Neutrophil Products Inhibit LLO Secretion and Activity, and Listeria Monocytogenes Intracellular Growth

Home , Defensin, Gangrene, Listeriolysin O, Listeriosis, Theta defensin

Neutrophil products inhibit LLO secretion and activity, and Listeria monocytogenes intracellular growth

Dissertation

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

Eusondia Arnett

Graduate Program in Microbiology

The Ohio State University

2013

Dissertation Committee:

Dr. Stephanie Seveau, Advisor

Dr. John Gunn

Dr. Mike Ibba

Dr. Larry Schlesinger

Eusondia Arnett

2013

Abstract

Listeria monocytogenes is a facultative intracellular pathogen that infects a large variety of host cells, including macrophages and diverse non-phagocytic cells. To avoid the phagosome microbicidal environment, L. monocytogenes secretes a pore-forming toxin (listeriolysin O; LLO) that releases the bacterium into the cytoplasm. Once in the cytosol, L. monocytogenes proliferates and infects adjacent cells through cell-to-cell spreading. Innate immune cells like neutrophils play an important role in the control of infection, yet the interaction between neutrophils, other host cells, and L. monocytogenes is not well understood. Neutrophils produce a high concentration and variety of antimicrobial molecules, including defensins and proteases; thus it is likely that these cells enhance the anti-listerial response of other host cells. This dissertation addresses if: i) human defensins, which can be released into the extracellular milieu by neutrophils, enable macrophages to control intracellular replication of L. monocytogenes; ii) L. monocytogenes is able to replicate in human neutrophils in a LLO-dependent manner as observed in macrophages; and iii) human neutrophils cooperate with macrophages to prevent L. monocytogenes replication in human macrophages.

Addressing i), we found that the α-defensin HNP-1 (one of the most abundant proteins in neutrophil primary granules) cooperates with macrophages to inhibit L.

ii monocytogenes phagosomal escape and intracellular growth. Importantly, HNP-1 is acquired by macrophages and trafficked to the phagocytosed bacteria. Finally, HNP-1 inhibits LLO secretion from the bacteria and directly blocks LLO activity. In conclusion, neutrophil defensins inhibit LLO function through two mechanisms (secretion and activity), and inhibit L. monocytogenes escape from macrophage phagosomes.

Addressing ii), we found that LLO enhances the phagocytic efficiency of neutrophils and does not protect L. monocytogenes from neutrophil intracellular killing.

L. monocytogenes produces multiple virulence factors, including LLO, that induce rapid neutrophil degranulation, even before closure of the phagosome. Intriguingly, degranulation protects neutrophils from LLO-mediated membrane damage. Neutrophils degranulate matrix metalloproteases, which degrade LLO, irreversibly blocking its activity. In summary, upon interaction with L. monocytogenes, neutrophils rapidly release matrix metalloproteases that degrade LLO, likely maintaining the bacterium in a bactericidal phagosome from which it cannot escape.

Addressing iii), we determined that co-incubating neutrophils with macrophages during L. monocytogenes infection does not significantly alter L. monocytogenes association with or phagocytosis by macrophages but does markedly reduce L. monocytogenes growth in macrophages.

In conclusion, human neutrophils produce molecules that inhibit LLO activity and intracellular replication of L. monocytogenes. Furthermore, neutrophils help limit L. monocytogenes replication in macrophages. We propose that during infection, macrophage internalization of neutrophils and/or neutrophil granule components enables

iii macrophages to limit L. monocytogenes replication. Thus, at the site of infection the cooperation between neutrophils and macrophages likely plays a critical role in the innate immune defense against L. monocytogenes.

Dedication

This document is dedicated to my family and friends for their support and encouragement. I would especially like to thank my husband, Matt Arnett, for all of his

help.

Acknowledgments

I would like to thank my advisor, Dr. Stephanie Seveau, for her encouragement, instruction, and guidance during my scientific training. I would also like to thank my committee members, Dr. Mike Ibba, Dr. John Gunn, and Dr. Larry Schlesinger for their comments and insights regarding my project, and their willingness to write recommendation letters at the last minute. I would especially like to thank Dr. Seveau and

Dr. Schlesinger for all the career advice they have provided.

I am grateful for past and present members of the Seveau lab, including Anne

Cecile-Haghighat, who trained me in many of these techniques, and Stephen Vadia for insightful conversations and help with experiments. I am also thankful for the undergraduate students in the lab who provided valuable assistance, particularly Colleen

Nackerman, Ben Foreman, and Eric Weber. Lastly, I am grateful for Dr. Chad Rappleye and his lab members, especially Dr. Jessica Edwards and Dr. Eric Holbrook for all of their help and endless advice.

I would like to thank all of our collaborators who provided materials and insightful conversations to help move the projects forward: Dr. Dan A. Portnoy

(University of California, Berkeley, CA) for the L. monocytogenes DP10403S wild type and DP-L2161 Δhly strains and the pET29b plasmid encoding native LLO; Dr. Pascale

Cossart (Pasteur Institute, Paris, France) for the L. monocytogenes L028 wild type and hly::Tn917 strains, Dr. Rodney K. Tweten (University of Oklahoma Health Sciences

Center, Oklahoma City, OK) for the pQE30 plasmid encoding PLY and Dr. Philipp C.

Hanna (University of Michigan Medical School, Ann Arbor, MI) for the pET15 plasmid encoding ALO. I also thank Tracy Tan and Grace Jung (David Geffen School of

Medicine at UCLA, Los Angeles, CA) for technical assistance with the defensins and

Sriram Satagopan (Ohio State University) for assistance with Chimera. I am grateful for

Dr. J.-Q. Wu (Departments of Molecular Genetics and Molecular and Cellular

Biochemistry) for use of the confocal microscope and I-Ju Lee (Dr. J.-Q. Wu’s lab) for assistance with confocal image acquisition and deconvolution.

I would like to acknowledge funding support through the OSU Presidential

Fellowship.

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Vita

2003 ...... High School Diploma with Honors

Fairborn High School

2003 to 2006 ...... National Commended Scholarship and

Honors Competitive Scholarship

Wright State University

2006 ...... Honors Research Grant

Wright State University

2006 ...... B.S. Biological Sciences

Magna Cum Laude

Wright State University

2007 to 2011 ...... Graduate Teaching Associate

Department of Microbiology

The Ohio State University

2010 ...... M.S. Microbiology

The Ohio State University

2011 to 2012 ...... Presidential Fellowship

The Ohio State University

2012 to present...... Graduate Teaching Associate

viii

Department of Microbiology

The Ohio State University

2012 ...... Travel Award

Public Health Preparedness for Infectious Diseases Annual Member Meeting

Ohio State University

2013 ...... Travel Award

Wexner Medical Center Annual Research Day

Ohio State University

Publications

Arnett E, Vadia S, Nackerman CC, Oghumu S, Satoskar A, McLeish KR, Uriarte SM, and Seveau S. 2013. The pore-forming toxin listeriolysin O is degraded by neutrophil proteases and fails to protect L. monocytogenes against intracellular killing. Manuscript submitted for publication.

Arnett E and Seveau S. 2011. The multifaceted activities of mammalian defensins.

Current Pharmaceutical Design, 17: 4254-4269.

Vadia S, Arnett E, Haghighat AC, Wilson-Kubalek EM, Tweten RK, Seveau S. 2011.

The pore-forming toxin listeriolysin O mediates a novel entry pathway of L. monocytogenes into human hepatocytes. PLoS Pathogens, 7: e1002356.

Arnett E, Lehrer RI, Pratikya P, Lu W, Seveau S. 2011. Defensins enable macrophages to inhibit the intracellular proliferation of Listeria monocytogenes. Cellular

Microbiology, 13: 635-651.

Fields of Study

Major Field: Microbiology.

Table of Contents

Abstract...... ii

Dedication ...... v

Acknowledgments ...... vi

Vita...... viii

Table of Contents ...... xi

List of Tables...... xvi

List of Figures...... xvii

List of Abbreviations...... xx

Chapter 1. Introduction ...... 1

1.1 Listeria monocytogenes ...... 1

1.1.1 Listeriosis...... 1

1.1.2 L. monocytogenes intracellular lifecycle...... 3

1.1.3 L. monocytogenes virulence factors: listeriolysin O ...... 4

1.1.4 Other major L. monocytogenes virulence factors ...... 9

1.2 L. monocytogenes and the host immune system ...... 10

1.2.1 Innate Immunity...... 11

1.2.2 Adaptive Immunity ...... 13

1.3 Neutrophils...... 14

1.3.1 Neutrophil oxidative killing mechanisms ...... 15

1.3.2 Neutrophil non-oxidative killing mechanisms...... 15

1.3.3 Neutrophil granules...... 16

1.3.4 Defensins ...... 18

1.4 Neutrophil cooperation with macrophages ...... 22

1.5 Research Hypothesis and Goals ...... 23

Chapter 2. Defensins enable macrophages to inhibit the intracellular proliferation of Listeria monocytogenes ...... 26

2.1 Introduction...... 26

2.2 Materials and Methods...... 28

2.2.1 Reagents...... 28

2.2.2 Cell culturing, bacterial strains and infection...... 29

2.2.3 Fluorescence labeling...... 31

2.2.4 Image Acquisition ...... 32

2.2.5 Image Analysis ...... 33

2.2.6 Gentamicin survival assay...... 35

2.2.7 Antimicrobial activity of the defensins...... 35

2.2.8 Hemolytic assay...... 37

2.2.9 Measurement of BMM perforation by LLO ...... 37

2.2.10 LLO release by L. monocytogenes ...... 38

2.2.11 Statistics ...... 38 xii

2.3 Results...... 39

2.3.1 Defensins inhibit L. monocytogenes proliferation in macrophages...... 39

2.3.2 HNP-1 interaction with macrophages and RC-1 interaction with L.

monocytogenes are sufficient to inhibit intracellular proliferation...... 42

2.3.3 Defensins have little effect on phagocytosis of L. monocytogenes...... 46

2.3.4 Defensins inhibit L. monocytogenes phagosomal escape...... 49

2.3.5 α- and θ-Defensins inhibit LLO-mediated macrophage membrane perforation

and bacterial release of LLO ...... 51

2.3.6 The α-defensin HNP-1 is taken up by macrophages and colocalizes with

intracellular L. monocytogenes ...... 55

2.4 Discussion...... 57

Chapter 3. The pore-forming toxin listeriolysin O is degraded by neutrophil proteases and fails to protect L. monocytogenes against intracellular killing. 62

3.1 Introduction...... 62

3.2 Materials and Methods...... 64

3.2.1 Recombinant proteins and reagents ...... 64

3.2.2 Isolation of human neutrophils and serum ...... 65

3.2.3 Macrophage and bacterial cell cultures ...... 66

3.2.4 L. monocytogenes association with, phagocytosis by, and replication in

neutrophils...... 67

3.2.5 L. monocytogenes association with, and phagocytosis by, macrophages ...... 68

3.2.6 L. monocytogenes viability following incubation with neutrophils or

macrophages ...... 68

3.2.7 Macrophage perforation following incubation with L. monocytogenes ...... 69 xiii

3.2.8 CD63 colocalization with L. monocytogenes...... 69

3.2.9 Kinetics of neutrophil degranulation following challenge with bacteria...... 70

3.2.10 Wide field and Confocal Microscopes...... 71

3.2.11 LDH release by cells challenged with LLO...... 72

3.2.12 Collection of Neutrophil Degranulation Products...... 73

3.2.13 Hemolysis assay...... 73

3.2.14 LLO treatment with NDP followed by Western blotting...... 74

3.2.15 LLO treatment with MMPs followed by Western blotting ...... 74

3.2.16 Statistics ...... 75

3.3 Results...... 75

3.3.1 LLO does not confer a survival advantage to L. monocytogenes phagocytosed by

neutrophils...... 75

3.3.2 L. monocytogenes and LLO activate rapid neutrophil degranulation...... 80

3.3.3 Degranulation protects neutrophils from perforation by LLO ...... 83

3.3.4 Degranulation products directly inhibit LLO activity ...... 86

3.3.5 A protease activity is responsible for LLO inhibition...... 86

3.4 Discussion...... 92

Chapter 4. Co-incubation of macrophages and neutrophils limits L. monocytogenes replication in macrophages...... 101

4.1 Introduction...... 101

4.2 Materials and Methods...... 103

4.2.1 Reagents...... 103

4.2.2 Isolation of human neutrophils ...... 103

4.2.3 Macrophage and L. monocytogenes culture...... 104 xiv

4.2.4 Infection assays ...... 104

4.2.5 Fluorescence labeling...... 105

4.2.6 Image acquisition and analysis...... 105

4.2.7 Statistics...... 106

4.3 Results...... 106

4.3.1 Co-culturing macrophages with neutrophils does not significantly alter

macrophage phagocytosis of L. monocytogenes...... 106

4.3.2 Co-culturing macrophages with neutrophils inhibits L. monocytogenes

replication in macrophages ...... 109

4.4 Discussion...... 113

Chapter 5. Synthesis...... 117

Bibliography...... 124

List of Tables

Table 3.1. L. monocytogenes is unable to proliferate in neutrophils, regardless of LLO expression...... 78

Table 3.2. LLO inhibition by neutrophil degranulation products involves heat-labile factor(s) and divalent cations...... 89

xvi

List of Figures

Figure 1.1. Human listeriosis...... 2

Figure 1.2. Schematic of L. monocytogenes intracellular life cycle...... 3

Figure 1.3. The structure of two CDCs...... 5

Figure 1.4. Neutrophil granules...... 17

Figure 1.5 Ribbon structures and primary sequences of human α- and β-defensins and humanized θ-defensins...... 19

Figure 1.6. Major activities of the defensins on mammalian cells...... 21

Figure 2.1. Defensins inhibit L. monocytogenes intracellular proliferation...... 40

Figure 2.2. Defensins inhibit L. monocytogenes intracellular proliferation in RAW 264.7 cells...... 41

Figure 2.3. HNP-1 interaction with macrophages and RC-1 interaction with L. monocytogenes are sufficient to inhibit bacterial intracellular proliferation...... 43

Figure 2.4. RC-1 is more listericidal than HNP-1 in DMEM...... 45

Figure 2.5. HNP-1 and RC-1 inhibit L. monocytogenes growth in TSB...... 47

Figure 2.6. Defensins do not affect phagocytosis of L. monocytogenes...... 48

Figure 2.7. Defensins inhibit L. monocytogenes-induced actin polymerization...... 50

Figure 2.8. Defensins increase L. monocytogenes colocalization with LAMP-1...... 52

xvii

Figure 2.9. Defensins inhibit LLO-mediated perforation of erythrocytes and macrophages and the release of LLO...... 54

Figure 2.10. HNP-1 is taken up by macrophages and colocalizes with L. monocytogenes.

...... 56

Figure 3.1. L. monocytogenes association with and phagocytosis by human neutrophils.77

Figure 3.2. Neutrophils kill L. monocytogenes, regardless of LLO expression...... 79

Figure 3.3. L. monocytogenes replication in macrophages is LLO-dependent...... 80

Figure 3.4. L. monocytogenes induces local degranulation of primary granules...... 82

Figure 3.5. L. monocytogenes and LLO induce neutrophil degranulation...... 83

Figure 3.6. Neutrophil degranulation is required for resistance to LLO-mediated perforation...... 85

Figure 3.7. Neutrophil degranulation products inhibit LLO-mediated perforation of macrophages and erythrocytes...... 87

Figure 3.8. Neutrophil degranulation products inhibit other CDCs...... 88

Figure 3.9. Neutrophil degranulation products degrade LLO...... 90

Figure 3.10. MMP-8 degrades LLO...... 91

Figure 3.11. Model of L. monocytogenes interaction with neutrophils...... 93

Figure 3.12. LLO does not enhance phagocytosis by macrophages...... 96

Figure 4.1. L. monocytogenes growth in Thp1 cells is LLO-dependent...... 107

Figure 4.2. Neutrophils do not significantly alter macrophage phagocytosis of L. monocytogenes...... 108

xviii

Figure 4.3. Co-culture of neutrophils with macrophages inhibits L. monocytogenes replication in macrophages...... 110

Figure 4.4. Neutrophil cooperation with macrophages is contact-dependent...... 112

xix

List of Abbreviations

ALO Anthrolysin O

AMP Antimicrobial peptide

ASM Acid sphingomyelinase

BHI Brain heart infusion

BMM Bone marrow-derived macrophages

CDC Cholesterol-dependent cytolysin

CFTR Cystic fibrosis transmembrane conductance regulator

CFU Colony forming units

CGD Chronic granulomatous disease

CS Control supernatant

DC Dendritic cell

DMEM Dulbecco’s modified Eagle’s medium

DS Donor serum

EthD Ethidium homodimer

FBS Fetal bovine serum fMLP N-Formyl-methionyl-leucyl-phenylalanine

GILT γ-interferon-inducible lysosomal thiol reductase

HBD Human β-defensin

HBSS Hank’s balanced salt solution

HD Human defensin

HGFR Hepatocyte growth factor receptor

HI Heat inactivated

HNP Human neutrophil peptide

HRP Horseradish peroxidase

IFN Interferon iNOS Inducible nitric oxide synthase

Inl Internalin

JNK c-Jun N-terminal kinase

LAMP Lysosome-associated membrane protein

Lm Listeria monocytogenes

LDH Lactate dehydrogenase

LLO Listeriolysin O

LLOpL Pre-pore locked listeriolysin O

LPS Lipopolysaccharide

MAPK Mitogen-activated protein kinase

MFI Mean fluorescence intensity

MMP Matrix metalloprotease

MOI Multiplicity of infection

MPO Myeloperoxidase

xxi

NADPH Nicotinamide adenine dinucleotide phosphate

NDP Neutrophil degranulation product

NET Neutrophil extracellular trap

NK Natural killer

OD Optical density

PC-PLC Broad-range phospholipase C

PFA Paraformaldehyde

PFO Perfringolysin O

PI Protease inhibitor

PI-PLC Phosphatidylinositol-specific phospholipase C

PKC Protein kinase C

PMA Phorbol myristate acetate

PMN Polymorphonuclear leuckocyte

PrI Propidium iodide

PLY Pneumolysin

RC Retrocyclin

RNI Reactive nitrogen intermediate

ROS Reactive oxygen species

RT Room temperature

SCID Severe combined immunodeficiency

SEM Standard error of the mean

SFI Sum of the fluorescence intensities

xxii

SGD Neutrophil-specific granule deficiency

SLAP Spacious Listeria-containing phagosome

SLO Streptolysin O

SNAP-23 Synaptosome-associated protein-23

TAT HIV transactivator of transcription

TMH Transmembrane β-hairpin

TNF Tumor necrosis factor

TSB Trypticase soy broth

UPR Unfolded protein response wt wild type

xxiii

Chapter 1. Introduction

1.1 Listeria monocytogenes

1.1.1 Listeriosis

Listeria monocytogenes is a Gram-positive foodborne intracellular pathogen.

Only two species of Listeria are pathogenic, L. monocytogenes and L. ivanovii. L. monocytogenes infects humans, while L. ivanovii primarily infects sheep and cattle, although human infections have been reported (1, 2). L. monocytogenes is a saprophyte that is able to grow at a wide range of temperatures (1-45°C), pH (4.4-9.6), and salt concentrations (up to 10%), making it a particular concern for infection through ready to eat foods like milk, cheeses, and deli meats (3, 4). Although healthy individuals typically control L. monocytogenes infection, outbreaks can be devastating. The 2011 multistate outbreak of listeriosis in the US (through contaminated cantaloupes) led to 99% hospitalization, 1 miscarriage, and 20% mortality, despite treatment (5). These statistics highlight the devastating outcomes of L. monocytogenes outbreaks that occur worldwide and the necessity for more efficient treatments.

After ingestion of contaminated food, L. monocytogenes can cross the intestinal barrier and is trafficked to the liver and spleen through the lymph and blood-stream (Fig

1.1). The host immune response typically controls infection at this point. In healthy

1 individuals, L. monocytogenes infection usually manifests as self-limiting gastroenteritis.

However, in susceptible populations including immunocompromised individuals and the elderly, L. monocytogenes can disseminate and cause disease, commonly bacteremia or septicemia. L. monocytogenes can also cross the blood-brain barrier, causing meningoencephalitis. In pregnant women, the bacteria can cross the placental barrier, causing miscarriage and neonatal septicemia (1, 2, 6, 7).

Figure 1.1. Human listeriosis. Upon ingestion of contaminated food, L. monocytogenes can cross the intestinal barrier, leading to gastroenteritis. L. monocytogenes is then trafficked to the liver and spleen. Healthy individuals typically mount an efficient immune response and control infection at this step; however, susceptible individuals may not control infection, and L. monocytogenes can re-enter the blood-steam, resulting in bacteremia. L. monocytogenes may then cross the blood-brain barrier, leading to meningoencephalitis, and the placental barrier, causing miscarriage and neonatal septicemia.

1.1.2 L. monocytogenes intracellular lifecycle

The ability of L. monocytogenes to cause disease and cross the intestinal, blood- brain, and placental barriers is mediated through its intracellular lifecycle (Fig 1.2). L. monocytogenes can induce internalization into typically non-phagocytic cells and is also internalized by phagocytic cells. L. monocytogenes escapes from the internalization vesicle within 15-30 min. Once in the cytosol, the bacterium proliferates and polymerizes host cell actin to mediate movement within the cell and cell-to-cell spreading. Spreading involves the formation of bacterial protrusions that extend into and are internalized by neighboring cells, leading to the formation of double membrane vacuoles. L.

Figure 1.2. Schematic of L. monocytogenes intracellular life cycle. 1) L. monocytogenes entry into non-phagocytic cells is mediated by InlA, InlB, and LLO. 2) LLO and the phosholipases PI-PLC and PC-PLC mediate escape from the internalization vesicle. 3) This releases the bacteria into the cytoplasm, where they can replicate and polymerize host cell actin (blue dots) via ActA. Actin polymerization mediates movement within the cell and 4) cell-to-cell spreading. 5) LLO, PI-PLC, and PC-PLC mediate escape from the secondary vesicle, releasing the bacteria into the cytosol where they replicate and continue infection.

3 monocytogenes escapes from these double membrane vacuoles and replicates in the cytosol, propagating infection without being exposed to the extracellular milieu (1, 8-10).

1.1.3 L. monocytogenes virulence factors: listeriolysin O

Listeriolysin O (LLO) was the first identified virulence factor of L. monocytogenes. LLO is important for intracellular replication and is indispensable for virulence (11-13). LLO is part of the largest family of bacterial pore-forming toxins, the cholesterol-dependent cytolysins (CDCs), which are produced by multiple pathogenic

Gram-positive bacteria, including Bacillus anthracis (anthrolysin O, ALO), Clostridium perfringens (perfringolysin O, PFO), Streptococcus pyogenes (streptolysin O, SLO), and

Streptococcus pneumoniae (pneumolysin, PLY) (14). CDCs are also produced by Gram- negative bacteria, like Desulfobulbus propionicus (desulfolysin) (15). The CDCs share high sequence homology and are secreted as water-soluble monomers that have highly similar four-domain structures (Fig 1.3). Domain 4 mediates binding to cell membranes and contains a conserved undecapeptide sequence and three loops (Loops 1-3), all of which insert into the membrane (14, 16). The undecapeptide sequence was initially thought to mediate binding to cholesterol, which is the receptor for most CDCs, but cholesterol binding actually occurs through the threonine-leucine pair located in loop 1

(17). The undecapeptide instead may help maintain the CDC structure such that the threonine-leucine pair is exposed for cholesterol binding. A few CDCs (intermedilysin, vaginolysin, and lectinolysin) use CD59 as their receptor instead of cholesterol, the binding motif for CD59 is also located in domain 4 (18-20). Upon monomer binding to

4 host cell membranes, the CDCs undergo conformational changes and oligomerize into a ring shaped pre-pore complex composed of 30-50 subunits. Subsequently, two sets of α- helices in domain 3 unfold to form amphipathic transmembrane β-hairpins (TMHs), which insert into the membrane and make up the β-barrel channel of the 50 nm diameter pore. Cholesterol is required for this transition from the pre-pore to pore complex, even for CDCs that use CD59 as their receptor instead of cholesterol (14, 16, 21).

Figure 1.3. The structure of two CDCs. Cartoon representations of PFO [PDB 1PFO, (317)] and ALO [PDB 3CQF, (318)] structures are shown. Domains 1-4 are labeled D1-4. The α-helices that unfurl to form the transmembrane β-hairpins (TMHs) are in teal, the undecapeptide sequence in green, Loops 1-3 in yellow, and the threonine-leucine (Thr-Leu) pair in blue. Structures were generated with PyMol.

1.1.3.1 LLO Intracellular Effects

LLO mediates multiple steps in the L. monocytogenes intracellular lifecycle. It was first appreciated to affect escape from the phagosome (Fig 1.2) (22). LLO is also important for escape from the secondary vesicle, which forms during cell-to-cell spreading (Fig 1.2) (23, 24). How LLO mediates escape is not well understood, but involves ion fluxes, protein kinase C (PCK) β activation, the cystic fibrosis transmembrane conductance regulator (CFTR), and γ-interferon-inducible lysosomal thiol reductase (GILT). Intriguingly, GILT is a host factor that reduces LLO, thereby increasing LLO activity (25-28). LLO also delays phagosome maturation, likely because it alters phagosomal calcium concentrations and decreases acidification, both of which are involved in phagosome maturation (26).

The amount of LLO that is secreted by L. monocytogenes may influence the L. monocytogenes intracellular lifecycle. It has been proposed that high amounts of LLO lead to phagosomal escape and that low amounts result in spacious Listeria-containing phagosomes (SLAPs) (29). SLAPs label with autophagy components but are not acidic and likely result from autophagosome formation around a damaged phagosome. L. monocytogenes appears to inhibit maturation of this autophagosome, resulting in a replicative niche in the cell termed a SLAP. However, SLAPs are rare and likely do not represent a typical intracellular niche for L. monocytogenes. Indeed, the majority of L. monocytogenes escapes the phagosome and replicates in the cytoplasm (29, 30).

Furthermore, although LLO induces autophagy, L. monocytogenes escapes capture into autophagosomes and are not typically labeled with autophagy markers (30, 31).

1.1.3.2 LLO Regulation

LLO activity and expression are tightly regulated in the cytosol, deregulated LLO activity leads to perforation of the plasma membrane, destroying the intracellular niche and exposing L. monocytogenes to the immune system (32, 33). Temperature and pH regulate LLO activity, LLO has an acidic pH optimum at 37°C and is more active at pH

5-6, like in the maturing phagosome (12). This pH sensitivity is mediated by an acidic triad in domain 3, which induces premature unfolding of the TMHs at neutral pH, leading to denaturation of LLO (34). Although LLO has a pH optimum, it is still highly active at neutral pH, and exerts a broad range of activity at the plasma membrane (33, 35-37).

Binding of LLO to membranes likely stabilizes the toxin, allowing it to maintain its activity at neutral pH. LLO expression is increased at 37°C and reduced at low pH (38,

39). LLO translation is repressed during exponential growth and in the host cytosol, L. monocytogenes mutants that do not negatively regulate translation are cytotoxic and less virulent (40). LLO is also degraded via proteases in the cytosol (33, 41).

1.1.3.3 LLO Extracellular Activities

Recently, our lab has shown that extracellular LLO is sufficient to mediate L. monocytogenes entry into non-phagocytic cells in a pore-dependent manner (Fig 1.2)

(35). Extracellular LLO also induces i) degradation of a component of the SUMOylation machinery (affecting post-translational modifications), ii) dephosphorylation of histone

H3, iii) mitochondrial fragmentation, iv) ER vesiculation, and v) activation of the unfolded protein response (UPR) (36, 42-46). The majority of these actions influence L. monocytogenes intracellular replication and are dependent on calcium influx or potassium efflux. Ion fluxes elicit signaling pathways, alert the immune system to cell damage and pathogen attack, and induce membrane repair. LLO initiated signaling cascades are not well characterized, but some components have been identified, they include MAPK,

JNK, and NFκB (36, 47).

Although mammalian cells are lysed by high concentrations of CDCs, they are able to withstand damage induced by moderate concentrations of CDCs (including LLO and SLO) (35, 48). Membrane repair in response to CDC-mediated damage is rapid and calcium dependent. Calcium influx through SLO pores induces lysosome release at the plasma membrane, which releases components that induce endocytosis of the membrane containing the SLO pore (49-51). Membranes containing SLO pores can also be released from the cell through blebbing, where the membrane undergoes exocytosis (52, 53). Both of these pathways are calcium dependent. As many of LLO’s activities are dependent on ion flux (entry, mitochondrial fragmentation, histone modification, etc), it seems likely that signaling pathways induced during membrane repair may overlap with LLO’s other activities. However, while signaling pathways may overlap, they are at least somewhat distinct; LLO-mediated entry is actin- and dynamin-dependent, while membrane repair in response to LLO is not (35).

1.1.4 Other major L. monocytogenes virulence factors

L. monocytogenes produces many internalins (Inls), two of which (InlA and InlB), are invasins that are sufficient to mediate entry into non-phagocytic cells (Fig 1.2) (54-

57). InlA specifically binds to E-cadherin and the primary host cell ligand for InlB is c- met, the hepatocyte growth factor receptor (HGFR) (58, 59). InlB also binds glycosaminoglycans and gC1qR, which is a receptor for the globular heads of the complement component C1q (60, 61). InlA recognizes human and guinea pig, but not murine or rat, E-cadherin due to a single amino acid difference at position 16, which is proline in humans and guinea pigs, and glutamic acid in mice and rats (62). Similarly,

InlB binding to HGFR is species specific, InlB interacts with human and murine, but not rabbit or guinea pig, HGFR (63). The internalins may have different roles during infection, InlA mediates crossing of the placental and intestinal barriers while InlB primarily mediates crossing of the placental and blood-brain barriers (8, 64). Neither

InlA, InlB, nor LLO are known to induce entry into phagocytic cells and in fact, LLO may delay entry into some macrophage-like cell-lines (65). L. monocytogenes phagocytosis by macrophages is mediated by complement receptors CR3 and C1q, heparan sulfate proteoglycans and scavenger receptors that recognize lipotechoic acid (1,

66). L. monocytogenes phagocytosis by neutrophils is presumably mediated by Fcγ and complement receptors as the bacteria can be opsinized by antibodies and complement

(67-69).

L. monocytogenes produces two phospholipases, a broad-range phospholipase C

(PC-PLC), encoded by plcB, which hydrolyzes phosphitdylcholine,

9 phosphatidylethanolamine, and phosphatidylserine (70, 71) and a phosphatidylinositol- specific phospholipase C (PI-PLC), encoded by plcA (72-74). PC-PLC and PI-PLC, along with LLO, mediate escape from the primary and secondary vesicles (Fig 1.2) (70, 75-78).

In human epithelial cells (Henle 407 and HeLa cell lines), PC-PLC can mediate L. monocytogenes escape from the primary vacuole in the absence of LLO, although escape is less efficient without LLO (76, 79).

ActA is a secreted protein which is anchored to the L. monocytogenes membrane.

This protein co-opts the host cell actin machinery, leading to formation of actin clouds around the bacteria and polar actin comet tails that mediate intracellular movement and cell-to-cell spreading (Fig 1.2) (80-83). ActA is also important for L. monocytogenes resistance to autophagy-mediated killing (30). These virulence factors, as well as LLO, are regulated by PrfA, which is a thermoregulator that positively controls transcription of the virulence factors upon switch to high temperature (9).

1.2 L. monocytogenes and the host immune system

Many host cells are important for control of L. monocytogenes infection. Innate immune cells, including macrophages, monocytes, and neutrophils, mount the initial response to infection while the adaptive immune response, primarily CD8+ T cells, mediates sterilizing immunity (2, 84-88).

1.2.1 Innate Immunity

Macrophages and monocytes are found at infectious foci and depletion of these cells at infectious foci exacerbates L. monocytogenes burden in mice (89, 90). However,

L. monocytogenes is able to replicate in macrophages both in vivo and in vitro (13, 91,

92). The ability of macrophages to control infection likely results from the activation state of the macrophage. Mice with tissue macrophages unable to respond to interferon

(IFN) γ activation are more sensitive to L. monocytogenes infection than wild type mice

(93). Accordingly, L. monocytogenes are unable to replicate in IFNγ activated macrophages, remain trapped in the phagosome, and are killed due to enhanced production of reactive oxygen species (ROS) and reactive nitrogen intermediates (RNI)

(94, 95). However, L. monocytogenes is able to replicate in non-activated macrophages

(13, 91, 92). In fact, macrophages may serve as a reservoir for L. monocytogenes and have been proposed to propagate infection by transporting live L. monocytogenes to the liver and brain through the blood (1, 87). Thus, macrophages may play dual roles during infection, benefiting both the bacteria and the host.

Neutrophils are important for initial control of L. monocytogenes infection in humans and animal models (96-98). They rapidly accumulate at infectious foci, where they can exert direct antibacterial actions, as well as produce chemokines and cytokines to recruit and activate other immune cells (98, 99). Depleting neutrophils in mice prior to inoculation increases susceptibility to L. monocytogenes infection (100-102). The importance of neutrophils in mice during L. monocytogenes infection has been questioned recently, as neutrophil depletions were previously performed with antibodies against

11 neutrophil receptor 1 (Gr-1), clone RB6-8C5, which is now appreciated to bind epitopes on neutrophils (Ly6G and Ly6C) and monocytes (Ly6C) (100-102). Recently, an antibody that targets Ly6G specifically, clone 1A8, has been identified and used to specifically deplete neutrophils. Two groups have used this antibody and confirmed that neutrophils are important for initial control of L. monocytogenes infection (97, 99).

Conversely, Shi et al. did not find enhanced bacterial burden after using this antibody

(89), likely because this group administered a low dose of antibody following infection while the former studies depleted neutrophils prior to infection with a higher dose of depleting antibodies. Thus, these results highlight the importance of neutrophils in a rapid response to L. monocytogenes infection. This is not unexpected, as L. monocytogenes are killed by neutrophils in vitro (68, 69, 103, 104). However, the mechanisms behind this are not well understood.

Natural killer (NK) cells and dendritic cells (DCs) have also been implicated in control of L. monocytogenes infection, although the role of NK cells is not clear. In rat models, NK cells are important for control of L. monocytogenes, whereas different murine models have indicated that NK cells mediate resistance and sensitivity to L. monocytogenes infection (88). These differences may reflect disparities in NK cell function in different animal species and strains. C57BL/6 mice depleted of NK cells clear

L. monocytogenes infection more rapidly than mice with normal NK levels (105) whereas

CB6F1 mice or rats depleted of NK cells are more susceptible to L. monocytogenes infection (106, 107). The discrepancy between these animal models may be due to different expression levels of NK cells (88). NK cells may help control infection through

12 the secretion of IFNγ (106, 108), which has long been appreciated to be an import cytokine in the immune response to L. monocytogenes (109-111), primarily because IFNγ activates macrophage bactericidal capabilities (95). DCs prime naive CD8+ T cells and induce an efficient CD8+ T-cell memory response (112, 113). A subset of DCs (Tip DCs) are a principal source of tumor necrosis factor (TNF) and inducible nitric oxide synthase

(iNOS) and are important for control of L. monocytogenes infection. The majority of Tip

DCs are not infected, thus the TNF and RNI that they produce likely limit L. monocytogenes growth in other cells (114, 115).

1.2.2 Adaptive Immunity

Although the innate immune system is critical for initial control of infection, T cells are required for sterilizing immunity (84, 110, 116, 117). Severe combined immunodeficiency (SCID) mice, which lack T and B cells, mount an initial response to, but do not resolve, L. monocytogenes infection (118). Lack of sterilizing immunity in these mice is due to a lack of T cells; adoptive transfer of T cells, but not antibodies in serum, protects mice from L. monocytogenes infection (85, 118-120). Differentiation of

CD4+ T cells into Th1 cells leads to production of Th1 cytokines, which are thought to enhance elimination of L. monocytogenes during infection (121). CD4+ T cells are not required for priming of naive CD8+ T cells but are required for generation of an optimal

CD8+ T cell memory response (122, 123). Conversely, CD8+ T cell proliferation during secondary infection is repressed by CD4+ regulatory T cells (124). CD4+ and CD8+ T cells are both important for elimination of L. monocytogenes, but CD8+ T cells play a

13 larger role in clearance of L. monocytogenes (125-128). The exact mechanisms used by

CD8+ T cells to control L. monocytogenes infection are unknown, but may involve lysis of infected cells via perforin and granzymes. However, perforin deficient CD8+ T cells still provide sterilizing immunity (129). Memory CD8+ T cells are an important source of

IFNγ, which, as mentioned above, is an important activator of macrophages (130, 131).

1.3 Neutrophils

Neutrophils constitute ∼70% of circulating white blood cells in humans and are the primary responders to the site of infection. They are granular phagocytic cells with multi-lobed nuclei and hence are often called polymorphonuclear leuckocytes (PMNs).

Neutrophil granules store high concentrations of antimicrobial molecules and proteases, which can be toxic to the host as well as invading organisms. Neutrophils can kill extracellular and intracellular bacteria through both oxidative-dependent and

-independent mechanisms. They are short-lived and typically undergo apoptotic cell death, thus maintaining the damaging granular components within intact membranes.

Apoptotic neutrophils are cleared by macrophages, thereby reducing neutrophil mediated tissue damage and enhancing macrophage antimicrobial responses, discussed in further detail below. Neutrophils exert direct antimicrobial activity and cooperate with other cells to enhance antimicrobial actions of other host cells (132-135).

1.3.1 Neutrophil oxidative killing mechanisms

Neutrophils have a robust respiratory burst, which is rapidly initiated upon activation. Components of the nicotinamide adenine dinucleotide phosphate (NADPH) oxidase are located in the cytosol (p47phox, p67phox, and p40phox) and granule membranes

phox phox (flavocytochrome b558 subunits gp91 and p22 ). Upon activation, the cytosolic and membrane components assemble, forming the NADPH oxidase at both the phagosome and plasma membrane. Thus, ROS produced by NADPH oxidase are released into the phagosome and the extracellular environment (136-138). NADPH oxidase reduces

- molecular oxygen to superoxide (O2 ), which dismutates to form hydrogen peroxide

(H2O2). Myeloperoxidase (MPO), stored in the primary granules, can oxidize chloride ions, yielding hypochlorous acid (HOCl). Other ROS species, including singlet oxygen

1 . ( O2) and hydroxyl radicals (OH ), are also generated (133, 139, 140). The importance of the NADPH oxidase during infection is emphasized in individuals with chronic granulomatous disease (CGD), who do not have a functional NADPH oxidase.

Individuals with CGD are highly sensitive to fungal and bacterial infections (141, 142).

1.3.2 Neutrophil non-oxidative killing mechanisms

Phagosome maturation in neutrophils is quite distinct from other phagocytic cells as it occurs rapidly, involves fusion with granules, and does not involve acidification

(133, 139, 143). In neutrophils, phagosomes fuse with granules as opposed to endosomes and lysosomes. Furthermore, granule fusion can begin before the phagosome seals (Fig.

1.4) (133, 143-145). Although vATPase is recruited to the neutrophil phagosome, 15 phagosomes do not acidify, instead they become slightly basic (pH 7.8), then return to neutral pH or slightly acidify (pH 6.5) during the maturation process (146, 147). This is thought to be due to the highly active NADPH oxidase, which removes protons from the phagosome during dismutation of superoxide into hydrogen peroxide, and passive leakage of hydrogen ions out of the phagosome (133, 139, 146, 147).

Neutrophils also form neutrophil extracellular traps (NETs), complexes of histones and antimicrobial peptides on a DNA scaffold, through a process called

NETosis. NETs entrap extracellular bacteria in close contact with toxic granular components, thus preventing bacterial spread and mediating extracellular killing of bacteria (148). While NET formation has long been thought to require neutrophil death, neutrophils can extrude mitochondrial DNA to form NETs while maintaining their viability (149). The steps leading to formation of NETs are not well known, but may involve ROS production (150).

1.3.3 Neutrophil granules

Neutrophils contain secretory vesicles and three granule subsets (Fig. 1.4), which are released at the plasma membrane in the following order: secretory vesicles, tertiary/gelatinase granules, secondary/specific granules, and lastly primary/azurophilic granules. Granule release into the phagosome mainly happens in the inverse order, and typically only the primary/azurophilc granules and the secondary/specific granules merge with the phagosome (132, 134, 151).

Neutrophil granules and vesicles contain a range of proteases and antimicrobial molecules (stored within the granule or vesicle matrix) and receptors (stored in the membrane) (Fig 1.4). Highlighting just a few of the granule contents, the primary granules store defensins, which constitute 30-50% of total protein in these granules (152), serine proteases, and MPO. Secondary granules contain cathelicidins, lactoferrin, matrix metalloproteases (MMPs), and adhesion molecules. The tertiary granules mainly contain

MMPs and adhesion molecules while secretory vesicles contain matrix proteins, phagocytic receptors and adhesion molecules. Thus, primary and secondary granules mostly contain antimicrobials to mediate control of infection while tertiary granules and secretory vesicles mediate neutrophil extravasation to the tissue (132, 134, 153, 154). The

Figure 1.4. Neutrophil granules. Neutrophils produce secretory vesicles and three types of granules (primary, secondary, and tertiary), which contain many proteases and antimicrobials, only some of which are listed here. Components listed in the first line below the granule or vesicle name are stored in the matrix, while membrane components are listed in the second line.

17 importance of neutrophil granules is highlighted in individuals with neutrophil-specific granule deficiency (SGD). These individuals lack secondary/specific granules and some proteins in primary granules, like the defensins. People with SGD are highly susceptible to fungal and bacterial infections (142).

1.3.4 Defensins

Defensins are amphiphilic, cationic, 2 to 5 kDa, antimicrobial peptides (AMPs), which are differentiated from other classes of AMPs based on their six cysteine residues that form three intramolecular disulfide bridges. The position of the cysteines and the pattern of cysteine connectivity divide the defensins into α-, β-, and θ-defensins (Fig. 1.5)

(155-157). The human genome encodes all three types of defensins, but humans do not produce θ-defensins due to a premature stop codon (158-161). However, human cells can produce θ-defensins; cells treated with aminoglycosides to read-through the premature stop codon produce a fully functional θ-defensin, retrocyclin 1 (RC-1) (162). Humans naturally produce α- and β-defensins, α-defensins are primarily produced by neutrophils

(human neutrophil peptides, HNPs) and Paneth cells (human defensins, HDs) while β- defensins (human β-defensins, HBDs) are mainly produced by epithelial cells (155, 157,

163-165). The θ-defensins are only found in Old World primate phagocytes, mainly in neutrophil granules (160, 161, 166, 167).

Defensins exert potent antimicrobial and immunomodulatory activities (157).

Their antimicrobial actions include: targeting bacterial, viral, and eukaryotic pathogens, neutralizing various bacterial toxins (155, 165, 168), and regulating the host-associated 18 microbiota (169-172). The antimicrobial action of defensins has long been presumed to be mediated through disruption of microbial membranes, but defensins also bind to and

Figure 1.5 Ribbon structures and primary sequences of human α- and β- defensins and humanized θ-defensins. In the ribbon structures, disulfide bonds are in gray, the β-sheets are in yellow and the α-helix is in red. The PDB accession codes are 1DFN [HNP3, (229)], 1IJV [HBD1, (319)], and 2ATG [RC2, (320)]. Structures were generated with Chimera. The primary sequences for the defensins are also shown. Only the four β-defensins isolated from human tissues are depicted. Cysteines are in red, the disulfide linkages for each class of defensin are in gray, and the circular backbone of the θ-defensins is depicted with the black dashed line. The θ-defensins are cyclic peptides formed by the head-to-tail covalent assembly of two nonapeptides, the arrows above the θ-defensin class indicate the first residue of the two nonapeptides. The θ-defensins are not produced in humans, the provided structure and sequences are deduced from the human pseudogene sequences.

19 inhibit the synthesis of DNA, RNA and proteins; however, it is unclear if such activity or the initial membrane perturbation is primarily responsible for cell death (173-175).

Furthermore, whether membrane perforation is required for the intracellular targeting of the defensins is unknown (176). Defensins also bind to the cell-wall precursor lipid II and inhibit cell-wall synthesis (175, 177-179). Defensins likely utilize multiple mechanisms to exert their antimicrobial actions.

Defensins inhibit numerous toxins, including lipopolysaccharide (LPS) (180-182),

Bacillus anthracis lethal factor (183, 184), the mono-ADP-ribosyltransferases diphtheria toxin and Pseudomonas aeruginosa exotoxin A (185), Clostridium difficile toxin B (186) and Staphylococcus aureus staphylokinase (187). Of note, α-defensins inhibit LLO and other CDCs. This is thought to result from defensin binding to the CDCs, blocking toxin oligomerization within host membranes (188).

In addition to their direct antimicrobial and antiviral activities, mammalian defensins interact with a variety of host cells to regulate the immune response and maintain tissue homeostasis (Fig. 1.6). Exactly how defensins interact with mammalian cells is not well understood and is quite complex, resulting in both pro- and anti- inflammatory activities. In general, the defensins antimicrobial actions, but not the immunomodulatory functions, are diminished at physiological concentrations of salts and in the presence of divalent cations or serum (3, 157, 165, 189); therefore, it is thought that the defensins’ immunomodulatory activities may prevail within host tissues (156, 157,

168, 190-192).

Defensins are potent chemotactic agents, β-defensins mediate chemotactism through the chemokine receptors CCR6 and CCR2 and the α-defensins through an unidentified G protein coupled receptor (193-195). Defensins regulate the production of inflammatory messengers through TLRs (β-defensins) and the purinergic receptor P2Y6

Figure 1.6. Major activities of the defensins on mammalian cells. Schematic illustrating the broad range of defensin activities on mammalian cells. See the text for more detail. Abbreviations are as follows: Mo, Monocyte; MΦ, Macrophage; DC, Dendritic cell; PMN, Polymorphonuclear leuckocyte, Neutrophil; Ep, Epithelial cell; En, Endothelial cell; K, Keratinocyte; F, Fibroblast.

(HNPs) (196-198). Defensins also potentiate phagocytosis and intracellular killing of pathogens. Multiple processes are thought to be involved in this, defensins can act as opsonins and activate phagocytic functions. In addition, the delivery of defensins into phagosomes participates in intracellular killing of the phagocytosed pathogen (199-201).

Defensins also activate antigen presenting cells, increasing membrane receptor expression on these cells in a TLR dependent manner (197, 198). Defensins enhance migration and proliferation of fibroblasts and epithelial cells and induce angiogenesis, all of which are involved in tissue remodeling during wound healing (157, 202, 203). Thus, defensins exert many immunomodulatory actions (Fig. 1.6).

1.4 Neutrophil cooperation with macrophages

Neutrophils contain substantially more antimicrobial molecules than other host cells and at infectious foci neutrophils are in close contact with other cells, including monocytes and macrophages (89). It is likely that neutrophils cooperate with other host cells to control infection. In fact, neutrophil/macrophage cooperation may be involved in the control of intracellular pathogens such as Mycobacterium tuberculosis and Legionella pneumophila (204-206), but this has not been studied in regards to L. monocytogenes.

Cooperation between neutrophils and macrophages enhances macrophage bactericidal activity (207-209), this can occur through neutrophil mediated activation of macrophages or macrophage internalization of neutrophils, neutrophil granules, or granule components (210, 211). These cooperation mechanisms can occur in a contact- dependent or -independent manner. For example, macrophage activation can occur

22 through contact with neutrophils or be induced by components released by neutrophils in a contact-independent manner. Neutrophils cooperate with macrophages to limit

Leishmania braziliensis infection in a contact-dependent manner than involves TNFα and

ROS production (212). Neutrophil granule proteins (defensins and heparin-binding protein) activate macrophages, inducing TNFα and IFNγ release, which act in an autocrine manner to enhance macrophage expression of Fcγ receptors and phagocytosis

(200). Neutrophil granule components also enhance the macrophage ROS burst (206).

Similarly, macrophage internalization of neutrophils is contact-dependent, but internalization of released neutrophil components does not require contact with neutrophils. Macrophage phagocytosis of apoptotic neutrophils or purified neutrophil granules decreases M. tuberculosis infection of macrophages, presumably due to internalization of defensins, as they colocalized with M. tuberculosis in the phagosome

(213). Several antimicrobial molecules produced by neutrophils (defensins, lactoferrin, and MPO) are taken up by macrophages in vivo and in vitro, enhancing the macrophage antimicrobial activity (200, 204, 205, 214).

1.5 Research Hypothesis and Goals

The central hypothesis of this dissertation is that neutrophils control L. monocytogenes infection through multiple processes, including killing of phagocytosed

L. monocytogenes and cooperating with other host cells to enable those cells to control L. monocytogenes intracellular replication. The main goal of this dissertation is to elucidate the interaction between L. monocytogenes, its main virulence factor LLO, and innate

23 immune cells (neutrophils and macrophages) and innate immune cell products. To address this goal, we: i) determined if neutrophil defensins inhibit L. monocytogenes replication in macrophages, ii) elucidated the interaction between L. monocytogenes,

LLO, and neutrophils, and iii) determined if neutrophils cooperate with macrophages to prevent L. monocytogenes replication in macrophages. Importantly, we used human neutrophils and human neutrophil products to address this hypothesis, as human and murine neutrophils are quite distinct. For example, in humans the most abundant proteins in neutrophil primary granules are defensins, which are lacking in murine neutrophils

(157).

i) Determine if neutrophil defensins inhibit L. monocytogenes replication in macrophages. Defensins are the most abundant protein in human neutrophil primary granules, and they exert potent antibacterial and anti-toxin activities. It is unknown if defensins inhibit LLO-mediated perforation of macrophages or L. monocytogenes intracellular replication. Using primary murine macrophages as a model, we determined if human defensins protect macrophages from LLO-mediated perforation and prevent L. monocytogenes intracellular replication in macrophages.

ii) Elucidate the interaction between L. monocytogenes, LLO, and neutrophils.

Neutrophils are important for control of L. monocytogenes infection, yet little is known regarding the interaction between neutrophils and L. monocytogenes. LLO plays an important role in the L. monocytogenes intracellular life cycle in other cell types but the role of LLO in the interaction between neutrophils and L. monocytogenes is unclear, both regarding how LLO affects neutrophils and if LLO protects L. monocytogenes from

24 neutrophil killing. We used primary human neutrophils to determine if LLO protects L. monocytogenes from neutrophil killing and elucidated the interactions between neutrophils and LLO.

iii) Determine if neutrophils cooperate with macrophages to prevent L. monocytogenes replication in macrophages. Neutrophils have been proposed to cooperate with macrophages to limit infection of intracellular pathogens like M. tuberculosis and L. pneumophila. However, it is unknown if neutrophils cooperate with macrophages during

L. monocytogenes infection. Using human neutrophils and a human monocyte cell line that was differentiated into macrophages (Thp1 cells), we determined if co-culturing macrophages with neutrophils alters L. monocytogenes phagocytosis by and/or replication in macrophages.

Chapter 2. Defensins enable macrophages to inhibit the intracellular proliferation of Listeria monocytogenes

2.1 Introduction

Antimicrobial peptides (AMPs) are structurally diverse molecules that are active

against bacteria, fungi, viruses, and parasitic protozoa (191, 215-219). AMPs combat

microbes by disrupting their membranes and/or interacting with various intracellular

targets to inhibit DNA and protein synthesis, protein folding, enzymatic activity, or cell

wall synthesis (199, 216, 218, 220). In higher organisms, AMPs also exert immuno-

modulatory functions (156, 199, 221). More recently, certain AMPs (α-defensins) were

shown to inhibit bacterial toxins such as anthrax lethal factor, diphtheria toxin,

Clostridium difficile toxin B and the cholesterol-dependent cytolysins (CDCs), including

listeriolysin O (LLO) (183-186, 188, 222).

In humans, defensins and other AMPs are localized on the skin, at mucosal

surfaces, in secretions, and in body fluids and tissues (221, 223). Significant amounts of

AMPs are delivered by neutrophils to infected tissues at early stages of infection (224-

226). Neutrophil azurophil granules contain high concentrations of AMPs that fuse

primarily with phagosomes but are also released in the extracellular environment (227).

Human α-defensins HNP-1, 2 and 3 are cationic, amphipathic and cysteine-rich peptides

26 with 29 or 30 amino acid residues and nearly identical sequences. Collectively, they comprise 30 to 50% of the total protein content of azurophil granules, wherein their concentrations approximate 50 mg/ml (199, 215, 228, 229). Several reports have suggested that a cooperation with neutrophils can enhance macrophage bactericidal activity (207, 208, 226). One potential mechanism of the neutrophil-macrophage cooperation involves the transfer of antimicrobial molecules from neutrophils to macrophages. Such a mechanism may be involved in controlling intracellular pathogens such as Mycobacterium tuberculosis and Legionella pneumophila (204-206). In favor of this idea, several antimicrobial molecules produced by neutrophils (defensins, lactoferrin, and myeloperoxidase) are taken up by macrophages in vivo and in vitro, enhancing macrophage antimicrobial activity (200, 204, 205, 213, 214).

Listeria monocytogenes is a facultative intracellular pathogen that crosses the intestinal, materno-fetal, and blood-brain barriers causing severe infections (1, 8). L. monocytogenes pathogenicity involves LLO, a CDC that acts collaboratively with bacterial phospholipases to disrupt the membrane of the phagosome, mediating bacterial escape into the cytoplasm (33, 230). The innate immune responses play an essential role in controlling L. monocytogenes growth and dissemination, preventing the spread into systemic, lethal infection (85). Neutrophils appear to have critical functions during the early stages of listeriosis, as their absence in humans and animal models exacerbates infection (66, 96, 98, 100-102, 224). Neutrophils may directly phagocytose and kill bacteria; however, little is known about L. monocytogenes interactions with neutrophils and a recent report indicates that L. monocytogenes are not efficiently killed by these

27 cells (231). The present study tested the hypothesis that the α-defensin HNP-1 could enhance the ability of macrophages to control L. monocytogenes infection. For comparison, we also tested retrocyclin-1 (RC-1), a humanized version of the cyclic octadecapeptides (θ-defensins) found in the neutrophils of nonhuman primates (161, 216,

232). Our findings strengthen the belief that neutrophils, and/or their products, can enhance the antimicrobial performance of monocytes (233) or macrophages (204, 205,

213).

2.2 Materials and Methods

2.2.1 Reagents

HNP-1 and RC-1 were chemically synthesized as described previously (160, 165,

234). Defensins were stored in 0.01% acetic acid at -80oC. Recombinant six-histidine- tagged LLO was purified from Escherichia coli BL21(DE3) harboring the pET29b plasmid encoding LLO (provided by Dr. D. A. Portnoy, University of California,

Berkeley, CA) as described previously (235). LLO was stored at -80oC in 1 M NaCL, 1 mM EDTA, 50 mM phosphate buffer, pH 6. The purity and integrity of LLO was assessed by SDS-PAGE followed by Coomassie blue staining and Western blotting.

Polyclonal rabbit against L. monocytogenes was from GeneTex, rabbit anti-LLO was from Abcam, monoclonal mouse anti-HNP was from Hycult, rat anti-LAMP-1 clone

1D4B (developed by J. T. August and stored at the Developmental Studies Hybridoma

Bank under the auspices of the NICHD and maintained by the University of Iowa) was a gift from Dr. A. Amer, Ohio State University, and secondary horseradish peroxidase- 28 conjugated antibody was from Cell Signaling. Secondary fluorescent antibodies, phalloidin conjugated to Alexa488, and ProLong Gold antifade containing DAPI were purchased from Molecular Probes. Antibodies were used at 5 µg/ml (10 µg/ml for anti-

HNP) and phalloidin at 3 Units/ml. Ethidium homodimer (EthD) was purchased from

Fluka. Enhanced chemiluminescence reagent for Western blotting was purchased from

Amersham. Dulbecco’s modified Eagle’s medium (DMEM) was purchased from

Invitrogen, fetal bovine serum (FBS) from Lonza and brain heart infusion (BHI) from BD

Biosciences. Paraformaldehyde (PFA) and glutaraldehyde were purchased from Sigma-

Aldrich.

2.2.2 Cell culturing, bacterial strains and infection

Bone marrow-derived macrophages (BMM) were obtained from tibias and femurs of 5-6 week old C3HOuJ mice (Jackson Laboratories) as previously described (236).

BMM were frozen in DMEM supplemented with 30% L-cell conditioned media, 20% heat inactivated-FBS (HI-FBS), 10 units penicillin/streptomycin (Pen/Strep), 50.3 µM β- mercaptoethanol, and 10% dimethylsulfoxide. L-cell conditioned media was obtained by collecting media from confluent NCTC clone 929 (ATCC # CCL-1) cells. The day before the experiment, BMM were thawed and plated (1.2 x 105 BMM/well) onto glass coverslips in 24-well tissue culture dishes in DMEM supplemented with 10 units

Pen/Strep and 10% HI-FBS. RAW 264.7 cells (ATCC TIB-71) were maintained in

DMEM supplemented with 100 units Pen/Strep and 10% HI-FBS. The day before the experiment, RAW cells were plated (1.2 x 105 cell /well) onto glass coverslips in 24-well 29 tissue culture dishes. The day of the experiment, cells were washed twice with DMEM to remove the antibiotics from the culture medium and incubated in 10% HI-FBS in

DMEM.

L. monocytogenes strains DP10403S (wt) and DP-L2161 (DP10403S Δhly) and the plasmid pAM401, which encodes hly with its entire regulatory region were provided by Dr. D. A. Portnoy (University of California, Berkeley, CA). L. monocytogenes over- expressing LLO was generated by electroporation of pAM401 into strain DP10403S and was grown under standard conditions, with the addition of 20 µg/ml chloramphenicol (32,

237). Unless otherwise indicated, bacteria were cultured overnight in BHI at 37°C, diluted 1:20 in BHI, and cultured at 37°C with shaking until bacteria reached OD 600 =

0.8. Bacteria were washed with PBS and added to the macrophages at MOI 0.1 in 0.5 ml for the cluster and gentamicin survival assays, and MOI 3 for all other assays. RAW cells were infected at MOI 0.4. Infection was carried out in serum-free DMEM. The cell culture plate was centrifuged for 3 min at 230 g. The cells were then incubated for 15 min at 37°C, washed to remove unbound bacteria, and fixed (phagocytosis assay) or further incubated with 15 µg/ml gentamicin and 10% HI-FBS in DMEM (all other assays).

When pre-exposing bacteria to defensins or gentamicin, bacteria were incubated with defensins or gentamicin in BSA-coated tubes in serum-free DMEM. After 15 min at

37°C, the bacteria were washed with PBS, then diluted to MOI 0.1 or 3 for infection of the BMM. When pre-incubating macrophages with defensins, BMM were pre-incubated with defensins for 15 min at 37°C in serum-free DMEM, washed with DMEM, then infected as described above.

2.2.3 Fluorescence labeling

Following bacterial infection, cells were washed and fixed with 4% PFA in PBS, pH 7.4 for 15 min at room temperature (RT). Cells were washed with 0.1 M glycine in

PBS and incubated in blocking solution (0.1 M glycine, 10% HI-FBS in PBS, pH 7.4) for

1 h at RT. Antibodies were diluted in PBS containing 10% HI-FBS (plus 0.2% saponin for LAMP-1, F-actin, and HNP-1 labeling) and incubated for 30 min at RT. Cells were washed twice with PBS between each antibody incubation. Coverslips were mounted with ProLong Gold antifade containing DAPI.

Cluster assay-Following blocking, macrophages were permeabilized with 0.2%

Triton X-100 in PBS for 5 min at RT, then total (extracellular + intracellular) bacteria were labeled with the anti-Listeria antibody and the secondary Alexa568 antibody.

Phagocytic assay- Following blocking, extracellular bacteria were labeled with the anti-Listeria antibody and the secondary Alexa568 antibody. Cells were permeabilized with 0.2% Triton X-100 in PBS for 5 min at RT, then total bacteria

(intracellular + extracellular) were labeled using the same primary antibody and the secondary antibody conjugated to Alexa488.

F-actin labeling- For F-actin labeling only, cells were fixed for 5 min at RT in

6.6% PFA and 0.05% glutaraldehyde in cytoskeleton stabilization buffer (138 mM KCl, 3 mM MgCl2, 2 mM EGTA, 10 mM MES, pH 6.1), washed with glycine and blocked as described above. Cells were permeabilized with 0.2% saponin in PBS for 30 min at RT.

Total bacteria (extracellular + intracellular) were labeled with the anti-Listeria antibody

31 and the secondary Alexa568 antibody. Actin was labeled with Alexa488 phalloidin in

10% HI-FBS and 0.2% saponin in PBS for 30 min at RT.

LAMP-1 labeling- Following blocking, cells were permeabilized with 0.2% saponin in PBS for 30 min at RT. Total bacteria (extracellular + intracellular) were labeled with the anti-Listeria antibody and the secondary Alexa488 antibody. LAMP-1 was labeled with the anti-LAMP-1 antibody and the Alexa568-conjugated secondary antibody.

HNP-1 labeling- Following blocking in blocking solution containing 1% BSA, cells were permeabilized with 0.2% saponin in PBS for 30 min at RT. HNP-1 was labeled with the anti-HNP antibody and the secondary Alexa488 antibody. In infected macrophages, total bacteria (extracellular + intracellular) were labeled with the anti-

Listeria and Alexa568 antibodies.

In the F-actin, LAMP-1 and HNP-1 colocalization experiments, the percentage of intracellular bacteria was determined in separate cell culture wells using the labeling protocol described for the phagocytic assay.

2.2.4 Image Acquisition

Images were acquired on a motorized inverted epi-fluorescence microscope (Axio

Observer D1, Zeiss) equipped with 20 X Plan Neofluar (N.A. = 0.5) and 100 X Plan Apo

(N.A. = 1.4) objectives. The high speed filter changer Lambda DG-4 (Xenon Arc bulb), an optical emission filter wheel Lambda 10-3 for the fluorescence imaging, and a Smart shutter that controls phase-contrast illumination were from Sutter Instrument Company.

The camera (back-illuminated, frame-transfer EMCCD Cascade II 512) was from

Photometrics. The filter sets for fluorescence were purchased from Chroma Technology

Corporation: DAPI (49000), Alexa488 (49002), Alexa568 (49005). The microscope equipment was controlled by MetaMorph imaging software.

2.2.5 Image Analysis

Cluster assay- 40 sets of fluorescence images (DAPI and Alexa568) were automatically acquired for each experimental condition using the 20 X objective. Clusters were counted through direct visualization and were defined as a minimum of four bacteria per cell. The number of BMM was automatically counted by the software based on DAPI fluorescence (238). About 4,500 BMM were counted in each experimental condition. The number of bacterial clusters /BMM was enumerated in each experimental condition and the results were expressed as intracellular proliferation relative to the control (BMM incubated with wild type bacteria without defensin treatment).

Phagocytic Assay- 40 sets of fluorescence (DAPI, Alexa488 and Alexa568) and phase-contrast images were automatically acquired for each experimental condition using the 20 X objective. BMM, extracellular and total bacteria were automatically counted by the imaging software. The number of intracellular bacteria [(total bacteria) –

(extracellular bacteria)] /BMM was calculated as described previously (238). About

4,500 BMM were counted in each experimental condition. The results were expressed as a phagocytic index corresponding to the percentage of intracellular bacteria per BMM in

33 the sample test relative to the control (BMM incubated with wild type bacteria without defensin treatment).

F-actin, LAMP-1 and HNP-1 colocalization with L. monocytogenes- Fluorescence and phase-contrast images were acquired using the 100 X objective. A minimum of 100 bacteria was counted by direct observation in each experimental condition. The total number of bacteria (extracellular + intracellular) per BMM (L.m. per cell) was enumerated (Fig. 2.7C and E). The percentage of total bacteria that colocalized with F- actin, LAMP-1 or HNP-1 was then enumerated (Total L.m.%). The percentage of intracellular bacteria (% intracellular L.m.) was enumerated in each experimental condition as described for the phagocytic assay. Finally, results were expressed as the percentage of intracellular bacteria that colocalized with F-actin, LAMP-1 or HNP-1 according to the formula: (100 x Total L.m.%) / (% intracellular L.m.) (Figs. 2.7B and D,

2.8B and C, and 2.10D).

HNP-1 uptake- 20 sets of fluorescence images (DAPI and Alexa488) were automatically acquired for each experimental condition using a 40 X objective. Images were automatically analyzed with the MetaMorph software. The number of BMM

(NBMM) was automatically counted based on DAPI images. To quantify the average fluorescence of HNP-1 per BMM, the Alex488 images were first background corrected and a threshold was applied such that only the sum of the fluorescence intensities (SFI) of the pixels located within the BMM was measured. The results were expressed as the mean fluorescence intensity (MFI) per cell according to: (MFI per cell) = (SFI) / NBMM.

About 500 BMM were counted in each experimental condition.

2.2.6 Gentamicin survival assay

BMM were incubated with defensins and L. monocytogenes (10403S) at MOI 0.1 in serum-free DMEM and the cell culture plate was centrifuged for 3 min at 230 g. After

15 min at 37oC, cells were washed and further incubated with 15 µg/ml gentamicin and

10% HI-FBS in DMEM. After 3 h of infection at 37oC, macrophages were washed with

PBS and lysed with 0.2% Triton X-100 in H2O. Serial dilutions of cell lysates were immediately performed in PBS and bacteria were plated on BHI agar. Plates were incubated at 37oC for 48 h and the colony-forming units (CFU) were enumerated.

2.2.7 Antimicrobial activity of the defensins

CFU Assay- L. monocytogenes (7.2 x 105/ml) were incubated in DMEM with the indicated concentrations of defensins or gentamicin for 15 min or 3 h at 37°C. Bacteria were then plated on BHI agar to enumerate the CFU (Fig. 2.4A and B). Alternatively, L. monocytogenes (7.2 x 105/ml) were incubated in DMEM with the indicated concentrations of defensins or gentamicin in BSA-coated tubes for 15 min, washed twice with warm PBS, and either directly plated on BHI agar (Fig. 2.4C) or incubated at 2.4 x

104/ml for 3 h in DMEM at 37°C, then plated on BHI agar (Fig. 2.4D).

Virtual Colony Counting Assay- L. monocytogenes was cultured overnight in trypticase soy broth (TSB) at 37°C, diluted 1:11 in TSB and cultured at 37oC in a shaking water bath until bacteria reached OD600 = 0.5. Bacteria were diluted 1:10 with cold

DMEM and placed on ice to delay further bacterial multiplication. A standard curve was calculated by incubating serial 10-fold dilutions of bacteria (2.5 x 106 CFU /ml - 2.5 x 102

CFU / ml) in DMEM (Fig. 2.5A). After 15 and 90 min at 37°C, the bacteria were washed twice, suspended in 100 µl PBS, and placed on ice. The samples were transferred to 96- well plates (Corning Costar No. 3596) containing 100 µl TSB and absorbance at 650 nm

(OD 650) was measured every 5 min for 16 h using a Spectra Max Gemini XS UV-Vis microplate spectrophotometer (Molecular Devices). The time required by each inoculum to attain a ΔOD of 0.1 absorbance unit (corresponding to an increase of ~8.6 x 107

CFU/ml) was determined and the least mean squares fit of the data points was calculated

(Fig. 2.5B). Once the standard curves were determined, L. monocytogenes (2.5 x 105

CFU/ml) was incubated for 15 or 90 min with the indicated concentrations of HNP-1 or

RC-1 in DMEM in pre-lubricated microcentrifuge tubes (Corning Costar No. 3207). The bacteria were washed twice, suspended in 100 µl PBS, and placed on ice. The samples were transferred to 96-well plates containing 100 µl TSB and the OD 650 was monitored for 16-20 h every 4 min with interval shaking. The least mean squares fit of the data points generated from the standard curve was used to calculate x, the number of untreated bacteria that would have attained a ΔOD of 0.1 in the same time. The results were expressed as percentage imputed survival, calculated as [x/ (initial number of bacteria) X

100] and are the mean of two independent experiments, performed in triplicate (Fig. 2.5C and D). Pratikhya Pratikhya (Dr. Lehrer’s lab; David Geffen School of Medicine at

UCLA, Los Angeles, CA) performed these experiments.

2.2.8 Hemolytic assay

LLO-mediated lysis of erythrocytes was measured with a kinetic assay (188) performed in 96-well plates with a final volume of 200 µl per well as follows: 20 µl LLO

(final concentration 0.075 nM), 20 µl RC-1 (final concentration 5-10 µg/ml) and 160 µl

7 erythrocytes (1 .7 x 10 /ml). Plates were placed in a PowerWavex340 spectrophotometer

(Bio-Tek) at 37°C for 30 min and optical density at 700 nm (OD 700) was measured every 5 min. Erythrocytes incubated with PBS or 0.1% Triton X-100 in PBS served as negative and positive controls, respectively.

2.2.9 Measurement of BMM perforation by LLO

Bone marrow-derived macrophages were seeded (1.2 x 105 cells/well) in 24-well cell culture dishes and incubated at 37°C for 24 h. BMM were washed twice with ice cold DMEM and incubated on ice in serum-free DMEM containing 4 µM ethidium homodimer (EthD). Defensins (0-20 µg/ml) and LLO (2.5 nM) were added and cells were incubated at 37°C for 15 min, washed with PBS and fixed with 4% PFA in PBS for 15 min. Coverslips were mounted with DAPI. To measure membrane perforation, 30 sets of images (DAPI, EthD, and phase-contrast) were automatically acquired for each experimental condition using the 20 X objective. Images were automatically analyzed with the MetaMorph software. The number of BMM was counted based on DAPI fluorescence (238), about 3,500 BMM were analyzed in each experimental condition. To quantify membrane perforation, we measured the average fluorescence intensity of EthD in the nuclear regions as follows. First, all EthD images were background corrected. 37

Second, based on the DAPI images, mask images were created and applied to the EthD images such that only the average fluorescence intensity of EthD within the nuclear regions was quantified. Results were expressed as perforation relative to the control

(BMM incubated with LLO and EthD without defensins).

2.2.10 LLO release by L. monocytogenes

L. monocytogenes that overexpress LLO from a plasmid were grown in BHI until

6 OD 600 = 0.8, washed three times with PBS and diluted (7.2 x10 bacteria/ml) in serum- free DMEM in BSA-coated tubes. Defensins were added and after 0, 30 and 60 min at

37°C, bacteria were centrifuged and bacterial supernatants were collected. Proteins were precipitated in 10% trichloroacetic acid for 60 min on ice, then samples were centrifuged

(11,000 g for 20 min at 4°C) and the precipitates were washed twice with cold acetone.

The protein pellets were air-dried and suspended in Laemmli’s sample buffer containing a reducing agent. LLO was detected by dot blot using rabbit anti-LLO and horseradish peroxidase-conjugated secondary antibodies.

2.2.11 Statistics

A minimum of three independent experiments were performed in duplicate for each assay (unless indicated otherwise). Data were expressed as mean ± standard error of the mean (SEM). P-values were calculated using a standard two-tailed t-test and

38 considered significant if lower than 0.05. Asterisks indicate a significant difference between the indicated experimental condition and control, * P < 0.05, ** P <0.01.

2.3 Results

2.3.1 Defensins inhibit L. monocytogenes proliferation in macrophages

We developed a fluorescence microscopy-based assay, called the cluster assay, to determine whether HNP-1 or RC-1 could influence the intracellular fate of L. monocytogenes in murine bone marrow-derived macrophages (BMM) and RAW 264.7 cells. In this assay, macrophages were infected with L. monocytogenes (multiplicity of infection; MOI 0.1) for 15 min at 37°C in the presence or absence of HNP-1 or RC-1.

The macrophages were washed and incubated for a total of 3 h at 37°C with gentamicin, a cell-impermeant antibiotic. The macrophages were then washed, fixed and permeabilized to fluorescently label bacteria and macrophage nuclei. The number of bacterial clusters, defined as a minimum of four bacteria per macrophage, was enumerated as a read-out of L. monocytogenes proliferation in macrophages. To validate this method, we verified that: i) no cluster was observed after 15 min of incubation at

37°C (Fig. 2.1A); ii) no extracellular cluster was observed at any time points (assessed by labeling bacteria in non-permeabilized macrophages, data not shown); and iii) after 3 h, clusters were observed in macrophages infected with wild type L. monocytogenes but not in macrophages infected with a LLO-deficient (Δhly) L. monocytogenes that is unable to replicate within macrophages (Fig. 2.1A and B) (13, 92). Together, these observations

Figure 2.1. Defensins inhibit L. monocytogenes intracellular proliferation. BMM were incubated at 37°C for 15 min with L. monocytogenes DP10403S (wt) or DP-L2161 (Δhly) at MOI 0.1 in the presence or absence of defensins. Cells were washed and incubated for a total of 3 h with gentamicin. (A and B) Cells were fixed and permeabilized. Bacteria and BMM nuclei were fluorescently labeled. (A) Representative images of BMM incubated with bacteria in the absence of defensin. The arrow indicates one of three clusters of bacteria. (B) The number of clusters/BMM was counted in each experimental condition and results were expressed relative to the control (BMM incubated with wt bacteria in the absence of defensin). * P < 0.05, ** P < 0.01. (C) Cells were lysed and lysates were plated on BHI agar to count the CFU. Results of one experiment, performed in triplicate, are provided. The control consists of untreated BMM.

demonstrate that bacterial clusters were formed as a result of intracellular bacterial proliferation. We observed that in both macrophage models, defensins prevented L. monocytogenes intracellular multiplication (Figs. 2.1B and 2.2). We selected BMM to pursue this study because they were more phagocytic than the RAW 264.7 cells.

Specifically, after 15 min of infection at 37°C, 58% of L. monocytogenes associated with

BMM were intracellular, whereas only 8% of RAW cell-associated L. monocytogenes were intracellular.

The inhibition of L. monocytogenes intracellular multiplication by HNP-1 and

RC-1 was concentration-dependent (Fig. 2.1B). RC-1 was more efficient than HNP-1 as

0.25 µg/ml RC-1 (0.13 µM) reduced bacterial proliferation by about 50%, whereas 10

µg/ml HNP-1 (approximately 2.9 µM) was required to exert a similar effect. We also performed a standard gentamicin survival assay that enumerates viable bacteria following

Figure 2.2. Defensins inhibit L. monocytogenes intracellular proliferation in RAW 264.7 cells. RAW 264.7 cells were incubated at 37°C for 15 min with the defensins and L. monocytogenes at MOI 0.4. Cells were washed and incubated for a total of 3 h with gentamicin. Cells were fixed and permeabilized. Bacteria and BMM nuclei were fluorescently labeled. The number of clusters/macrophage was counted and results were expressed as intracellular bacterial proliferation relative to the control macrophages (C=control) incubated with bacteria in the absence of defensin.

41 lysis of the BMM and obtained similar results (Fig. 2.1C). Together, these results clearly demonstrate that HNP-1 and RC-1, when added during phagocytosis, inhibit subsequent

L. monocytogenes proliferation in BMM and RAW 264.7 macrophages. Three hypotheses can explain these results. First, the defensins target the bacteria, inhibiting their division and/or blocking the bacterial virulence factors. Second, defensins activate the macrophage bactericidal activity. Finally, a synergy between the macrophage and defensins leads to efficient control of pathogen proliferation.

2.3.2 HNP-1 interaction with macrophages and RC-1 interaction with L.

monocytogenes are sufficient to inhibit intracellular proliferation

To further elucidate the mechanism(s) of action used by the defensins to inhibit L. monocytogenes intracellular proliferation, we determined whether the defensins acted on the bacterium or on the macrophage. We pretreated the bacteria or the macrophages with the defensins for 15 min at 37°C, washed the cells to remove the unbound defensins and conducted the cluster assay in the absence of defensin. As presented in Figure 2.3A, pre- exposure of the bacteria to HNP-1 did not prevent subsequent bacterial multiplication in macrophages. However, bacteria pre-exposed to RC-1 were unable to multiply intracellularly, as were bacteria pre-exposed to the bactericidal antibiotic gentamicin (Fig.

2.3C). We then observed that bacterial proliferation was inhibited in BMM pre-incubated with HNP-1, but not RC-1 (Fig. 2.3B and D). These data indicate that HNP-1 and RC-1 prevent L. monocytogenes growth within macrophages through distinct mechanisms.

Based on these observations, we hypothesized that RC-1 is a more potent anti-listerial

42 peptide than HNP-1, acting primarily by targeting the bacteria; whereas HNP-1 acts at the macrophage/bacterial interface to protect the macrophage against subsequent L. monocytogenes infection.

We next determined whether the defensins interaction with L. monocytogenes during (Fig. 2.1B) or before (Fig. 2.3A and C) phagocytosis was sufficient to kill the bacteria in the extracellular environment and restrict their subsequent intracellular

Figure 2.3. HNP-1 interaction with macrophages and RC-1 interaction with L. monocytogenes are sufficient to inhibit bacterial intracellular proliferation. (A and C) L. monocytogenes (7.2 x 105/ml) was pre-exposed to 30 µg/ml gentamicin (Gent.) or the indicated concentrations of defensins in DMEM at 37°C. After 15 min, bacteria were washed and added to BMM at MOI 0.1 (2.4 x 104/ml). (B and D) BMM were pre-incubated with the defensins for 15 min at 37°C, washed, then infected with L. monocytogenes at MOI 0.1 (2.4 x 104/ml). (A - D) After 15 min of infection, BMM were washed, incubated for a total of 3 h with gentamicin, fixed and permeabilized. Bacteria and macrophages were fluorescently labeled. The number of clusters/BMM was counted and results were expressed as the percentage of bacterial proliferation relative to the control (C, no defensin treatment). * P < 0.05, ** P < 0.01.

43 proliferation. We incubated the bacteria with the defensins for 15 min at 37°C in the macrophage culture medium (DMEM), as performed in the cluster assay (Fig. 2.1).

Bacteria were then plated on BHI agar to determine bacterial survival. Figure 2.4A shows that the concentrations of HNP-1 and RC-1 used in the cluster assay were not bactericidal within 15 min. We repeated this experiment while increasing the incubation time or the concentration of peptides to determine whether these defensins could exert an anti- listerial activity. Assuming that bacteria are phagocytosed with the defensins in the cluster assay (Fig. 2.1), we tested if the defensins could affect bacterial growth over a longer period of time. We observed that HNP-1 (20 µg/ml) and RC-1 (0.5-1 µg/ml) inhibited bacterial growth in DMEM after 3 h of incubation at 37°C (Fig. 2.4B), resulting in approximately 30% (20 µg/ml HNP-1, 0.5 µg/ml RC-1) and 70% (1 µg/ml RC-1) bacterial killing. We also noted that RC-1 is a more potent bactericidal peptide than

HNP-1. Indeed, within 15 min 4 µg/ml RC-1 killed 100% of L. monocytogenes, whereas

HNP-1 had little effect on bacterial viability, even at concentrations as high as 180 µg/ml

(Fig. 2.4C). Together, these results indicate that at the concentrations used in the cluster assay (Figs. 2.1 and 2.3A and C), viable bacteria were phagocytosed by the macrophages and that HNP-1 and RC-1 could inhibit bacterial growth in DMEM when present during the entire time of the experiment. We then determined whether the absence of HNP-1, but not RC-1, activity on bacteria pre-exposed to the peptides for 15 min and washed before

BMM infection (Fig. 2.3A and C), was due to a difference between HNP-1 and RC-1 antimicrobial activity. Bacteria were pre-exposed to HNP-1 or RC-1 for 15 min, washed and diluted as performed in the cluster assay. The bacteria were then incubated in DMEM

44 at 37°C for 3 h and the CFU were enumerated on BHI agar. As presented in Fig. 2.4D, bacteria pre-exposed to RC-1, but not HNP-1, failed to grow in DMEM, indicating that

RC-1 is a more efficient anti-listerial peptide than HNP-1 and/or that HNP-1, but not RC-

1, was efficiently washed away from the bacterial surface.

Figure 2.4. RC-1 is more listericidal than HNP-1 in DMEM. L. monocytogenes (7.2 x 105/ml) were incubated in DMEM at 37°C in the presence of the indicated concentrations of defensins or gentamicin (Gent.). (A and B) After 15 min (A) or 3 h (B), bacteria were plated on BHI agar to enumerate the CFU. (C and D) After 15 min, bacteria were washed twice and either immediately plated on BHI agar (C) or incubated at 2.4 x 104/ml for an additional 3 h in DMEM at 37°C, then plated on BHI agar (D). Results were expressed as the percentage of CFU relative to untreated bacteria at T = 15 min. In (B) and (D), the dashed lines indicate the CFU of the starting inoculum. In (A – D), * P < 0.05, ** P < 0.01.

We also assessed the antimicrobial activity of the defensins using the “virtual colony counting assay” (Fig. 2.5) (239). Bacteria were incubated with up to 35 µg/ml defensins in DMEM for 15 and 90 min at 37°C, washed, and incubated at 37°C in trypticase soy broth (TSB) for 16 h. The bacterial growth curves were determined by measuring the optical absorbance (OD 650) every 4 min. The time required to achieve an increase in 0.1 unit of OD 650 was used to calculate the imputed bacterial survival based upon the assumption that the defensins bactericidal activity was responsible for decreasing bacterial growth (Fig. 2.5C and D). While this assay cannot distinguish between bacteriostatic and bactericidal activities of the defensins, it shows that both

HNP-1 and RC-1, when incubated with bacteria in DMEM, retarded subsequent bacterial growth in TSB. Together, these results indicate that in the extracellular environment,

HNP-1 and RC-1 inhibition of L. monocytogenes multiplication is time-, concentration-, and culture medium-dependent. However, these results cannot predict how defensins behave within the phagosomal environment.

2.3.3 Defensins have little effect on phagocytosis of L. monocytogenes

To gain insight into the mechanism(s) used by the defensins to prevent L. monocytogenes growth in the macrophage, we determined which step of the intracellular infectious cycle was inhibited: phagocytosis, vacuolar escape or cytosolic multiplication.

To evaluate if the defensins affect phagocytosis, BMM were incubated with the defensins before (pre-incubation) or during (co-incubation) infection. BMM were infected for 15 min at 37°C (MOI 3), washed and fixed. Extracellular bacteria were labeled with an

Figure 2.5. HNP-1 and RC-1 inhibit L. monocytogenes growth in TSB. (A) A standard curve was calculated by incubating serial 10-fold dilutions of bacteria (2.5 x 106 CFU /ml - 2.5 x 102 CFU / ml) in DMEM. After 15 and 90 min at 37°C, the bacteria were washed, suspended in PBS/TBS (1:1), and OD at 650 nm was measured every 5 min for 16 h. (B) This standard curve was derived from the data in (A) and shows the time required by each inoculum to attain a ΔOD of 0.1 absorbance unit (corresponding to an increase of ~8.6 x 107 CFU/ml). The equation shown is from a least mean squares fit of the data points. (C and D). L. monocytogenes (2.5 x 105 CFU/ml) was incubated for 15 (C) or 90 (D) min with the indicated concentrations of HNP-1 or RC-1 in DMEM. The bacteria were washed, suspended in PBS/TBS (1:1), and OD 650 was measured every 4 min for 16-20 h. The results were expressed as percentage imputed survival (see materials and methods) and are the mean of two independent experiments, performed in triplicate. These experiments were performed by Pratikhya Pratikhya.

Alexa568-conjugated antibody, macrophages were permeabilized to label total bacteria

(extracellular + intracellular) with an Alexa488-conjugated antibody, and macrophage nuclei were labeled with DAPI (Fig. 2.6A). The phagocytic index was similar for wild type (100) and LLO-deficient bacteria (100 ± 27, 98 ± 2) (Fig. 2.6B and C, respectively).

Figure 2.6. Defensins do not affect phagocytosis of L. monocytogenes. BMM were incubated with the indicated concentrations of defensins before (pre- incubation) or during (co-incubation) infection. BMM were infected with L. monocytogenes (MOI 3 for 15 min at 37°C), washed and fixed. Extracellular bacteria were labeled with Alexa568-conjugated antibodies (red). Following permeabilization of the BMM, total bacteria were labeled with Alexa488-conjugated antibodies (green) and BMM nuclei were stained with DAPI (blue). (A) Representative images of BMM infected with L. monocytogenes without defensin treatment. The arrow indicates one of eight intracellular bacterium labeled as “total” but not “extracellular”. (B and C) The number of intracellular bacteria /BMM was automatically enumerated and the results were expressed as a phagocytic index relative to the control.

Concentrations of HNP-1 (20 µg/ml) and RC-1 (1 µg/ml) that inhibited L. monocytogenes intracellular multiplication (Figs. 2.1 and 2.3) had no effect on phagocytosis (Fig. 2.6B and C). We obtained similar results with RAW 264.7 macrophages (data not shown).

2.3.4 Defensins inhibit L. monocytogenes phagosomal escape

LLO is known to mediate bacterial escape from the phagosome within 15-30 min after phagocytosis (230). In the cytosol, the bacterial surface protein ActA activates actin polymerization as a means to induce bacterial motility (64). Therefore, the polymerization of F-actin around bacteria is commonly used as a read-out of L. monocytogenes cytosolic localization (29, 32, 240). We determined whether HNP-1 and

RC-1 could affect bacterial escape from the phagosome by enumerating bacteria that polymerize actin. BMM were incubated with the defensins before (pre-incubation) or during (co-incubation) infection. BMM were infected for 15 min at 37°C (MOI 3), washed and further incubated with gentamicin as performed in the cluster assay (Figs. 2.1 and 2.3). Cells were fixed, permeabilized, and bacteria and F-actin were labeled with fluorescent antibodies and phalloidins, respectively. As a negative control, we used LLO- deficient (Δhly) L. monocytogenes that are known to be trapped in the macrophage phagosome where they are unable to recruit actin (Fig. 2.7A, B, and D) and to multiply

(Fig. 2.7A, C and E) (13, 92). In contrast, wild type L. monocytogenes induces actin polymerization and replicates in the cytoplasm (Fig. 2.7). In BMM pre-incubated with

HNP-1, L. monocytogenes polymerization of actin was significantly reduced (Fig. 2.7B), 49

Figure 2.7. Defensins inhibit L. monocytogenes-induced actin polymerization. BMM were incubated with defensins (20 µg/ml HNP-1 or 1 µg/ml RC-1) before (15 min pre-incubation) or during (co-incubation) infection (MOI 3). After 15 min of infection at 37°C, cells were washed and either fixed (T=15 min) or further incubated with gentamicin at 37°C. After a total of 75, 135 or 180 min, the cells were fixed and permeabilized. Bacteria, F-actin, and BMM nuclei were fluorescently labeled. (A) Representative images of BMM infected with wt or Δhly L. monocytogenes in the absence of defensin at T=75 min. The arrow indicates a bacterium surrounded by a cloud of F-actin, the arrowhead an F-actin comet tail. (B and D) Percentage of intracellular bacteria that colocalize with F- actin clouds or comet tails. (B) RC-1 was not tested. (C and E) Total number of bacteria per BMM. (B – E) The asterisks apply to HNP-1, RC-1 and Δhly data points. * P < 0.05, ** P < 0.01.

50 as was bacterial proliferation (Fig. 2.7C). In contrast, L. monocytogenes was able to grow in BMM pre-incubated with RC-1 (Fig. 2.7C), in accordance with figure 2.3D. HNP-1 and RC-1 applied to the macrophage during phagocytosis (co-incubation) also markedly inhibited L. monocytogenes-induced actin polymerization (Fig. 2.7D) and intracellular proliferation (Fig. 2.7E). To rule out the possibility that bacteria were cytosolic but unable to polymerize actin, it was necessary to demonstrate that the bacteria were located in phagosomes. To accomplish this, we measured the colocalization of bacteria with lysosome-associated membrane protein-1 (LAMP-1). It is known that L. monocytogenes escapes from the phagosome before or soon after LAMP-1 recruitment, whereas LLO- deficient L. monocytogenes remains in a LAMP-1-positive phagosome (Fig. 2.8A) (230).

In macrophages pre-incubated with HNP-1, bacteria were confined in a LAMP-1-positive phagosome (Fig. 2.8B). HNP-1 and RC-1 applied to the macrophage during phagocytosis

(co-incubation) also induced bacterial retention in a LAMP-1-positive phagosome (Fig.

2.8C). Taken together, these results demonstrate that defensins do not affect the efficiency of phagocytosis but inhibit L. monocytogenes escape from the phagosome.

2.3.5 α- and θ-Defensins inhibit LLO-mediated macrophage membrane

perforation and bacterial release of LLO

We hypothesized that, in addition to their antimicrobial activity, the defensins could decrease vacuolar escape by inhibiting the release of LLO and/or its activity.

Recently, intestinal (HD-5) and neutrophil (HNP-1 to 3) α-defensins were shown to inhibit the hemolytic activity of the CDCs, including LLO (188). LLO hemolytic activity

Figure 2.8. Defensins increase L. monocytogenes colocalization with LAMP-1. BMM were incubated with defensins (20 µg/ml HNP-1 or 1 µg/ml RC-1) before (15 min pre-incubation) or during (co-incubation) infection (MOI 3). After 15 min of infection at 37°C, cells were washed and either fixed (T=15 min) or further incubated with gentamicin. After a total of 75, 135 or 180 min, the cells were fixed and permeabilized. Bacteria, LAMP-1, and BMM nuclei were fluorescently labeled. (A) Representative images of BMM infected with Δhly or wt L. monocytogenes in the absence or presence of RC-1 (co-incubation), T = 75 min. The arrows indicate LAMP- 1-positive bacteria and the arrowheads indicate bacteria that do not colocalize with LAMP-1. (B and C) Percentage of intracellular bacteria that colocalize with LAMP-1, * P < 0.05, ** P < 0.01. (B) RC-1 was not tested. (C) The asterisks apply to HNP-1 and RC-1 treatment.

52 was measured as a decrease in optical density (OD 700) of a solution of erythrocytes upon lysis. Here, we show the first evidence that the θ-defensin RC-1 efficiently inhibits

LLO-mediated hemolytic activity (Fig. 2.9A). As BMM and erythrocyte membranes may display different susceptibility to pore-forming toxins and the defensin’s anti-toxin activity, we tested whether α- and θ-defensins prevented the perforation of BMM membranes by LLO. BMM were incubated with LLO and ethidium homodimer (EthD) in the presence or absence of defensins. EthD is a membrane-impermeant dye that only penetrates permeabilized BMM and its fluorescence increases upon binding to nucleic acids (Fig. 2.9B). We measured the fluorescence of EthD as a read-out of BMM permeabilization by LLO and observed that HNP-1 and RC-1 markedly reduced LLO- induced BMM perforation (Fig. 2.9C). These results demonstrate that defensins inhibit

LLO-mediated perforation of macrophage membranes. We then tested if the defensins could inhibit LLO release from the bacteria. We used L. monocytogenes that overexpressed LLO from a plasmid encoding hly and its entire regulatory system, as it was difficult to detect the release of LLO from wild type bacteria within the time frame of the experiment. Bacteria were incubated for 30 and 60 min in DMEM at 37°C with 20

µg/ml HNP-1 or 0.5 µg/ml RC-1 and LLO was precipitated from the bacterial culture supernatants. We found that both HNP-1 and RC-1 markedly decreased the amount of

LLO recovered from the supernatants (Fig. 2.9D). These results were not due to bacterial killing as enumerating the CFU showed no loss of bacteria over time (data not shown).

These results highlight that HNP-1 and RC-1 efficiently inhibit LLO release and pore- forming activity. For HNP-1, which is less listericidal than RC-1 (Fig. 2.4), the anti-

Figure 2.9. Defensins inhibit LLO-mediated perforation of erythrocytes and macrophages and the release of LLO. (A) Erythrocytes were incubated with 0.075 nM LLO and RC-1 (µg/ml). Triton X-100 (TX-100) was used as a positive control and PBS as a negative control. OD 700 was acquired every 5 min. A representative experiment of three (performed in triplicate) is shown. (B and C) BMM were incubated at 37°C for 15 min with 4 µM EthD, 2.5 nM LLO and defensins in DMEM. Cells were washed, fixed and their nuclei were stained with DAPI. Phase-contrast and fluorescence images of nuclei (blue) and EthD (red) were acquired. (B) Representative images of BMM incubated in the absence or presence of LLO (no defensin was added). (C) The average fluorescence intensity of EthD within the nuclei was measured in each experimental condition. Results were expressed relative to BMM incubated with LLO in the absence of defensin. ** P < 0.01. (D) L. monocytogenes were incubated with HNP-1 (20 µg/ml) or RC-1 (0.5 µg/ml) in DMEM at 37°C. At the indicated times, samples were centrifuged and proteins were precipitated from the bacterial supernatants. LLO was detected by dot blot analysis. The samples corresponding to the 60 min time points were analyzed undiluted and at a 1/10 dilution. 54 toxin activity likely plays a critical role in preventing phagosomal escape.

2.3.6 The α-defensin HNP-1 is taken up by macrophages and colocalizes with

intracellular L. monocytogenes

For HNP-1 to prevent LLO release and/or activity within the phagosome, it is expected that this peptide interacts with the bacterium, the toxin or the phagosomal membrane, individually or collectively. In the co-incubation experiments (Fig. 2.1),

HNP-1 is in contact with the bacteria and the forming phagosome and is likely internalized with the bacteria; however, in the macrophage pre-incubation experiments

(Fig. 2.3B), we do not know if HNP-1 is taken-up by the macrophage and trafficked towards the phagosome and bacteria. We therefore determined whether HNP-1 is taken up by macrophages. Macrophages were incubated with HNP-1 for 15 min at 37°C, washed and fixed. HNP-1 was fluorescently labeled in non-permeabilized and permeabilized cells. As shown in permeabilized cells, the distribution pattern of HNP-1 delineates the plasma membrane, some internal vesicles and the cytoplasm (Fig. 2.10A).

The average fluorescence intensity of the cells was measured by quantitative fluorescence microscopy. HNP-1 was mainly localized within the cells, as higher levels of fluorescence were detected in permeabilized cells in comparison with non-permeabilized cells (Fig. 2.10B). To determine whether HNP-1 could be trafficked to the bacteria, we performed colocalization experiments in macrophages incubated with HNP-1 before (pre- incubation) or during (co-incubation) infection. As presented in Fig. 2.10C and D, about

25% intracellular bacteria colocalized with HNP-1. It is possible that our detection

Figure 2.10. HNP-1 is taken up by macrophages and colocalizes with L. monocytogenes. (A and B) BMM were incubated with the indicated concentrations of HNP-1 at 37°C in DMEM. After 15 min, cells were washed and fixed. HNP-1 was labeled with a secondary Alexa488-conjugated antibody (green) in permeabilized and non-permeabilized macrophages. (A) Representative image of HNP-1 (20 µg/ml) fluorescence in permeabilized macrophages. (B) MFI of Alexa488-labeled HNP-1 per BMM. (C and D) BMM were incubated with 20 µg/ml HNP-1 before (15 min pre-incubation) or during (co- incubation) infection (MOI 3). After 15 min of infection at 37°C, cells were washed and either fixed (T=15 min) or further incubated with gentamicin. After a total of 75 or 180 min, the cells were fixed and permeabilized. Bacteria were labeled with an Alexa568- conjugated antibody (red) and HNP-1 was labeled with an Alexa488-conjugated antibody (green). (C) Representative images of BMM incubated with HNP-1 during infection (co- incubated), T=180 min. The arrow indicates a bacterium that colocalizes with HNP-1. (D) Percentage of intracellular bacteria that colocalize with HNP-1. * P < 0.05, ** P < 0.01.

56 method underestimated the amount of intracellular HNP-1 and HNP-1 colocalization with bacteria. Indeed, HNP-1 association with the host cell structures, such as membranes, may prevent antibody accessibility to HNP-1 or mask epitope(s) that the antibody recognizes. Nonetheless, these results support the idea that HNP-1 is taken up by macrophages and targeted to the phagosome and bacteria.

2.4 Discussion

This study demonstrates that the cooperation between macrophages and the human α-defensin HNP-1 or the humanized θ-defensin RC-1 prevents the intracellular multiplication of L. monocytogenes in macrophages. The defensins did not affect the efficiency of phagocytosis but inhibited subsequent L. monocytogenes vacuolar escape and intracellular proliferation. Similar to our observations, α-defensins (HNP-1 to 3 and

HD-5) also block the escape of human papillomaviruses from endocytic vesicles but not virion binding or internalization (241).

Incubating the macrophages with HNP-1 before or during infection efficiently prevented bacterial proliferation. We propose that the HNP-1/macrophage cooperation operates via HNP-1-mediated inhibition of LLO release and activity in the phagosome and/or the combined anti-listerial activity of HNP-1 and the macrophage. The α-defensins are multifunctional peptides that can exert immunomodulatory functions; therefore, we do not exclude that HNP-1 also stimulates the macrophage bactericidal activity (204, 206,

226). The θ-defensin RC-1, whose gene arose by the mutation of pre-existant α-defensin genes (158), was included in this study because it is a proposed therapeutic agent due to

57 its potent anti-bacterial and anti-viral activities (216). Intriguingly, RC-1 and HNP-1 appear to inhibit L. monocytogenes intracellular proliferation though different mechanisms. While HNP-1 interaction with the macrophage is important for control of L. monocytogenes growth, pre-incubating macrophages with RC-1 before infection had no effect. Indeed, bacterial exposure to RC-1 was required to inhibit subsequent intracellular proliferation. We found that RC-1 is a more potent anti-listerial peptide than HNP-1, in accordance with the reported anti-listerial activities of HNP-1 (40-60 µg/ml) and RC-1

(<1 µg/ml) (242-244). Therefore, it is possible that RC-1 acts primarily through its anti- listerial activity. Nonetheless, RC-1 also inhibits LLO activity and release from the bacteria.

While RC-1 ability to inhibit LLO hemolytic activity is a novel finding, several α- and θ-defensins are known to inactivate bacterial toxins. HNP-1 to 3 inhibit anthrax lethal factor, protecting murine macrophages and mice from otherwise lethal doses of toxin (183). RC-1 inhibits anthrax lethal factor in addition to killing germinating spores and vegetative bacilli (184). The diphtheria toxin, pseudomonas exotoxins and C. difficile toxin B are inhibited by HNP-1 to 3 (185, 186). Most recently, HNPs and HD-5 were shown to inhibit the hemolytic activity of the CDCs, including LLO (188). Finally, our study demonstrates for the first time that HNP-1 and RC-1 inhibit macrophage membrane perforation by LLO. How α- and θ-defensins exert such a broad-spectrum anti-toxin activity, inhibiting toxins that are unrelated in structure and function, remains to be determined. Defensins anti-toxin activity likely involves specific peptide/toxin interactions (185). For example, HNP-1 neutralizes C. difficile toxin B but not toxins that

58 are similar in structure such as the C. difficile toxin A or C. sordellii lethal toxin (186).

Conversely, the cathelicidin LL-37 that has a similar net charge to the HNPs has no effect on lethal factor, diphtheria toxin or clostridial toxins (183, 185, 186). It is thought that defensins bind to the catalytic site, preventing the toxin’s activity or association with the substrate (185). Among the toxins inhibited by the defensins, the CDCs are the only non- catalytic toxins and they act by forming pores in host membranes. It was proposed that α- defensins bind to multiple sites on the CDCs, thereby preventing perforation of erythrocyte membranes (188). In addition to inhibiting LLO-mediated host membrane perforation, we found that HNP-1 and RC-1 efficiently prevented LLO release from the bacteria. Whether the defensins prevent LLO synthesis, secretion, or both, remains to be determined. Similar to our observations, it has been suggested that α-defensins prevent adenovirus escape from endosomes by inhibiting the release of protein VI, which is responsible for the lysis of the endosomes (245).

Neutrophils are found at infectious foci during listeriosis and are important effectors of the innate immune defense against L. monocytogenes at the early stages of infection, as demonstrated in animal models (66, 101, 246). In humans, neutropenic patients display higher susceptibility to bacterial infections, including listeriosis (96). As evidenced in this study, neutrophil HNPs prevent intracellular multiplication of L. monocytogenes in macrophages. Macrophages may acquire neutrophil antimicrobial molecules from the extracellular environment as previously proposed (191, 204, 205,

213, 247) and in accordance with our observation that HNP-1 is taken up by macrophages and colocalizes with intracellular bacteria. Macrophages may also acquire

59 antimicrobial molecules by phagocytosis of apoptotic neutrophils (213). Indeed, macrophage phagocytosis of apoptotic neutrophils led to decreased viability of intracellular M. tuberculosis and HNP-1 was found to localize to the macrophage phagosomes following uptake of apoptotic neutrophils (213). The α-defensins likely impact several stages of L. monocytogenes infection. The α-defensin HD-5, produced by

Paneth cells in the small intestine, and HNPs may limit L. monocytogenes crossing of the intestinal barrier. At later stages of infection, defensins may limit infection in the liver and other organs. In support of this, treating HepG2 cells (a human hepatocyte cell line) with HNP-1 prior to or during infection reduces L. monocytogenes internalization into hepatocytes ~4 to 5-fold (data not shown). LLO is required for efficient L. monocytogenes entry into hepatocytes (35), therefore, it seems likely that HNP-1 inhibits

L. monocytogenes internalization into hepatocytes by inhibiting LLO activity. Finally, the defensins anti-toxin activity may decrease LLO-mediated cell injury in infected tissues.

α-Defensins are not encoded by human cells of the monocyte/macrophage lineage and neither are they encoded by murine neutrophils (248, 249). Therefore, while murine macrophages afford useful models, mice are imperfect experimental surrogates for humans in models of infection in which neutrophil defensin-macrophage cooperation is significant. In humans, the α-defensins HNP-1 to 3 are present in the blood and tissues where neutrophils circulate, migrate and release their granule contents. Plasma concentrations of HNPs range from ~50 ng/ml in healthy individuals to multi-µg/ml in septic patients and local concentrations of HNPs in inflamed or infected tissues are likely to be much higher (250). Therefore, the concentrations of HNP-1 used in the present

60 study are within physiological range. It is likely that bacteria face various concentrations of defensins in vivo, such that the combination of the defensins anti-toxin, microbicidal and immunomodulatory activities ensure a robust innate immune defense.

In conclusion, we demonstrate for the first time that α- and θ-defensins enable macrophages to control L. monocytogenes intracellular proliferation. We speculate that the cooperation between defensins, and possibly other AMPs, and macrophages plays an important role in the innate immune defenses against L. monocytogenes. The understanding of the multiple anti-infective strategies of AMPs is expected to help with developing new antibiotics. We have studied two classes of defensins: α-defensins produced by human neutrophils and a circular θ-defensin that is a proposed therapeutic agent. The potent anti-bacterial and toxin-neutralizing activities of RC-1 described here provide additional support for developing θ-defensins as therapeutics.

Chapter 3. The pore-forming toxin listeriolysin O is degraded by neutrophil proteases and fails to protect L. monocytogenes against intracellular killing

3.1 Introduction

L. monocytogenes is a Gram-positive bacterial pathogen responsible for the

foodborne disease listeriosis (1, 6), discussed in section 1.1.1. The L. monocytogenes

intracellular life cycle in professional phagocytes and in cells that are normally non-

phagocytic, such as epithelial cells, is essential for disease progression (1, 8-10).

Following internalization of L. monocytogenes into a host cell, the bacterium escapes

from its internalization vesicle within 15-30 min to reside in the cytosol, where it

proliferates. Cytosolic bacteria then undergo F-actin-based motility to form extracellular

protrusions that are internalized by adjacent cells. This leads to the ingestion of the

bacterium in a vesicle made of two membranes, which are later disrupted by the

bacterium to repeat the intracellular life cycle (251). Disruption of the endocytic vesicles

containing L. monocytogenes is central to intracellular survival and requires the secreted

pore-forming toxin listeriolysin O (LLO) (13, 33, 230). Indeed, LLO-deficient L.

monocytogenes strains remain trapped in the endocytic vesicle, are unable to divide

intracellularly, and are completely non-virulent in vivo (11).

LLO was the first identified virulence factor of L. monocytogenes (11, 12, 47) and is a member of the largest family of bacterial pore-forming toxins, the cholesterol- dependent cytolysins (CDCs) (14, 16, 21), discussed in section 1.1.3. The mechanisms by which LLO pores halt phagosomal maturation and facilitate phagosome disruption remain incompletely understood, and are discussion in section 1.1.3.1. It was previously thought that LLO could only perforate membranes at low pH in the context of acidifying phagosomes; however, a large body of evidence established that LLO is also a potent pore-forming toxin when it is released in the extracellular compartment before bacterial internalization (35, 37). Host cell plasma membrane perforation by extracellular LLO is also likely to play an important role in regulating the bacterial intracellular life cycle. As recently shown, plasma membrane perforation by extracellular LLO is sufficient to induce L. monocytogenes internalization into epithelial cell lines (35). Extracellular LLO also controls post-translational modifications, mitochondrial remodeling, and histone modifications in epithelial cell lines; all of these activities importantly regulate host cell invasion in vitro (42, 43, 45). How host cell perforation by extracellular LLO has such diverse effects on target cells is not well understood. It is known that host cells can recover from perforation by moderate concentrations of pore-forming toxins (35, 49,

252). Fluxes of ions and small molecules across toxin pores elicit multiple signaling pathways that control cell repair and alert the immune system to cell damage and pathogen attack (48). It is reasonable to propose that those ionic fluxes similarly play an important role in the activities of extracellular LLO (44, 253-255). Indeed, an increase in intracellular Ca2+ upon neutrophil exposure to CDCs was shown to trigger the exocytosis

63 of their numerous granules (256-260). Neutrophils are known to exert anti-listerial activity, but it was unclear if and how LLO i) affects the biology of neutrophils and ii) confers any survival advantage to L. monocytogenes in those cells.

The objective of the present study was to ascertain the role of LLO in the interplay between L. monocytogenes and neutrophils. Our data establish that LLO fails to protect L. monocytogenes from intracellular killing in neutrophils. Furthermore, L. monocytogenes virulence factors, such as LLO, initiate exocytosis of secondary and primary neutrophil granules at the plasma membrane before closure of the phagosome.

We propose that early degranulation leads to the release of LLO-neutralizing molecules that protect the neutrophil plasma membrane from damage caused by extracellular LLO and prevents perforation of the phagosome by intracellular LLO.

3.2 Materials and Methods

3.2.1 Recombinant proteins and reagents

Recombinant six-histidine-tagged LLO, pre-pore locked LLO (LLOpL), and cysteine-free LLO (LLO C484A) were purified from Escherichia coli BL21(DE3) harboring the pET29b plasmid coding for those LLO derivatives (35). Pneumolysin

(PLY) and anthrolysin O (ALO) were purified from E. coli BL21(DE3) harboring the pQE30 plasmid encoding PLY (a gift from Dr. R.K. Tweten, University of Oklahoma

Health Sciences Center, Oklahoma City, OK) or pET15 plasmid encoding ALO (a gift from Dr. P.C. Hanna, University of Michigan Medical School, Ann Arbor, MI), respectively. All toxins were purified as described previously and stored at -80oC (35). 64

The following antibodies were used: anti-L. monocytogenes (GeneTex), anti-LLO

(Abcam), horse radish peroxidase (HRP)-conjugated secondary antibodies (Cell

Signaling), FITC-conjugated anti-CD63 (clone 46-4-5) from Ancell, and FITC- conjugated anti-CD66b (clone CLB-B13.9) and FITC-conjugated isotype control (clone

MOPC-21) from Accurate Chemical and Scientific Corporation. Fc Receptor Blocking

Solution (Human TruStain FcX) was purchased from Biolegend. ProLong Gold antifade containing DAPI, secondary Alexa Fluor antibodies, Dulbecco’s modified Eagle’s medium (DMEM), Hank’s Balanced Salt Solution (HBSS) without Ca2+ and Mg2+

(HBSS-), and HBSS with 1.26 mM Ca2+ and 0.9 mM Mg2+ (HBSS+) were from Life

Technologies. Brain heart infusion (BHI) and fibronectin were from BD Biosciences. The protease inhibitor cocktails (PI) ± EDTA were from Roche. The human matrix metalloproteases (MMPs) pro-MMP-8 and active recombinant MMP-9 were from

Calbiochem (Millipore). Fetal bovine serum (FBS), paraformaldehyde (PFA), latrunculin

A, N-Formyl-methionyl-leucyl-phenylalanine (fMLP), and the lactate dehydrogenase

(LDH) detection kit were purchased from Sigma-Aldrich. Heparinized and thrombin- treated Vacutainers were from Fisher and Polymorphprep from Axis-Shield.

3.2.2 Isolation of human neutrophils and serum

All studies were approved by the Institutional Review Board at The Ohio State

University. Human peripheral blood was collected from healthy donors into heparinized and thrombin-treated Vacutainer blood collection tubes for neutrophil and serum isolation, respectively. Donor serum (DS) was isolated at room temperature, then heat-

65 inactivated for 30 min at 56°C and stored at 4°C until needed. Neutrophils were isolated at room temperature by a one-step density gradient centrifugation on Polymorphprep as previously described (261). Residual erythrocytes were lysed by hypotonic shock then neutrophils were washed in HBSS- and suspended in HBSS- or HBSS+ (±10% autologous

DS) immediately before use. For all experiments with adherent neutrophils, 4.5x105 neutrophils (9x105/ml) were plated in 24-well tissue culture treated dishes coated with 30

µg fibronectin and pre-incubated for 10 min at 37°C to allow the neutrophils to attach.

3.2.3 Macrophage and bacterial cell cultures

The RAW264.7 macrophage-like cell line (TIB-71, ATCC) was cultured in

DMEM supplemented with 100 Units Penicillin/Streptomycin and 10% heat inactivated-

FBS (HI-FBS). Bone marrow-derived macrophages (BMM) were obtained from tibias and femurs of 5- to 6-week-old C3HOuJ mice (Jackson Laboratories) as described in

Chapter 2 (236). BMM were incubated in DMEM supplemented with 10 units

Penicillin/Streptomycin and 10% HI-FBS. RAW264.7 cells and BMM were seeded at

1x105 cells per well, in 24-well tissue culture treated dishes coated with 30 µg fibronectin, incubated for 24 h, then washed before use.

Bacterial strains used include L. innocua (ATCC 33090) and the following L. monocytogenes strains (which were used unless indicated otherwise): wild type (wt,

DP10403S), isogenic LLO over-expressing (LLO+, DP10403S harboring the pAM401 plasmid encoding LLO; described in Chapter 2), and isogenic LLO-deficient (DP10403S

Δhly, DP-L2161). L. monocytogenes wt (L028) and the isogenic LLO-deficient (L028 66 hly::Tn917) strains were used in Fig. 3.12. DP10403S and DP-L2161 were provided by

Dr. D. A. Portnoy (University of California, Berkeley, CA). L. monocytogenes L028 and hly::Tn917 strains were provided by Dr. P. Cossart (Pasteur Institute, Paris, France).

LLO+ secreted 5.5 ± 1.3 fold more LLO than wt L. monocytogenes, as determined with hemolytic activity assays. Bacteria were cultured overnight in BHI at 37°C with shaking, diluted 1/20 in BHI and grown at 37°C until optical density at 600nm (OD 600) = 0.7-

0.8.

3.2.4 L. monocytogenes association with, phagocytosis by, and replication in

neutrophils

L. monocytogenes were added at multiplicity of infection (MOI) 1 or 10 to adherent neutrophils in HBSS+ (± 10% DS), and cell culture plates were centrifuged (230 g) at room temperature (RT) for 5 min. After 30 min at 37°C, cells were washed, fixed, and labeled or treated for 30 min with 15 µg/ml gentamicin, washed, and further incubated for 4 h at 37°C in 10% DS/HBSS+. Cells were fixed with 3.5% PFA in PBS, pH 7.4 for 15 min at RT, washed with 0.1 M glycine in PBS, and blocked for 45 min in

10% FBS/0.1 M glycine. Extracellular bacteria were labeled with anti-L. monocytogenes and secondary Alexa Fluor-conjugated antibodies. Cells were permeabilized with 0.2%

Triton X-100 in PBS for 2 min, then intracellular + extracellular (total) bacteria were labeled using the same primary antibody and a different secondary antibody. Antibodies were diluted in 10% FBS/PBS and incubated with cells for 30 min at RT. After labeling, coverslips were mounted with ProLong Gold antifade containing DAPI. Forty sets of

67 fluorescence and phase contrast images were automatically acquired for each experimental condition using the 20X or 100X objective. Neutrophils, extracellular, and total bacteria were enumerated. The percentage of intracellular bacteria [(total bacteria – extracellular bacteria) / total bacteria X 100] was calculated as described previously

(238); 100-300 bacteria were counted in each experimental condition.

3.2.5 L. monocytogenes association with, and phagocytosis by, macrophages

BMM or RAW 264.7 cells were infected with wt (L028) and isogenic LLO- deficient (hly::Tn917) L. monocytogenes at MOI 1 (BMM) or MOI 3 (RAW264.7) in

DMEM. MOI was adjusted such that L. monocytogenes association with RAW264.7 and

BMM would be similar. After 30 min at 37°C, cells were washed, fixed, and total and extracellular L. monocytogenes and nuclei were labeled as described above. L. monocytogenes association with macrophages and the percentage of intracellular L. monocytogenes was calculated as described above.

3.2.6 L. monocytogenes viability following incubation with neutrophils or

macrophages

L. monocytogenes at MOI 1 and 10 in 10% DS/HBSS+ were added to adherent neutrophils or macrophages and cell culture plates were centrifuged (230 g) for 5 min at

RT. After 1.5, 3, and 5 h at 37°C, mammalian cells were lysed by addition of Triton X-

100 (0.2% final concentration) to enumerate the total number of viable L. monocytogenes

(Lm + PMNs / RAW, Total). Alternatively, 15 µg/ml gentamicin was added for the last hour of incubation and cells were washed before the addition of Triton X-100 to only enumerate intracellular viability of L. monocytogenes (Lm + PMNs / RAW, Intra.). As a control, L. monocytogenes were incubated in the absence of neutrophils (Lm). Cell lysates were diluted and plated on BHI agar to enumerate colony forming units (CFU). Results were expressed as fold increase in L. monocytogenes growth relative to the inoculum.

3.2.7 Macrophage perforation following incubation with L. monocytogenes

RAW264.7 macrophages were incubated with L. monocytogenes at MOI 1 and 10 in 10% Serum/DMEM for 4 h, then 20 µM propidium iodide (PrI) was added and cells were incubated for 1 h (T = 5 h). Cells were fixed and the mean fluorescence intensity

(MFI) of PrI in the nuclear regions was determined by quantitative fluorescence microscopy, as we previously described (Section 2.2.9) (35).

3.2.8 CD63 colocalization with L. monocytogenes

L. monocytogenes were added to adherent neutrophils at MOI 4 (non- permeabilized cells) or MOI 2 (permeabilized cells) in 10% DS/HBSS+ and centrifuged

(230 g) for 1 min at RT. The MOIs were adjusted such that individual bacteria could be clearly seen for colocalization determination. Cells were incubated for 1 min at 37°C, washed and incubated for a total of 3 and 10 min at 37°C in 10% DS/HBSS+. Cell were washed, fixed, and blocked as described above. L. monocytogenes were fluorescently

69 labeled as described above and CD63 was labeled with FITC-conjugated anti-CD63 antibodies in permeabilized (0.2% saponin for 30 min) or non-permeabilized cells.

Fluorescence and phase contrast images were acquired with a 100X objective.

3.2.9 Kinetics of neutrophil degranulation following challenge with bacteria

Measured by quantitative microscopy: L. monocytogenes were added to adherent neutrophils at MOI 4 and 40 in 10% DS/HBSS+ and centrifuged (230 g) for 1 min at RT.

Cells were incubated for 1 min at 37°C, washed and incubated for a total of 3, 5, 10, and

30 min at 37°C in 10% DS/HBSS+. Non-permeabilized cells were washed, fixed, blocked, and labeled as described above. Fluorescence and phase contrast images were acquired with a 40X objective. Images were background corrected, then cell perimeters were traced and the MFI of cells was determined. Results were expressed as the MFI of

CD63 labeled cells.

Measured by flow cytometry: To measure degranulation in response to challenge with L. monocytogenes and L. innocua, suspended neutrophils (2x107/ml) were incubated

+ in 10% DS/HBSS at 37°C for 10 min. Bacteria were added to neutrophils at MOI 10 and incubated at 37°C with rotation to maximize contact with bacteria. After 30 min, neutrophils were transferred to ice and maintained at 4°C during subsequent steps.

Neutrophils were centrifuged (17,000 g, 5 min), washed to remove unbound bacteria, then labeled as described below. To measure degranulation in response to challenge with

LLO, suspended neutrophils (9x105/ml) were incubated at 37°C for 10 min in HBSS+.

LLO (0.5-5 nM) or LLOpL (50 nM) was added and cells were incubated at 37°C in

HBSS+. After 15 min, neutrophils were transferred to ice and maintained at 4°C during subsequent steps. After fixation, cells were incubated with Fc Receptor Blocking

Solution in 0.1 M glycine/ 5% FBS/ 0.05% NaN3/ PBS, pH 7.4 for 30 min. Neutrophils were then incubated with fluorescent anti-CD63, anti-CD66b, or isotype control antibodies for 30 min. Cells were washed and stored in 0.3% PFA/ 0.05% NaN3/ PBS.

Fluorescence intensities were measured with a FACSCalibur flow cytometer (BD

Biosciences) and data analysis was performed with FlowJo software (Tree Star, Inc.).

Gating, based on forward and side-scatter, was used to remove cell debris from the analysis. Average background intensity (based on cells labeled with the isotype control antibody) was subtracted from the MFI of CD63 or CD66b labeled cells. Results were expressed as the corrected MFI relative to untreated cells.

3.2.10 Wide field and Confocal Microscopes

Images were acquired with a motorized inverted epi-fluorescence microscope

(Axio Observer D1 from Zeiss) equipped with 20X Plan Neofluar (N.A. = 0.5), 40 X

Plan Neofluar (N.A. = 1.3), and 100 X Plan Apo (N.A. = 1.4) objectives. The camera

(back-illuminated, frame-transfer EMCCD Cascade II 512) was from Photometrics. The microscope equipment was controlled by MetaMorph imaging software. Labeling of intracellular CD63 was visualized with a spinning-disk confocal microscope (UltraVIEW

ERS from PerkinElmer) equipped with a 100 X objective lens (N.A. = 1.4). The camera

(cooled CCD ORCA-AG) was from Hamamatsu. Images were excited with 488-, 568-, and 647-nm argon ion lasers and z-slices were acquired every 0.05 - 0.1 µm.

Representative deconvolved images (Volocity software from PerkinElmer) are from a single z-slice.

3.2.11 LDH release by cells challenged with LLO

LLO (5-500 nM in HBSS ± Ca2+) was added to adherent neutrophils and cells were incubated at 37°C for 15 min (Fig. 3.6A). Cells were transferred to ice, then supernatants were collected, centrifuged (5 min, 230 g, 4°C) and the LDH assay was performed according to the manufacturer’s instructions. Maximum LDH release was determined by treating cells with 1% Triton X-100 for 15 min. Percentage of LDH release was calculated as: [(LDH released from LLO treated cells)/(LDH released from

Triton X-100 treated cells) x 100]. To inhibit neutrophil degranulation prior to challenge with LLO (Fig. 3.6B), adherent neutrophils were incubated with 0, 1.5, and 2 µg/ml

TAT-Inhibitor or TAT-GST (negative control) in HBSS+ for 10 min. The TAT-Inhibitor mixture consisted of 0.75 µg/ml of TAT-SNAP-23 and TAT-syntaxin-4 (1.5 µg/ml TAT-

Inhibitor) or 1 µg/ml of TAT-SNAP-23 and TAT-syntaxin-4 (2 µg/ml TAT-Inhibitor).

After incubation with the TAT-conjugates, cells were challenged with 5 nM LLO in

HBSS+ for 15 min at 37°C and supernatants were collected to measure LDH release. For macrophage protection experiments (Fig. 3.7A and B), RAW264.7 macrophages were incubated with neutrophil degranulation products (NDP), control supernatant (CS), chemical controls (fMLP + latrunculin A) or protein controls (BSA) and 5 nM LLO in

HBSS+ for 15 min at 37°C, then supernatants were collected to measure LDH release.

Chemicals (fMLP + latrunculin A) and BSA controls were used at concentrations found 72 in comparable volumes of NDP. For example, a solution of 24% NDP contains 72 nM fMLP + 0.24 µM latrunculin A and 82 µg/ml protein. Thus, 24% fMLP + latrunculin A or 24% BSA contain 72 nM fMLP + 0.24 µM latrunculin A or 82 µg/ml BSA, respectively.

3.2.12 Collection of Neutrophil Degranulation Products

Suspended neutrophils (2x107/ml in HBSS+) were incubated with 1 µM latrunculin A for 30 min, followed by a 5 min incubation with 300 nM fMLP.

Neutrophils were centrifuged (21,000 g) at 4°C for 5 min and the supernatant (neutrophil degranulation products, NDP) was collected and stored at -20°C. Non-induced, control supernatant (CS) was collected from neutrophils incubated for 35 min in the absence of latrunculin A and fMLP. The mean protein concentration of NDP and CS were 341 ± 60 and 129 ± 48 µg/ml, respectively, as determined with a Bio-Rad DC protein assay.

3.2.13 Hemolysis assay

Erythrocytes (2.8x107/ ml), CDCs (LLO, PLY, or ALO), NDP (3-24% v/v), CS

(24% v/v), chemical controls (fMLP + latrunculin A; 24% v/v), or protein controls (BSA,

24% v/v) were added in a final volume of 210 µl /well, in a 96-well plate on ice. CDCs were diluted to 0.24 nM (LLO, PLY, and ALO) or 2.4 nM (LLO C484A). Chemicals

(fMLP + latrunculin A) and BSA controls were used at concentrations found in comparable volumes of NDP. When indicated, 1.2 mM EDTA or protease inhibitor 73 cocktail with EDTA (PIEDTA; inhibits serine, cysteine, and metalloproteases) or without

EDTA (PI; inhibits serine and cysteine proteases) were added. PI ± EDTA was used at

1X according to the manufacturer’s instructions. Plates were incubated at 37°C in a

PowerWavex340 spectrophotometer (Bio-Tek) and OD 700 was measured every min for

30 min. Data were shown as kinetic curves (Figs. 3.7C and D and 3.8) or percent hemolysis at 30 min (Table 3.2). Erythrocytes were incubated with 0.05% Triton X-100 to calculate 100% lysis.

3.2.14 LLO treatment with NDP followed by Western blotting

LLO (38 nM) was incubated with heat-inactivated NDP (75°C for 30 min; HI-

NDP; 88% v/v) or NDP (11-88% v/v). Alternatively, LLO (38 nM) and NDP (88% v/v) were incubated in the presence or absence of protease inhibitor cocktails (PI±EDTA) or 2 mM EDTA. As a control, LLO was either incubated alone or in the presence of PIEDTA or

EDTA. After 1 and 5 min at 37°C, reduced Laemmli’s sample buffer was added and samples were boiled. The equivalent of 25 ng of LLO was loaded in each well of a 10%

SDS-PAGE gel, followed by western blotting analysis using anti-LLO and HRP- conjugated antibodies.

3.2.15 LLO treatment with MMPs followed by Western blotting

MMP-8 activation was achieved by incubating MMP-8 with 10 mM APMA in

MMP assay buffer (50 mM Tris-HCl pH 7.6, 200 mM NaCl, 5 mM CaCl2, 20 µM ZnCl,

74 and 0.05% Brij L23) for 1.5 h at 37°C. MMP-9 was purchased already active. LLO (38 nM) was incubated with active MMP-8 (100 nM), MMP-9 (100 nM), or a mixture of

MMP-8 and MMP-9 (100 nM each) in the presence or absence of 5 mM EDTA in MMP assay buffer for 5 min at 37°C. Reduced Laemmli’s sample buffer was added and samples were boiled. The equivalent of 8.5 ng of LLO was loaded in each well of a 10%

SDS-PAGE gel, followed by western blotting analysis using anti-LLO and HRP- conjugated antibodies.

3.2.16 Statistics

At least three independent experiments were performed in duplicate for each assay, unless indicated otherwise. Neutrophils, NDP, and CS from at least three different donors were used. Data were expressed as mean ± SEM. P-values were calculated using a standard two-tailed t-test and considered significant if lower than 0.05, * P<0.05 and **

P<0.01.

3.3 Results

3.3.1 LLO does not confer a survival advantage to L. monocytogenes phagocytosed

by neutrophils

We first determined if LLO could affect L. monocytogenes association with or phagocytosis by human neutrophils. Neutrophils were incubated with wild type (wt) or

LLO-deficient (Δhly) L. monocytogenes (MOI 1 and 10) for 30 min in the presence or

75 absence of heat-inactivated autologous serum. Cells were washed, chemically fixed, fluorescently labeled, and imaged by fluorescence microscopy to enumerate bacterial association with and phagocytosis by neutrophils (Fig. 3.1A). L. monocytogenes association and phagocytosis were significantly increased in the presence of serum, regardless of LLO production and multiplicity of infection (Fig. 3.1B and C). LLO did not affect L. monocytogenes association with neutrophils in any experimental conditions

(Fig. 3.1B). However, LLO significantly increased the efficiency of phagocytosis at MOI

1 (Fig. 3.1C). This effect was not observed at MOI 10 or in the presence of serum, likely because phagocytic efficiency was already optimal under those conditions, masking the role of LLO.

We next determined if LLO could facilitate L. monocytogenes proliferation in neutrophils, as previously observed in macrophages (Fig 2.1A and B) (13). Neutrophils were infected for 30 min with wt or Δhly L. monocytogenes in the presence or absence of heat-inactivated serum. Extracellular bacteria were killed by a brief treatment with the cell impermeant antibiotic gentamicin, and cells were incubated for a total of 5 h. The total number of bacteria associated with neutrophils was enumerated by fluorescence microscopy. As presented in Table 3.1, the number of L. monocytogenes per neutrophil was constant over time in all conditions tested, indicating that L. monocytogenes is unable to proliferate in neutrophils, regardless of LLO expression.

Such immunofluorescence analysis does not provide information about the viability of intracellular bacteria. To determine bacterial viability, wt and Δhly L. monocytogenes (MOI 1 and 10) were incubated for up to 5 h in three distinct

Figure 3.1. L. monocytogenes association with and phagocytosis by human neutrophils. Neutrophils were incubated with wt or Δhly L. monocytogenes (Lm) at MOI 1 or 10 in HBSS+ ± 10% donor serum. After 30 min at 37°C, cells were washed and fixed. Extracellular, total bacteria and neutrophil nuclei were fluorescently labeled. (A) Representative images of neutrophils infected with wt Lm at MOI 10 in HBSS+ ± 10% Serum were acquired with a 20X (two upper panels) and a 100X objective (bottom panel). (B) Lm association with neutrophils was calculated as the total number of bacteria per neutrophil (Lm/PMN), and (C) the percentage of intracellular Lm was calculated as [(intracellular bacteria) / (total bacteria) x 100]. Results are the mean ± SEM of at least three independent experiments, performed in duplicate, * P < 0.05; ** P < 0.01. Colleen Nackerman performed some of these experiments.

experimental conditions (Fig. 3.2). First, bacteria were incubated in cell culture medium without neutrophils (Lm). Second, bacteria were co-incubated with neutrophils and total bacterial viability was measured (Lm + PMNs, Total). Third, bacteria were co-incubated with neutrophils and extracellular bacteria were killed by a 1 h treatment with gentamicin before enumeration of viable intracellular bacteria (Lm + PMNs, Intra). Results clearly established that in the presence of neutrophils, most bacteria (about 70% of the inoculum) were killed within 1.5 h (Fig. 3.2A and B). Viable bacteria were extracellular as no bacteria were recovered from gentamicin-treated cells, regardless of LLO production. To confirm that LLO does not confer a survival advantage to L. monocytogenes, we compared survival of wt and LLO over-expressing (LLO+) L. monocytogenes incubated with neutrophils for 3 h. Both bacterial strains were killed to a similar extent and did not

Table 3.1. L. monocytogenes is unable to proliferate in neutrophils, regardless of LLO expression.

Neutrophils were incubated with wt or Δhly L. monocytogenes (Lm) at MOI 10 in HBSS+ ± 10% Donor Serum. After 0.5 h, cells were fixed or incubated with 15 µg/ml gentamicin for 0.5 h, washed and incubated for 4 h (T = 5 h) in HBSS+/10% Serum then fixed. Total Lm were labeled with fluorescent antibodies and neutrophil nuclei were labeled with DAPI. The total number of Lm per neutrophil (Lm / PMN) was determined. Results are the mean ± SEM of at least three independent experiments, performed in duplicate. aLm / PMN was significantly different in the presence vs absence of serum for both Lm strains and both times, P < 0.02.

78 grow intracellularly (Fig. 3.2C). For comparison, we repeated those assays with a murine macrophage-like cell line (RAW264.7 cells). As expected, L. monocytogenes grew intracellularly in a LLO-dependent fashion (Fig. 3.3A). Indeed, wt L. monocytogenes grew so well in the presence of macrophages that after 5 h at MOI 10, macrophages were extensively damaged (Fig. 3.3B). In conclusion, LLO, which is important for L. monocytogenes replication in multiple cell types, including macrophages, does not protect L. monocytogenes from intracellular killing in neutrophils.

Figure 3.2. Neutrophils kill L. monocytogenes, regardless of LLO expression. L. monocytogenes (Lm: wt, Δhly, and LLO+) were incubated alone (Lm) or with neutrophils (Lm + PMNs) in 10% DS/HBSS+ at MOI 1 (A) and MOI 10 (B and C). At the indicated times, Triton X-100 was added and cell lysates were plated to enumerate CFU. This measured the viability of Lm in the absence of neutrophils (Lm) and the viability of extracellular plus intracellular Lm in the presence of neutrophils (Lm + PMNs, Total). To measure viability of intracellular bacteria (Lm + PMNs, Intra.), cells were incubated with gentamicin for the last hour of incubation and then washed before lysis and CFU enumeration. (A-C) At least three independent experiments were performed in triplicate. Results are expressed relative to the inoculum. Asterisks indicate a statistically significant difference between Lm viability in the absence (Lm) and presence (Lm + PMNs, Total) of neutrophils for all Lm strains at all time points, P < 0.05. Colleen Nackerman generated the data shown in A and B.

3.3.2 L. monocytogenes and LLO activate rapid neutrophil degranulation

In neutrophils, granules rapidly fuse with the phagosome (133). We hypothesized that L. monocytogenes enhances neutrophil degranulation and that neutrophil granules

Figure 3.3. L. monocytogenes replication in macrophages is LLO-dependent. (A) L. monocytogenes (Lm: wt and Δhly) were incubated alone (Lm) or with RAW264.7 macrophages (Lm + RAW) in 10% Serum/DMEM at MOI 1. At the indicated times, Triton X-100 was added and cell lysates were plated to enumerate CFU. This measured the viability of Lm in the absence of macrophages (Lm) and the viability of extracellular plus intracellular Lm in the presence of macrophages (Lm + RAW, Total). To measure viability of intracellular bacteria (Lm + RAW, Intra.), cells were incubated with gentamicin for the last hour of incubation and then washed before lysis and CFU enumeration. Results are expressed relative to the inoculum and are the mean ± SEM of at least three independent experiments, performed in triplicate. Asterisks indicate a statistically significant difference between viable Δhly Lm in the absence (Lm) or presence of macrophages (Lm + RAW, Total), P < 0.05. (B) RAW264.7 macrophages were incubated with wt or Δhly Lm at MOI 1 and 10 in 10% Serum/DMEM for 4 h, then 20 µM propidium iodide (PrI) was added and cells were incubated for 1 h (T = 5 h). Cells were fixed and the mean fluorescence intensity (MFI) of PrI in the nuclear regions was determined by quantitative fluorescence microscopy, as we previously described (35). Colleen Nackerman performed some of these experiments.

80 contain LLO-neutralizing agents that prevent L. monocytogenes intracellular survival. To test this idea, we analyzed granule exocytosis in neutrophils incubated with L. monocytogenes, the non-pathogenic Listeria innocua, and recombinant LLO. We first visualized and quantified primary granule exocytosis by fluorescence microscopy.

Neutrophils were incubated at 37°C for 3, 5, 10, and 30 min with L. monocytogenes. We fluorescently labeled L. monocytogenes and CD63, a membrane-associated primary granule marker, in non-permeabilized and permeabilized neutrophils. After 3 min and at the later times, L. monocytogenes induced localized primary granule exocytosis at the bacterial interaction site (Fig. 3.4A). Fusion of primary granules with formed phagosomes was also visualized in permeabilized cells at the different times (Fig. 3.4A).

Quantitative fluorescence microscopy analysis performed over the entire cell surface, which tends to attenuate the impact of local degranulation, confirmed that L. monocytogenes induces substantial degranulation at the plasma membrane (Fig. 3.4B).

As expected, degranulation increased with time and MOI, indicating that L. monocytogenes activates neutrophils in a dose-dependent fashion (Fig. 3.4B).

We next compared the extent of neutrophil degranulation induced by L. monocytogenes and L. innocua, which does not express virulence factors (262), by flow cytometry. Cells were incubated for 30 min with bacteria at MOI 10 then labeled with anti-CD63 and anti-CD66b fluorescent antibodies to report exocytosis of both primary and secondary granules (Fig. 3.5A). L. monocytogenes was a more potent inducer of degranulation of both types of granules. This result is in accordance with a previous study reporting that L. monocytogenes induces neutrophil degranulation in a LLO-dependent

81 fashion (256). Indeed, we found that recombinant LLO added exogenously to neutrophils induced dose-dependent degranulation of primary and secondary granules at the plasma membrane (Fig. 3.5B). This induction was pore-dependent as the LLO variant LLOpL that cannot form pores was unable to induce degranulation. This was not unexpected, as

Figure 3.4. L. monocytogenes induces local degranulation of primary granules. Neutrophils were incubated with L. monocytogenes (Lm) for 1 min, washed and incubated in 10% DS/HBSS+. Cells were fixed and Lm and CD63 were fluorescently labeled in non-permeabilized and permeabilized cells to distinguish granule fusion with the plasma membrane from fusion with formed phagosomes. (A) Representative images of extracellular and intracellular Lm associated with CD63 at 3 and 10 min. For permeabilized cells, extracellular and total Lm were fluorescently labeled, only intracellular Lm are shown. (B) Mean fluorescence intensity (MFI) of non- permeabilized neutrophils was determined with fluorescence microscopy. Results are expressed as the MFI of CD63 labeling ± SEM, SEM depicts variation of labeling within the experiment. Data from a representative experiment of two are shown.

LLO pores are well-known to allow the influx of extracellular Ca2+ inside host cells (254,

255) and a rise in intracellular Ca2+ is responsible for degranulation of all granule subsets in neutrophils (263). In conclusion, L. monocytogenes virulence factors such as LLO facilitate degranulation at the plasma membrane even before completion of the phagosome.

3.3.3 Degranulation protects neutrophils from perforation by LLO

In the hypothesis that degranulation products inhibit LLO, we expect that granule

Figure 3.5. L. monocytogenes and LLO induce neutrophil degranulation. (A) Suspended neutrophils were incubated for 30 min with L. monocytogenes or L. innocua at MOI 10, washed, and labeled with FITC-conjugated anti-CD66b, anti-CD63, or isotype control antibodies. (B) Suspended neutrophils were incubated for 15 min with LLO (0.5-5 nM) or LLOpL (50 nM), fixed, and labeled with FITC-conjugated anti- CD66b, anti-CD63, or isotype control antibodies. (A and B) Neutrophil mean fluorescence intensity (MFI) was measured by flow cytometry. Data are expressed as MFI relative to untreated cells. Data from a representative experiment of three are shown.

83 fusion with the plasma membrane would protect the membrane from further LLO- induced damages. Therefore, we analyzed the role of degranulation in maintaining neutrophil plasma membrane integrity despite attack by LLO. Neutrophils were incubated with increasing concentrations of recombinant LLO (5 to 500 nM) in the presence or absence (to inhibit degranulation) of extracellular Ca2+, for 15 min. Cell culture supernatants were collected to measure the release of lactate dehydrogenase

(LDH). LDH is a cytosolic enzyme, thus the presence of LDH in cell supernatant reflects cell damage. We observed that in the presence of extracellular Ca2+, neutrophils were highly resistant to LLO, as 200 nM LLO was required to induce 50% LDH release (Fig.

3.6A). All cells that we have analyzed in our laboratory, including macrophage and epithelial cell lines, were totally lysed by 10 nM LLO (data not shown). In Ca2+-free medium, which prevents neutrophil degranulation, only 5 nM LLO led to 50% LDH release by neutrophils.

To further demonstrate that degranulation is required for neutrophil resistance to perforation by LLO, neutrophils were pre-incubated with the TAT fusion proteins TAT-

SNAP-23 and TAT-syntaxin-4 to inhibit degranulation of all granule subsets (264, 265).

These fusion proteins contain the HIV transactivator of transcription (TAT) cell- penetrating sequence and the N-terminal SNARE domain of synaptosome-associated protein-23 (SNAP-23) or the SNARE domain of syntaxin-4. As shown in figure 3.6B, treatment with the TAT-Inhibitors increased cell damage caused by LLO to similar levels as observed in Ca2+-free medium (43.3 ± 1.2% and 50.2 ± 3.5% LDH release, respectively), whereas treatment with the TAT negative control (TAT-GST) did not (12.2

± 1.7% LDH release). This result confirms that the Ca2+-dependent protection process observed in figure 3.6A corresponds to the exocytosis of granules.

Figure 3.6. Neutrophil degranulation is required for resistance to LLO- mediated perforation. (A) Neutrophils were incubated with 5-500 nM LLO or cell culture medium only for 15 min in HBSS ± Ca2+. Asterisks indicate a statistically significant difference between neutrophils incubated in HBSS with and without Ca2+, P < 0.05. (B) Neutrophils were incubated with 0, 1.5, or 2 µg/ml TAT-Inhibitors (Inhib, equal mixture of TAT-SNAP-23 and TAT-syntaxin-4) or control TAT-GST (GST) for 10 min in HBSS+ to inhibit degranulation. Cells were then incubated with media alone (Control, C) or 5 nM LLO in HBSS+ for 15 min, P < 0.01 (A and B) LDH release into the cell culture supernatant was measured. Data are expressed as the mean percentage of maximal LDH release ± SEM of at least three independent experiments, performed in duplicate.

3.3.4 Degranulation products directly inhibit LLO activity

We next determined if neutrophil degranulation products could directly inhibit

LLO activity. Neutrophils were treated with latrunculin A and fMLP to induce exocytosis of all granule subsets (266), and the resultant supernatant containing neutrophil degranulation products (NDP) was collected. Alternatively, control supernatant (CS) was collected from untreated neutrophils. We determined if NDP could protect RAW264.7 macrophages and erythrocytes from perforation by LLO. As shown in figure 3.7A-C,

NDP, but not CS, protected macrophages and erythrocytes from perforation by LLO, in a dose-dependent manner. We verified that latrunculin A and fMLP alone do not affect

LLO activity (Fig. 3.7B and D). We also verified that LLO inhibition by NDP was not the result of an increase in protein in the cell culture medium (Fig. 3.7B and D).

We determined if NDP could also protect host cells from other cholesterol- dependent cytolysins, such as anthrolysin O (ALO) or pneumolysin (PLY) produced by

Bacillus anthracis and Streptococcus pneumoniae, respectively. Erythrocytes were incubated with ALO and PLY along with CS or NDP. Similar to LLO, ALO and PLY activity was abolished by NDP (Fig. 3.8). Collectively, these data demonstrate that neutrophil degranulation products are protective against CDC-mediated perforation of host cells.

3.3.5 A protease activity is responsible for LLO inhibition

We further analyzed the molecular basis of LLO inhibition by NDP. Oxidation of the unique cysteine located in the undecapeptide sequence is known to block LLO 86

Figure 3.7. Neutrophil degranulation products inhibit LLO-mediated perforation of macrophages and erythrocytes. (A and B) RAW264.7 macrophages were incubated with media alone (Control, C), 5 nM LLO, NDP (6-24%), CS (6-24%), fMLP + latrunculin A (fMLP + Lat; 6-24%), and BSA (24%) for 15 min at 37°C. LDH release into the cell culture supernatant was measured. Data are expressed as the mean percentage of maximal LDH release ± SEM of at least three independent experiments, performed in duplicate, P < 0.01. (C and D) Erythrocytes were incubated with PBS, 0.24 nM LLO, NDP (3-24%), CS (24%), fMLP + Lat (24%), or BSA (24%) for 30 min at 37°C. PBS and 0.05% Triton X-100 are negative and positive controls, respectively. Data are expressed as mean OD 700 ± SEM of at least three independent experiments, performed in duplicate. (B and D) Chemical (fMLP + Lat) and BSA controls were used at concentrations found in comparable volumes of NDP. For example, a solution of 24% NDP contains 72 nM fMLP + 0.24 µM Lat and 82 µg/ml protein. Thus, 24% fMLP + Lat or 24% BSA contain 72 nM fMLP + 0.24 µM Lat or 82 µg/ml BSA, respectively.

87 activity (12). As indicated in Table 3.2, we found that NDP also inhibited a cysteine-free

LLO variant, ruling out an oxidative inhibition mechanism. The inhibitory role of NDP was lost after heating at 75°C for 30 min, indicating that proteins were responsible for

LLO inhibition. We then tested a cocktail of protease inhibitors that blocks metalloproteases, cysteine proteases, and serine proteases (PIEDTA) as well as a second cocktail that did not contain the metalloprotease inhibitor EDTA but contained identical amounts of all the other inhibitors (PI). Only the EDTA-containing inhibitor prevented

Figure 3.8. Neutrophil degranulation products inhibit other CDCs. Erythrocytes were incubated with 0.24 nM ALO (A) or PLY (B) and NDP (6-24%) or CS (24%) for 30 min at 37°C. OD 700 was measured every min for 30 min. Results are from a representative experiment of two, performed in duplicate. Stephen Vadia performed these experiments.

88 the activity of NDP. Furthermore, the addition of EDTA alone abolished NDP activity, suggesting the involvement of a metalloprotease. Importantly, addition of the inhibitors without NDP did not affect LLO hemolytic activity (Table 3.2).

To test the hypothesis that NDP proteolytically degrade LLO, we incubated LLO with increasing concentrations of NDP for 1 and 5 min at 37°C and assayed for LLO degradation by western blotting. LLO was degraded within 1 min as we observed a clear decrease in full length LLO together with the appearance of lower molecular weight LLO fragments. By 5 min, LLO was degraded to the point that it was almost undetectable (Fig.

3.9A). PIEDTA abolished NDP-mediated LLO degradation, whereas PI did not. EDTA alone was sufficient to inhibit NDP-mediated LLO degradation (Fig. 3.9B). Importantly,

EDTA alone did not lead to proteolysis of LLO (Fig. 3.9B). In accordance with the

Table 3.2. LLO inhibition by neutrophil degranulation products involves heat- labile factor(s) and divalent cations.

Erythrocytes were incubated with 0.24 nM LLO or 2.4 nM LLO C484A, 6% NDP, protease inhibitor cocktail that contains 1.2 mM EDTA (PIEDTA) or not (PI), or 1.2 mM EDTA. Results are the mean percent hemolysis at 30 min ± SEM of at least three independent experiments, performed in duplicate. 100% lysis was determined by incubating erythrocytes with 0.05% Triton X-100. Asterisks indicate a significant difference between LLO and LLO + the indicated treatments, P < 0.01. Stephen Vadia performed these experiments.

89 functional assays in Figure 3.7 and Table 3.2, NDP-mediated LLO degradation was concentration dependent, mediated by heat-labile component(s), and divalent cation-

Figure 3.9. Neutrophil degranulation products degrade LLO. (A) LLO (38 nM) was incubated with various amounts of NDP (88, 44, and 11%), or heat-inactivated NDP (HI, 88%) for 1 and 5 min at 37°C. (B) LLO (38 nM) was incubated with NDP (88%) and protease inhibitor cocktail that contains 2.6 mM EDTA (PIEDTA) or not (PI), or 2 mM EDTA for 5 min at 37°C. (A and B) Laemmli’s sample buffer was added and the equivalent of 25 ng LLO were loaded in each well of a 10% SDS-PAGE for western blot analysis. Non-degraded LLO is indicated by an arrow. Note the higher concentration of toxin required for detection with this assay compared to the functional hemolytic assay, the LLO/NDP ratio is higher for this assay. Stephen Vadia generated some of these blots.

90 dependent. Importantly, the LLO/NDP ratio was much higher in the experimental conditions used to analyze LLO degradation than in the functional assay.

These results indicate that a matrix metalloprotease (MMP) may be responsible for NDP-mediated LLO degradation. Human neutrophils produce three MMPs, MMP-8

(collagenase), MMP-9 (gelatinase), and MMP-25 (leukolysin or MT6-MMP). The first two are soluble, whereas MMP-25 is membrane-associated (267, 268). We determined if the soluble MMPs, MMP-8 and MMP-9, degrade LLO. Incubation of LLO with MMP-8, but not MMP-9, resulted in degradation of LLO, as observed by the appearance of lower molecular weight LLO fragments (Fig. 3.10). Co-incubating MMP-8 with MMP-9 did not appear to alter MMP-8 mediated cleavage of LLO, however, incubating this mixture with EDTA did inhibit MMP activity, as expected (Fig. 3.10). Of note, some of the LLO degradation fragments were different than what was observed when LLO was incubated

Figure 3.10. MMP-8 degrades LLO. LLO (38 nM) was incubated with 100 nM MMP-8, 100 nM MMP-9, 100 nM MMP-8 + 100 nM MMP-9, or 100 nM MMP-8 + 100 nM MMP-9 + 5 mM EDTA for 5 min at 37°C. Laemmli’s sample buffer was added and the equivalent of 8.5 ng LLO were loaded in each well of a 10% SDS-PAGE gel for western blot analysis. Non-degraded LLO is indicated by an arrow. A representative blot from two experiments is shown.

91 with NDP (Fig. 3.9). This is not unexpected as neutrophil granules contain serine proteases and other components that may alter MMP activity (134). We propose that initial cleavage of LLO is mediated by the cation-dependent MMP-8 and that other proteases produced by neutrophils further degrade LLO fragments.

3.4 Discussion

This study demonstrates that L. monocytogenes is efficiently phagocytosed and killed intracellularly by human neutrophils regardless of LLO expression. Rapid fusion of intracellular granules with the plasma membrane at the bacterial entry site releases diverse antimicrobial molecules, including proteases that degrade LLO during the formation of the phagosome. These data not only provide a molecular basis to account for the lack of LLO activity in the neutrophil phagosome, but also show that degranulation is critical for maintaining neutrophil plasma membrane integrity despite perforation by LLO

(Fig 3.11). This anti-toxin response is observed with other cholesterol-dependent cytolysins (anthrolysin O and pneumolysin) and may apply to additional families of pore- forming toxins.

Immunity against L. monocytogenes requires the innate and adaptive immune responses. Although sterilizing immunity requires CD8+ and CD4+ T-cell responses (84,

110, 116, 117), innate immune cells, including neutrophils and monocytes, are important for initial control of L. monocytogenes infection (96-98). Neutrophils are recruited to the various organs infected with L. monocytogenes including the intestines, liver, spleen, placenta, and central nervous system (97, 98, 269-272). Several studies have previously

92 concluded that neutrophils play a critical role in the control of L. monocytogenes infection in mice (100-102). However, a caveat in those studies is that they used a neutrophil-depleting antibody that was later shown to target both monocytes (Ly6C+) and neutrophils (Ly6G+). Using anti-Ly6G antibodies to specifically deplete neutrophils, recent studies confirmed that neutrophils are protective in regards to L. monocytogenes infection (97, 99, 116), as discussed in Section 1.2.1. Supporting the role of neutrophils in efficient killing of L. monocytogenes in vivo, analysis of infected murine liver tissue

Figure 3.11. Model of L. monocytogenes interaction with neutrophils. 1) Upon secretion by L. monocytogenes, LLO monomers bind to host cell membranes and oligomerize into pre-pore complexes. 2) LLO pre-pore complexes undergo conformational changes to form pores, which induce Ca2+ influx. 3) An increase in intracellular Ca2+ levels leads to rapid degranulation at the phagocytic cup. Neutrophil granular components, like the MMPs, rapidly degrade extracellular LLO, irreversibly inhibiting its activity. Degranulation also occurs into the phagosome, which would lead to degradation of intracellular LLO. These actions inhibit LLO-mediated escape, trapping L. monocytogenes in a bactericidal phagosome. 4) Degranulation is linked to increased endocytosis, endocytosis of membrane containing LLO pores likely contributes to the neutrophils resistance to LLO-mediated membrane damage.

93 showed that L. monocytogenes was degraded within neutrophil phagosomes and proliferated in the cytosol of hepatocytes (273). Furthermore, in zebrafish, neutrophils sequestered L. monocytogenes within bactericidal phagosomes, whereas bacteria proliferated in macrophages (274). In light of those various studies, it is reasonable to conclude that neutrophils efficiently ingest and kill L. monocytogenes in vivo, as we observed in vitro. In addition to their role in eliminating extracellular bacteria, neutrophils may cooperate with infected cells, such as macrophages and epithelial cells, to limit their infection. For example, neutrophils facilitate bacterial clearance by releasing cytokines and inflammatory mediators (98), and antimicrobial molecules that can directly target extracellular bacteria or be taken up by other cells to reinforce their antimicrobial response (224, 275, 276). Similarly, phagocytosis of apoptotic neutrophils enhances macrophage resistance to infection (213). Finally, it has been proposed that neutrophils can directly lyse infected cells (277-279).

In vitro studies revealed that human neutrophils kill a large portion of L. monocytogenes via both oxidative-dependent and -independent processes (68, 280-283).

However, those studies did not determine if L. monocytogenes could survive within neutrophils and if LLO could promote bacterial intracellular survival. Also, a recent manuscript argued that L. monocytogenes bacilli could resist killing by neutrophils (231).

Therefore, we compared association, phagocytosis, and intracellular survival of wild type and LLO-deficient L. monocytogenes by fluorescence microscopy and gentamicin survival assays. In accordance with previous reports, we observed that opsonization of L. monocytogenes with heat-stable serum components greatly enhances bacterial

94 phagocytosis (67-69, 284). Importantly, we report that in the presence or absence of serum, L. monocytogenes is killed intracellularly, regardless of LLO expression.

Furthermore, LLO, which facilitates L. monocytogenes intracellular survival in many cell types, appears to disadvantage the bacterium in neutrophils. Indeed, at a low multiplicity of infection, LLO increases bacterial phagocytosis. We previously reported that LLO can induce L. monocytogenes uptake by non-phagocytic cells (35). The role of LLO in bacterial uptake is not observed in all cell types, as LLO does not increase bacterial uptake by murine macrophages (25) (Fig. 3.12). The molecular basis for LLO-induced bacterial uptake is currently under investigation in our laboratory and it is unknown if the same mechanism is involved in neutrophils and non-phagocytic cells. It is tempting to propose that in neutrophils, LLO increases phagocytosis by facilitating degranulation, described in further detail below. Host cell perforation by LLO and other pore-forming toxins elicits Ca2+ influx, which then activates exocytosis of lysosomes or other intracellular granules (211, 285, 286). Neutrophils contain secretory vesicles and three classes of granules, which are secreted at the plasma membrane in a hierarchical fashion as the cytosolic Ca2+ concentration increases (134, 287). Intriguingly, release of primary granules into the phagosome is not dependent on intracellular Ca2+ concentrations (287,

288). The most readily mobilized vesicles or granules to the plasma membrane are secretory vesicles, followed by the tertiary/gelatinase granules, secondary/specific granules, and the primary/azurophilic granules. Tertiary granules and secretory vesicles contain phagocytic and adhesion receptors as well as enzymes for breakdown of the extracellular matrix. Secondary granules potentiate neutrophil responses by providing

95 diverse adhesion molecules, as well as releasing opsonins, proteases, and antimicrobial molecules. Primary granules contain serine proteases and anti-microbial molecules, mainly defensins (Fig 1.4) (132, 134, 275). Therefore, LLO-induced granule exocytosis likely potentiates diverse neutrophil functions, including phagocytosis, the production of reactive oxygen species and the release of defensins and other toxic molecules. In addition to LLO, other L. monocytogenes virulence factors such as the secreted

Figure 3.12. LLO does not enhance phagocytosis by macrophages. Macrophages were infected with L. monocytogenes (Lm) strain L028 (wt or the isogenic LLO-deficient hly::Tn917) at MOI 1 (A, BMM) or MOI 3 (B, RAW264.7). MOI was adjusted such that Lm association with RAW264.7 and BMM would be similar. After 30 min at 37°C, cells were washed, fixed, and total and extracellular Lm and nuclei were fluorescently labeled. Lm association with macrophages and the percentage of intracellular Lm was calculated as described for Fig. 3.1. Results are the mean ± SEM of at least three independent experiments, performed in duplicate.

96 phospholipases induce neutrophil degranulation (256). In accordance with this, L. innocua, which does not express virulence factors, is a weaker activator of degranulation than L. monocytogenes. We observed that the efficiency of L. monocytogenes phagocytosis increases with the MOI. This is likely due to the increase in degranulation in response to higher concentrations of L. monocytogenes virulence factors.

In cultured macrophages, LLO induces phagosomal escape within 20 min of phagocytosis (26). Our data show that granule fusion with the forming phagosome is initiated at the plasma membrane within 3 minutes (earlier time points were not studied).

Therefore, the rapid release of LLO-neutralizing molecules should quickly inhibit LLO in the forming phagosome. Although our data favor the notion that LLO is detrimental to L. monocytogenes in neutrophils, it should be noted that during severe infection, high concentrations of LLO and other virulence factors may have deleterious effects on the host by causing excessive degranulation, leading to tissue damage.

Three non-mutually exclusive hypotheses could explain LLO inhibition by neutrophil degranulation products. First, defensins are known to exert LLO-neutralizing activity (157, 275, 289). Second, LLO is a substrate for MMP-8, and likely additional neutrophil proteases. Finally, reactive oxygen species produced by neutrophils may oxidize LLO. Indeed, LLO contains a unique cysteine located within a conserved undecapeptide sequence in the C-terminus that acts as a redox switch. Once oxidized, the cysteine prevents the formation of the pore complex (12); however, a cysteine-free LLO variant (LLO C484A) is still able to form pores (290). We found that neutrophil degranulation products inhibited LLO C484A-mediated hemolysis, indicating that

97 inhibition of LLO is independent of the oxidation state of the toxin. We then observed that inhibition of LLO-mediated hemolysis and LLO degradation by neutrophil products require heat-labile component(s) and divalent cations. Together, these data suggest that a metalloprotease (MMP) is responsible for LLO degradation. Neutrophils produce two soluble MMPs, MMP-8 and MMP-9 (267, 268). We show that MMP-8 degrades LLO.

Of note, incubating LLO with MMP-8 or neutrophil degranulation products yielded slightly different LLO degradation fragments. We propose that initial cleavage of LLO is mediated by MMP-8, and that other neutrophil proteases degrade LLO fragments.

Similar to our observations, previous studies have reported that lysosomal cathepsin-D degrades LLO and that L. monocytogenes intracellular survival was increased in cathepsin-D-deficient cells in vitro. Also, cathepsin-D-deficient mice were more susceptible to L. monocytogenes infection (291, 292). Therefore, enzymatic degradation of LLO in the extracellular environment and in the phagosome is an important and general mechanism for the anti-listerial innate defense.

We propose that neutrophils efficiently kill L. monocytogenes because of their redundant anti-bacterial and anti-toxin mechanisms. Indeed, in addition to proteases, the human neutrophil α-defensins exert potent-anti-LLO activities, both inhibiting activity and secretion of this toxin (275). The present study focused on LLO degradation by neutrophil metalloproteases, but it will be important to dissect how defensins, in conjunction with metalloproteases, affect LLO activity. These molecules may act independently of each other or may cooperate for more efficient proteolysis of the toxin.

Pore-forming toxins are commonly produced by bacterial and eukaryotic pathogens. These toxins display a large array of activities from host cell killing to more subtle subversion of host cell functions at sublytic concentrations. Neutrophils are specialized in the uptake and elimination of pathogens and are therefore exposed to those toxins. It is tempting to speculate that neutrophils have evolved to counteract perforation by pore-forming toxins. All eukaryotic cells undergo membrane repair in response to mechanical and biochemical disruption of their plasma membrane (293, 294). Those repair mechanisms are activated by the influx of extracellular Ca2+ consecutive to membrane damage. In response to pore-forming toxins, it was recently shown that epithelial cells undergo Ca2+-dependent exocytosis of lysosomes leading to the release of the lysosomal enzyme acid sphingomyelinase (ASM) (49, 50). In this model, ceramide synthesis by ASM changes the curvature of the lipid bilayer, facilitating endocytosis of the plasma membrane, thereby removing toxin pores (51). The requirement for extracellular Ca2+ and degranulation to maintain neutrophil membrane integrity in the presence of LLO (Fig. 3.6) is reminiscent of the membrane repair process of other cells.

Therefore, neutrophil degranulation may exert a protective effect via activation of the membrane repair response. Indeed, previous reports have established a link between degranulation and increased fluid phase endocytosis, which likely facilitates removal of

LLO pores (295, 296). Also, our data show that degranulation products directly inhibit

LLO. Indeed, erythrocytes which cannot undergo membrane repair are protected from

LLO by neutrophil degranulation products. Therefore, neutrophils likely display two redundant mechanisms to maintain plasma membrane integrity in response to the pore-

99 forming toxin LLO: direct degradation of the toxin combined with increased membrane turnover to remove pore complexes from the plasma membrane (Figs. 3.9-3.11).

Reciprocally, it can be hypothesized that other cell types also directly inactivate toxins, through release of lysosomal cathepsin-D or MMP-8.

We observed that neutrophil degranulation products protect erythrocytes from lysis by the CDCs anthrolysin O, produced by Bacillus anthracis, and pneumolysin, produced by Streptococcus pneumonia. Therefore, it is tempting to speculate that the neutrophil’s response to LLO characterized in this study also applies to other CDCs. In accordance with this idea, neutrophils efficiently kill numerous CDC-producing bacteria

(297-301). It would be of considerable interest to study neutrophil degranulation and membrane repair mechanisms in response to various pore-forming toxins as well as to evaluate the anti-toxin activity of neutrophil degranulation products. This is especially interesting in regards to pore-forming toxins produced by pathogens known to resist killing by neutrophils, such as Staphylococcus aureus (302, 303).

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Chapter 4. Co-incubation of macrophages and neutrophils limits L. monocytogenes replication in macrophages

4.1 Introduction

L. monocytogenes is a foodborne intracellular pathogen and the causative agent of

listeriosis (1, 6), discussed in section 1.1.1. L. monocytogenes intracellular life cycle is

critical for infection. After internalization, listeriolysin O (LLO), the main virulence

factor of L. monocytogenes, disrupts the phagosomal membrane to mediate L.

monocytogenes escape from the phagosome. This releases the bacterium into the cytosol,

where it replicates and polymerizes the host cell actin cytoskeleton to mediate cell-to-cell

spreading, thereby propagating infection without being exposed to the extracellular

environment (1, 8, 10).

L. monocytogenes is able to replicate in macrophages, but not in neutrophils

(Chapters 2 and 3) (13, 91, 92). Neutrophils and macrophages are located in close

proximity at infectious foci (89) and cooperation between these cells has been proposed

to enhance macrophage bactericidal activities (207-209) and may be involved in

controlling infections of intracellular pathogens such as M. tuberculosis and L.

pneumophila (204-206). However, this has not been studied in regards to L.

monocytogenes infection. Neutrophil macrophage cooperation can occur through

101 neutrophil mediated macrophage activation and/or macrophage internalization of neutrophils, neutrophil granules, or neutrophil antimicrobial molecules (210, 211), as discussed in section 1.4. For example, neutrophil granule products enhance macrophage

ROS production and phagocytosis (200, 206). We have found that exogenously added neutrophil defensins enable macrophages to prevent L. monocytogenes intracellular replication (Figs. 2.1 and 2.2).

In the present study, we determined if neutrophils could cooperate with macrophages to prevent L. monocytogenes growth in macrophages. We co-cultured neutrophils and macrophages during infection and observed that macrophage phagocytosis of L. monocytogenes was not altered in the presence of neutrophils. L. monocytogenes replication in macrophages was markedly decreased in the presence of neutrophils, in a concentration dependent manner. These results indicate that during L. monocytogenes infection, when macrophages and neutrophils are located at infectious foci together, neutrophils likely enable macrophages to limit L. monocytogenes intracellular growth. Thus, neutrophils may help control L. monocytogenes infection through direct killing of bacteria (Chapter 3) and by cooperating with other cells to prevent L. monocytogenes replication in those cells. Indeed, this has already been proposed in regards to hepatocytes, it is suspected that neutrophils lyse infected hepatocytes to limit infection (277-279).

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4.2 Materials and Methods

4.2.1 Reagents

Rabbit anti-L. monocytogenes antibodies were purchased from GeneTex, ProLong

Gold antifade containing DAPI, secondary Alexa Fluor antibodies, HBSS without Ca2+ and Mg2+ (HBSS-), HBSS with 1.26 mM Ca2+ and 0.9 mM Mg2+ (HBSS+), and RPMI

1640 Medium were from Life Technologies. Brain heart infusion (BHI) and fibronectin were from BD Biosciences. Fetal bovine serum (FBS), paraformaldehyde (PFA), and phorbol myristate acetate (PMA) were purchased from Sigma-Aldrich. Heparinized

Vacutainers were from Fisher and Polymorphprep from Axis-Shield. Polycarbonate transwell filters (0.4 µm) were from Corning.

4.2.2 Isolation of human neutrophils

All studies were approved by the Institutional Review Board at The Ohio State

University. Human peripheral blood was collected from healthy donors into heparinized

Vacutainer blood collection tubes. Neutrophils were isolated at room temperature by a one-step density gradient centrifugation on Polymorphprep as previously described

(Section 3.2.2) (261). Residual erythrocytes were lysed by hypotonic shock then neutrophils were washed in HBSS- and suspended in RPMI immediately before use.

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4.2.3 Macrophage and L. monocytogenes culture

Thp1 macrophages (ATCC TIB-202) were cultured in RPMI supplemented with

100 U penicillin/streptomycin and 10% HI-FBS (RPMI+). Thp1 cells in RPMI+ were seeded at 0.7x105 cell per well in a 24-well tissue culture treated dish coated with 30 µg fibronectin. Thp1 cells were incubated with 160 nM PMA for 48 h to induce differentiation into macrophages (304, 305) then washed 3 times before infection. L. monocytogenes wild type (wt, DP10403S) and isogenic LLO-deficient (Δhly, DP-L2161) strains were cultured overnight at 37°C in BHI, diluted 20 times in BHI, and cultured until OD 600 = 0.8. Bacteria were washed three times with HBSS+ and diluted to multiplicity of infection (MOI) 1 in RPMI.

4.2.4 Infection assays

Neutrophils (1.4 and 2.1x105 / well, corresponding to 2:1 and 3:1 ratio of neutrophil:Thp1 cell) in RPMI were added to Thp1 cells and incubated for 10 min at

37°C to allow the neutrophils to attach. Thp1 cells and neutrophils were infected with L. monocytogenes at MOI 1 in RPMI and centrifuged 5 min (230 g, RT). For experiments using transwell filters, Thp1 cells were infected with L. monocytogenes at MOI 1 in the absence of neutrophils, then centrifuged (5 min, 230 g, RT). Transwell filters (0.4 µm) were added to wells containing infected macrophages, then neutrophils (2.1 and 7x105 / well, corresponding to 3:1 and 10:1 neutrophil:Thp1 cell) and bacteria at MOI 1 were added above the filter. Bacteria did not pass through the 0.4 µm filter during the time frame of the experiment. For all experiments, infected cells were incubated for 30 min at 104

37°C. Cells were washed and fixed or incubated for 30 min with 15 µg/ml gentamicin in

10% HI-FBS/RPMI, washed, and incubated for an additional 4 h in 10% HI-FBS/RPMI

(5 h final) before being washed and fixed.

4.2.5 Fluorescence labeling

Cells were fixed with 4% PFA in PBS, pH 7.4 for 15 min, washed 3 times with

0.1 M glycine in PBS, then incubated for 1 h in blocking solution (10% HI-FBS/ 0.1 M glycine / PBS). After blocking, extracellular bacteria were incubated with rabbit anti-L. monocytogenes antibodies diluted in 10% HI-FBS in PBS for 30 min at RT, followed by labeling with Alexa488 secondary antibodies. Cells were permeabilized with 0.2% Triton

X-100 in PBS for 5 min at RT, then total bacteria (intracellular + extracellular) were labeled using the same primary antibody and a secondary antibody conjugated to

Alexa568. After labeling, coverslips were mounted with ProLong Gold antifade containing DAPI.

4.2.6 Image acquisition and analysis

Images were acquired with a motorized inverted epi-fluorescence microscope

(Axio Observer D1 from Zeiss) equipped with a 20X Plan Neofluar (N.A. = 0.5) objective. The camera (back-illuminated, frame-transfer EMCCD Cascade II 512) was from Photometrics. The microscope equipment was controlled by MetaMorph imaging software. Forty sets of fluorescence and phase contrast images were acquired for each

105 experimental condition using the 20X objective. Macrophages, neutrophils, extracellular, and total bacteria were enumerated. Thp1 cells and neutrophils were differentiated based on nuclei morphology. The percentage of intracellular bacteria [(total bacteria – extracellular bacteria) / total bacteria X 100] was calculated as described previously

(238); 100-200 bacteria were counted in each experimental condition. The percentage of

L. monocytogenes growth was calculated as: [(total intracellular bacteria at 5 h) / (total intracellular bacteria at 30 min) X 100]. L. monocytogenes growth in macrophages incubated without neutrophils was set to 100%.

4.2.7 Statistics

At least two independent experiments were performed in duplicate for each assay.

Data were expressed as mean ± SEM. P-values were calculated using a standard two- tailed t-test and considered significant if lower than 0.05, * P<0.05, ** P<0.01.

4.3 Results

4.3.1 Co-culturing macrophages with neutrophils does not significantly alter

macrophage phagocytosis of L. monocytogenes

We first determined if the presence of neutrophils affects L. monocytogenes phagocytosis by macrophages. As a human macrophage model, we used a human monocyte cell line, Thp1 cells, which were differentiated into macrophages with PMA

(304, 305). L. monocytogenes can grow in Thp1 cells in a LLO-dependent manner (Fig.

106

4.1) (306). This is similar to murine macrophages, but in contrast to neutrophils, which kill L. monocytogenes (Chapters 2 and 3) (13, 92).

Neutrophils (at a ratio of 2 and 3 neutrophils per Thp1 cell) were applied to cultured Thp1 cells and incubated for 10 min to allow neutrophils to attach to the coverslips. Cells were then infected with wild type (wt) or Δhly L. monocytogenes at

MOI 1 such that there was one bacterium present per one mammalian cell (number of L. monocytogenes = number of neutrophils + Thp1 cells). After 30 min at 37°C, cells were washed and fixed. Extracellular and total L. monocytogenes were labeled with fluorescent antibodies. Thp1 cells and neutrophils were labeled with DAPI and differentiated based upon nuclear morphology (Fig. 4.2A). Total and intracellular L. monocytogenes per macrophage and neutrophil were enumerated. The presence of neutrophils did not

Figure 4.1. L. monocytogenes growth in Thp1 cells is LLO-dependent. Thp1 macrophages were incubated with wt or Δhly L. monocytogenes (Lm) at MOI 1 in RPMI. After 30 min at 37°C, cells were washed and incubated with 15 µg/ml gentamicin. After 5 h, cells were fixed then extracellular and total bacteria, and macrophage nuclei were fluorescently labeled. Total Lm per macrophage (Lm/Thp1) was enumerated. Results are the mean ± SEM of two (Δhly) or four (wt) independent experiments, performed in duplicate, ** P < 0.01.

107 significantly alter L. monocytogenes association with or phagocytosis by Thp1 cells (Fig

4.2B and C). This resulted in a similar number of intracellular L. monocytogenes per

Figure 4.2. Neutrophils do not significantly alter macrophage phagocytosis of L. monocytogenes. Neutrophils (PMNs) were added to Thp1 macrophages at the indicated ratio of neutrophil to Thp1 cell (PMN: Thp1). After 10 min, cells were infected with wt or Δhly L. monocytogenes (Lm) at MOI 1 in RPMI. After 30 min at 37°C, cells were washed and fixed. Extracellular and total bacteria, and macrophage and neutrophil nuclei were fluorescently labeled. (A) Representative images of macrophages infected in the absence and presence of neutrophils (3 PMN: 1 Thp1). The empty arrowheads indicate a macrophage and the filled arrowheads indicate a neutrophil, as indicated by the distinctive multi-lobed nuclei. The empty and filled arrows indicate bacteria that have been phagocytosed by macrophages and neutrophils, respectively. Total Lm per macrophage (B), percent intracellular Lm (C), and intracellular Lm / Thp1 cell (D) were enumerated. Results are the mean ± SEM of two to four independent experiments, performed in duplicate, * P < 0.05.

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Thp1 cell after 30 min of infection, regardless of the neutrophil to macrophage ratio (Fig.

4.2D). Thus, co-culture of neutrophils with macrophages does not significantly alter macrophage phagocytosis of L. monocytogenes. Of note, unlike murine macrophages

(Fig. 3.12), but similar to human hepatocytes, epithelial cells (35, 307), and neutrophils

(Fig. 3.1), LLO can facilitate entry into Thp1 macrophages (Fig. 4.2C).

4.3.2 Co-culturing macrophages with neutrophils inhibits L. monocytogenes

replication in macrophages

We next determined if co-culturing macrophages with neutrophils prevented L. monocytogenes replication in macrophages. Neutrophils and Thp1 cells were infected as described above. However, after 30 min at 37°C instead of fixing cells, they were washed and incubated with gentamicin to kill extracellular bacteria. After a total of 5 h, cells were washed and fixed then bacteria, macrophages, and neutrophils were fluorescently labeled (Fig. 4.3A). Total and intracellular L. monocytogenes per macrophage and neutrophil were enumerated. The percentage of L. monocytogenes growth in macrophages and neutrophils was calculated as: [(intracellular L. monocytogenes at 5 h) /

(intracellular L. monocytogenes at 30 min) X 100]. As shown in Fig. 4.3B, L. monocytogenes intracellular growth in Thp1 cells was significantly reduced in the presence of neutrophils, in a concentration dependent manner. L. monocytogenes fate in neutrophils was unaltered in the presence of Thp1 cells (data not shown).

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Figure 4.3. Co-culture of neutrophils with macrophages inhibits L. monocytogenes replication in macrophages. Neutrophils (PMNs) were added to Thp1 macrophages at the indicated ratio of neutrophil to Thp1 cell (PMN: Thp1). After 10 min, cells were infected with wt or Δhly L. monocytogenes (Lm) at MOI 1 in RPMI. After 30 min at 37°C, cells were fixed or washed and incubated with gentamicin to kill extracellular bacteria. Cells were incubated for a total of 5 h then fixed. Extracellular and total bacteria, and macrophage and neutrophil nuclei were fluorescently labeled. (A) Representative images of Thp1 macrophages incubated in the absence or presence of neutrophils (3 PMN: 1 Thp1), T = 5 h. The empty arrowheads indicate a macrophage and the filled arrowheads indicate an apoptotic neutrophil. (B) Intracellular Lm / Thp1 cell was enumerated at 30 min and 5 h. Percent Lm growth was calculated as: [(intracellular Lm per Thp1 cell at 5 h) / (intracellular Lm per Thp1 cell at 30 min) x 100], relative to macrophages infected with wt Lm in the absence of neutrophils. Results are the mean ± SEM of two to four independent experiments, performed in duplicate, ** P < 0.01.

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After 5 h of infection, extracellular DNA (presumably NETs) was observed and neutrophils exhibited condensed nuclei, indicative of apoptotic neutrophils (Fig. 4.3A).

Formation of NETs was donor dependent (observed with 2 out of 4 donors) and highly variable between experiments (data not shown). Unlike NET formation, most neutrophils appeared apoptotic by 5 h, regardless of the donor. Neutrophil apoptosis appeared to be macrophage dependent as apoptotic neutrophils were only observed in the presence of

Thp1 cells (data not shown). However, the PMA used to differentiate the Thp1 cells may also induce neutrophil apoptosis (308, 309). This is unlikely with these experimental conditions as macrophages were washed three times before neutrophils were added, thus diluting the PMA to less than 20 pM, which is not sufficient to induce neutrophil apoptosis (308, 309). After 30 min, cells were further washed before incubating for 5 h, diluting the PMA even more. Thus, it is likely that the Thp1 cells, and not the PMA, induced neutrophil apoptosis. In conclusion, these results show that L. monocytogenes growth in macrophages is limited if neutrophils are present during infection, this may be linked to neutrophil apoptosis.

We next determined if neutrophils cooperate with macrophages in a contact- dependent or -independent manner. Macrophages and neutrophils were infected as above, but were separated by a transwell filter to inhibit mammalian cell contact. This transwell filter contains 0.4 µm pores, which should allow neutrophil and macrophage products to pass through the membrane, but prevent the passage of L. monocytogenes, neutrophils, and macrophages. In these experiments, infected macrophages were below the filter and infected neutrophils were above the filter. Preliminary experiments (n=1) indicate that L.

111 monocytogenes association with and phagocytosis by macrophages is unaffected by the presence of neutrophils above the filter (Fig. 4.4A-C). L. monocytogenes replication in macrophages is also unaffected by the presence of neutrophils above the filter, even if additional neutrophils are added (Fig. 4.4D). These results indicate that neutrophil

Figure 4.4. Neutrophil cooperation with macrophages is contact-dependent. Thp1 macrophages were infected with wt L. monocytogenes (Lm) at MOI 1 in RPMI. Transwell filters (0.4 µm) were added to wells then neutrophils and Lm at MOI 1 in RPMI were applied above the filter. After 30 min at 37°C, macrophages were washed and fixed (T= 30 min) or incubated with gentamicin to kill extracellular bacteria. These cells were incubated for 4 h, then washed and fixed (T= 5 h). Extracellular and total bacteria, and macrophage nuclei were fluorescently labeled. Total Lm per macrophage (A), percent intracellular Lm (B), and total intracellular Lm / macrophage (C) at 30 min was determined. Percent Lm growth after 5 h (D) was calculated as described for Fig. 4.3B. Results are from one experiment.

112 cooperation with macrophages may be contact-dependent, although this experiment needs to be repeated to confirm these findings.

4.4 Discussion

Our data establish that co-culture of human macrophages with human neutrophils does not alter macrophage phagocytosis of L. monocytogenes but does limit L. monocytogenes replication in macrophages. There are a few different hypotheses that may explain this. Firstly, neutrophils may kill extracellular L. monocytogenes, thereby preventing growth following phagocytosis by Thp1 cells. Secondly, neutrophils may cooperate with macrophages in a contact-independent manner. This may include activation of macrophages via mediators released by the neutrophil and/or macrophage internalization of neutrophil products. Thirdly, neutrophils may cooperate with macrophages in a contact-dependent manner. This may include contact-dependent macrophage activation or macrophage internalization of neutrophils (200, 206, 212, 213,

226).

In support of the first hypothesis, L. monocytogenes induces NET formation and degranulation, mechanisms used by neutrophils to kill extracellular bacteria (Figs. 3.4 and 3.5) (153, 256, 310). However, neutrophil mediated killing of extracellular bacteria should not be altered by the presence of a transwell membrane, and thus L. monocytogenes replication in macrophages should still be limited under these conditions.

This was not the case in preliminary experiments, as observed in figure 4.4D. However, it is possible that the transwell filter hinders diffusion of neutrophil products and that

113 bacteria separated from the neutrophils by this filter would not be killed efficiently. It is also possible that macrophage products activate neutrophils to enhance neutrophil killing of extracellular L. monocytogenes, and that diffusion of both the macrophage and neutrophil products are not efficient when separated (209). Therefore, we do not rule out the possibility that extracellular killing of L. monocytogenes plays a role in the results seen here.

In support of the second hypothesis, that cooperation is contact-independent, L. monocytogenes induces neutrophil degranulation (Figs. 3.4 and 3.5) and neutrophil granules contain many antimicrobial molecules and proteases, including the defensins and matrix metalloproteases, which inhibit LLO activity (Figs. 2.9 and 3.10) (188).

Furthermore, exogenously added neutrophil defensins inhibit L. monocytogenes intracellular replication in macrophages (Figs. 2.1 and 2.2) and enter macrophages, where they colocalize with L. monocytogenes (Fig 2.10); defensins can also activate macrophage bactericidal activities (157). However, contact-independent cooperation may not occur under these experimental conditions. Preliminary experiments separating macrophages and neutrophils with a transwell filter allowed L. monocytogenes to replicate in macrophages as efficiently as in the absence of neutrophils (Fig. 4.4).

Nonetheless, neutrophil macrophage cooperation may be contact-independent; as mentioned above, neutrophil products may not efficiently diffuse across the transwell membrane, and thus may not access the macrophage. It would be interesting to transfer neutrophil supernatant to macrophages during L. monocytogenes infection and determine if this limits L. monocytogenes replication in macrophages to confirm the above results.

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The preliminary results shown in figure 4.4 indicate that neutrophil cooperation with macrophages may be contact-dependent, supporting the third hypothesis.

Macrophages induce neutrophil apoptosis through membrane TNF in a contact-dependent manner (311, 312) and macrophage internalization of apoptotic neutrophils limits M. tuberculosis replication in macrophages (213). Thus, it is possible that the apoptotic neutrophils that we observed help macrophages control L. monocytogenes infection in a contact-dependent manner (Fig. 4.3). L. monocytogenes also induces apoptosis of neutrophils (313). In our experimental conditions, bacteria alone were not sufficient to induce neutrophil apoptosis and apoptotic neutrophils were only observed if neutrophils were incubated with macrophages, regardless of L. monocytogenes infection (data not shown), although L. monocytogenes may contribute to macrophage induced neutrophil apoptosis. Apoptotic neutrophils may be internalized by macrophages and targeted to the

L. monocytogenes containing phagosome to mediate bacterial degradation. Of concern for this hypothesis is the time required to induce neutrophil apoptosis and if this would happen quickly enough to inhibit L. monocytogenes replication. However, neutrophils can undergo apoptosis after just a 20 min incubation with stimulus (314). L. monocytogenes and macrophages may cooperate to induce rapid neutrophil apoptosis.

Therefore, it is of interest to elucidate the time frame of neutrophil apoptosis in these experimental conditions and determine if preventing apoptosis abrogates the effects seen when neutrophils and macrophages are co-incubated. It also needs to be determined if macrophages phagocytose neutrophils in these experiments, and if so, what the kinetics of phagocytosis are. It would also be interesting to know if L. monocytogenes escape

115 from the phagosome or growth in the cytosol is prevented during macrophage and neutrophil co-culture, as this could indicate how quickly the L. monocytogenes life cycle must be inhibited. For example, if L. monocytogenes are unable to escape from macrophage phagosomes when macrophages are co-cultured with neutrophils, the cooperative action between these cells must occur within the first 30 min of infection, before the bacteria escape from the phagosome.

In conclusion, co-incubation of neutrophils with macrophages limits L. monocytogenes replication in macrophages, which may occur through an unknown contact-dependent manner. It is of much interest to characterize this cooperation as similar mechanisms may limit replication of other intracellular pathogens in macrophages as well as other cells. As macrophages, and other cells, are in close contact with neutrophils at infectious foci, it seems likely that this cooperation occurs in vivo to impact the course of infection.

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Chapter 5. Synthesis

L. monocytogenes is a foodborne intracellular pathogen and the causative agent of listeriosis, a severe disease with a 30% fatality rate among susceptible populations such as the elderly and immunocompromised individuals. Pregnant women are also susceptible, as L. monocytogenes can infect the developing fetus, leading to miscarriage and stillbirth. However, healthy individuals typically control infection and may only present with mild gastroenteritis (1).

The innate immune system is important for initial control of many infections, including L. monocytogenes. Neutrophils and macrophages are an important component of the innate immune system and are among the first immune cells to come in contact with invading microorganisms. Although macrophages are important for control of L. monocytogenes infection, they may also serve as a reservoir for the bacteria (84, 85, 87).

While activated macrophages efficiently kill L. monocytogenes, the bacteria are able to grow in non-activated macrophages (94). The L. monocytogenes intracellular life cycle in macrophages has been extensively characterized (95, 315). However, L. monocytogenes interactions with neutrophils are not well understood. Neutrophils are important for control of L. monocytogenes infection, and are able to kill a large proportion of the L. monocytogenes inoculum (69, 96, 97, 99, 103). However, it is unknown if any L.

117 monocytogenes can survive within neutrophils and the mechanisms neutrophils use to kill

L. monocytogenes are not well known. Neutrophils produce a robust ROS burst, and can kill L. monocytogenes in a ROS-dependent manner, however, when this is inhibited, neutrophils still kill a large proportion of the L. monocytogenes inoculum (68, 280-282).

Thus, unknown non-oxidative mechanisms are used by neutrophils to kill L. monocytogenes. Neutrophils have been shown to enhance macrophage bactericidal actions, although the mechanisms behind this are not well understood and have not been studied in regards to L. monocytogenes infection (210, 211).

We addressed if neutrophil products, like the defensins, enable macrophages to control intracellular replication of L. monocytogenes (Chapter 2). We next wanted to determine if neutrophils cooperate with macrophages to limit L. monocytogenes replication in macrophages. However, little is known regarding L. monocytogenes interaction with neutrophils, so we first determined if L. monocytogenes is able to replicate in neutrophils (Chapter 3). After gaining more insight into L. monocytogenes interaction with neutrophils, we assessed if neutrophils can inhibit L. monocytogenes replication in macrophages (Chapter 4).

We found that defensins enable macrophages to control intracellular proliferation of L. monocytogenes (Figs. 2.1 and 2.2). The α-defensin HNP-1 interacted with macrophages and the humanized θ-defensin RC-1 interacted with bacteria to inhibit intracellular proliferation (Fig. 2.3). The defensins did not alter macrophage phagocytosis of L. monocytogenes (Fig. 2.6), but inhibited phagosomal escape (Figs. 2.7 and 8), presumably due to their ability to inhibit LLO release and activity (Fig. 2.9).

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Furthermore, HNP-1 is taken up by macrophages and colocalizes with intracellular L. monocytogenes (Fig. 2.10). This study indicates that neutrophil products can enter macrophages and inhibit bacterial intracellular replication. This supports the notion that during infection, cooperation between macrophages and neutrophils limits L. monocytogenes replication in macrophages.

We next studied the interaction between L. monocytogenes, LLO, and human neutrophils. We found that neutrophils kill intracellular L. monocytogenes and degrade

LLO (Figs. 3.2 and 3.9). Interestingly, LLO enhances neutrophil phagocytosis of L. monocytogenes, but does not protect L. monocytogenes from neutrophil killing (Figs. 3.1 and 3.2, Table 3.1). L. monocytogenes induces rapid neutrophil degranulation, which is mediated through multiple virulence factors, including LLO (Figs 3.4 and 3.5) (256).

Degranulation protects neutrophils from LLO-mediated perforation (Figs. 3.6 and 3.7), likely because neutrophil granule products (like MMP-8) degrade LLO (Figs. 3.9 and

3.10, Table 3.2). This study shows that neutrophils rapidly degranulate in response to L. monocytogenes and LLO, releasing MMP-8 into the extracellular environment. We propose that LLO degradation by MMP-8 happens before the phagosome has sealed and following phagocytosis, thus preventing LLO-mediated escape from the phagosome and trapping the bacteria in a toxic phagosome (Fig. 3.11).

After determining that LLO does not protect L. monocytogenes from neutrophil intracellular killing, and that L. monocytogenes activates neutrophils (Figs. 3.2, 3.4, and

3.5), we determined if human neutrophils cooperate with macrophages to limit L. monocytogenes replication in human macrophages. Co-incubation of neutrophils and

119 macrophages does not alter macrophage phagocytosis but does limit L. monocytogenes replication in macrophages (Figs. 4.2 and 4.3).

Together, these results show that neutrophils efficiently kill L. monocytogenes and are resistant to LLO-mediated perforation, likely due to multiple mechanisms as neutrophil products inhibit LLO secretion and activity (defensins), and degrade LLO

(MMP-8). These results also establish that macrophages limit L. monocytogenes intracellular replication if treated with neutrophil defensins or co-cultured with neutrophils during infection. It is of considerable interest to elucidate how neutrophil defensins inhibit LLO activity and if defensins cooperate with MMP-8 to enhance inactivation of LLO. The Lehrer group showed that HNP-1 does not inhibit LLO binding to erythrocytes and proposed that defensins inhibit LLO oligomerization in host cell membranes (188). It is of interest to confirm this hypothesis and determine if the θ- defensin RC-1 inhibits LLO activity in a similar manner as the α-defensin HNP-1. It would also be interesting to determine where MMP-8 cleaves LLO and which neutrophil proteases cooperate with MMP-8 to degrade LLO. Inhibiting LLO activity is expected to prevent L. monocytogenes escape from the phagosome, trapping the bacteria in a degradative compartment. As such, inhibiting either the defensins and/or MMP-8 may restore LLO activity and allow L. monocytogenes to escape the phagosome and replicate within neutrophils. However, neutrophils contain many antimicrobial molecules that are rapidly mobilized and that may exert redundant activities. Also of interest is how HNP-1 enters macrophages, what activities HNP-1 exerts once inside macrophages, and if HNP-

1 entry is important for subsequent macrophage activation. Results in chapters 2 and 3

120 indicate that neutrophils could cooperate with macrophages through the release of defensins and/or MMP-8, and presumably other antimicrobial molecules. Indeed, data in chapter 4 shows that neutrophils can cooperate with macrophages to limit L. monocytogenes infection of macrophages. It is of interest to determine if macrophages induce the neutrophil apoptosis observed in chapter 4, what the kinetics of apoptosis are, and if this mediates the observed cooperation between neutrophils and macrophages. It would also be interesting to determine if macrophages internalize neutrophils during co- incubation and if so, if neutrophils and/or their products colocalize with intracellular L. monocytogenes. Furthermore, it needs to be determined if neutrophils kill extracellular L. monocytogenes under those experimental conditions. The results described here, and future projects stemming from these results, should further the understanding of the interaction between invading pathogens like L. monocytogenes and the host immune system.

In conclusion, we show that neutrophils degrade LLO and kill intracellular L. monocytogenes, two novel findings that have broad impact for the fields of microbe host interaction and immunology. We identified multiple mechanisms for neutrophil mediated anti-LLO activity, inhibition of LLO secretion and activity via HNPs, and degradation of

LLO via MMP-8. Our results also indicate that neutrophil resistance to LLO-mediated perforation is likely not limited to LLO, but representative of neutrophil interaction with other CDCs, as neutrophil granule products inhibit erythrocyte perforation by two other

CDCs, ALO and PLY. Furthermore, neutrophils kill many CDC producing organisms

(297-301). Results described herein may apply to neutrophil interaction with other CDC

121 producing organisms and may indicate an important role for neutrophils in controlling

CDC-mediated damage.

We also show that neutrophils (and their products) cooperate with macrophages to limit L. monocytogenes replication in macrophages, something that was not previously shown in regards to L. monocytogenes. As neutrophils and macrophages are seen in close proximity at infectious foci, it seems likely that neutrophils also cooperate with macrophages in vivo. Our results provide potential mechanisms for how neutrophils may cooperate with macrophages to limit L. monocytogenes infection. Application of neutrophil defensins limits L. monocytogenes intracellular replication in macrophages, through mechanisms independent of direct antibacterial action. For example, HNP has little anti-listerial activity but still limits L. monocytogenes replication in macrophages, presumably through inhibiting LLO secretion and activity and cooperating with macrophage antimicrobial responses. Also, we hypothesize that macrophage treatment with MMP-8 will lead to LLO degradation and thus limit L. monocytogenes replication in macrophages. Treatment of L. monocytogenes, and likely other bacterial infections could be mediated by application of neutrophil products (like defensins and/or MMP-8) to limit

L. monocytogenes intracellular replication in host cells. Alternatively, inducing neutrophil recruitment to infectious sites, even during later stages of disease, may lead to better control of infection. Although these studies focused on L. monocytogenes replication in macrophages, it seems likely that neutrophils also cooperate with other host cells at infectious foci. For example, defensins inhibit L. monocytogenes entry into hepatocytes

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(data not shown) and neutrophils have been proposed to lyse infected hepatocytes to help control infection (277-279).

In closing, human neutrophils are critical for control of bacterial infections, including those caused by L. monocytogenes. Capitalizing on these natural defenses, either through upregulation of neutrophil production or mobilization, or application of neutrophil defensins or MMP-8, may provide viable treatment options. However, this needs to be approached cautiously, as deregulated neutrophil, defensin, and MMP-8 expression are implicated in various diseases (132, 157, 316).

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