherichia coli encoding

recombinant N-terminal-tagged His-LLO, as described (Gedde et al., 2000). Bacteria were

grown in Brain-Heart Infusion (BHI) broth 14-16 h at 30oC in a standing incubator. A 1:10

dilution of the culture was grown for 2 h at 37oC in a shaking incubator prior to infection. LLO

and PFO expression in iLLO and iPFO strains were induced by IPTG, as previously described

(Alberti-Segui et al., 2007).

7 Macrophage generation and culture conditions

RAW 264.7 macrophages were purchased from American Type Culture Collection (Rockville,

MD). All experimental protocols involving mice were approved by the Animal Care Committee

of The Hospital for Sick Children. Mice were euthanized by cervical dislocation. The femur and

tibia were removed, cleansed of muscle fibers and cut distally. The bone marrow was then

removed via a 10 sec pulse of centrifugation at 2000 rpm. The resulting cells were centrifuged at

1500 rpm for 5 min, washed with growth medium and plated onto 10 cm tissue culture dishes.

Medium was replaced with fresh RPMI growth media (see below) every 3 days. Typically, 108

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bone marrow-derived macrophages (BMDM) were typically recovered after 7 days. Human

macrophages were prepared as previously described (McGilvray et al., 2000). RAW 264.7 cells

were maintained in DMEM growth medium (HyClone) supplemented with 10% FBS (Wisent) at

37oC in 5% CO2 without antibiotics. Murine macrophages were maintained in RPMI-1640

medium (Wisent) supplemented with 10% FBS (Wisent), 5% sodium pyruvate (Invitrogen), 5%

antibiotics (Invitrogen), 5% non-essential amino acids (Invitrogen) and 0.5 µM β-

mercaptoethanol (Invitrogen). BMDMs were differentiated in 30% L929 conditioned medium.

L929 conditioned medium was generated by growing L929 cells (ATCC) in 150-cm2 flasks at an

initial density of 1x108 cells per flask in growth medium as described above for use with RAW

264.7 cells. After 3 days, confluency was reached and the growth medium was substituted with

DMEM alone. After 7-10 days, culture supernatant was collected and centrifuged at 1,500 rpm

for 5 mins, aliquoted and stored at -20oC.

8 Plasmids and transfections

Transfections were performed 24 h prior to infection. Transfection reagents FuGene 6 and

FuGene 6 HD (Roche Applied Sciences) were used according to the manufacturer’s instructions.

BMDM were transfected using the Amaxa Primary Murine Macrophage Transfection kit

according to the manufacturer’s instructions (Amaxa). Constructs used were GFP-LC3 (provided

by Tamotsu Yoshimori, Osaka University, Japan) (Kabeya et al., 2000), RFP-LC3 (provided by

Walter Beron, Universidad Nacional de Cuyo, Argentina), PKCδ-C1 (provided by Sergio

Grinstein, Hospital for Sick Children, Toronto, Canada) (Tse et al., 2005) and HA-PLD1 K898R

and HA-PLD2 K758R (provided by Michael Frohman, SUNY, NY, USA) (Denmat-Ouisse et

al., 2001).

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9 Neutrophil purification and recombinant LLO (rLLO)

treatment

Neutrophils were isolated from healthy human volunteers, as described (Tole et al., 2009).

Neutrophils were stimulated with 10 µg/ml PMA along with treatment of 1µg/ml, 100ng/ml or

10ng/ml of rLLO for 1 h. ROS production was assessed using the NBT quantification assay

(described below).

10 In vivo infection

Three to five week old C57BL/6J and gp91phox-/- (Cybb/tm1d) mice were purchased from Jackson

Laboratory. Mice were infected via intravenous injection in the lateral tail vein with wild type L.

monocytogenes at 5x104 CFU in 200 µl of PBS, and ΔLLO bacteria at 1x109 CFU in 200 µl of

PBS. Mice were sacrificed at indicated time points and the livers excised. The right lobes of the

livers were fixed with neutral-buffered formalin, and embedded sections stained with

hematoxylin and eosin (H&E). The left lobes were homogenized in sterile PBS for CFU

quantification from serial dilutions on BHI-agar plates.

11 Macrophage replication assay and infection

After 7-10 days of differentiation, BMDM were washed twice and detached with ice cold

Versene Buffer (0.8 mM EDTA, 1 mM glucose in PBS) for 20 min at 4oC and plated at 5x105

cells per well in 24-well tissue culture plates, 24 h prior to infection. All strains of L.

monocytogenes were infected at a multiplicity of infection (MOI) of 1. After 30 min of invasion

at 37oC, cells were washed three times with phosphate buffered saline (PBS) followed by the

addition of DMEM. At 1 h post-infection, medium was changed and growth medium containing

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50 µg/ml gentamicin was added. Cells were then lysed at 2, 4, 8, 12, and 24 h post-infection with

0.2% TritonX-100 in PBS. Serial dilutions of the lysates were plated on BHI-agar plates and

incubated 14-16 h for subsequent quantification of colony forming units (CFUs).

12 Reagents and Antibodies

The following drugs were used as indicated: 0.3% v/v 1-butanol (Sigma), 0.3% v/v tert-butanol

(Sigma), 250 µM propranolol hydrochloride (Biomol International), 10 µg/ml PMA (Sigma) and

10 µM DPI (Sigma) were used. Staining for p22phox, p47phox and p67phox was completed using

mouse anti-GFP, rabbit anti-p22phox (gift from Dr. Mark T. Quinn, Montana State University,

Montana, USA), rabbit anti-p47phox and rabbit anti-p67phox (gifts from Nathalie Grandvaux,

McGill University, Montreal, Canada). Rabbit polyclonal antibodies against L. monocytogenes

were a gift from Dr. Pascale Cossart (Institut Pasteur), mouse monoclonal antibodies against

GFP were purchased from Invitrogen; rat monoclonal antibodies against Lamp-1 (clone ID4B)

from Developmental Studies Hybridoma Bank (University of Iowa); and antibodies against

mono- and poly-ubiquitinated protein were from Biomol International (FK2). All fluorescent

secondary antibodies were AlexaFluor conjugates from Molecular Probes (Invitrogen).

13 Immunostaining and Immunofluorescence

Immunostaining was conducted, as previously described (Brumell et al., 2001). In brief, after

infections, human macrophages were fixed using 2.5% paraformaldehyde for 10 min at 37oC.

Extracellular L. monocytogenes was detected by immunostaining prior to permeabilization. Cells

were then permeabilized and blocked using 0.2% saponin with 10% normal goat serum 14-16 h

at 4oC and stained for intracellular bacteria and various host proteins. All colocalization

quantifications and image acquisitions of sRBC fed cells were done using a Leica DMIRE2

epifluorescence microscope equipped with a 100X oil objective, 1.4 numerical aperture. Images

of WT or ΔLLO infected cells are confocal z slices taken using a Zeiss Axiovert confocal

81

microscope and LSM 510 software. Volocity software (Improvision) was used to analyse

images. Images were imported into Adobe Photoshop and assembled in Adobe Illustrator.

14 Dynamic Luminol based assays of ROS production

1x105 macrophages in PBS-g (PBS plus 7.5mM glucose) in the presence of 125µM luminol and

100U/ml horseradish peroxidase were pre-warmed for 10 min at 37oC. 15µg of superoxide

dismutase (SOD) was added to quench extracellular production of ROS. Cells were then infected

with WT, ΔLLO or ΔLLOΔPI-PLCΔPC-PLC bacteria at MOI 10 or treated with either 10 µg/ml

PMA alone or 10 µM DPI upon ΔLLO bacterial infection. After centrifugation at 500rpm for 1

min, luminescence was recorded every 10 min for a total of 90 min at 37oC using a Envision

Multimode Plate reader (Perkin Elmer). The same protocol, without the addition of SOD, was

used for the isoluminol assay.

15 Nitroblue tetrazolium (NBT) quantification of

superoxide production

NBT is a compound that when reduced, forms a blue insoluble precipitate called formasan. NBT

quantification of global intracellular superoxide formation was conducted, as previously

described (Keith et al., 2009). In brief, RAW 264.7 macrophages were cultured as described and

infected with wild type, ΔLLO or iLLO or iPFO, with or without IPTG, for 30 min. For controls,

10 µg/ml PMA and 10 µM DPI were used. Cells were washed and 300 µl of freshly prepared and

pre-warmed DMEM with 1.25 mg/ml NBT was added per well. Medium was replaced with

growth media with 10 µg/ml gentamicin and NBT. At 4 h, cells were washed with RPMI, twice

with methanol and allowed to air dry. Reduced formasan particles were solublized with 240 µl

2M KOH and 280 µl DMSO and the resulting solution analyzed spectrophotometrically at 570

nm. All readings were normalized against an uninfected control with NBT treatment. A standard

curve was generated using serial dilutions of a 1 mg/ml NBT solution in 1.2:1.4 2M

KOH/DMSO solution.

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16 Flow Cytometric Quantification of ROS

RAW 264.7 cells were infected with L. monocytogenes strains or treatment conditions for 30 min

as described and cells were detached from 24-well plates using 0.25% tryspin and 1 mM EDTA.

Cells were then stained with CM-H2DCFDA dye (Invitrogen) as per the manufacturer’s protocol.

1x104 macrophages were analyzed per condition in a FACS-Calibur flow cytometer (Becton-

Dickinson). Live macrophages were identified based on a characteristic forward and side

scattering property. Analysis of the flow cytometry data was performed with Flow Jo software

(Tree Star Flow Jo). Sequential gating of the live population, followed by background ROS

fluorescence gating on an uninfected stained control, was performed to calculate the percentage

of macrophages that had an increase in ROS+ fluorescence.

17 Cerium perhydroxide precipitate quantification of

phagosomal ROS

RAW 264.7 cells were infected with L. monocytogenes strains for 30 min as described and cells

washed with 0.1M Tris maleate buffer, pH 7.5 at 37oC, followed by the addition of 0.1 M Tris

maleate, 7% sucrose (BioShop), 1 mM aminotriazole buffer, pH 7.5 for 10 min at 37oC. Next,

cells were treated with 0.1 M Tris maleate, 7% surcrose, 1 mM aminotriazole, 1 mM CeCl3, 0.71

mM NADH, 0.71 mM NADPH buffer pH 7.5 for 20 min at 37oC (all reagents were from Sigma-

Aldrich unless otherwise specified). Cells were subsequently washed with 0.1 M Tris maleate,

7% sucrose and fixed in 2.5% glutaraldehyde in PBS 14-16 h, postfixed in 1% OsO4, stained

with 1% aqueous uranyl acetate, dehydrated in graded series of ethanols and embedded in Epoxy

resin. Samples were then sectioned, stained with 2% uranyl acetate, then 0.2% lead citrate, and

examined on a Tecnai 20 transmission electron microscope (FEI) operating at 200 kV.

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18 Statistical Analysis

Statistical analyses were conducted using GraphPad Prism v4.0a. In all figures, data are

expressed as the mean ± standard error of the mean (s.e.m) from three separate experiments. P

values were calculated using two-tailed two-sample equal variance Student’s t-test unless

specified otherwise. A p-value of less than 0.05 was considered statistically significant and is

denoted by *. p < 0.01 is denoted by ** and p < 0.005 is denoted by ***.

84

Chapter 4

Results

19 Host and bacteria factors regulate autophagic targeting of L. monocytogenes during early stages of infection in macrophages

Overview: Listeria monocytogenes is a bacterial pathogen that can escape the phagosome and

replicate in the cytosol of host cells during infection. These bacteria can be targeted by

autophagy, with up to 35% becoming LC3+ at 1 h p.i. (Birmingham et al., 2007a). Here, I show

that LC3+ L. monocytogenes are present within phagosomes at this time, and not the cytosol.

Diacylglycerol (DAG) production by both bacterial and host phospholipases is required for LC3

recruitment to bacteria. Reactive oxygen species (ROS) production by the NOX2 NADPH

oxidase, a downstream consequence of DAG production, is required for LC3 recruitment to

bacteria. Protein ubiquitination was recently shown to play a role in targeting cytosolic L.

monocytogenes to autophagy (Yoshikawa et al., 2009). However, I found that protein

ubiquitination is not associated with LC3+ bacteria within phagosomes at early stages of

infection. My data demonstrate that distinct signals mediate targeting of L. monocytogenes to the

autophagy pathway at different stages of infection within host cells.

19.1 Autophagy targets wild type L. monocytogenes within

macrophage phagosomes prior to phagosomal escape.

Consistent with previous work (Birmingham et al., 2007a; Meyer-Morse et al., 2010; Py et al.,

2007), I found that LC3 recruitment to L. monocytogenes peaks at 1 h p.i. in an LLO-dependent

manner (Figure 19.1.1 A and 19.1.1 B). Therefore, I sought to characterize the mechanisms that

regulate autophagic targeting at this time point. Based on published reports of L. monocytogenes

phagosome escape kinetics (Beauregard et al., 1997; de Chastellier and Berche, 1994; Henry et

al., 2006), it is unclear whether L. monocytogenes is in the phagosome or cytosol during the peak

of autophagic targeting.

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Figure 19.1.1. Autophagy targets wild type L. monocytogenes during macrophage infection.

(A) Confocal images of RAW 264.7 macrophages transfected with GFP-LC3 and infected for 1

h with wild type L. monocytogenes. (B) Quantification of LC3 colocalization with intracellular

wild type or ΔLLO bacteria over time.

86

To determine whether L. monocytogenes is present inside phagosomes or the cytosol at 1 h p.i., I

first quantified the percentage of L. monocytogenes that were both LC3+ and LAMP-1+. I found

that the majority (78%) of LC3+ bacteria were also LAMP-1+ (Figure 19.1.2 A and 19.1.2 B),

suggesting that L. monocytogenes within phagosomes are targeted by autophagy.

To confirm this, I employed a recently described probe constructed from the cell wall binding

domain (CBD) of a Listeria bacteriophage protein (Henry et al., 2006). A yellow fluorescent

protein fusion to CBD (CBD-YFP) was developed by Swanson and colleagues to track

phagosome escape by L. monocytogenes (Henry et al., 2006). CBD-YFP expressed within host

cells binds to cytosolic bacteria, resulting in an accumulation of YFP fluorescence around the

bacteria (Henry et al., 2006). Upon escape from the phagosome, L. monocytogenes also

polymerizes actin via the ActA virulence factor. Thus, I first confirmed the ability of the CBD-

YFP probe to detect cytosolic L. monocytogenes by quantifying the number of CBD-YFP+

bacteria that were also positive for polymerized actin at 4 h p.i. when bacteria are known to be in

the cytosol. As expected, the majority (60%) of CBD-YFP+ L. monocytogenes were also actin+,

confirming that the CBD-YFP can detect cytosolic L. monocytogenes after their escape from

phagosomes (Figure 19.1.2 C and 19.1.2 D). In contrast, I did not observe any accumulation of

CBD-YFP to LC3+ L. monocytogenes at 1 h p.i. (Figure 19.1.2 E and 19.1.2 F). Together, this

data demonstrates that L. monocytogenes is targeted by autophagy within phagosomes at early

stages of infection.

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Figure 19.1.2. Autophagy targets wild type L. monocytogenes within macrophage

phagosomes during early stages of infection. (A) Confocal images of RAW 264.7

macrophages transfected with GFP-LC3 and infected for 1 h with wild type L. monocytogenes.

Cells were then stained for Lamp-1. (B) Quantification of the percentage of LC3+ bacteria that

are Lamp-1+ or Lamp-1-. (C) Confocal images of RAW 264.7 macrophages transfected with

GFP-LC3 and infected for 1 h with wild type L. monocytogenes. Cells were then stained for

ubiquitinated proteins (Ub+). (D) Quantification of the percentage of LC3+ L. monocytogenes

that are Ub+ or Ub-. The inner panels represent a higher magnification of the boxed areas. (E)

Confocal images of RAW 264.7 macrophages expressing RFP-LC3 and CBD-YFP and infected

for 1 h with wild type L. monocytogenes. (F) Quantification of the percentage of LC3+ L.

monocytogenes that are CBD-YFP+ or CBD-YFP-.

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Previous studies suggest that protein ubiquitination may mediate autophagic targeting of L.

monocytogenes in the cytosol (Yoshikawa et al., 2009). LC3 targeting of S. Typhimurium and S.

flexneri within phagosomes has also been shown to involve protein ubiquitination (Birmingham

et al., 2006; Dupont et al., 2009; Thurston et al., 2009; Zheng et al., 2009). These observations

suggested the possibility that protein ubiquitination may initiate autophagic targeting of L.

monocytogenes within phagosomes. However, I observed that the majority (98%) of LC3+ L.

monocytogenes did not colocalize with ubiquitinated proteins at 1 h p.i. (Figure 19.1.2 C and

20.1.3 B). These results indicated that autophagic targeting of L. monocytogenes within

phagosomes is regulated by signals distinct from those in the cytosol.

19.2 Host and bacterial factors promote DAG accumulation on

phagosomes containing L. monocytogenes

DAG is known to regulate autophagy of S. Typhimurium (Shahnazari et al., 2010). However, the

role of DAG in autophagy of L. monocytogenes has not been examined. Previous studies have

shown that L. monocytogenes infection induces DAG production (Sibelius et al., 1996a; Smith et

al., 1995) in a manner dependent on the activity of LLO (Sibelius et al., 1996a), bacterial PLCs

(Smith et al., 1995) and possibly host phospholipases (Goldfine et al., 2000). Therefore, it was

conceivable that DAG production on the phagosome may play a role in autophagic targeting of

L. monocytogenes at early stages of infection.

To visualize DAG during L. monocytogenes infection I employed a fluorescent probe

constructed from the DAG binding C1 domain of PKCδ fused to green fluorescent protein

(PKCδ-C1-GFP). The majority (90%) of LC3+ L. monocytogenes were observed to be DAG+

(Figure 19.2.1 A). However, only 25% of DAG+ L. monocytogenes were observed to be LC3+

(Figure 19.2.1 B). Furthermore, DAG colocalization with L. monocytogenes was observed prior

to the peak of autophagy at 1 h p.i. (Figure 19.2.1 C and 19.2.1 D). Together, these observations

suggest that DAG may serve as an upstream signal for autophagy.

90

Figure 19.1.3. CBD-YFP+ L. monocytogenes is also actin+. (A) Confocal images of RAW

264.7 macrophages transfected with CBD-YFP and infected for 1 h with wild type L.

monocytogenes. Cells were then stained with phalloidin to label F-actin. The brightness and

contrast for the image (but not the inset) was enhanced to allow visualization of fluorescence

inside the cell. (B) Quantification of the percentage of CBD-YFP+ bacteria that are actin+. Size

bars = 5 µm.

92

Figure 19.2.1. Host and bacterial factors promote DAG accumulation on phagosomes

containing L. monocytogenes. (A) Confocal images of RAW 264.7 macrophages co-transfected

with RFP-LC3 and PKCδ-C1-GFP and infected for 1 h with wild type L. monocytogenes. PKCδ-

C1 is a specific probe for DAG. Size bar = 5 µm. Quantification at 1 h p.i. of the percentage of

LC3+ L. monocytogenes that are PKCδ-C1+ or PKCδ-C1-. (B) Cells treated as (A).

Quantification at 1 h p.i. of the percentage of PKCδ-C1+ L. monocytogenes that is also LC3+.

(C) Confocal images of RAW 264.7 macrophages transfected with PKCδ-C1-GFP and infected

for 1 h with wild type or ΔLLO bacteria. Size bar = 5 µm. (D) Quantification of the percentage

of intracellular wild type, ΔLLO or ΔPI-PLCΔPC-PLC L. monocytogenes that are DAG+ over

time.

93

The kinetics of DAG colocalization with L. monocytogenes lacking both PLCs (ΔPI-PLCΔPC-

PLC) was examined. As shown in Figure 19.2.1 D, the percentage of DAG+ ΔPI-PLCΔPC-PLC

bacteria was approximately 50% less than wild type bacteria at 30 min and 45 min p.i.. This

suggests that bacterial PLCs contribute to DAG production on phagosomes containing L.

monocytogenes. However, the decrease in DAG localization to ΔPI-PLCΔPC-PLC bacteria was

not complete, raising the likely possibility that host factors also contribute to DAG production.

Indeed, host phospholipases were previously shown to contribute to DAG production during L.

monocytogenes infection (Smith et al., 1995).

I determined the effect of inhibiting host enzymes that produce DAG. Recent studies of S.

Typhimurium infection indicate that DAG is produced by host phospholipase D (PLD) and

phosphatidic acid phosphatase (PAP) (Shahnazari et al., 2010). I, therefore, employed inhibitors

of PLD (1-butanol) and PAP (propranolol hydrochloride) to determine changes in DAG

production on phagosomes containing wild type or ΔPI-PLCΔPC-PLC bacteria. Inhibition of

PLD or PAP resulted in a decrease in DAG colocalization to the ΔPI-PLCΔPC-PLC mutants

(Figure 19.2.2). This effect was not observed with tert-butanol, an isomer of 1-butanol that has

no inhibitory effect on PLD. This finding suggests that host factors PLD and PAP contribute to

the accumulation of DAG at L. monocytogenes containing phagosomes. Surprisingly, DAG

colocalization to phagosomes containing wild type L. monocytogenes was not altered upon

addition of PLD or PAP inhibitors. This observation strongly suggests that bacterial PLCs

compensate for inhibition of host DAG production. These results indicate that both host and

bacterial factors contribute to DAG accumulation on phagosomes containing L. monocytogenes.

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Figure 19.2.2. Inhibition of host DAG producers can further decrease DAG localization to

ΔPI-PLCΔPC-PLC L. monocytogenes. Quantification at 45 min p.i. of intracellular wild type or

ΔPI-PLCΔPC-PLC L. monocytogenes that are DAG+ upon treatment with growth media (GM),

DMSO, 1-butanol, propranolol, or tert-butanol.

95

19.3 DAG production is required for autophagic targeting of L.

monocytogenes at early stages of infection.

To investigate whether DAG plays a role in autophagic targeting of L. monocytogenes, I assessed

LC3 recruitment to both wild type and ΔPI-PLCΔPC-PLC mutants following treatment with

PLD and PAP inhibitors (Figure 19.3 A). Similar to what I observed with DAG localization in

Figure 19.2.2, LC3 targeting to wild type L. monocytogenes did not change with PLD or PAP

inhibition. In contrast, LC3 targeting of the ΔPI-PLCΔPC-PLC mutant was significantly

inhibited upon treatment with either the PLD inhibitor (90% decrease) or the PAP inhibitor (73%

decrease). This suggests that DAG is important for LC3 recruitment to L. monocytogenes

containing phagosomes.

To confirm the contribution of PLD-mediated generation of DAG to autophagy, dominant

negative constructs for both isoforms of PLD (PLD1 and PLD2) were employed. Similar to the

pharmacological evidence, dominant negative forms of PLD reduced DAG recruitment (40-62%

decrease) (Figure 19.3 B) as well as LC3 recruitment (60-61% decrease) (Figure 19.4 C) to ΔPI-

PLCΔPC-PLC mutants. Taken together, both bacterial and host mediated production of DAG

contribute to autophagic targeting of phagosomes containing L. monocytogenes.

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Figure 19.3. DAG production is required for autophagic targeting of L. monocytogenes

during early stages of infection. (A) Quantification at 1 h p.i. of intracellular wild type or ΔPI-

PLCΔPC-PLC L. monocytogenes that are LC3+ following treatment with DMSO, 1-butanol,

propranolol, or tert-butanol. (B) Quantification at 1 h p.i. of DAG+ intracellular ΔPI-PLCΔPC-

PLC L. monocytogenes in RAW 264.7 macrophages that were co-transfected with LC3-GFP and

either PLD1 DN or PLD2 DN constructs. (C) Quantification at 1 h p.i. of LC3+ intracellular wild

type or ΔPI-PLCΔPC-PLC L. monocytogenes in RAW 264.7 macrophages that were co-

transfected with LC3-GFP and either PLD1 dominant negative (DN) or PLD2 DN constructs.

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19.4 ROS production by the NOX2 NADPH oxidase is required

for autophagic targeting of L. monocytogenes at early stages

of infection.

PLD-mediated generation of DAG is upstream of ROS production via the NOX2 NADPH

oxidase (Usatyuk et al., 2009). I previously showed that NOX2-generated ROS mediates LC3

targeting to phagosomes containing S. Typhimurium and IgG-coated latex beads (Huang et al.,

2009). Therefore, it was conceivable that DAG promotes autophagy of L. monocytogenes via

ROS production.

To test this hypothesis, I inhibited ROS production via the addition of the NOX2 NADPH

oxidase inhibitor, diphenyliodonium (DPI), which at 1 h p.i. significantly reduced LC3

colocalization with L. monocytogenes (Figure 19.4 A). The same reduction in autophagy was

also seen when antioxidants, resveratrol and α-tocopherol, were added to decrease ROS levels

(Figure 19.4 A). These findings are consistent with a role for ROS in regulating autophagy of L.

monocytogenes.

I next evaluated autophagic targeting of L. monocytogenes in bone-marrow macrophages

deficient in NOX2 NADPH oxidase activity (gp91phox-/-) (Figure 19.4 B and 19.4 C). I found that

gp91phox-/- macrophages displayed a significant reduction in LC3 colocalization with wild type L.

monocytogenes (Figure 19.4 B and 19.4 C). Furthermore, consistent with previous studies

(Birmingham et al., 2007a; Meyer-Morse et al., 2010; Py et al., 2007), LC3 recruitment to

bacteria was dependent on LLO expression. Colocalization of LC3 with ΔLLO mutant bacteria

was restored upon LLO complementation (Figure 19.4 C). Taken together, ROS production by

the NOX2 NADPH oxidase appears to play a key role in autophagic targeting of phagosomes

containing L. monocytogenes.

99

Figure 19.4. ROS production by the NOX2 NADPH oxidase is required for autophagic

targeting of L. monocytogenes during early stages of infection. (A) RAW 264.7 cells were

infected with wild type L. monocytogenes for 1 h in the presence or absence of DPI, resveratrol

or α-tocopherol, as indicated. The percentage of intracellular bacteria that are LC3+ upon

treatment with each agent is shown. (B) Confocal images of wild type or gp91phox-/- bone

marrow-derived macrophages transfected with LC3-GFP and infected for 1 h with wild type

bacteria. Size bar = 5 µm. (C) Quantification at 1 h p.i. of the percentage of intracellular L.

monocytogenes that are LC3+ in bone marrow-derived macrophages isolated from wild type or

gp91phox-/- mice. Wild type bacteria were compared to ΔLLO bacteria (negative control), as well

as a ΔLLO mutant complemented with LLO.

100

19.5 LC3 targeting is dependent on LLO pore-formation

Given that LLO is necessary and sufficient to induce LC3 targeting, I next examined whether

LLO-mediated signaling, pore-formation or both is responsible for facilitating LC3 recruitment.

To address this question, PFA or heat-killed wild type L. monocytogenes was fed to

macrophages. Regardless of opsonization with IgG, non-viable L. monocytogenes did not recruit

LC3 at 1 h post addition (Figure 19.5 A). Given that LC3 is not recruited to ΔLLO L.

monocytogenes (Figure 19.1 and 19.4), these results suggest that active LLO production by

viable L. monocytogenes is required for LC3 targeting.

To directly test the contribution of LLO pore-formation to LC3 targeting, a series of L.

monocytogenes LLO point mutants exhibiting varying degrees of hemolytic activity were next

employed. The C484A mutant has 75% of wild type hemolytic activity, the C484S mutant has

20%, the W491A mutant has 5% while the W492A mutant has 0.1% (Michel et al., 1990). These

mutants are identical other than their point mutation and thus they likely share LLO signaling

properties, but not the pore-forming property. At 1 h p.i., roughly 30% of wild type L.

monocytogenes were LC3+ while the mutant bearing a transposon insertion in hly (Tn916) did

not colocalize with LC3 (Figure 19.5 B). However, roughly 20% of C484A mutants were LC3+,

17% of C484S mutants were LC3+, 5% of W491A mutants were LC3+, while 3% of W492A

mutants were LC3+. These observations indicate that LC3 targeting is dependent on LLO

hemolytic or pore-forming activities.

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Figure 19.5. LC3 targeting is dependent on LLO pore-forming ability secreted from live L.

monocytogenes. (A) PFA fixed or heat killed wild type L. monocytogenes (PFA-WT or HK-WT)

with or without opsonization with IgG (IgG PFA-WT or IgG HK-WT) were added to LC3-GFP

expressing RAW 264.7 macrophages. Percentage LC3 colocalization to intracellular bacteria at 1

h post addition was quantified. (B) RAW 264.7 macrophages were infected with a panel of LLO

point mutants: C484A, C484S, W491A, and W492A. The wild type is the background strain

used to generate the point mutants and the Tn916 mutant is an LLO disrupted mutant. The

brackets following each mutant denote the percentage of wild type L. monocytogenes hemolytic

activity exhibited by the mutant. LC3 targeting of intracellular mutants was quantified at 1 h p.i..

102

19.6 ROS production by the NOX2 NADPH oxidase is required

for SLAP formation at later stages of infection.

Here I focused on autophagy during the early stages of infection prior to phagosomal escape into

the cytosol. The outcome of this autophagic targeting event depends on many factors. Previously,

our laboratory showed that one outcome of this early autophagic targeting event is the formation

of spacious Listeria containing phagosomes (SLAPs), compartments that are non-degradative,

LAMP-1+ and LC3+ (Birmingham et al., 2007b). Formation of SLAPs was found to require

autophagy in the host cell, and expression of LLO by bacteria. It was proposed that SLAPs

represent a “stalemate” between the host and bacteria, allowing slow bacterial replication in

SLAPs that allows chronic L. monocytogenes infection in a host (Birmingham et al., 2007b).

Indeed, compartments resembling SLAPs have been observed in a mouse model of L.

monocytogenes chronic infection (Bhardwaj et al., 1998). Consistent with the notion that early

autophagic targeting leads to SLAP formation, pharmacological inhibition of ROS production by

the NOX2 NADPH oxidase reduced the formation of SLAPs (Figure 19.6). SLAP formation was

decreased in gp91phox-/- bone marrow-derived macrophages compared to wild type macrophages.

These findings demonstrate that ROS-mediated autophagy at 1 h p.i. is required for the later

formation of SLAPs. Thus, signals that mediate early autophagic targeting of L. monocytogenes

may also are likely to be important in determining events that lead to establishment of chronic

infection.

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Figure 19.6 ROS production by the NOX2 NADPH oxidase is required for the generation

of SLAPs. (A) Confocal images of RAW 264.7 macrophages infected for 4 h with wild type L.

monocytogenes, with or without DPI. Cells were stained with LAMP-1 antibodies. Size bar = 5

µm. White arrowheads indicate Spacious Listeria phagosomes (SLAPs) while the white arrow

indicates LAMP-1 colocalization with bacteria that are not in SLAPs (B) Quantification at 4 h

p.i. of the percentage of infected macrophages that form SLAPs in the presence or absence of

DPI, resveratrol or a-tocopherol, as indicated. (C) Confocal images of wild type or gp91phox-/-

bone marrow-derived macrophages infected for 8 h with wild type L. monocytogenes. Cells were

stained with LAMP-1 antibodies. Size bar = 5 µm. White arrowheads indicate SLAPs while the

white arrows indicate LAMP-1 colocalization and not SLAPs. (D) Quantification at 8 h p.i. of

the percentage of infected wild type or gp91phox-/- bone marrow-derived macrophages that form

SLAPs from C.

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19.7 Summary

In my graduate studies, I examined the signals that mediate autophagic targeting of L.

monocytogenes during the early stages of infection in host cells. I showed that L. monocytogenes

is largely confined within LAMP-1+ phagosomes at the peak of autophagy (1 h p.i.). A recent

study by Sasakawa and colleagues indicates that cytosolic L. monocytogenes deficient in ActA

are coated with ubiquitinated proteins and subsequently targeted by autophagy 4 h p.i.

(Yoshikawa et al., 2009). Within the phagosome, however, my studies indicate that autophagic

targeting is not dependent on protein ubiquitination but is instead dependent on DAG and ROS

mediated signaling. Thus, L. monocytogenes can be targeted by autophagy by distinct

mechanisms at different stages of infection within host cells. These findings highlight the

complex nature of autophagy regulation and its versatility in targeting pathogens within different

compartments in eukaryotic cells.

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20 Listeriolysin O suppresses PLC-mediated activation of the microbicidal NADPH oxidase to promote Listeria monocytogenes infection

Overview: Listeria monocytogenes is an intracellular bacterial pathogen that utilizes two

phospholipases C (PI-PLC and PC-PLC) and a pore-forming cytolysin (listeriolysin O, LLO) to

escape the phagosome and replicate within the host cytosol. Here, I show that PLCs are capable

of activating the phagocyte NOX2 NADPH oxidase during L. monocytogenes infection, a

response that would adversely affect pathogen survival. However, secretion of LLO inhibits the

NADPH oxidase by preventing its localization to phagosomes. LLO-deficient bacteria can be

complemented by perfringolysin O, a related cytolysin, suggesting that other pathogens may also

use pore-forming cytolysins to inhibit the NOX2 NADPH oxidase. My studies demonstrate that

while the PLCs induce antimicrobial NOX2 NADPH oxidase activity, this effect was alleviated

by the pore-forming activity of LLO. Therefore, the combined activities of PLCs and LLO on

membrane lysis and their opposing effects on the NOX2 NADPH oxidase allow L.

monocytogenes to efficiently escape the phagosome while avoiding the microbicidal respiratory

burst.

20.1 LLO inhibits ROS production by the NOX2 NADPH oxidase

I performed dynamic measurements of intracellular and extracellular ROS production in bone

marrow-derived macrophages (BMDM) using luminol or isoluminol, respectively (Figure 20.1.1

A-D). During infection by wild type L. monocytogenes, both intracellular and extracellular ROS

production peaked at 10 min p.i. and decreased at later time points (Figure 20.1.1 A and 20.1.1

C). Treatment of cells with phorbol 12-myristate 13-acetate (PMA), a known activator of the

NOX2 NADPH oxidase, served as a positive control (Figure 20.1.1 B and 20.1.1 D). These

findings demonstrate that L. monocytogenes can induce rapid activation of ROS in leukocytes,

consistent with previous observations (Sibelius et al., 1999).

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Next, I examined ROS production in cells infected with LLO-deficient bacteria (ΔLLO, lacking

the hly gene). I observed higher intracellular ROS production in ΔLLO infected cells compared

to wild type infected cells at 40-70 min post p.i. (Figure 20.1.1 A and 20.1.1 C; Table 20.1 A).

Addition of the flavoprotein inhibitor diphenyliodonium (DPI, a known NADPH oxidase

inhibitor) prevented ROS production in ΔLLO infected cells (Figure 20.1.1 B and 20.1.1 D;

Table 20.1 A and 20.1 B). The kinetics of extracellular ROS production were similar upon

infection with wild type or ΔLLO L. monocytogenes (Figure 20.1.1 C, Table 20.1 B). These

observations suggest that LLO inhibits intracellular ROS production.

Since I observed a significant difference in ROS production between wild type and ΔLLO

infected cells during 40-70 min p.i. (Figure 20.1.1 A and 20.1.1 C; Table 20.1.1 A and 20.1.1 B),

I further examined intracellular ROS production in this window of infection using the 5-(and-6)-

chloromethyl-2-7-dichlorodihydrofluorescein diacetate (CM-H2DCFDA) dye. RAW 264.7

macrophages were infected with either wild type or ΔLLO bacteria and CM-H2DCFDA dye was

added 30 min p.i. and examined at 1 h p.i. As expected, treatment of cells with PMA resulted in

robust ROS production (Figure 20.1.2 A and 20.1.2 B). Consistent with the luminol assay, I

observed little ROS production in wild type infected cells but high levels of ROS production in

ΔLLO infected cells, comparable to those attained by stimulation with PMA (Figure 20.1.2 A

and 20.1.2 B).

I also assessed NOX2 NADPH oxidase activity by specifically measuring its product,

superoxide, using the nitroblue tetrazolium (NBT) reduction assay (described in (Keith et al.,

2009)). ΔLLO bacteria induced a marked production of superoxide, while infection with wild

type bacteria did not (Figure 20.1.3). These results indicate that LLO inhibits ROS production

during infection. To further define the role of LLO, I employed an LLO-deficient strain that

inducibly expresses LLO (iLLO) in response to isopropyl β-D-1-thiogalactopyranoside (IPTG).

In the absence of IPTG, iLLO bacteria triggered ROS production. However, in the presence of

IPTG, iLLO bacteria did not trigger ROS production upon infection (Figure 20.1.3).

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Figure 20.1.1. ΔLLO mutants induce greater ROS production compared to WT or

ΔLLOΔPI-PLCΔPC-PLC L. monocytogenes during infection in macrophages. (A-B)

Intracellular ROS production was assessed via the luminol assay. BMDMs were (A) infected at

an MOI of 10 with wild type (WT) or ΔLLO bacteria or (B) treated with PMA or DPI upon

ΔLLO bacteria infection. Background ROS production from uninfected cells (BKGD) was also

assessed. Time 0 represents the first reading shortly after the addition of bacteria or drugs. The

figure shows data from one representative experiment done in duplicate where each data point

represents the average and range. The experiment was performed a total of three independent

times (see Table 20.1 A). (C-D) Extracellular ROS production was assessed via the isoluminol

assay. BMDMs were (C) infected at an MOI of 10 with wild type (WT) or ΔLLO bacteria or (D)

treated with PMA or DPI in presence of ΔLLO bacteria infection. ROS production from

uninfected cells (BKGD) was also assessed. Time 0 represents the first reading shortly after the

addition of bacteria or drugs. The figure shows data from one representative experiment done in

duplicate where each data point represents the average and range. The experiment was performed

a total of three independent times (see Table 20.1 B).

109

Table 20.1. Mean and SEM of luminol and isoluminol readings from primary BMDM

infected with WT or ΔLLO bacteria. (A) Mean and SEM of three independent luminol assays

done on BMDM that were infected at MOI 10 with WT or ΔLLO bacteria, or treated with PMA

or DPI upon ΔLLO infection. (B) Mean and SEM of three independent isoluminol assays done

on BMDMs that were infected with WT or ΔLLO bacteria, or treated with PMA or DPI upon

ΔLLO infection.

110

Figure 20.1.2. LLO inhibits PLC-mediated ROS production by the NOX2 NADPH oxidase.

(A) RAW 264.7 macrophages were treated with PMA or infected with the indicated bacterial

strain (MOI 10) with or without DPI. ROS production was assessed via flow cytometry with

CM-H2DCFDA. The CM-H2DCFDA dye was added 30 min p.i. and cells were analyzed at 60

min p.i.. Flow cytometry analysis was conducted on live populations, gated against a stained

unstimulated control (red). Percentages indicated on the plots refer to the portion of cells that are

ROS+. Representative plots are shown. (B) Quantification of the percentage of ROS+ cells that

were infected with the indicated bacterial strains. Graph represents mean ± SEM from five

independent experiments.

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Figure 20.1.3. LLO inhibits ROS production by NOX2 NADPH oxidase. RAW 264.7

macrophages were either treated with PMA or infected at a MOI of 10:1 with the indicated

bacterial strain. Superoxide production was assessed using NBT, a compound that upon

reduction, forms a black formasan precipitate. Formasan formation was assessed

spectrophotometrically in cell lysates (see Methods). NBT was added 30 min p.i..

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20.2 LLO can inhibits ROS production in other cell types

Addition of purified LLO to human neutrophils was sufficient to inhibit PMA-induced ROS

production (Figure 20.2 A) at concentrations that did not cause cell toxicity (Figure 20.2 B).

Therefore, LLO is both necessary and sufficient for inhibition of intracellular ROS production by

the NADPH oxidase.

Figure 20.2. LLO is capable of inhibiting ROS production by PMA activated human

neutrophils. (A) NBT quantification assay of PMA stimulated human neutrophils treated with

varying doses of recombinant LLO (rLLO) for 60 min. (B) Percentage neutrophil viability was

assessed using the trypan blue exclusion assay at 60 min post treatment with the various doses of

rLLO.

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20.3 LLO inhibits phagosomal production of ROS

To visualize the localization of ROS production during L. monocytogenes infection, I employed

transmission electron microscopy (TEM) with cerium chloride (Vazquez-Torres et al., 2000).

IgG-coated 3.8 µm latex beads were fed to RAW 264.7 macrophages as positive controls of

phagosomal ROS production. A dark cerium precipitate was observed in phagosomes containing

latex beads, indicative of ROS production in this compartment (Figure 20.3 A, 20.3 B and 20.3

G), which was inhibited by DPI treatment (Figure 20.3 C, 20.3 D and 20.3 G). ROS production

was not observed in phagosomes containing wild type L. monocytogenes (Figure 20.3 F and 20.3

G). However, marked production of ROS was observed in phagosomes containing ΔLLO

bacteria (Figure 20.3 E and 20.3 G), but not in phagosomes containing ΔLLO ΔPI-PLC ΔPC-

PLC bacteria (Figure 20.3 G). These results indicate that LLO inhibits PLC-mediated NOX2

NADPH oxidase-dependent ROS production in the phagosome.

20.4 L. monocytogenes PLCs contribute to the induction of ROS

production by the NOX2 NADPH oxidase

The observation that ΔLLO bacteria induced high levels of ROS production, comparable to those

observed in cells treated with PMA, suggested that bacterial products have the capacity to

promote NOX2 NADPH oxidase activity in the absence of LLO. It was previously shown that

PLCs from Clostridium perfringens, Pseudomonas aeruginosa and Bacillus cereus are linked to

activation of the NOX2 NADPH oxidase (Styrt et al., 1989; Titball, 1993). These enzymes

generate diacylglycerol, a cofactor for activation of protein kinase C (PKC), which contributes to

NOX2 NADPH oxidase activation via the phosphorylation of p47phox (Fontayne et al., 2002).

Similarly, L. monocytogenes PLCs generate diacylglycerol and activate PKC during infection of

macrophages (Camilli et al., 1993; Goldfine et al., 1993; Wadsworth and Goldfine, 2002).

Therefore I hypothesized that L. monocytogenes PLCs could activate NOX2 NADPH oxidase in

the absence of LLO.

114

Figure 20.3. LLO inhibits ROS production in phagosomes. (A, B) Phagosomal production of

ROS was assessed via transmission electron microscopy (TEM). RAW 264.7 macrophages were

treated with IgG-coated latex beads in the presence of cerium chloride. Reduction of cerium

chloride results in the formation of cerium perhydroxide precipitate. The presence of these

electron-dense products in TEM was indicative of NOX2 NADPH oxidase production of ROS.

(B) is a higher magnification view of the boxed region in (A). (C, D) Experiment conducted as

in (A) and (B) but in the presence of DPI. (D) is a higher magnification view of the boxed region

in (C). RAW 264.7 macrophages were infected with either WT or ΔLLO bacteria at MOI 10. (E)

Representative TEM of a ΔLLO-containing phagosome at 60 min p.i.. (F) Representative TEM

of a wild type-containing phagosome at 60 min p.i.. (G) The percentage of cerium perhydroxide+

phagosomes at 60 min p.i. were quantified. One hundred phagosomes were assessed per sample

from four independent experiments. Bars represent mean ± SEM. All scale bars are 1 µm.

115

To test this hypothesis, I examined the effect of L. monocytogenes PLCs on ROS production. In

contrast to ΔLLO bacteria, a strain lacking LLO and both PLCs (ΔLLO ΔPI-PLC ΔPC-PLC)

produced little ROS (Figure 20.1.1 A, Table 20.1 A and 20.1.2 B). Infection of RAW 264.7

macrophages with a strain that lacks both PLCs (ΔPI-PLC ΔPC-PLC, lacking plcA and plcB

genes) resulted in ROS production comparable to that of uninfected controls and cells infected

with wild type bacteria (Figure 20.1.2 B). Thus, these observations suggest that PLCs promote

low levels of ROS production during infection by wild type bacteria.

Next, I treated cells with purified PI-PLC and observed a significant induction of ROS

production in macrophages that was inhibited by the broad spectrum PKC inhibitors Rottlerin

and GÖ6983 (Figure 20.4 A). ROS production by PI-PLC was also inhibited by concurrent

treatment with purified LLO. These results suggest that PI-PLC is sufficient to drive NOX2

NADPH oxidase activation by stimulating PKC and that LLO is sufficient to inhibit this

deleterious consequence of PLC activity. This is consistent with a previous finding that L.

innocua expressing PI-PLC induces greater ROS production than wild type L. innocua in

neutrophils (Sibelius et al., 1999). However, L. monocytogenes mutants that only express PI-PLC

(and not PC-PLC or LLO) or PC-PLC (and not PI-PLC or LLO) do not induce significant ROS

production (Figure 20.4 B). One possible explanation for these observations is that

physiologically both PLCs are needed to mediate ROS production. However, higher doses of PI-

PLC may be sufficient to drive NOX2 NADPH oxidase activation by stimulating PKC.

Collectively, my results indicate that bacterial PLCs induce ROS production and that LLO

inhibits this deleterious consequence of PLC activity.

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Figure 20.4. Both PLCs are required to induce ROS production by macrophages during

infection. (A) RAW 264.7 macrophages were treated with the indicated strains with or without

15µM Rottlerin or 10nM Go6983, PKC inhibitors, for 60 min and CM-H2DCFDA dye was used

to measure ROS production. Quantification of the percentage of ROS+ cells was done using flow

cytometry. Bars represent mean ± SEM from three independent experiments. (B) RAW 264.7

macrophages were infected of MOI 10:1 with WT, ΔLLO, ΔLLOΔPC-PLC, ΔLLOΔPI-PLC

bacteria for 30 min. Quantification of the percentage of ROS+ cells that were infected with the

indicated bacterial strains. Graph represents mean ± SEM from five independent experiments.

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20.5 LLO deficient bacteria survive in NOX2 NADPH oxidase

deficient macrophages

The relevance of LLO-mediated inhibition of the NOX2 NADPH oxidase on intracellular L.

monocytogenes survival/replication was determined by gentamicin protection assays using bone

marrow-derived macrophages (BMDM) isolated from C57BL/6 and gp91phox-/- mice. The number

of intracellular ΔLLO bacteria decreased over time in C57BL/6 BMDM (Figure 20.5 A),

consistent with previous observations (Alberti-Segui et al., 2007). In contrast, ΔLLO bacteria

displayed extended survival in gp91phox-/- BMDM. Wild type L. monocytogenes replicated at a

similar rate in both C57BL/6 and gp91phox-/- BMDM (Figure 20.5 B). ΔLLO bacteria that were

complemented by expression of LLO (ΔLLO + LLO) displayed a similar replication profile as

that of wild type bacteria (Figure 20.5 C).

20.6 LLO deficient bacteria survive in NOX2 NADPH oxidase

deficient mice

The in vivo relevance of LLO-mediated inhibition of the NOX2 NADPH oxidase was examined

in C57BL/6 and gp91phox-/- mice via infection by wild type L. monocytogenes or ΔLLO bacteria.

Wild type L. monocytogenes replicated significantly over the course of 24 h in both C57BL/6

and gp91phox-/- mice (Figure 20.6 A). The bacterial load at 24 h was higher in gp91phox-/-

compared to C57BL/6 mice (Figure 20.6 B), as previously reported (Dinauer et al., 1997).

Histological examination of the liver revealed a greater number of granulomas in the gp91phox-/-

mice (Figure 20.6 C). However, clearance of ΔLLO bacteria was significantly impaired in the

gp91phox-/- mice as compared to C57BL/6 mice (Figure 20.6 D and 20.6 E). Consistent with this

result, granulomas were observed at 24 h post-infection in the livers of gp91phox-/- mice infected

with ΔLLO bacteria, but not C57BL/6 mice (Figure 20.6 F). These results indicate that LLO-

mediated inhibition of NOX2 NADPH oxidase activity is required for L. monocytogenes survival

both in vitro and in vivo.

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Figure 20.5. ΔLLO bacteria survive in NOX2 NADPH oxidase-deficient macrophages in

vitro. (A, B, C) BMDM from C57BL/6 or gp91phox-/- mice were infected with WT, ΔLLO or

ΔLLO + LLO bacteria. The number of intracellular bacteria (colony forming units) was

quantified by gentamicin protection assay at the indicated times. Data represent the mean ± SEM

for three independent experiments.

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Figure 20.6. ΔLLO bacteria survive in NOX2 NADPH oxidase-deficient macrophages in

vitro and in vivo. (A) C57BL/6 and gp91phox-/- mice were intravenously infected with wild type

L. monocytogenes. Mice were sacrificed at the indicated times and livers removed for

quantification of bacterial load. Graphs represent bacterial CFUs from three independent

experiments for a total of six mice per condition per time point. Statistical analyses were

performed using a non-parametric Mann-Whitney test to assess significance. (B) Comparison of

bacterial CFUs (mean ± SEM) recovered at 24 h post-infection from C57BL/6 and gp91phox-/-

mice from three independent experiments were compared using the non-parametric Mann-

Whitney test. (C) Liver sections from both C57BL/6 and gp91phox -/- mice 24 h post-infection

were embedded and mounted. Tissues were stained with H&E and representative images were

acquired. Scale bar represents 100 µm. (D) C57BL/6 and gp91phox-/- mice were intravenously

infected with 109 ΔLLO bacteria in a volume of 200µl. Mice were sacrificed at the indicated

times and livers removed for quantification of bacterial load. Data represent results from three

independent experiments with a total of six mice per condition per time point. Statistical analyses

were performed using a non-parametric Mann-Whitney test to assess significance. NS = not

significant. (E) Bacterial CFU in the liver (mean ± SEM) recovered at 48 h post-infection from

C57BL/6 and gp91phox-/- mice from three independent experiments were compared using the non-

parametric Mann-Whitney test. (F) Liver sections from C57BL/6 and gp91phox-/- mice 24 h post-

infection were embedded and mounted. Tissues were stained with hematoxylin and eosin (H&E)

and representative images are shown. Scale bar, 100 µm.

121

20.7 LLO inhibits NOX2 NADPH oxidase assembly at the

phagosome

The NOX2 NADPH oxidase is composed of a transmembrane heterodimer of gp91phox and

p22phox, and 4 regulatory cytosolic subunits: p40phox, p47phox, p67phox and the small GTPase,

Rac2. I hypothesized that LLO inhibits ROS production by preventing proper localization of

NOX2 NADPH oxidase components to the phagosome. To test this hypothesis, human

peripheral blood mononuclear cells (PBMCs) were infected with GFP-expressing wild type or

ΔLLO bacteria and immunostained for p22phox (Figure 20.7.1), p47phox or p67phox (Figure 20.7.2).

IgG-opsonized sheep red blood cells were employed as a positive control for localization of the

NOX2 NADPH oxidase to phagosomes (Figure 20.7.1A, 20.7.2A and 20.7.2 C). I observed

minimal colocalization between intracellular wild type bacteria and the NOX2 NADPH oxidase

components tested (Figure 20.7.1 B, 20.7.1C, 20.7.2 B and 20.7.2 D). In contrast, I observed

more than a three-fold increase in co-localization between intracellular ΔLLO bacteria and all of

the NOX2 NADPH oxidase components tested (Figure 20.7.1 B, 20.7.1 C, 20.7.2 B and 20.7.2

D). These observations demonstrate that LLO prevents proper NOX2 NADPH oxidase assembly

at the phagosome.

20.8 Perfringolysin O (PFO) expression can complement NOX2

NADPH oxidase inhibition by LLO deficient bacteria

I next determined whether other bacterial pore-forming cytolysins could also inhibit the NOX2

NADPH oxidase, similar to LLO. I used ΔLLO bacteria expressing perfringolysin O (PFO) from

C. perfringens under a tightly controlled IPTG-inducible promoter (iPFO). In the absence of

IPTG, iPFO infected cells generated significant amounts of superoxide, similar to ΔLLO (Figure

20.8 A). However, upon IPTG induction, superoxide production was inhibited.

The same effect was observed in bacteria-containing phagosomes via TEM analysis of cerium

perhydroxide precipitates (Figure 20.8 B). Since PFO can complement LLO for inhibition of

NOX2 NADPH oxidase, these results suggest that the inhibition of ROS production may be a

common strategy used by other pore-forming cytolysin secreting bacteria.

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Figure 20.7.1. LLO inhibits NOX2 NADPH oxidase assembly at the phagosome. (A) Human

macrophages were treated with IgG-opsonized sheep red blood cells (sRBC) for 60 min and

fixed. Cells were then stained with anti-p22phox antibodies. Differential interference contrast

(DIC) microscopy and epifluorescence images were acquired. Arrows indicate colocalization of

p22phox with phagosomes. (B) Human macrophages were infected at MOI 10 with either wild

type or ΔLLO bacteria for 60 min and fixed. Cells were then stained with anti-p22phox antibodies

in red and for bacterial-expressed GFP in green (to ‘boost’ this signal). Representative confocal z

slices are shown. Insets are higher magnifications of the boxed areas. (C) Quantification of

p22phox, p47phox or p67phox colocalization with sRBC or bacteria containing phagosomes. One

hundred phagosomes were assessed from three independent experiments. Bars represent mean ±

SEM.

123

Figure 20.7.2. Wild type L. monocytogenes inhibit ROS production through LLO to

mislocalize p47phox and p67phox to the phagosome. (A, C) Human macrophages were treated

with IgG-opsonized sheep red blood cells (sRBC) for 1 h and then fixed. Cells were then stained

with either anti-p47phox (A) or anti-p67phox antibodies (C). Differential interference contrast

(DIC) microscopy and epifluorescence images were acquired. (B, D) Human macrophages were

infected with either wild type or ΔLLO bacteria for 1 h and fixed. Cells were then stained with

either anti-p47phox (B) or anti-p67phox antibodies (D), in red. Bacteria expressing GFP were

visualized with anti-GFP antibodies. Representative confocal z slices are shown. Insets in (B)

and (D) are higher magnifications of the boxed areas. All scale bars are 10 µm.

124

Figure 20.8. LLO inhibition of ROS production is complemented by perfringolysin O

(PFO). (A) Superoxide production by RAW 264.7 macrophages was measured by NBT

reduction assay. Cells were either treated with PMA or infected at MOI 10 with wild type or ΔLLO bacteria for 30 min, followed by NBT treatment. Cells were also infected with iPFO, a

complemented strain of ΔLLO that expresses PFO under an IPTG inducible promoter. Graph

represents mean ± SEM from three independent experiments. (B) Cells were either treated with

10 µg/ml PMA or infected at MOI 10 with wild type or ΔLLO bacteria for 30 min, followed by

cerium chloride treatment. TEM analysis was employed to quantify the percentage of

phagosomes that were cerium perhydroxide precipitate positive. One hundred phagosomes were

assessed from four independent experiments. Bars represent mean ± SEM.

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20.9 LLO inhibition of NOX2 NADPH oxidase assembly is not

mediated through altering key signaling events that mediate

NOX2 NADPH oxidase assembly

It remains to be determined how LLO can selectively block delivery of NADPH oxidase

components to the phagosome. To begin to explore this question, I examined signaling events

linked to activation of the NADPH oxidase in L. monocytogenes infected cells. I observed

phosphorylation of the p40phox subunit of the NADPH oxidase occurs in macrophages infected

with both wild type and ΔLLO bacteria (Figure 20.9 A). Given that p38, AKT and ERK1/2

activation results in ROS production (Chen et al., 2003; Dang et al., 2003; Dewas et al., 2000), I

next examined the activation of these signaling kinases during wild type and ΔLLO bacterial

infection. I observed no major differences in activating phosphorylation between wild type or

ΔLLO infected macrophages (Figure 20.9 B-D). Finally, I examined the phosphorylation state of

a number of protein kinase C (PKC) isoforms since PKCα, PKCβ, and PKCδ have been shown

to activate NOX2 NADPH oxidase via phosphorylation of the regulatory subunits (Bey et al.,

2004; Dekker et al., 2000; Fontayne et al., 2002). However, no significant differences in the

phosphorylation states of any PKC isoform examined were observed between wild type or ΔLLO

L. monocytogenes infected cells (Figure 20.9 E).

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Figure 20.9. LLO mediated inhibition of ROS production is not dependent on altering the

phosphorylation states of key signaling molecules responsible of NOX2 NADPH oxidase

activation. BMDMs were infected with WT or ΔLLO bacteria for 15, 30, 45, 60 or 120 min.

Cells were lysed and western blot analysis was performed to examine changes in (A) phospho-

p40phox, (B) phospho-p38, (C) phospho-AKT, (D) phospho-ERK1/2 and (E) phospho-PKCα/β

or phospho-PKCδ.

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20.10 Summary

Here I demonstrate, for the first time, that L. monocytogenes modulates NOX2 NADPH oxidase

activity during infection. Upon invasion, L. monocytogenes produces PLCs and LLO, which

mediate bacterial phagosome escape. Production of diacylglycerol by PLCs has been implicated

in phagosome escape by L. monocytogenes (Grundling et al., 2003). However, diacylglycerol can

also activate the NOX2 NADPH oxidase through the activation of PKC (Dang et al., 2001b).

Therefore, the induction of antibacterial ROS by PLCs is a potentially deleterious consequence

of the mechanism that these enzymes utilize to promote phagosome escape in host cells.

Countering this deleterious effect, LLO plays a key role in inhibiting the NOX2 NADPH oxidase

by blocking localization to phagosomes. Therefore, one consequence of LLO pore formation is

inhibition of PLC-driven ROS production, in addition to its role in phagosome escape and other

virulence functions (Goebel and Kuhn, 2000).

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Chapter 5

Discussion

The host and L. monocytogenes interactions are a complex series of events that can be either

directed by the host to clear the infection or by the bacteria to evade toxic host defenses. Here, I

provide insight to further current understanding of this dynamic interplay during L.

monocytogenes intracellular infection. Autophagy is a key host defense against a multitude of

pathogens. Past work has shown that autophagy can target L. monocytogenes at both early and

late stages of infection. While a number of studies have shed light into the mechanism of

autophagy targeting during the late stages of infection, the mechanism of how early targeting

occurs has remained elusive. In this thesis, I provide evidence indicating that LC3 targeting of L.

monocytogenes during early and late stages of infection is mediated via two different

mechanisms, depending on where the bacterial is located inside the cell. LC3 targeting of

cytosolic L. monocytogenes is mediated via ubiquitination of the bacteria and p62 binding

(Figure 21.1 A) while early LC3 targeting to L. monocytogenes containing phagosomes is

mediated via ROS and DAG (Figure 21.1 B).

Given that ROS can induce autophagy and is in itself bactericidal, I next examined the possibility

of bacterial evasion of ROS. Here, I show that L. monocytogenes, via the activity of LLO,

inhibits local ROS production by NOX2 NADPH oxidase in bacteria containing phagosomes. In

addition, I found that other LLO related pore forming cytolysins are also capable of inhibiting

ROS production, suggesting that this mechanism of host defense subversion may be a common

bacterial strategy for intracellular survival.

Together, this thesis provides a clearer model of the “push and pull” of host defense versus

bacterial evasion (Figure 21.2). Host defensive strategies, which include autophagy and ROS,

target intracellular pathogens, resulting in their clearance in the phagolysosome. However, L.

monocytogenes encodes a number of virulence factors, including LLO, which is capable of

shutting down host defenses, such as ROS production, resulting in bacterial survival (Figure

21.2). While this thesis provides new insights into L. monocytogenes pathogenesis, some

intriguing observations have generated a number of new hypotheses that are worthy of further

discussion and future exploration.

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Figure 21.1. Model of autophagy targeting to L. monocytogenes at early and late stages of

infection, which are mediated by different mechanisms. (A) LC3 targeting to cytosolic L.

monocytogenes is mediated via ubiquitination of the bacteria and p62 binding to recruit

autophagy. (B) LC3 targeting to L. monocytogenes containing phagosomes at 1 h p.i. is

dependent upon NOX2 NADPH oxidase production of ROS as well as both bacterial and host

PLCs production of DAG.

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Figure 21.2. The “Push and Pull” dynamics of host and bacteria interaction. (A) Host

defenses, such as autophagy and ROS, target invading pathogens, resulting in bacterial clearance

in the phagolysosome. (B) Bacteria, such as L. monocytogenes, encode virulence factors that are

capable of shutting down host defenses (such as LLO inhibition of ROS production), resulting in

intracellular bacterial survival.

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21 Deciphering the complexity of L. monocytogenes and

autophagy interaction

My work described in this thesis sheds light into the signals that mediate autophagic targeting of

L. monocytogenes at 1 h p.i. (Figure 21.1). While future studies will be needed to fully

understand the complexity of autophagic induction upon L. monocytogenes infection, my studies

provide novel insight into the unique phagosomal targeting of bacteria that is likely relevant to a

number of other intracellular organisms.

21.1 Different downstream signaling consequences of

DAG production by bacteria or host

From my studies, I found evidence to suggest that DAG is produced by both the bacteria and

host. This observation suggests that DAG production has beneficial consequences for both

bacteria and host. DAG production by bacterial PI-PLC and PC-PLC has been extensively

documented (Camilli et al., 1991; Goldfine et al., 1993; Poussin et al., 2009; Sibelius et al., 1999;

Smith et al., 1995). PLCs production of DAG at the phagosome mediates PKC recruitment and

activation, which in turn is capable of recruiting host phospholipases, including host PLCs and

PLDs. Since both host phospholipases are implicated in allowing more efficient bacterial escape

from the phagosome, the production of DAG by the bacteria allows ultimately for phagosome

lysis and escape. Here, I provide evidence that in addition to its role in mediating escape, DAG

production may be also responsible for the recruitment of LC3.

Since LLO is required for DAG generation (Sibelius et al., 1996), it has been hypothesized that

LLO allows bacterial PLCs to travel from the lumen of the phagosome to the outer leaflet of the

phagosomal membrane. Here, bacterial PLCs drive the production of DAG, which in turn

recruits host phospholipases to generate more DAG. This model remains controversial since it is

unclear whether bacterial PLCs, which are 30-35kDa, can freely passage through an LLO pore

(~350Å) (Gilbert et al., 2000).

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Another potential model can be proposed based on the findings of Goldfine and colleagues. In

this model, LLO, even in absence of bacterial PLCs, recruits host PKCs, which in turns recruits

host PLCs and PLDs (Goldfine et al., 2000). As host PLCs convert phosphatidylinositides into

inositide triphosphate and DAG, the phagosomal membrane becomes more porous, thus allowing

the diffusion of bacterial PLCs from the lumen to the outer leaflet of the phagosomal membrane.

However, my results argue against this model since wild type L. monocytogenes displayed

unaltered DAG production at the phagosome despite treatment with various host PLC inhibitors.

While previous studies suggest that host PLCs are upstream of PLD recruitment, it is possible

that bacterial PLCs can also recruit PLD. Given that PI-PLC specifically recruits and activates

PKCβ (Poussin et al., 2009), which is specific for the recruitment and activation of host PLD

(Dieter and Fitzke, 1995), it can be argued that PI-PLC activity can recruit and activate host

PLD.

Here, I propose a new model based on my findings. During wild type L. monocytogenes

infection, LLO recruits and activates PKC, which recruits host PLCs. The concerted activity of

both bacterial and host PLCs break down the phagosomal membrane to allow bacterial PLCs to

travel from the lumen of the phagosome to the outer leaflet. Here, PI-PLC production of DAG

recruits host PKCβ, which then recruits host PLD. Together, bacterial and host phospholipases

continue to lyse the phagosomal membrane, generating large amounts of DAG, which recruits

LC3.

21.2 Induction of Autophagy – Advantage to the cell or

bacteria?

In light of my proposed model, it is possible that the recruitment of host PLCs and DAG

production is part of an autophagic response that is activated in presence of membrane damage.

Thus, host PLCs are recruited to sites of membrane damage in an effort to remodel and repair the

membrane via a concerted process of breakdown (by the PLCs) and rebuild (by DAG mediated

targeting of autophagy). The fact that host PLCs assist in bacterial escape from the phagosome

may be an unintended outcome of membrane degradation during the remodeling process.

Previously, membrane damage induced by Vibrio cholera cytolysin, Vcc, been reported to

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induce autophagy in an attempt to protect cells from Vcc intoxication (Gutierrez et al., 2007).

More recently, alpha-toxin (secreted by S. aureus), streptolysin O (secreted by S. pyrogenes), and

hemolysin (secreted by E. coli) were all shown to induce autophagy via the AMP-activated

protein kinase (AMPK) pathway (Kloft et al., 2010). AMPK is a pathway that monitors cellular

ATP levels. Pore-formation can result in leakage of ATP into the extracellular milieu, thus

creating “energy stress”. This then activates the AMPK pathway, resulting in the inhibition of the

mTOR complex. The work presented here suggests that the degree to which LLO can form pores

influences autophagic targeting at 1 h p.i. (Figure 19.5 B). Thus, these observations combine to

argue for the possibility of autophagy targeting to L. monocytogenes containing phagosomes in

response to LLO pore-formation, which then damages the phagosomal membrane. In the absence

of membrane damage, ΔLLO mutants therefore do not trigger an autophagy response (Figure

19.1 and Figure 19.4 A).

As mentioned above, other pore-forming cytolysins have the ability to induce autophagy via

activation of AMPK, which activates protein kinase A (PKA), resulting in subsequent

phosphorylation of its substrate, cyclic-AMP response element binding (CREB). Therefore, I

examined changes in phospho-CREB levels via western blot analysis of cells infected with wild

type or ΔLLO L. monocytogenes. Preliminary studies indicate that wild type infected cells

showed greater levels of phospho-CREB over ΔLLO mutant infected cells at 1 h p.i.. Thus, this

result suggests that LLO may also activate AMPK. This could explain, at least in part, the

induction of autophagy at 1 h p.i.. However, just how LLO pore-formation targets LC3 to the

phagosomal membrane is less clear.

21.3 How is LC3 targeted to L. monocytogenes containing

phagosomes?

The question of how LLO recruits LC3 is a complex one and requires examination of the origin

of the LC3. Knowing how LC3 is recruited would then give insight into how LLO may be

triggering this process. There are two models of LC3 recruitment. The classical model of

autophagy targeting involves the fusion of LC3+ vesicles with the damaged organelle to create a

double-membraned compartment. It is theorized that de novo conjugation of LC3-I to PE to form

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LC3-II, mediated by the Atg12-Atg5-Atg16L1 complex, is possible at the phagosomal

membrane (Figure 21.3). This would give rise to a LC3+ single-membraned compartment.

However, the issue of how LC3 is recruited to phagosomes remains controversial in the field.

Here, I propose that LC3 targeting to L. monocytogenes is mediated by a de novo mechanism.

Examining wild type L. monocytogenes containing phagosomes at 1 h, I observed mainly single-

membraned phagosomes and thus it appears unlikely that LC3+ phagosomes are double-

membraned. However, to confirm this observation, CLEM should also be performed to examine

if LC3+ L. monocytogenes is likewise contained in a single-membraned phagosome.

21.4 L. monocytogenes possesses the ability to limit

autophagic targeting

It stands to reason that if bacterial replication is enhanced when autophagy is inhibited, then

autophagic induction is a host-mediated event since the host benefits from the induction of

autophagy. Conversely, if bacterial replication is decreased when autophagy is inhibited, then

autophagic induction is likely a bacterial-mediated event since the bacteria benefits from the

induction of autophagy. This distinction is less clear in the case of autophagic targeting of L.

monocytogenes. Previous reports of L. monocytogenes in vitro survival in mouse embryonic

fibroblasts (MEFs) show no significant difference in L. monocytogenes replication between

infection in wild type or Atg 5-/- MEFs (Birmingham et al., 2007a; Py et al., 2007). These

findings suggest that autophagy does not play a role in either limiting or enhancing bacterial

infections or that L. monocytogenes has a mechanism of overcoming autophagic clearance.

Certainly, in absence of LLO, bacterial replication is limited in both wild type and Atg5-/- MEFs.

However, it is hard to ascribe autophagy evasion activity to LLO since LLO is required for

bacterial escape and thus, the drop in bacterial replication may not necessarily be due to

inhibition of autophagy but an inability to escape into the cytosol. As such, the search for this

putative L. monocytogenes encoded factor which limits autophagic targeting is hampered by the

limitation that a number of factors can influence bacterial replication in addition to autophagic

targeting. Thus, to fully appreciate the role of LLO on autophagy evasion, LC3 targeting of wild

type and ΔLLO L. monocytogenes must be examined in a cell type that does not require LLO for

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Figure 21.3. Proposed model of autophagic targeting of latex beads or bacteria. FcgR/TLR

signaling mediates NOX2 NADPH oxidase assembly and activation, resulting in phagosomal

production of ROS. ROS acts as a signal for the recruitment of autophagic machinery, mediating

the de novo conversion of LC3-I to LC3-II directly at the phagosome. Maturation of the LC3+

phagosome results in the fusion with lysosome. Reprinted from NADPH oxidase contribute to

autophagy regulation. Huang et al. Autopahgy 5:6 887-9,2009, with authors’ permission.

137

escape (ie: in Hela or Henle-407 cells). In addition to LLO, L. monocytogenes PLCs have also

been implicated in autophagy evasion. ΔPI-PLC mutants exhibit increased replication in Atg5-/-

MEFs when compared to control MEFs suggesting that PI-PLC may play either a direct or

indirect role in the evasion of autophagy targeting (Birmingham et al., 2007a). A kinetic study of

ΔPI-PLC mutants indicated that 30% of these bacteria remain LC3+ up to 8 h p.i. while only 10%

of wild type L. monocytogenes remains LC3+. This finding suggests that PI-PLC plays a role in

the evasion of autophagy targeting. However, since LC3 targeting to either wild type or ΔPI-PLC

mutants never increases beyond 30% of the population, it is possible that additional L.

monocytogenes factors may be involved to limit autophagy targeting. Thus, to overcome

activation of autophagy, I hypothesize that L. monocytogenes may encode an as yet unknown

factor, in addition to PI-PLC, which can inhibit autophagy to promote bacterial survival.

21.4.1 Future Directions

Previously, my work and work by others in the Brumell laboratory demonstrated that roughly

20-30% of wild type (strain named 10403S) and ΔactA L. monocytogenes are LC3+ at 2 h p.i.

(Birmingham et al., 2007a). Intriguingly, work done by Yoshikawa and colleagues using L.

monocytogenes strains from a different background (EGDe) indicated that 10% of wild type L.

monocytogenes are LC3+ and 60% of ΔactA bacteria are LC3+ at 2 h p.i.. Collectively, these

observations suggest the possibility that in absence of ActA, EGDe L. monocytogenes is targeted

by LC3 while 10403S L. monocytogenes expresses an unknown factor to evade LC3 targeting.

Thus, a comparison of genes expressed in 10403S but not in EGDe may reveal the putative

autophagy-inhibiting factor. In collaboration with John Parkinson of the University of Toronto,

we have compared the genomes of both wild type strains of L. monocytogenes and compiled a

list of 62 genes that are unique to 10403S. Of these genes, 12 are expressed only in virulent

strains of Listeria. Thus, future efforts to search for the putative autophagy-inhibiting factor

should begin with the generation of 10403S L. monocytogenes knock outs of each of these 62

genes and LC3 targeting should then be examined. It is predicted that a greater percentage of

mutants deficient in the autophagy-inhibiting factor would be targeted by LC3 compared to the

wild type. Conversely, EGDe strains expressing each of these 12 genes should also be

constructed and LC3 targeting then examined. It is my prediction that a lower percentage of the

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mutant EGDe strain expressing the autophagy-inhibiting factor would be targeted by LC3

compared to the wild type.

21.5 Multiple signals acting in the same pathway could

mediate autophagy targeting

Given that DAG, ROS and LLO pore-formation all exert significant individual impact on

autophagic targeting of L. monocytogenes at 1 h, these three signals of autophagy may act in the

same pathway. DAG is required for the activation of protein kinase Cs (Mellor and Parker,

1998), which have been reported to activate NOX2 NADPH oxidase activity (Raad et al., 2009).

Thus, DAG may act upstream of ROS.


To test the hypothesis is that DAG is upstream of ROS production, I quantified DAG localization

to L. monocytogenes containing phagosomes in the presence or absence of ROS inhibition by

DPI. In cells infected with ΔPI-PLCΔPC-PLC L. monocytogenes, DAG localization to the

phagosome was unaltered in presence of DPI inhibition, suggesting that host production of DAG

is upstream of ROS production. Surprisingly, localization of DAG to wild type L.

monocytogenes-containing phagosomes decreased upon treatment of DPI, suggesting that

bacterial production of DAG is downstream of ROS production. Given the confusing nature of

these preliminary data, more work is needed to fully understand the difference between host and

derived and bacterial derived DAG.

Furthermore, I hypothesize that bacterial-derived DAG is located on the inner leaflet of the

phagosomal membrane immediately post infection, whereas host-derived DAG is located on the

outer leaflet of the phagosomal membrane. This localization would, presumably, change as the

membrane becomes permeabliized. DAG specific antibodies derived from PKCδ and PKCε

DAG binding regions have been employed in the context of EM to study DAG production

(Babazono et al., 1998). Thus, I propose that DAG production on both leaflets of the phagosomal

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membrane should be examined using immuno-gold electron microscopy. This technique can be

employed to study DAG production over time. Cells infected with wild type or ΔPI-PLCΔPC-

PLC L. monocytogenes can be fixed at 15, 30, 45 and 60 mins p.i. and DAG production on the

inner versus outer leaflet of the phagosomal membrane assessed at each time point. Presumably,

in cells infected with wild type bacteria, DAG production would occur on both leaflets of the

membrane. In cells infected with ΔPI-PLCΔPC-PLC L. monocytogenes, DAG production would

occur first in the outer leaflet of the phagosomal membrane, followed by production on the inner

leaflet of the membrane.

22 Walking a tight rope: The importance of L.

monocytogenes virulence factor antagonism in

presence of NOX2 NADPH oxidase activity

The work presented in this thesis describes the previously unknown function of LLO to inhibit

local ROS production by NOX2 NADPH oxidase inside L. monocytogenes containing

phagosomes. Furthermore, I provide evidence to suggest that the inhibition of ROS production

may be a general function of other LLO related pore-forming cytolysins. As such, this thesis

provides new insights that further advance current understanding L. monocytogenes

pathogenesis.

22.1 Multiple peaks of ROS production may be due to

different activation signals

Previous studies have reported that at 10 min p.i., both wild type and ΔLLO bacteria induce ROS

production. Here, I confirm these findings of an early wave of ROS production and further show

that there is a second wave of ROS production from 40 min – 70 min p.i.. The dynamic studies

with luminol indicate that these two waves are induced by different mechanisms, where the first

wave is PLCs-independent induction of ROS while the second wave is PLCs-dependent

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induction of ROS. Furthermore, the second wave of ROS production can be inhibited by LLO,

whereas the first wave at 10 min cannot. These observations suggest that a third bacterial factor

can trigger the early wave of ROS production. Given that TLRs have been shown to be upstream

of NOX2 NADPH oxidase activation (Bae et al., 2009; Elsen et al., 2004; Lee et al., 2008;

Suliman et al., 2005; Vulcano et al., 2004), a potential source of this initial wave of ROS

production may be TLR activation in response to an unidentified L. monocytogenes PAMP.

Thus, I hypothesize that TLR activation mediates both intra- and extracellular ROS production at

10 min p.i. where LLO cannot effectively inhibit ROS production. In contrast, PLC-mediated

ROS production at 40 – 70 min p.i. occurs locally inside the phagosome, where LLO effectively

mislocalizes the NOX2 NADPH oxidase to inhibit ROS production.

An alternate explanation to consider is that two different populations of phagosomes are

responsible for generating the first and the second wave of ROS production. It has been observed

that there is a degree of heterogeneity in phagosome formation within a given cell (Henry et al.,

2004). Henry and colleagues showed there appears to be two distinct populations of phagosomes

that are formed upon addition of IgG-coated latex beads or sheep red blood cells to RAW 264.7

macrophages. One population loses PI3P localization at the phagosome after 10 min while the

other population continues PI3P acquisition for up to 20 min. Consistent with this, other

laboratories have observed NOX2 NADPH oxidase assembly on only 50% of phagosomes

(personal communication with Dr. Mary Dinauer). Thus, given that NOX2 NADPH oxidase

localization to the phagosome is dependent on its binding to PI3P (Lambeth, 2004), it is possible

that the wave of ROS production at 10 min p.i. is derived from all phagosomes prior to the loss

of PI3P, while ROS production at 40 – 70 min p.i. occurs in the population of phagosomes that

retain PI3P over time.


To examine the possibility that TLR recognition of bacterial PAMP results in ROS production at

10 min p.i., cells incapable of mounting TLR-mediated signaling should be examined. Myeloid

differentiation primary response gene 88 (MyD88) is a critical adapter protein in the signaling

pathway of all TLRs (with the exception of TLR3) (Arancibia et al., 2007). Thus, mice deficient

in MyD88 would exhibit an inability to mount the majority of TLR mediated responses. BMDM

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from wild type and MyD88 knockout mice should be compared for their ability to produce ROS

upon L. monocytogenes infection. A luminol assay can be used to compare dynamic ROS

production. If ROS production is MyD88 dependent, then the burst of ROS at 10 min p.i. would

be absent in the MyD88 knockout. If ROS production is dependent on MyD88, BMDM from

specific TLR knockout mice can be examined to determine which TLR is responsible for ROS

production.

Phagosomal heterogeneity may also be responsible for ROS production by different phagosomes

at different times. Thus, this possibility should also be tested. Given that only half of

phagosomes remain PI3P+ after 10 min, immunofluorescence microscopy analysis can be

employed to quantify the percentage of PI3P+ and PI3P- L. monocytogenes containing

phagosomes that are also p22phox, p40phox or p67phox +. This experiment will give an indication of

whether the NOX2 NADPH oxidase complex is present exclusively on PI3P+ phagosomes.

However, the presence of the NOX2 NADPH oxidase complex on PI3P+ phagosomes will not

directly confirm the presence of ROS in those specific phagosomes. One technically challenging

but highly informative experiment that can directly show ROS production at PI3P+ phagosomes

is correlative light microscopy – electron microscopy (CLEM). To do this, human peripheral

blood mononuclear cells (PBMCs) can be cultured and transfected with PI3P specific probes

(either PX-GFP or 2-FYVE-GFP constructs). 24 h later, cells can be infected with either wild

type or ΔLLO L. monocytogenes and cerium chloride treatment can be employed as described in

section 17. After fixation, cells could imaged via using confocal microscopy to identify PI3P+ L.

monocytogenes containing phagosomes followed by TEM imaging to quantify the percentage of

PI3P+ L. monocytogenes containing phagosomes that have cerium perhydroxide precipitates. It is

predicted that PI3P+ L. monocytogenes containing phagosomes are cerium perhydroxide+ while

PI3P- L. monocytogenes containing phagosomes are cerium perhydroxide-.

22.2 Relationship between ROS and LLO provides insight

into L. monocytogenes pathogenesis in different cell types

Interestingly, it must be noted that while LLO is a critical virulence factor to ensure L.

monocytogenes survival in murine macrophages, it is the PLCs that are required for bacterial

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survival in other cell types, such as in human laryngeal cancer cells (Hep-2), human cervical

epithelial cells (HeLa), human embryonic intestinal epithelial cells (Henle-407) (Grundling et al.,

2003) and human embryonic kidney cells (Burrack et al., 2009). It is currently unclear whether

the difference in virulence factor importance is due to a difference of species (mouse vs. humans)

or of cell type (macrophages vs non-phagocytes). Furthermore, it is also unclear why this

distinction of virulence factor dependence exists. In light of my work, one possible explanation

emerges. Given that macrophages produce a substantially higher level of ROS than epithelial

cells, it is possible that a virulence factor that can decrease the level of oxidative stress (ie: LLO)

may be critical for L. monocytogenes survival, while virulence factors that inadvertently induce

more oxidative stress (ie: PLCs) may be dispensable in this cell type. Conversely, in an

environment with little ROS production (ie: epithelial cells), counteracting oxidative stress is not

a priority, so LLO is dispensable. Thus, the evolution of LLO and PLCs may be important for

mediating bacterial survival in different cell types.


To test whether LLO dependency differs between cell type or host species, a replication assay of

wild type and ΔLLO L. monocytogenes in human PBMCs can be performed. Presumably, if LLO

is required for bacterial survival in macrophages regardless of species, then ΔLLO L.

monocytogenes in human PBMCs should be effectively cleared in the same fashion as it is in

murine BMDMs (Figure 20.5).

An alternate experiment would be to perform replication assays in both murine epithelial cells

and BMDMs. If LLO is critical for bacterial survival in macrophages but not epithelial cells,

then ΔLLO replication should exhibit wild type L. monocytogenes-like growth kinetics in murine

epithelial cells. If LLO is not needed for survival in murine epithelial cells, then this would

support the hypothesis that L. monocytogenes survival in high ROS producing cell types require

LLO to inhibit ROS while survival in low ROS producing cell types do not require LLO.

Recently, Clark and colleagues created a K562 human erythroleukemia cell line that stably

expresses functional NOX2 NADPH oxidase (Clark and Valente, 2004). Thus, to directly test

this hypothesis, K562 cells with or without NOX2 NADPH oxidase can be infected with either

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wild type or ΔLLO L. monocytogenes in replication assays. If the hypothesis is valid, then K562

cells expressing NOX2 NADPH oxidase should rapidly clear ΔLLO L. monocytogenes in a

fashion that is similar to macrophages while K562 cells without NOX2 NADPH oxidase would

permit similar replication kinetics by both wild type and ΔLLO L. monocytogenes.

22.3 Understanding the mechanism of LLO mislocalization

of NOX2 NADPH oxidase

The question of how LLO mislocalizes NOX2 NADPH oxidase remains unanswered. While

LLO may not be influencing the phosphorylation states of PKCs, p38, AKT, p40 or ERK1/2, I

cannot rule out the possibility that LLO is altering the activation of other signaling molecules

upstream of NOX2 NADPH oxidase activation. However, it can be concluded that pore

formation is critical in mediating NOX2 NADPH oxidase mislocalization. It remains to be

determined whether pore formation has a direct or indirect effect on NOX2 NADPH oxidase

mislocalization. Certainly, previous studies have indicated that LLO can modify the phagosomal

compartment, resulting in a delay in phagosomal maturation (Alvarez-Dominguez et al., 2008;

Alvarez-Dominguez et al., 1997; Henry et al., 2006; Prada-Delgado et al., 2005). In particular,

studies conducted by Prada-Delgado and colleagues found that LLO excludes Rab5a recruitment

to the L. monocytogenes containing phagosome. Rab5a was found to participate in the

recruitment of Rac2 to the phagosome (Prada-Delgado et al., 2005). Since activated Rac2 is one

of the key NOX2 NADPH oxidase components that mediate proper localization to the

phagosome, it is possible that LLO-dependent exclusion of Rab5a prevents Rac2 recruitment and

therefore prevents NOX2 NADPH oxidase localization (Prada-Delgado et al., 2001). Certainly,

in the context of Yersinia pseudotuberculosis, an outer membrane protein, YopE inactivates

Rac2, thus preventing ROS production during Y. pseudotuberculosis infection in macrophages

(Songsungthong et al., 2010). To address this hypothesis, I compared levels of activated Rac2

between wild type and ΔLLO infected RAW 264.7 macrophages using a pulldown assay. PAK-

PBD, which binds specifically to activated Rac, was used the bait. However, I did not detect any

differences between wild type and ΔLLO infected cells. This may be due to the fact that

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pulldowns reflect the level of activated Rac2 in the entire cell and the phenotype may be

localized and limited to the phagosome.

If LLO is mediating NOX2 NADPH oxidase mislocalization via the exclusion of Rab5a, it

remains unclear how a pore-forming toxin can be excluding a phagosomal protein. Under normal

conditions, Rab5 is transiently recruited to the phagosome via binding to phosphatidylinositol (3)

phosphate (PI3P), where it subsequently recruits a number of effectors required for the

maturation of the early phagosome (Duclos et al., 2000; Fratti et al., 2001). Since PI3P is also

required for the proper localization of NOX2 NADPH oxidase components (Lambeth, 2004), I

examined if there are differences in PI3P colocalization to wild type or ΔLLO L. monocytogenes

containing phagosomes. At 1 h p.i., I observed similar recruitment of PI3P to wild type and

ΔLLO L. monocytogenes, suggesting that LLO-dependent NOX2 NADPH oxidase

mislocalization is not mediated via alterations in PI3P enrichment at the phagosome.

There are few clues that exist in the field to direct the search for the mechanism behind Rab5

exclusion by a pore-forming toxin. Legionella pneumophila is another intracellular

microorganism that induces a similar exclusion of Rab5c (Clemens et al., 2000). However, the

mechanism is likewise unknown. Mycobacterium tuberculosis also has the ability to alter

phagosomal maturation by modifying the bacteria containing phagosome. Unlike L.

monocytogenes, M. tuberculosis compartments are Rab5+ but Rab7-. Despite the presence of

Rab5, both Rab5 and Rab7 are inactive upon M. tuberculosis infection, in a manner that is

dependent on the bacterial GTPase activating protein, nucleoside diphosphate kinase (Ndk) (Sun

et al., 2010). Ndk mediates the conversion of the active GTP-bound Rab protein to the inactive

GDP-bound form. Interestingly, M. tuberculosis mislocalizes both the NOX2 NADPH oxidase

and the inducible nitric oxide synthase (iNOS) (Katti et al., 2008). Thus, exclusion of NOX2

NADPH oxidase by L. monocytogenes may also be mediated by the inactivation of Rab5a or

other Rab proteins.

It must be noted that there are a number of different functions already attributed to LLO and

there likely exists many more. Thus, it is possible that LLO inhibits NOX2 NADPH oxidase via

multiple mechanisms. The difficulty in studying these other roles of LLO lies in the fact that

LLO is required for phagosomal escape and is critical for survival in macrophages. Thus, the

other roles of LLO can be hidden since bacteria lacking LLO are rapidly cleared. Furthermore,

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cytosolic roles or consequences of LLO activity are difficult to assess since efficient escape does

not occur in absence of LLO. To get around this limitation, a number of studies have utilized

exogenous addition of LLO or LLO point mutants to understand the biochemical changes that

occur upon exposure to LLO.

Another potential method to study LLO function is to study non-phagocytic cells infected with

wild type and ΔLLO L. monocytogenes. Since LLO is not required for efficient phagosomal

escape in these cells, the limitation of LLO dependent escape can be eliminated. Therefore,

future studies should be directed towards a better understanding of LLO function. Given that

current understanding of the many potential roles of LLO is quite limited, I expect that the

mechanism by which LLO inhibits NOX2 NADPH oxidase may be much more complex than

what we can currently appreciate.


Further examination of whether differential Rac2 recruitment could explain differences in ROS

production between wild type and ΔLLO infected cells should be pursued. My attempts to

examine the levels of activated Rac2 at the whole cell level were not fruitful. Thus, Rac2

localization and activation at the level of the phagosome should be tested. Previously, fluorescent

fusion constructs for murine Rac1 (YFP-Rac1), Rac2 (YFP-Rac2) and activated Rac (CFP-PBD)

have been successfully employed in RAW 264.7 macrophages to study red blood cell

phagocytosis (Hoppe and Swanson, 2004). Therefore, one can express these constructs in RAW

264.7 macrophages and infect for 1 h with either wild type or ΔLLO bacteria. If the inhibition of

NOX2 NADPH oxidase assembly by wild type bacteria is mediated via Rac2 mislocalization, the

percentage of Rac2+ or PBD+ wild type L. monocytogenes-containing phagosomes should be

significantly lower than that of ΔLLO L. monocytogenes-containing phagosomes.

If Rac2 is found to be mislocalized, then it is possible that Rab5a mislocalization in wild type L.

monocytogenes containing phagosomes might be cause of Rac2 mislocalization and ROS

inhibition. To examine a possible link between Rab5a and Rac2, immunofluorescence co-

staining with Rab5a and Rac2 should be done to determine if Rab5a+ ΔLLO L. monocytogenes

containing phagosomes are also Rac2+. If Rab5a+ compartments are also Rac2+, then the link

146

between Rab5a and ROS production should next be examined using CLEM. If Rab5a allows the

recruitment of Rac2, which mediates the assembly of the rest of the NOX2 NADPH oxidase

components, then Rab5a+ L. monocytogenes-containing phagosomes should be cerium

perhydroxide+. Wild type and Rab5a deficient BMDM can be infected with either wild type or

ΔLLO bacteria followed by the examination of ROS production using each of the ROS

quantification methods that are described in this thesis. If Rab5a is required for NOX2 NADPH

oxidase assembly at the phagosome, then infection with ΔLLO L. monocytogenes would not

induce ROS in the Rab5a deficient BMDM.

In addition to Rab5a and Rac2, other mechanisms of LLO mediated mislocalization of the NOX2

NADPH oxidase should also be examined. My preliminary work, as well as a recent publication,

demonstrate that LLO mediates protein kinase A (PKA) activation as evidenced by increases in

the phosphorylation level of the PKA substrate, cyclic-AMP response element biding (CREB)

(Gonzalez et al., 2011). Given that PKA also phosphorylates the inhibitory site of p47, resulting

in the inhibition of ROS production, it is possible that LLO mediates NOX2 NAPDH oxidase

mislocalization via the activation of PKA. I propose that further work should be done to fully

understand how LLO activates PKA.

Previously, pore-forming cytolysins have been shown to activate PKA by increasing cellular

cAMP levels (Gonzalez et al., 2011). Thus, the effect of LLO on cAMP levels should be

examined. To do so, western blotting techniques can be applied to compare cAMP levels

between macrophages infected with wild type or ΔLLO L. monocytogenes. Next, the question of

how LLO can influence cAMP levels should be examined. cAMP is produced by adenyl cyclase

and degraded by phosphodiesterase (PDE). In vitro alterations of cAMP levels can be achieved

either by inhibition its production (inhibiting adenyl cyclase) or decreasing its degradation

(inhibiting PDE). The modulation of cAMP levels or PKA activity should thus be examined in

context of macrophages infected with wild type or ΔLLO L. monocytogenes. If LLO inhibition of

NOX2 NADPH oxidase is mediated by PKA activation, treatment with pharmacological

inhibitors of PKA, such as H-89 dihydrochloride or KT5720, or compounds that inhibit cAMP

production, such as 2’, 5’-dideoxyadenosine, should increase ROS production during wild type

L. monocytogenes infection. Conversely, drugs that either enhance PKA activity directly, such as

forskolin or 6-bnz-cAMP (cAMP mimetic), or inhibit cAMP degradation, such as 3-isobutyl-1-

methylxanthine (IBMX), should inhibit ROS production during ΔLLO L. monocytogenes

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infection. My preliminary studies involving the NBT quantification assay demonstrate that

treatment with either 6-bnz-cAMP, forskolin or IBMX can significantly reduce ROS production

in macrophages infected with ΔLLO L. monocytogenes. Thus, these observations suggest that

LLO modulation of PKA activity may be a key mechanism of NOX2 NADPH oxidase

inhibition.

Lastly, the functional effect of PKA modulation on bacterial replication and survival should be

examined. In vitro replication assays of wild type and ΔLLO infected macrophages infected with

wild type and ΔLLO L. monocytogenes should be conducted in the presence of both PKA

activating and inhibiting drugs. It is predicted that PKA activation should inhibit ROS production

in macrophages infected with ΔLLO bacteria, thus enhancing bacterial survival and replication

over time. Conversely, it is predicted that PKA inhibition should allow ROS production in

macrophages infected with wild type bacteria, thus decreasing bacterial survival and replication

over time. The rate of survival of both L. monocytogenes strains in wild type or PKA deficient

animals should be examined. I hypothesize that survival of wild type L. monocytogenes should

decrease in absence of PKA.

22.4 Multiple mechanisms may be involved in NOX2

NADPH oxidase inhibition

Given that all of the NOX2 NADPH oxidase components examined were mislocalized from the

wild type L. monocytogenes-containing phagosome, the assumption was made that a single

mechanism universal to all components might be responsible for the inhibition of ROS

production. However, it is possible that L. monocytogenes may have multiple mechanisms to

mediate NOX2 NADPH oxidase mislocalization. In the case of F. tulerensis, a number of

different mechanisms of NOX2 NADPH oxidase inhibition have been reported. ROS production

by neutrophils in response to F. tulerensis infection is inhibited by decreasing phosphorylation of

p47phox and mislocalizing gp91phox from phagosomes containing F. tulerensis. Furthermore, this

bacterium also encodes a phosphatase, AcpA, which contributes to ROS inhibition. Finally, cells

infected with the F. tulerensis mutants deficient in the virulence operon regulator, MigR, display

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a partial increase in ROS production. Together, these observations suggest that F. tulerensis

employs multiple mechanisms to downregulate NOX2 NADPH oxidase activity. Given the

importance of ROS as an antibacterial defense for all intracellular pathogens, it is likely that L.

monocytogenes also has multiple mechanisms of ROS inhibition. Since different PKCs are

responsible for phosphorylation of different NOX2 NADPH oxidase subunits, it is possible that

specific bacterial virulence factors may target a specific PKC. Furthermore, L. monocytogenes

encodes a manganese superoxide dismutase (MnSOD) as well as a catalase. Mutants deficient in

MnSOD display attenuated survival in vitro in macrophages and in vivo in a manner than is

similar to ΔLLO mutants (Archambaud et al., 2006). Furthermore, ΔMnSOD L. monocytogenes

also displays a deficiency in phagosomal escape. Thus, L. monocytogenes possesses at least three

different mechanisms to defend against ROS, acting at different steps in the pathway to produce

ROS (Figure 22.5). It remains to be seen if these ROS inhibition mechanisms are synergistic or

cooperative in nature.

22.5 Opposing consequences of LLO and PLCs activity

allows for ROS production in the phagosomal

compartment

Why would L. monocytogenes have evolved two virulence factors that have antagonistic effects

on NOX2 NADPH oxidase activity? There are two potential explanations for this conundrum.

One possible explanation would be that PLCs activation of NOX2 NADPH oxidase is an

unintended consequence of PLCs activity. Given that PLCs play an important role in L.

monocytogenes escape from the phagosome, the deleterious effect of ROS induction can thus be

counterbalanced by LLO to decrease local oxidative stress. The opposing consequences of LLO

and PLCs activity can be termed “virulence factor antagonism”. Thus, while these two virulence

factors are seemingly antagonistic to each other, they function together to allow for bacterial

survival. An alternate explanation could be because limited host production of ROS is somehow

beneficial for L. monocytogenes survival. Thus, bacterial PLCs may drive ROS production while

LLO ensures that ROS in the phagosome, or in the immediate vicinity of the bacteria, is kept to a

minimum. It is known that ROS participate in a number of signaling events and perhaps the

149

induction of these events may be advantageous for bacterial survival. However, since L.

monocytogenes displays an increased replication rate in mice deficient in gp91phox compared to

wild type mice, it does not appear that ROS improve bacterial survival.

150

Figure 22.5. Methods of ROS inhibition employed by L. monocytogenes. L. monocytogenes

possesses three different mechanisms against the deleterious effects of ROS. LLO acts to inhibit

local production of ROS by mislocalizing the NOX2 NADPH oxidase. Residual ROS is

eliminated by bacterial superoxide dismutase (SOD) and catalase.

151

22.6 NOX2 inhibition by pore forming cytolysins and

virulence factor antagonism may be common strategies for

intracellular survival

The observation that PFO, a pore-forming cytolysin secreted by C. perfringens, can also mediate

a similar inhibition of NOX2 NADPH oxidase as LLO suggests that inhibition of ROS

production may be a conserved consequence of pore-forming cytolysin activity. A number of

pathogens that encode pore-forming cytolysins can also inhibit ROS production, including C.

perfringens, Y. pestis, B. anthracis, H. pylori among others. Thus, further examination of the

mechanism of LLO inhibition of NOX2 NADPH oxidase activity may reveal a common function

for all pore-forming cytolysins.

Furthermore, the work presented in this thesis reveals how L. monocytogenes initiates a dynamic,

and in this case antagonistic, set of signals in host cells via its virulence factors to allow for

optimal bacterial growth and survival during infection. Like L. monocytogenes, C. perfringens is

capable of escaping from the phagosome (O'Brien and Melville, 2000). This process is

dependent on the activity of the bacterial PLC in addition to PFO (O'Brien and Melville, 2004).

Another intracellular organism that also produces both a bacterial PLC and a pore-forming

cytolysin is Bacillus anthracis. As mentioned, this bacteria can inhibit NOX1 generated ROS in

colonic epithelia in a manner that is dependent on the pore-forming cytolysin, edema toxin (Kim

and Bokoch, 2009). While edema toxin belongs to a different class of pore-forming cytolysins

from either LLO or PFO, the potential for virulence factor antagonism between B. anthracis PLC

and pore-forming cytolysin remains a viable possibility. Finally, both H. pylori and F. tulerensis

have the ability to inhibit ROS production (Table 2.3), and encode bacterial PLCs. Given that a

number of pathogens secrete PLCs and are also able to inhibit ROS production, this model of

virulence factor antagonism may be applicable to several pathogenic species.

While perhaps counterintuitive at first glance, virulence factor antagonism is potentially an

extremely powerful mechanism that would allow two opposing responses to be regulated both

spatially and temporally. The induction of ROS production by PLCs is detrimental to the

bacteria. However, the presence of LLO allows for local inhibition of ROS while global ROS

152

production may continue. In this light, LLO can modulate the effect of PLC spatially such that L.

monocytogenes inside the phagosome can be protected from the production of ROS.

22.7 Understanding LLO pore-formation and ROS in the

targeting of autophagy

Given that LLO can inhibit ROS production and that ROS mediates autophagy targeting, cells

infected with ΔLLO should produce significant level of ROS, thus recruiting LC3. However, my

work (Figure 19.1.1), and that of others (Birmingham et al., 2007a; Meyer-Morse et al., 2010; Py

et al., 2007) clearly indicates that LLO is required for autophagy targeting. How then can LLO

inhibit ROS yet the lack of ROS prevents autophagy targeting? There are a number of possible

explanations for this apparent paradox. First, it is possible that ROS need to be transported from

the phagosomal lumen to the outer membrane of the phagosome (perhaps via the LLO pores) in

order to be detected by the autophagic system. Thus, while ROS is needed for autophagic

targeting, production of ROS inside the lumen of the ΔLLO containing phagosome will not be

detected by the autophagic machinery in the cytosol. Thus, ΔLLO is not targeted by autophagy.

In cells infected with wild type L. monocytogenes, low levels of ROS production was observed

(Figure 20.1.2 B). Thus, it may be possible that this low level of ROS can traverse the

phagosomal membrane via LLO pores and be detected by the autophagic machinery.

A second possibility is the idea of bacterial virulence heterogeneity. Indeed, it has been theorized

that not every single wild type L. monocytogenes express the same level of LLO (Birmingham et

al., 2008). Previously, work from the Brumell laboratory suggests that there are three possible

fates that can occur upon L. monocytogenes infection. One population of internalized L.

monocytogenes can escape from the phagosome, replicate inside the cytosol and achieve

secondary spread into neighbouring cells via the formation of the actin rocket tail. A second

population of L. monocytogenes fails to escape from the phagosome and is degraded in the

lysosome in the conventional phagolysosomal pathway. A third population of bacteria is targeted

by autophagy inside a non-degradative compartment. These compartments, termed spacious

Listeria associated phagosomes (SLAPs), cannot fuse with the lysosome but the bacteria can also

153

not escape. SLAP are non-degradative, LAMP-1+ and LC3+ (Birmingham et al., 2007b).

Formation of SLAPs was found to require autophagy in the host cell, and expression of LLO by

the bacteria. Modulation of LLO expression via an inducible promoter which expresses a

moderate level of LLO (compared to wild type expression level of LLO) can increase SLAP

formation (Birmingham et al., 2007b). Birmingham and colleagues proposed that SLAPs allow

slow bacterial replication that may allow chronic L. monocytogenes infection in a host

(Birmingham et al., 2007b). Indeed, compartments resembling SLAPs have been observed in a

mouse model of L. monocytogenes chronic infection (Bhardwaj et al., 1998). Thus, these

observations suggest that there may be variable LLO expression even in the wild type L.

monocytogenes population. Bacteria that express high level of LLO can efficiently escape from

the phagosome while those that express low levels of LLO are degraded in the lysosome.

However, the population of bacteria expressing a moderate level of LLO may be targeted by

autophagy and form SLAPs.

Incorporating my observations into this model, it is possible that the population of bacteria

expressing high levels of LLO is effectively inhibiting ROS production (and thus preventing

autophagic targeting), thus allowing for escape into the cytosol. The population of bacteria

expressing low levels of LLO cannot effectively inhibit ROS production and thus is cleared. The

population of bacteria expressing moderate levels of LLO does not inhibit ROS production

efficiently and thus recruits autophagy (Figure 22.7). This moderate level of ROS production is

likely insufficient to clear the bacteria but is sufficient to recruit autophagy. Consistent with this

model, my preliminary work indicates that SLAPs are also dependent on ROS production

(Figure 19.6).


One direct way of testing this model is via the use of the iLLO mutant. Using the iLLO mutant,

one can control the expression level of LLO by altering the concentration of IPTG.

Immunofluorescence microscopy and cerium chloride TEM methods or CLEM can then be

employed to examine ROS production and autophagic targeting. I predict that in the absence of

IPTG, the iLLO mutant would be predominately cleared in the phagolysosomal pathway with

154

most phagosomes exhibiting cerium perhydroxide precipitates. Moderate expression of iLLO

would show greater autophagic targeting at 1 h with moderate cerium perhydroxide precipitation

inside the phagosome. High or over expression of LLO would result in enhanced autophagic

targeting at 1 h with no cerium perhydroxide precipitation inside the phagosome. The same study

should be conducted with the iPFO mutant. This experiment would then confirm the importance

of pore formation on modulating the intracellular fate of L. monocytogenes.

155

Figure 22.7. Proposed model of L. monocytogenes interaction with early host defenses

against phagocytosed pathogens. Upon phagocytosis, L. monocytogenes encounters a number

of host defenses and depending on the level of LLO, one of three potential fates can occur. One

population of bacteria with low LLO expression cannot inhibit NOX2 NADPH oxidase

production of ROS, and is thus readily degraded in the phagolysosome. Another population of

bacteria with medium LLO expression mediates a moderate inhibition of ROS production. It is

hypothesized that this moderate level of ROS is insufficient to kill the bacteria but does prevent

phagosomal escape and mediate LC3 targeting to the L. monocytogenes containing phagosome.

Over time, this “stalemate” compartment becomes a SLAP where neither the host can clear the

bacteria or nor the bacteria escape. Finally, the population of bacteria with high LLO expression

efficiently inhibits ROS production, allowing the bacteria to escape into the cytosol. Modified

from Listeria evades killing by autophagy. Birmingham et al. Autopahgy 4:3 368-71, 2008, with

authors’ permission.

156

23 Concluding Remarks

In this thesis, I have provided insight into the signals that mediate autophagic targeting of L.

monocytogenes at 1 h p.i.. In addition, my work provides evidence for a novel function of LLO

in the inhibition of ROS production by the NOX2 NADPH oxidase. These observations led to

the proposal of a new model that furthers the current understanding of L. monocytogenes

intracellular survival and targeting by host defenses. Despite these advances, more questions

remain to be answered before we will fully understand L. monocytogenes pathogenesis. Future

research should continue to focus on understanding the intracellular lifestyle of L.

monocytogenes, as this bacterium is a useful model pathogen to study the host and bacterial

processes that are activated upon infection. Given the rise of antibiotic resistance, new

approaches are necessary in combating infections. Insights into how host defenses are activated

in response to intracellular infections may allow the creation of novel therapeutics that enhance

host defenses to target pathogens. Such therapies could be less susceptible to bacterial resistance,

since they would act on host proteins and not bacterial factors. Thus, administration of drugs that

enhance host immunity, such as autophagy inducers, and those that weaken bacterial attack, such

as pore-forming cytolysin inhibitors, may together provide a solution to a wide range of

infections.

157

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Copyright Acknowledgements

Reprint and adaptation permission were received from various journals and cited after each

figure. Text reprint permissions were granted by Mary Ann Liebert, Inc: Antioxidants Redox

Signaling, Huang J et al. Autophagy Signaling Through Reactive Oxygen Species. 14(11): 2215-

31 and by Springer Science + Business Media: Seminars in Immunopathology, Lam G et al. The

many roles of NOX2 NADPH oxidase-derived ROS in immunity, 32(4): 415-30. The text and

data presented in Chapter 4, Section 20 is reprinted from Cell Host and Microbe, 10(6), Lam GY

et al. “Listeriolysin O Suppresses Phospholipase C-Mediated Activation of the Microbicidal

NADPH Oxidase to Promote Listeria monocytogenes Infection”, p627-34 with permission from

Elsevier.

interactions of l. monocytogenes with host cellular defenses

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