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Pathogen-induced inflammation in immunocompromised conditions McDonald, Allison 2014

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                 PATHOGEN-INDUCED INFLAMMATION IN IMMUNOCOMPROMISED CONDITIONS  by Allison McDonald  B.Sc., The University of New Brunswick, 2009  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY in The Faculty of Graduate and Postdoctoral Studies (Microbiology and Immunology)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  July 2014  © Allison McDonald, 2014   ii Abstract Primary immunodeficiencies arise from genetic anomalies and can cause aberrant inflammation, tissue destruction, and uncontrolled infections. Defects in immunity often first present as cases of recurrent or severe infections with unusual pathogens. Patients with chronic granulomatous disease (CGD) have mutations in the NADPH oxidase that prevent the formation of reactive oxygen species (ROS) critical for effective immunity. We investigated the phagocyte response to Burkholderia cenocepacia, which causes severe lung infections in patients with CGD. B. cenocepacia survived within ROS-deficient neutrophils and induced rapid cell death that was not observed by infection with other bacteria, including other CGD pathogens. Increased cell death was associated with enhanced caspase-3 activation and phosphatidylserine surface exposure compared to healthy neutrophils, both hallmarks of apoptotic programmed cell death. Caspase activation was required to induce increased neutrophil death in the absence of ROS. B. cenocepacia-induced apoptotic neutrophils were pro-inflammatory to macrophages, especially in the absence of ROS, inducing the secretion of IL-6, IL-8, and TNF-α but not the anti-inflammatory cytokines IL-10 or TGF-β from human macrophages. Surprisingly, macrophages enhanced the growth of B. cenocepacia by providing a nutrient-rich intracellular replication niche that allowed the bacteria to escape from normal neutrophil killing.  We also report here two sisters with an underlying immunodeficiency who presented with fatal neurodegeneration triggered by infections. Whole exome sequencing identified mutations in perforin1, involved in the granule-mediated cytotoxicity pathway for eliminating infected host cells. Perforin1 mutations are associated with familial hemophagocytic lymphohistiocytosis (FHL). In contrast to FHL, these girls did not have hematopathology or elevated cytokine overproduction. However, 3 years after disease onset, the proband had markedly deficient   iii cytokine production, including IL-1β production following stimulation with a panel of bacterial ligands. These observations extend the spectrum of disease associated with perforin mutations to immune-mediated neurodegeneration triggered by infection.  Studies that investigate how opportunistic pathogens cause disease aim to decrease morbidity and mortality in immunocompromised patients and also provide the opportunity to better understand the molecular mechanisms that allow healthy individuals to combat disease.    iv Preface Ethics approval was obtained for collection of blood samples from UBC C&W Research Ethics Board (ethics certificate H04-70193) and Biohazard Approval certificate UBC (protocol B11-0012). Patient family members in Chapter 4 gave informed consent/assent for UBC C&W Research Ethics Board protocol H09-01228 and for National Institutes of Health protocol 76-HG-0238. Written consent was obtained for the use of photos in Figure 4.1.   Permission for reproduction of Figure 1.1 and Table 1.1 were obtained from the Annual Review of Medicine. Chapters 2 and 3 were designed by A McDonald with assistance from DP Speert. TJ Hird and R Hickman assisted with ELISAs and microscopic cell and bacteria counting. A McDonald designed and performed all of the experiments, analyzed the data, and wrote the chapters with assistance from DP Speert. A version of Chapter 2 and a version of Chapter 3 are each being prepared as separate manuscripts to be submitted for publication in the Summer of 2014.   A version of Chapter 4 has been published: Dias C*, McDonald A*, Sincan M, Rupps R, Markello T, Salvarinova R, Santos RF, Menghrajani K, Ahaghotu C, Sutherland DP, FortunoIII ES, Kollmann TR, Demos M, Friedman JM, Speert DP, Gahl WA, and Boerkoel CF. Recurrent subacute post-viral onset of ataxia associated with a PRF1 mutation. European Journal of Human Genetics. 2013 Nov;21(11):1232-9. * Joint co-first authors. Permission to reproduce Chapter 4 in whole has been obtained from the authors. The study was designed by A McDonald and C Dias, with assistance from DP Speert and CF Boerkoel. C Dias performed the clinical analysis and whole exome sequencing and assembled Figure 4.1 of this thesis. A McDonald directed the immunological investigations and performed the PMBC stimulations and IL-1β ELISAs (Figure 4.2), which formed the basis of the paper. DP Sutherland performed the Luminex assay and prepared Figure 4.3. A McDonald contributed to the analysis of all data, the writing of the manuscript, and additional experiments during revisions as required by reviewers.     v Table of Contents Abstract.......................................................................................................................................... ii!Preface........................................................................................................................................... iv!Table of Contents ...........................................................................................................................v!List of Tables ..................................................................................................................................x!List of Figures............................................................................................................................... xi!List of Symbols and Abbreviations .......................................................................................... xiii!Acknowledgements ................................................................................................................... xvii!Chapter 1: Introduction ................................................................................................................1!1.1! Inflammatory disease ......................................................................................................... 2!1.1.1! Hemophagocytic lymphohistiocytosis ........................................................................ 2!1.1.2! Chronic granulomatous disease .................................................................................. 6!1.1.3! Cystic fibrosis ............................................................................................................. 9!1.2! Phagocyte biology............................................................................................................ 12!1.3! Cell death ......................................................................................................................... 14!1.3.1! Apoptosis .................................................................................................................. 14!1.3.2! Granule-mediated cytotoxicity.................................................................................. 17!1.3.3! Necrosis..................................................................................................................... 19!1.3.4! Alternative modes of programmed cell death........................................................... 20!1.4! The Burkholderia cepacia complex................................................................................. 23!1.5! Evasion of host defenses by the Bcc................................................................................ 25!1.5.1! Evasion of ROS and oxidative killing ...................................................................... 25!1.5.2! Resistance to non-oxidative killing........................................................................... 26!  vi 1.6! Interactions of the Bcc with phagocytes .......................................................................... 28!1.6.1! Survival and trafficking within macrophages ........................................................... 28!1.6.2! Neutrophils and the Bcc............................................................................................ 29!1.7! Hypothesis and overarching aims .................................................................................... 30!Chapter 2: Burkholderia cenocepacia induces rapid neutrophil apoptosis in the absence of reactive oxygen species that promotes a pro-inflammatory macrophage response. .............31!2.1! Introduction...................................................................................................................... 31!2.2! Materials and methods ..................................................................................................... 34!2.2.1! Isolation of cells........................................................................................................ 34!2.2.2! Bacterial strains and growth conditions.................................................................... 35!2.2.3! B. cenocepacia growth curves .................................................................................. 36!2.2.4! Neutrophil challenge with bacteria ........................................................................... 37!2.2.5! Association and intracellular killing assay ............................................................... 37!2.2.6! Flow cytometric analysis of neutrophil cell death .................................................... 38!2.2.7! Caspase activity by FLICA staining ......................................................................... 38!2.2.8! Immunoblots ............................................................................................................. 39!2.2.9! Lactate dehydrogenase assay .................................................................................... 40!2.2.10! Cytokine release by macrophages........................................................................... 40!2.2.11! Statistics .................................................................................................................. 41!2.3! Results.............................................................................................................................. 41!2.3.1! Impact of ROS on neutrophil death following challenge with B. cenocepacia clinical isolates................................................................................................................................... 41!  vii 2.3.2! Impact of ROS on neutrophil death following challenge with a panel of pathogenic and non-pathogenic bacteria in CGD.................................................................................... 43!2.3.3! Intracellular killing of B. cenocepacia by neutrophils.............................................. 47!2.3.4! Composition of live and apoptotic neutrophils containing B. cenocepacia.............. 48!2.3.5! B. cenocepacia-induced caspase activation .............................................................. 51!2.3.6! ROS-deficient neutrophils challenged with B. cenocepacia induce a pro-inflammatory response in macrophages................................................................................ 54!2.4! Discussion ........................................................................................................................ 57!Chapter 3: Human macrophages provide a rich replication niche that allows B. cenocepacia to escape neutrophil killing and enhances replication.........................................63!3.1! Introduction...................................................................................................................... 63!3.2! Materials and methods ..................................................................................................... 66!3.2.1! Isolation of cells........................................................................................................ 66!3.2.2! Bacterial strains and growth conditions.................................................................... 67!3.2.3! B. cenocepacia growth curves .................................................................................. 68!3.2.4! Neutrophil challenge with B. cenocepacia and incubation with macrophages ........ 68!3.2.5! Co-culture of macrophages, neutrophils, and B. cenocepacia and cytokine analysis….............................................................................................................................. 69!3.2.6! Phagocytosis of B. cenocepacia in co-culture with macrophages and neutrophils .. 69!3.2.7! Gentamicin protection assay ..................................................................................... 69!3.2.8! Bacterial culture with macrophages.......................................................................... 70!3.2.9! Bacterial culture with lysed macrophages ................................................................ 71!3.2.10! Transwell assays ..................................................................................................... 71!  viii 3.2.11! Lactate dehydrogenase assay .................................................................................. 72!3.2.12! Statistics .................................................................................................................. 72!3.3! Results.............................................................................................................................. 72!3.3.1! Macrophages promote bacterial survival from B. cenocepacia-infected neutrophils72!3.3.2! Effect of macrophages and neutrophils on survival of B. cenocepacia .................... 73!3.3.3! Modulation of B. cenocepacia-induced inflammatory mediators by neutrophils..... 75!3.3.4! Phagocytosis of B. cenocepacia by macrophages and neutrophils........................... 78!3.3.5! Intracellular growth of B. cenocepacia in human monocyte-derived macrophages. 81!3.3.6! Effect of macrophage-released factors on growth of B. cenocepacia ...................... 82!3.3.7! Role of phagocytosis in macrophage-induced growth of B. cenocepacia ................ 84!3.4! Discussion ........................................................................................................................ 87!Chapter 4: Recurrent subacute post-viral onset of ataxia associated with PRF1 mutation .93!4.1! Introduction...................................................................................................................... 93!4.2! Materials and methods ..................................................................................................... 95!4.2.1! Patient data and standard protocol approvals............................................................ 95!4.2.2! Exome sequencing and analysis................................................................................ 95!4.2.3! Cell stimulation and measurement of cytokines ....................................................... 96!4.2.4! Statistics .................................................................................................................... 97!4.3! Results.............................................................................................................................. 97!4.3.1! Clinical features ........................................................................................................ 97!4.3.2! Defective cytokine production by the proband....................................................... 100!4.3.3! Exome sequencing .................................................................................................. 103!4.4! Discussion ...................................................................................................................... 103!  ix Chapter 5: Conclusion...............................................................................................................107!5.1! Future directions ............................................................................................................ 111!Bibliography ...............................................................................................................................114!Appendix.....................................................................................................................................164!Appendix A  Growth curves of B. cenocepacia isolates K56-2 and MH1K .......................... 164!   x List of Tables Table 1.1 Current diagnostic criteria of HLH................................................................................. 3!Table 1.2 Species of the Burkholderia cepacia complex and their clinical relevance ................. 25!Table 2.1 Bacterial strains used in this study................................................................................ 36!   xi List of Figures Figure 1.1 Mechanism of cytotoxic granule trafficking in CTL/NK cells ................................... 19!Figure 2.1 Neutrophil apoptosis and necrosis following challenge with clinical B. cenocepacia isolates........................................................................................................................................... 43!Figure 2.2 Neutrophil apoptosis and necrosis following challenge with a panel of pathogenic and non-pathogenic CGD bacteria....................................................................................................... 46!Figure 2.3 Intracellular survival of B. cenocepacia in neutrophils............................................... 48!Figure 2.4 Composition of live and apoptotic neutrophils with detectable B. cenocepacia......... 50!Figure 2.5 B. cenocepacia-induced caspase-3 activation in neutrophils ...................................... 52!Figure 2.6 Caspase activation is required for enhanced cell death in DPI-treated neutrophils .... 53!Figure 2.7 Cytokines produced by macrophages cultured with B. cenocepacia-challenged neutrophils..................................................................................................................................... 56!Figure 3.1  Macrophages enhance survival of B. cenocepacia following phagocytosis by neutrophils..................................................................................................................................... 73!Figure 3.2  Effect of macrophages and neutrophils on the survival of B. cenocepacia................ 75!Figure 3.3 B. cenocepacia-induced inflammatory mediators. ...................................................... 77!Figure 3.4 Relative phagocytosis of B. cenocepacia by neutrophils and macrophages ............... 79!Figure 3.5 B. cenocepacia-induced phagocyte lysis. .................................................................... 80!Figure 3.6 Intracellular growth of B. cenocepacia within macrophages. ..................................... 82!Figure 3.7 Effect of macrophage-released factors on B. cenocepacia growth. ............................ 84!Figure 3.8 Uptake of B. cenocepacia by intact macrophages is necessary for enhanced bacterial replication. .................................................................................................................................... 86!Figure 4.1 Familial pedigree and photographs of the affected girls. .......................................... 100!  xii Figure 4.2 IL-1β production is defective in PBMCs from the proband...................................... 101!Figure 4.3 Cytokine and chemokine levels detected by Luminex. ............................................. 102!Figure A.1 B. cenocepacia isolates K56-2 and MH1K display similar growth curves .............. 164!   xiii List of Symbols and Abbreviations α: alpha β: beta γ: gamma ALT: alanine aminotransferase APC: antigen-presenting cell AST: aspartate aminotransferase ATP: adenosine triphosphate BC: British Columbia Bcc: Burkholderia cepacia complex BcCV: Bcc-containing vacuole BCG: Bacille Calmette-Guérin  CBCCRRR: Canadian Burkholderia cepacia Complex Research and Referral Repository CFU: colony forming units CGD: chronic granulomatous disease CF: cystic fibrosis CFTR: cystic fibrosis transmembrane conductance regulator CNS: central nervous system CO2: carbon dioxide CSF: cerebrospinal fluid CT: computed tomography CTL: cytotoxic lymphocyte DAMPs: danger-associated molecular patterns   xiv DMEM: Dulbecco’s modified Eagle medium DNA: deoxyribonucleic acid DPI: diphenyleneiodonium DISC: death-inducing signaling complex EEA-1: early endosome antigen 1 EDTA: ethylenediaminetetraacetic acid eGFP: enhanced green fluorescent protein ELISA: enzyme-linked immunosorbant assay ER: endoplasmic reticulum ERK: extracellular-signal-regulated kinase FCS: fetal calf serum FHL: familial hemophagocytic lymphohistiocytosis FLICA: fluorescent-labeled inhibitors of caspases IRF: interferon regulatory transcription factor KRG: Krebs-Ringer phosphate buffer with glucose GAPDH: glyceraldehyde 3-phosphate dehydrogenase GM-CSF: granulocyte-macrophage colony-stimulating factor HCT: hematopoietic stem cell transplantation HLH: hemophagocytic lymphohistiocytosis HMGB1: high mobility box protein B1 IFN: interferon IAP: inhibitor of apoptosis protein IL: interleukin   xv LB: Luria-Bertani LDH: lactate dehydrogenase LPS: lipopolysaccharide MAP: mitogen-activated protein MDM: monocyte-derived macrophage MH: Mueller Hinton MHC: major histocompatability complex MIC: minimum inhibitory concentration MOI: multiplicity of infection MPO: myeloperoxidase MRI: magnetic resonance imaging MRSA: methicillin-resistant Staphylococcus aureus Mtb: Mycobacterium tuberculosis NADPH: nicotinamide adenine dinucleotide phosphate NETs: neutrophil extracellular traps NF-κB: nuclear factor κB NK: natural killer NLR: NOD-like receptor OD600: optical density at 600 nm PAMP: pathogen-associated molecular pattern PBMCs: peripheral blood mononuclear cells PBS: phosphate buffered saline PCR: polymerase chain reaction   xvi PIC: poly(I:C) PGE2: prostaglandin E2 PGN: peptidoglycan PMN: polymorphonuclear cells PNHS: pooled normal human serum PRF1: perforin1 PRR: pattern recognition receptor PS: phosphatidylserine R10: RPMI 1640 medium containing 10% (v/v) fetal calf/bovine serum  RIC: reduced-intensity conditioning RIPA: radio immunoprecipitation assay RNA: ribonucleic acid ROS: reactive oxygen species RPMI 1640 media: Roswell Park Memorial Institute media SDS PAGE: sodium dodecyl sulfate polyacrylamide gel electrophoresis SEM: standard error of the mean SOD: superoxide dismutase TCR: T cell receptor TGF: transforming growth factor TLR: toll-like receptor TNF: tumor necrosis factor UV: ultraviolet   xvii Acknowledgements The work done for this thesis would not have been possible without the tremendous support from those around me. I would like to thank my supervisor, Dr. David Speert, for the opportunity to work in his laboratory. His guidance has allowed me to develop as a scientist and to be involved in extremely rewarding translational research. I thank my supervisory committee members, Dr. Laura Sly, Dr. Bob Hancock, and Dr. Jan Dutz for their ongoing support and encouragement. Their help in experimental design and critical thinking has been a vital part of my development as a trainee. I am especially grateful for Laura’s guidance: for always having an open door for me and for exceptionally helpful discussions. I would like to thank Dr. Miguel Valvano and his laboratory for providing B. cenocepacia strains MH1K and K56-2 carrying a pDA42-eGFP construct. I am also grateful for the healthy blood donors who participated in these studies as well as my funding through NSERC, the University of British Columbia, and the Rare Disease Foundation. I would like to thank the current and past members of the Speert laboratory. Dr. Rebecca Malott, for her mentorship that extended long past her time at CFRI. Her help and guidance in tissue culture, Burkholderia biology, graduate school, and life in general has been invaluable. I thank Dr. Alex Persson and Dr. James Zlosnik for their expertise in phagocyte biology and microbiology and for many insightful discussions over coffee. Dr. Agatha Jassem and Dr. Billie Velapatino, my lab sisters, were instrumental in helping me through the laughter and tears of graduate school. Dr. Kelly Brown has also been integral in helping me navigate through experimental and program challenges and I thank her for her support. Trevor Hird and Becky Hickman were especially helpful during the last few months of experiments. I am so grateful for all the cell counting and ELISAs that you helped with; I owe you new sets of eyes and wrists if I   xviii ever get rich. I also thank Maureen Campbell and Carolyn Smith for their administrative help and for always keeping me smiling during this rollercoaster ride.  I owe a lot of thanks to my friends and family for their love and support throughout this journey. My mother, who is my rock, and has listened throughout all of my breakthroughs and low points. Thank you for supporting every step. I thank my grandmother, whose phone calls are the highlight of my commute to work every week. To my west coast family, Barb and Dave, thank you for including me in every holiday and for your endless enthusiasm. I am also extremely grateful for my friends who remind me how much fun life is and what is truly important. To my partner in crime, Troy, you are my sanity. Thank you for your flexibility, your love, and your devotion to my science, even when it meant late night drives to the laboratory and getting tricked into counting CFUs.      1 Chapter 1: Introduction Human immunity is a tightly regulated system that responds to foreign material in a finely tuned manner to prevent infection and disease. Primary immunodeficiencies arise from genetic anomalies and can cause aberrant inflammation, tissue destruction, and uncontrolled infections (1). They often first present as cases of recurrent or severe infections with unusual pathogens (2).  Inflammation is a rapid innate response triggered by foreign particles that delivers leukocytes to the site of tissue injury in order to restore homeostasis (3). Phagocytes such as monocytes and neutrophils of the blood and tissue-resident macrophages are equipped with potent antimicrobial weaponry such as reactive oxygen species (ROS), antimicrobial peptides, pattern recognition receptors (PRRs) and degradative enzymes to combat disease (4). Phagocytes are also adept at phagocytosing pathogens and activating the adaptive arm of immunity through the release of inflammatory mediators and chemoattractants, and the presentation of antigens on major histocompatability complex (MHC) molecules. After phagocytosing pathogens, many immune cells undergo programmed cell death and are engulfed by macrophages in a non-inflammatory process that prevents cells from lysing and releasing their toxic intracellular contents (5). Innate defects in inflammatory and antimicrobial pathways and in cell death mechanisms can have a profound effect on infection and inflammation and cause chronic or recurrent diseases in humans (6, 7). The Burkholderia cepacia complex (Bcc) is a group of phenotypically similar opportunistic pathogens that cause respiratory infection in immunocompromised patients, particularly in patients with defective ROS production, as in chronic granulomatous disease (CGD), or in patients with cystic fibrosis (CF) leading to extensive mucus buildup and    2 inflammation in the lung (8, 9). Studies that investigate how the Bcc and other uncommon opportunistic pathogens cause disease can aim to decrease morbidity and mortality in patients but also provide the opportunity to gain a greater understanding of the molecular mechanisms that allow healthy individuals to combat disease.   1.1 Inflammatory disease Inflammation is a finely tuned response that is generally considered beneficial for the host by providing protection against infection. Aberrant inflammation, as is seen in certain primary immunodeficiencies, can become detrimental and cause adverse outcomes such as uncontrolled infection and/or septic shock (3). Although the genetic defects underlying various immunodeficiencies have been identified, the molecular mechanisms that lead to uncontrolled inflammation and/or infection often remain unclear. 1.1.1 Hemophagocytic lymphohistiocytosis Hemophagocytic lymphohistiocytosis (HLH) is an uncommon systemic inflammatory clinical syndrome associated with a number of genetic, infectious, and autoimmune diseases that can lead to multi-organ failure if left untreated. HLH is characterized by immune dysregulation with uncontrolled activation of T cells and macrophages and associated hypercytokinemia (10). Non-malignant activated leukocytes infiltrate the bone marrow, spleen, liver, lymph nodes, and central nervous system. Clinically, patients typically present with cardinal symptoms of high-grade fever, hepatosplenomegaly, progressive cytopenia, and variable neurological symptoms (11, 12).  HLH is a clinical syndrome characterized by hyperinflammation rather than a specific disease (13). Current diagnostic guidelines for HLH were described by the HLH Study Group of    3 the Histiocyte Society in 2004 (Table 1.1) (14). Eight laboratory and clinical criteria were listed, five of which are required for the diagnosis of HLH in the absence of a known genetic defect or family history. Other abnormal findings consistent with HLH include cerebromeningeal symptoms, elevated spinal fluid protein, lymph node enlargement, edema, skin rash, and infiltration of leukocytes and histiocytes into the periportal space of the liver causing jaundice (14). Although these conditions are characteristic of HLH, none are specifically diagnostic. HLH thus exists as a group of related disorders of pathological immune activation, often with widely variable clinical presentation.  Table 1.1 Current diagnostic criteria of HLH.  (© 2012 G.E. Janka, adapted with permission, (13)) 1. Genetic defect or family history consistent with HLH 2. Clinical and laboratory criteria: Five of eight must be fulfilled • Fever • Splenomegaly • Cytopenia (affecting ≥ 2 of 3 lineages in peripheral blood) - Hemoglobin < 90 g/l (in infants < 4 weeks: hemoglobin < 100 g/l) - Platelets < 100 x 109 /L - Neutrophils < 1.0 x 109 /L • Hypertriglyceridemia and/or hypofibrinogenemia  - Fasting triglycerides ≥ 3 mmol/L  - Fibrinogen ≤ 1.5 g/l • Hemophagocytosis in bone marrow, spleen, or lymph nodes - No evidence of malignancy • Low or absent natural killer (NK) cell activity  • Ferritin ≥ 500 µg/L • Soluble CD25 (sIL-2R) ≥ 2,400 U/ml sIL-2R, soluble interleukin (IL)-2 receptor     4 The prevalence of HLH is reported to range between 0.12 in 100,000 children per year in Sweden (15) to 0.34 in 100,000 in Japan (16). A recent study calculated the prevalence of HLH to be 1 in 100,000 children in Texas (17). HLH is classified as either primary or familial HLH (FHL), when there is an identified underlying genetic cause or a family history, or secondary or acquired HLH, which tends to manifest at an older age and is generally associated with an underlying infection, rheumatic illness, or malignancy. Acquired HLH associated with a rheumatic illness is often called macrophage activation syndrome. Overall, a genetic diagnosis can be made in 40-80% of patients with HLH (10, 18, 19) FHL is an autosomal recessive disorder subcategorized into 5 subtypes based on mutations in 5 genetic loci. All of the genes identified encode for proteins involved in the cytotoxic lymphocyte (CTL/NK)-cell granule-dependent cytotoxic pathway of cell death, which is discussed below. The locus responsible for FHL-1 is mapped to chromosome 9q21.3 but the specific mutated gene has not been identified (20). FHL-2 is caused by mutations in the gene encoding perforin (PRF1) (12, 21) and accounts for 20-40% of all FHL cases worldwide and approximately 50% of FHL cases in North America (21, 22). Perforin co-localizes with granzyme B in granules of cytotoxic NK and T cells and is secreted upon conjugation between target and effector cells. Once secreted, perforin functions with granzyme to mediate apoptosis through a mechanism that remains unclear (23). Mutations in Munc-13-4 (UNC13D) (24), syntaxin 11 (STX11) (25), and syntaxin-binding protein 2 (STXBP2) (26, 27) are associated with FHL-3, FHL-4, and FHL-5 respectively and are all involved in regulating the exocytosis of cytotoxic granules. Primary HLH is also described for genetic disorders involving defective intracellular granule trafficking, particularly in Griscelli syndrome type II (RAB27A), Chédiak-Higashi syndrome (LYST), and Hermansky-Pudlak syndrome type II (AP3B1). Epstein-Barr virus    5 (EBV)-driven HLH is associated with a number of primary immunodeficiencies such as X-linked lymphoproliferative disorder, and patients with mutations in genes encoding SAP or XIAP (18).  Untreated, FHL is rapidly fatal (28). Patients die of overwhelming bacterial or fungal infections associated with cytopenia, multiorgan failure, or cerebral dysfunction. Treatment is two-pronged, requiring immediate treatment to suppress life-threatening hyperinflammation and a long-term strategy to definitively correct the underlying genetic defect by replacing the patient’s immune system by allogenic hematopoietic stem cell transplantation (HCT). Management of the hyperinflammation typically involves controlling any identified infectious trigger and arresting CTL and macrophage activation. The first widely-accepted treatment strategy was published by the Histiocyte Society in 1994 (referred to as HLH-94) and aimed at reducing the mortality rate of HLH prior to intended HCT using dexamethasone, etoposide, and, after 8 weeks, adding cyclosporine A (29). In 2004, the standard was revised to include cyclosporine A at the beginning of therapy to prevent reactivations of HLH (14). A regimen of antithymocyte globulin and corticosteroids followed by HCT results in an overall survival identical to HLH-94 at 55% survival (30).  Ultimately, FHL must be corrected by HCT for long-term survival. The standard of care has long been HCT with myeloablative conditioning using bisulfan, cyclophosphamide, and etoposide, however this procedure still has a high mortality rate of 30-50% (31). Reduced-intensity conditioning (RIC) in HLH is associated with lower toxicity and increased survival and its use is increasing (32, 33). However, this procedure results in mixed chimeras in two thirds of children, which requires frequent donor lymphocyte infusions. While RIC HCT is promising, more follow-up is required to determine the long-term outcome (10).     6 1.1.2 Chronic granulomatous disease CGD is a rare primary immunodeficiency that affects approximately 1 in 200,000 live births and is caused by loss-of-function mutations in any one of 5 structural genes of the nicotinamide adenine dinucleotide phosphate (NADPH) oxidase complex (34, 35). The NADPH oxidase is formed on cell plasma and phagosomal membranes and is responsible for initiating the phagocyte oxidative burst that produces ROS upon interaction with microorganisms (36). When activated, the NADPH oxidase catalyses the production of superoxide anion from molecular oxygen. Superoxide is reduced to more toxic ROS such as hydrogen peroxide, hydroxyl radicals, and hypochlorite in the phagosome (37). In addition to the formation of toxic ROS, the oxidative burst leads to the activation of catalytic serine proteases contained within neutrophil granules (38). Collectively, the antimicrobial weaponry activated following NADPH oxidase activation is extremely efficient at killing invading pathogens.  The structural complex of the NADPH oxidase is composed of 2 membrane-bound subunits (gp91phox and p22phox, which together form cytochrome b558), and 3 cytosolic components, (p67phox, p47phox, and p40phox) that assemble into the complex upon activation. Upon pathogen recognition, the cytosolic subunits are phosphorylated and bind together. Secondary (specific) granules, which contain cytochrome b558 in their membrane, proceed to fuse with the phagolysosome. The cytosolic NADPH components and the GTP-binding protein Rac migrate to the membrane to associate with gp91phox and p22phox and form the assembled NADPH complex (39). The oxidase initiates the oxidative burst with the transfer of an electron from NADPH to molecular oxygen to produce superoxide (39). Genetic mutations in humans have been identified in all 5 of the structural genes of the NADPH oxidase (40). Mutations in gp91phox account for approximately 65% of CGD cases in    7 North America, mutations in gp47phox account for approximately 25% of cases, and mutations in p67phox and p22phox together account for approximately 10 % of cases (34). The first patient with a mutation in p40phox was recently described (41). The gp91phox gene is found on the short arm of the X chromosome, and is thus an X-linked disease; all other forms of CGD are autosomal recessive. Patients with X-linked CGD are generally male, present earlier in life, have more severe infections, and a worse prognosis compared to patients with autosomal recessive CGD. Females with mutations in gp91phox have two populations of neutrophils, one that produces ROS and one that does not, with ratios that vary depending on the degree of X chromosome inactivation (42). Various types of mutations in the NADPH oxidase cause CGD, including missense mutations, nonsense mutations, and splicing defects. The location and type of mutation is associated with the amount of residual ROS that a person with CGD can produce and even low levels of ROS seems to be extremely protective for patients (43). CGD is thus a heterogeneous group of disorders with varying degrees of severity and clinical outcomes. CGD is characterized by severe, often recurrent infections with unusual pathogens that are difficult to treat. However, increased susceptibility to infections in patients with CGD is limited to a select class of microorganisms. Many microbes endogenously produce oxygen metabolites such as hydrogen peroxide that CGD neutrophils can modify to use for microbial killing. Patients with CGD are most sensitive to organisms that produce catalase, which degrades hydrogen peroxide produced by the bacteria in the phagosome (44). CGD phagocytes have no means of generating potent ROS to combat catalase-positive organisms and must rely on non-oxidative mechanisms of killing, such as with antimicrobial peptides and catalytic proteases within granules. Thus, in addition to being catalase-positive, common CGD pathogens are also resistant to non-oxidative killing (44). In the United States, the majority of infections are caused by    8 Aspergillus spp, Staphylococcus spp, the Bcc, Serratia spp, and Nocardia spp, (34). Aspergillus spp and the Bcc are the first and second leading causes of deaths in CGD, respectively (34). Elsewhere in the world, infections with Salmonella, Bacille Calmette-Guérin (BCG), and Mycobacterium tuberculosis (Mtb) are also problematic (35, 45). In a United States registry of CGD patients, infections manifested most commonly as pneumonia (occurring at least once in 79% of patients), abscesses of the skin, liver, lung, or perirectum (in 68% of patients), supperative adenitis (in 53% of patients), osteomyelitis (in 25% of patients), and sepsis (in 18% of patients) (34). A significant number of patients also experienced osteomyelitis, sepsis, cellulitis, and meningitis. Patients with CGD suffer from a variety of non-infectious hyper-inflammatory complications, most prominently in the gastrointestinal and genitourinary tracts, as well as lupus-like syndromes (34, 40). The underlying reasons for the hyperinflammatory disorders associated with CGD remain a poorly understood aspect of the disease. In addition to their role in microbial killing, ROS are also critical signaling molecules involved in the regulation of various physiological activities, including metabolism, cell death, and oxidative signaling (Reviewed in (46). Both impaired apoptosis and removal of apoptotic cells, a process called efferocytosis, have been implicated in CGD (47-49). This may cause an imbalance in homeostasis, with associated lysis of dying cells and release of intracellular proteases and oxidants that cause exaggerated inflammation. Furthermore, CGD leukocytes express higher levels of pro-inflammatory mediators both at resting state or upon Toll-like receptor (TLR) stimulation compared to controls (50, 51), which is associated with elevated expression of nuclear factor (NF)-κB and ERK1/2 MAP kinase signaling (50). Overall, the role of ROS in modulating inflammation is likely multifaceted and extends far beyond a simple predisposition to infections.    9 Since CGD was first described in 1954 (52) it has progressed from a disease of childhood fatality to one of effective management and long-term survival. Current effective management of CGD includes prophylaxis for infectious disease with interferon (IFN)-γ (53, 54) and appropriate antimicrobials, most often with the antibiotic trimethoprim-sulfmethoxazole (55) and antifungal agent itraconazole (56), as well as treatment for acute infections when needed. Inflammatory complications such as granulomas are commonly treated with corticosteroids (57). The introduction of potent antimicrobials, particularly the antifungal azole agents, is attributed to the profound decrease in mortality in CGD (54, 58). However despite significant progress, patients still develop serious infections at a rate of approximately 0.15 to 0.3 per year (58). HCT with full myeloablation or nonmyeloablative conditioning is currently the only cure for CGD (59). Although HCT is an attractive option, the procedure continues to be associated with a high risk of morbidity and mortality that limits its use to some centers.  1.1.3 Cystic fibrosis  CF is the most common, lethal genetic disease that affects the Caucasian population, with an estimated prevalence of approximately 1:2500-3200 live births in North America (60). CF is an autosomal recessive disease caused by mutations in the cystic fibrosis transmembrane conductance regulator (CFTR) protein, which functions mainly as a cAMP-regulated chloride ion channel in epithelial and blood cells (61, 62). More than 1500 CFTR mutations have been identified, among which approximately 70% are caused by a deletion of phenylalanine at codon 508 (Phe508del or ΔF508). The phe508del mutation causes a CFTR trafficking defect that results in ubiquitination and degradation of the protein in the endoplasmic reticulum and Golgi of the cell (60). Different mutations in CFTR result in variable function of the protein and lead to different clinical presentation and prognosis.    10  CF is characterized by airway disease thought to be caused by poor chloride and sodium ion transport that results in reduced volume of airway surface fluid. This inhibits mucociliary clearance within the lung that is crucial in clearing inhaled bacteria and preventing colonization of the lung. Patients suffer from buildup of mucus in the lungs that contributes to chronic infections and inflammation with resulting lung fibrosis. In addition to lung pathology, mutations in CFTR cause disease in other epithelial-lined organs. Patients often experience gastro-oesophagael reflux, pancreatic insufficiencies, intestinal obstruction, celiac disease, Crohn’s disease, and constipation (63). Exaggerated inflammation is a common theme of CF, both in the presence and absence of high bacterial loads (64, 65). Abnormally high concentrations of inflammatory mediators and regulators such as interleukin (IL)-6, IL-8, and NF-κB have been found in CF cell lines, patients with CF, and in uninfected ex-vivo tissue samples (66-68).   Lung disease is an early manifestation of CF and the major cause of morbidity and mortality, causing approximately 80% of deaths. Infants develop viral infections that are more likely to be symptomatic compared to healthy children (69) and acquire bacterial infections early in life. Bacterial infections often result in long-term or permanent colonization of the lung, likely due to defective CF mucociliary clearance and an ineffective host defense (70). Infections are caused by Staphylococcus aureus, including methicillin-resistant strains (MRSA) and Haemophilus influenzae as well as opportunistic pathogens such as Pseudomonas aeruginosa and the Bcc. Less commonly, the late stages of lung disease may include Stenotrophomonas maltophilia, Streptococcus anginosus, Burkholderia gladioli, Achromobacter xylosoxidans, Aspergillus spp, and nontuberculosis mycobacteria (71).  P. aeruginosa is a major contributor to lung function decline in CF. A recent study in the United States found that 72.3% of CF children under the age of 3 were infected with culturable    11 P. aeruginosa (72). According to the 2011 Canadian and United States CF patient registries, 44% and 50.6% of CF patient sputum is culture positive for P. aeruginosa, respectively (73, 74).  S. aureus infections are found in 50% of Canadian CF patients and 67.9% of US patients. While the incidence of Burkholderia spp is far less than other pathogens in CF (occurring in 2.6% of the Canadian and 5% of US population), several species can cause chronic, severe respiratory tract infections and colonization with Burkholderia is associated with poor prognosis (75).  Lung disease in children with CF is characterized by recurrent infections that are often treatable with antibiotics. Later in life, infections become chronic and bacteria, commonly P. aeruginosa with or without co-infection with other pathogens, form biofilms in the lung that are difficult to treat and associated with an accelerated decline in pulmonary function (70). Infections stimulate a persistent inflammatory response, and neutrophil and bacterial products lead to epithelial surface damage, progressively impairing airway conductance. Bacterial infection persists and occasionally exacerbates lung disease, requiring intensive treatment (60).  Maintenance for CF involves a combination of strategies to decrease morbidity and mortality. Nutritional repletion regimens using pancreatic enzyme supplements and high caloric diet enable patients to thrive. Relief of airway obstruction is achieved through physiotherapy and aerobic exercise together with therapies to improve sputum clearance including inhaled dornase alfa (recombinant deoxyribonuclease) and hypertonic saline (76, 77).  Lung infections are treated using appropriate oral or intravenous antibiotics. Mild exacerbations are commonly treated with inhaled tobramycin or oral antibiotics, including amoxicillin-clavulanate, trimethoprim-sulfamethoxasole, and cephalexin, whereas more severe exacerbations are typically treated intravenously (77, 78). Ultimately, lung transplantation is the last-line therapeutic for patients with end stage lung disease and can offer a true survival benefit when available (79).    12 1.2 Phagocyte biology When infectious agents pass the body’s epithelial barrier, they encounter a variety of potent antimicrobial effectors. These include innate immune cells centered on the activity of dedicated phagocytes: neutrophils, monocytes, and macrophages. Phagocytes play a crucial role in regulating inflammation and infection and are important first responders to pathogens. Their main function is ultimately to ingest pathogens to control infections. Phagocyte killing can be accomplished by the innate arm of immunity through the fusion of toxic lysosomal granules to the phagosome as well as through the adaptive arm of immunity by acting as antigen presenting cells (APCs) that stimulate T and B cells.  Neutrophils are the most abundant leukocyte in humans and are early responders to sites of infection. Upon detection of microbes, neutrophils are recruited from the peripheral blood and infiltrate the tissue. They are extremely efficient at fighting pathogens using a combination of ROS and antimicrobial peptides and proteases from intracellular granules. Neutrophils can also release highly inflammatory neutrophil extracellular traps (NETs) composed of DNA, histones, and granule proteases that are effective at binding to and killing microbes (80). Activated neutrophils recruit other immune cells to the site of infection by releasing chemotactic mediators and can influence macrophage differentiation to an anti- or pro-inflammatory phenotype (81). Neutrophils are short-lived cells that are programmed to undergo apoptosis upon phagocytosis of microorganisms as a method of clearing tissue of spent cells and digested microbes (82). Apoptotic neutrophils are generally phagocytosed by tissue-resident macrophages in a non-inflammatory process called efferocytosis (5). Monocytes are an abundant and heterogeneous cell type in the peripheral blood that migrates to sites of infection or injury, albeit slower and in lower numbers compared to    13 neutrophils (83). Monocytes express a large repertoire of PRRs that detect conserved PAMPs including TLRs on their cell surface and intracellular NOD-like receptors. They are extremely efficient phagocytes and have a prominent role in producing inflammatory mediators including ROS and in presenting antigens and activating the adaptive arm of immunity (84). Unlike neutrophils, whose contents are mostly pre-determined, monocytes are capable of tremendous de novo synthesis of granule proteins, cytokines, and chemokines (83). Monocytes also retain the ability to proliferate after leaving the bone marrow and can differentiate into a variety of tissue resident cells including macrophages, dendritic cells, and osteoclasts (85, 86). Compared to neutrophils, monocytes exhibit longer survival in the blood, which provides some protection to the host during neutropenia caused by infection or chemotherapy (83).  Macrophages behave as a double-edged sword in maintaining host homeostasis: they can phagocytose pathogens and promote innate and adaptive immune responses and can also contribute to the repair of injured tissue during inflammation (87). Macrophages can assume at least two different phenotypes: M1 (classical) macrophages are inflammatory and produce a variety of pro-inflammatory cytokines, ROS, and have strong antimicrobial properties, whereas M2 (alternative) macrophages exhibit a regulatory phenotype that produce anti-inflammatory mediators and are important in tissue repair and wound healing (88, 89). M1 macrophages are usually associated with acute infection control against intracellular bacteria, however an M1-M2 switch may occur during chronic infection and provide protection against uncontrolled inflammation that can lead to sepsis (90).  Macrophages are abundant in many tissues and are proficient at phagocytosing pathogens and containing bacteria until the adaptive immune system is initiated (91). Although they are not as effective as neutrophils at inducing a potent ROS response, macrophages are longer lived, can    14 produce a larger repertoire of inflammatory mediators, and can act as APCs in tissues (91). Like monocytes, macrophages posses a large variety of PRRs that upon binding to PAMPs trigger downstream signaling through transcription factors including NF-κB and IRF3 that leads to the production of inflammatory mediators (92). Furthermore, macrophages can directly kill bacteria by phagocytosing and trafficking them through the phagosomal maturation pathway. This antimicrobial pathway consists of a lowering the pH of the bacteria-containing phagosome and accumulation of antimicrobials and proteolytic enzymes, assembly of NADPH oxidase for ROS production, and eventual fusion with destructive lysosomes (93).  1.3 Cell death 1.3.1 Apoptosis  Apoptosis is an innate form of programmed cell death that allows organisms to eliminate unwanted cells in a non-inflammatory fashion. Apoptosis occurs naturally during development and aging as a homeostatic method of maintaining appropriate cell populations (94). It can also act as a defense mechanism for damaged cells following exposure to pathogens or toxic agents (95, 96).  Apoptosis can occur through two main mechanisms depending on the cell stimulus. The extrinsic pathway of apoptosis recognizes external stimuli through death receptors on the cell surface and leads to the cleavage and activation of caspase-8, whereas the intrinsic/mitochondrial pathway recognizes intracellular stimuli and activates caspase-9 (Reviewed in (97)). The CTL-mediated perforin-granzyme pathway also induces apoptosis by activating effector caspases and is discussed below. Although the extrinsic and intrinsic pathways were traditionally considered    15 distinct and independent, there is evidence that there is crosstalk between pathways and that molecules from one pathway can influence the other (98, 99). Extrinsic apoptosis signaling pathways are activated by transmembrane death receptors on the cell surface. Death receptors are members of the tumor necrosis factor (TNF) receptor family and are responsible for transmitting death signals from outside the cell to intracellular signaling pathways through their cytoplasmic domains (100). The best-described ligands and corresponding death receptors are Fas ligand and Fas receptor, TNF-α and TNFR1, and TRAIL and DR4/DR5 (101). When death receptors bind to their specific ligand, they cross-link and activate their cytosolic domains. The adaptor proteins TRADD and/or FADD are recruited to active cytosolic domains and recruit caspase-8, forming what is known as the death-inducing signaling complex (DISC) (102-104). Aggregation and dimerization of procaspase-8 leads to its auto-activation and caspase-8 proceeds to activate downstream effector caspases (105, 106). The intrinsic pathway of apoptosis can be triggered by a range of diverse intracellular stimuli that lead to permeabilization of the mitochondrial membrane. Permeabilization of the outer mitochondrial membrane allows the release of mitochondrial proteins from the intermembrane space into the cytosol, including cytochrome c, Smac/DIABLO, and HtrA2/Omi. In the cytosol, cytochrome c, Apaf-1, and pro-caspase-9 form an apoptosome complex. Pro-caspase-9 is catalytically cleaved to active caspase-9, which acts as an initiator caspase for downstream signaling (107-109). Smac/DIABLO and Omi/HtrA2 facilitate caspase activation by neutralizing numerous endogenous caspase inhibitors called inhibitor of apoptosis proteins (IAPs) by binding through their N-terminal IAP-binding motifs and displacing target caspases (110). In addition to antagonizing IAPs, HtrA2/OMI acts as a protease and can degrade a number of IAPs in vitro, including XIAP, cIAP1, cIAP2, and Apollon to promote apoptosis in a caspase-   16 independent fashion (111). A second group of pro-apoptotic proteins are released from the mitochondria late in apoptosis and include AIF, endonuclease G, and CAD. These proteins translocate to the nucleus and induce DNA fragmentation and chromatin condensation (112). A number of proteins regulate intrinsic apoptosis, notably proteins of the Bcl-2 family, which function by altering the permeability of the outer mitochondrial membrane to govern the release of cytochrome c (113). The intrinsic and extrinsic pathways of apoptosis converge on the same terminal effector caspases, mainly caspase-3, caspase-7, and caspase-6 (99, 114). Effector caspases selectively cleave cellular proteins that lead to the morphological and biochemical features associated with apoptosis (115). Morphologically, apoptotic cells shrink in size and their chromatin condenses and gives rise to uniformly dense nuclei (94). Apoptotic cells undergo degradation of hundreds of cytoskeletal and nuclear proteins with extensive protein crosslinking, hydrolysis of DNA, fragmentation of the Golgi, endoplasmic reticulum (ER), and mitochondrial networks, and membrane blebbing leading to the formation of apoptotic bodies (Reviewed in (116)). A hallmark of apoptosis is the surface exposure of ligands that distinguish apoptotic cells from viable cells for recognition and engulfment of cells by phagocytes (112). The best-studied and most universal “eat me” signal is phosphatidylserine (PS) and its oxidized forms, which face inward in the lipid bilayer of healthy cells but are externalized to the outer layer of the plasma membrane following activation of effector caspases (117, 118). Other eat-me signals include changes in the glycosylation of glycoproteins and lipids and the binding of collectins such as mannose-binding lectin and C1q of the complement system to the apoptotic cell surface (119-121).     17 Intact apoptotic bodies expressing eat-me signals are engulfed by neighboring phagocytes through efferocytosis and are degraded with the phagolysosome (122). Efferocytosis is considered an anti-inflammatory process that has a profound influence on resolution of inflammation (123, 124). Macrophages produce IL-10, TGF-β, and prostaglandin E2 (PGE2) during the engulfment of apoptotic cells that inhibit inflammatory mediator production (123, 125, 126). Macrophages recognizing PS on apoptotic cells can accelerate the resolution of LPS-induced lung inflammation in mice in a TGF-dependent manner (127). Interestingly, engulfment of the apoptotic cell is not necessarily required for the release of anti-inflammatory mediators since the presence of apoptotic cell membrane alone could diminish the production of IL-12 by macrophages exposed to LPS in vitro (128) and PS vesicles can stimulate the production of TGF-β (127). Efferocytosis of intact apoptotic bodies limits the likelihood that they will progress to secondary necrosis and release their cytotoxic and/or imunogenic cellular constituents. Necrotic cells expose self-antigens that lead to inflammation and can potentially induce a break in self-tolerance. Indeed, impaired clearance of apoptotic cells has been described in both autoimmune and chronic inflammatory diseases (5, 129, 130). 1.3.2 Granule-mediated cytotoxicity The granule-mediated cytotoxic pathway is the primary means of eliminating abnormal cells arising from intracellular pathogens or malignant transformation. These target cells are recognized by CTLs and NK cells, which represent the adaptive and innate immune systems, respectively. NK cells act as the first line of defense against intracellular viruses and malignant cells, by detecting abnormal or missing MHC class 1 molecules on the surface of infected cells. Unlike NK cells, which do not require priming for cytotoxic activity, CTLs require T cell    18 receptor (TCR) activation by antigen-presenting MHC-1 molecules and cytokine activation such as by IL-12 produced from professional APCs (131). Despite these fundamental differences, both NK cells and CTLs engage the same granule-mediated cytotoxicity pathway to initiate target cell apoptosis (132).  Secretory (cytotoxic) granules of CTLs and NK cells are composed primarily of perforin and serine proteases such as granzyme, which when delivered to a target cell can induce rapid apoptosis in as little as 20 minutes (Figure 1.1) (13, 133). Perforin mediates the delivery of granzymes released into the intracellular space into the cytosol of target cells, where granzymes cleave specific substrates involved in activating apoptosis, such as BID and effector caspases (134, 135). The mechanism of perforin has been the focus of much debate. It was classically thought that perforin formed pores in the target cell that allowed passage of granzyme into the target cell, however more recent studies suggest that granzymes are endocytosed by the target cell and that perforin aids in their escape to the cytosol (136). The protein Munc13-4 and its partner proteins syntaxin11 and Munc18-2 (in which mutations cause FHL3, FHL4, and FHL5, respectively), play a role in the trafficking of secretory granules. CTL/NK cells from patients with these mutations fail to deliver functional perforin and granzyme to the immune synapse. Munc13-4 plays a crucial role in priming granules that dock to the plasma membrane for vesicle-membrane fusion and as a result, CTLs from FHL3 patients fail to degranulate (24). Mutations in syntaxin11 and Munc18-2 result in defective degranulation in NK cells but not CTLs, but the underlying mechanism remains unknown (26, 27, 137). Interestingly, mutations in granzyme do not lead to HLH despite the protein’s crucial role in granule-dependent cytotoxicity. This is likely due to the redundancy of multiple granzyme genes found in mammals (135).     19                   Figure 1.1 Mechanism of cytotoxic granule trafficking in CTL/NK cells  (© 2012 G.E. Janka, adapted with permission (13))  1.3.3 Necrosis Necrosis was formerly considered the alternative to apoptosis: an inflammatory energy-independent form of death characterized by cell swelling, rupture of the plasma membrane, and the release of cellular constituents into the interstitial tissue. However, since necrosis is not a set of biochemical processes occurring during cell death but rather refers to morphological alterations secondary to a number of cell death pathways, many have opted to use the term “primary necrosis” to describe the toxic uncontrolled cell death marked by cellular swelling     20 (138, 139). Some microorganisms can induce primary necrosis by escaping from phagosomes and/or secreting toxins into the cytosol that disrupt membrane integrity (140). Unlike apoptotic cells, primary necrotic cells release their toxic contents before being recognized by phagocytes. Cells that undergo a programmed cell death pathway such as apoptosis but are not cleared by efferocytosis undergo “secondary necrosis”, which exhibits similar morphological features and inflammatory outcomes as primary necrosis (141).  The main morphological features that characterize necrosis include cell swelling, formation of cytoplasmic vacuoles, ruptured mitochondria and lysosomes, disrupted organelle membranes, and rapid disruption of the cell membrane (94, 139). Various danger-associated molecular patterns (DAMPs or alarmins) are passively released by necrotic cells, notably ATP, high mobility box protein B1 (HMGB1), certain heat shock proteins, genomic DNA, and uric acid that stimulate PRRs on innate immune cells (142-144). The activation of PRRs on APCs can activate T cells, thus alarming the adaptive immune system (145). Overall, necrosis causes direct cell damage by the release of cytoplasmic contents and activates both the innate and adaptive immune systems, causing high levels of inflammation.  1.3.4 Alternative modes of programmed cell death While apoptosis and necrosis are the traditional forms of cell death, a number of alternative programmed cell death mechanisms can be initiated in response to various stimuli, notably to pathogens (146). For most alternative cell death mechanisms, a defining feature is the lack of observable hallmarks of apoptosis, including the loss of mitochondrial membrane integrity, apoptotic caspase activation (exception is caspase-1), membrane blebbing, DNA fragmentation within the nucleus, and response to pan-caspase inhibitors such as z-vad-fmk. Some of the more common alternative forms of phagocyte cell death are discussed here.    21 Autophagy is a catabolic process whereby unwanted or damaged cytosolic proteins are degraded (147). Autophagy-induced cell death occurs when the autophagic capacity is overwhelmed. Autophagy occurs under conditions of starvation, ER stress, or in the presence of damaged organelles with the absence of markers of apoptosis (148). Classic features of autophagy include the activation of autophagy-related genes that mediate the formation of autophagosomes, which are double membrane vesicles that engulf self and foreign components for bulk degradation (149, 150). Autophagosomes proceed to fuse with lysosomes to form autolysosomes, where their contents are degraded (151). Autophagy can also target intracellular bacteria for lysosomal degradation in a form of autophagy termed xenophagy (152, 153). Autophagosomes can form around phagosomes containing bacteria or can engulf bacteria that have escaped from vacuoles to the cytoplasm and target them for degradation, acting as a back-up defense mechanism against intracellular microorganisms (154, 155). The recognition of bacteria can also trigger signaling cascades that lead to protein ubiquitination and antigen-presenting mechanisms, thus playing an integral role in both innate and adaptive immunity (156). Autophagy is well described in macrophages and while evidence exists to support autophagy in neutrophils (157-159), there is a lack of published reports (155). Pyroptosis is a caspase-1-dependent inflammatory form of programmed cell death triggered by various pathogens and non-infectious stimuli such as cholesterol crystals, uric acid, and anthrax lethal toxin (160-162). Caspase-1 is activated by NOD-like receptors (NLRs) in the cytosol in response to intracellular stimuli (163). It is responsible for processing IL-1β and IL-18 precursors (pro-IL-1β and pro-IL-18) to mature, highly inflammatory cytokines. Caspase-1 activation also mediates pyroptosis, characterized by caspase-1-dependent pore formation in the plasma membrane that results in water influx, swelling, and rupture of the cell with release of    22 inflammatory contents (164). Caspase-1 activation helps to clear pathogens including Salmonella (165), Shigella (166), Francisella (167), and Listeria (168) in vivo. This effect is not due to inflammatory cytokines alone since mice deficient in caspase-1 are more susceptible to several pathogens compared to mice deficient in both IL-1β and IL-18 (167, 169, 170). Pyroptosis is thought to benefit the host by alarming the immune system and restricting pathogen replication in vivo (161). One study provided evidence that the role of pyroptosis is to allow the bacteria to escape from lysed macrophages to expose it to recruited neutrophils, which are much more efficient at killing bacteria (170). Some studies have shown that DNA fragmentation occurs in response to some pyroptosis-inducing pathogens, however the pathway leading to DNA cleavage is unique and the nuclei of cells pyroptotic cells do not undergo fragmentation as is seen during apoptosis (164, 171). Pyroptosis is described in macrophages and dendritic cells, however its relevance in neutrophils, which do not express some NLRs, appears doubtful (161).  Neutrophils and eosinophils can trigger a unique form of cell death called NETosis that can be triggered upon pathogen interaction (172). In addition to their ability to release ROS and cytotoxic proteases and antimicrobial peptides within intracellular granules, neutrophils can also produce highly inflammatory NETs (80). NETs are composed of multiple cellular components, including nucleic acids bound to histones and granule proteases that are very effective at binding to and killing a wide variety of microbes (80, 173). NET release can occur through rupture of the neutrophil plasma membrane resulting in cell death (lytic NETosis) or through nuclear budding and vesicular release, which allows the neutrophil to continue to function (vital NETosis) (174). One defining difference between lytic NETosis and pathogen-induced lysis (primary necrosis) is the disintegration of nuclear and granular membranes before plasma membrane rupture during NETosis that allows chromatin to mix with the granule’s antimicrobial machinery and greatly    23 enhance the DNA’s killing potential (175). However, both cell death processes result in the release of toxic cellular contents. Vital NETosis may in fact be the desired outcome of NET release because it kills microbes via the release of histone-coated DNA while allowing the cell to continue to undergo leukocyte recruitment, chemotaxis, and phagocytosis (176, 177). Further investigations are required to differentiate lytic NETosis, vital NETosis, and primary necrosis and to draw physiological relevance (174).  1.4 The Burkholderia cepacia complex  The Bcc comprises a group of at least 18 genetically distinct but phenotypically similar species of opportunistic pathogens that cause lung infections in immunocompromised patients, specifically those with CF or CGD (178). Species of the Bcc are also recognized as an emerging group of nosocomial pathogens causing bacteremia and their spread in hospital settings is associated with cross-transmission, frequent pulmonary procedures, and central vein access (179). Table 1.2 outlines the various Bcc species and their clinical relevance. Pseudomonas cepacia was first identified in 1949 by Dr. Walter Burkholder as the cause of onion skin rot (180). Subsequently renamed Burkholderia, bacteria from this genus were later found have a number of ecologically beneficial properties through colonizing plant rhizospheres and promoting growth, producing antimicrobial compounds that protect plants, and degrading pollutants (181-183). Their large and complex genome, consisting of at least two large chromosomal replicons and one or more plasmids containing multiple types of insertion sequences and genomic islands allows for remarkable ecological and nutritional versatility that allows it to survive in diverse environmental niches (181).   The adaptability of Bcc bacteria likely contributes to their ability to act as opportunistic pathogens in humans. Bcc species can survive in inhospitable environments and have been    24 identified in disinfectants, intravenous solutions, and contaminated medical devices, leading to nosocomial infections and septicemia (184, 185). Their ability to form biofilms and intrinsic resistance to host antimicrobials, together with other virulence traits, allows the Bcc to cause severe infections in immunocompromised conditions, such as those with CF and CGD. Bcc infections in CF and CGD usually initially present as lung infections characterized by excessive infiltration of neutrophils. In CF, Bcc infections are most often chronic and caused by persistence of a single strain that is difficult to eliminate. Bcc are often acquired by older CF patients already colonized by other pathogens, notably P. aeruginosa (186) and are associated with early fatality compared to patients without Bcc (74). B. cenocepacia and B. multivorans are capable of causing “cepacia syndrome”, where the bacteria escape from the lung to the blood stream and causes sepsis in patients with CF (187, 188). In CGD, infections with Bcc are most often acute and are more susceptible to antibiotics compared to those in CF. This could be due to the high level of microbial colonization and biofilm formation in the CF lung that is not seen in CGD (9). Furthermore, the acute nature of infections in CGD does not require long course antibiotics that select for resistant strains. Bcc infections in CGD are either cleared with appropriate antibiotics or are rapidly fatal. Re-infections are common and can involve multiple distinct strains (9). In CGD Bcc can cause necrotizing pneumonia and sepsis and is the leading bacterial cause of death (34). While B. cenocepacia and B. multivorans are responsible for the majority of Bcc infections in CF and CGD, the spectrum of Burkholderia species that infect CGD patients appears to be broader (9, 189). B. gladioli has also emerged as an important species of Burkholderia in CGD, though it is not considered part of the Bcc (9).      25 Table 1.2 Species of the Burkholderia cepacia complex and their clinical relevance Species Relevant characteristics  Reference B. cepacia Limited prevalence in CF, important in CGD, evidence of transmission, causes onion rot (8, 9, 190) B. multivorans Major pathogen in CF and CGD, evidence of transmission, epidemic strains described in CF (9, 189-192) B. cenocepacia Major pathogen in CF and CGD, highly transmissible, epidemic strains described in CF (8, 9, 189, 191, 192)  B. stabilis Rare in CF  (189) B. vietnamiensis Limited prevalence in CF, important in CGD, plant growth promoter (9) B. dolosa Almost exclusively cultured in CF, evidence of transmission, some epidemic strains described (189, 190) B. ambifaria Rare in CF, important in CGD  (9, 189) B. anthina Rare in CF  (193) B. pyrrocinia Rare in CF (189, 193) B. ubonensis Only one clinical strain (nosocomial infection) and one environmental strain described (194) B. latens Rare in CF (194) B. diffusa Rare in CF (194) B. arboris Rare in CF (194) B. seminalis Rare in CF (194) B. metallica Rare in CF, identified in CGD  (9, 194)  B. contaminans Rare in CF (195) B. lata Rare in CF (195) B. pseudomultivorans Recently described species, 9 isolates from CF sputum (196)  1.5 Evasion of host defenses by the Bcc 1.5.1 Evasion of ROS and oxidative killing Oxidative killing by host phagocytes is crucial for potent immunity against fungi and bacteria. Although the Bcc is susceptible to ROS, it does exert multiple mechanisms to avoid oxidative killing. Species of the Bcc produce periplasmic superoxide dismutase (SOD) and catalase or a bifunctional catalase-peroxidase that catalyzes the breakdown of toxic superoxide    26 and hydrogen peroxide produced during the oxidative burst (197, 198). Several species of Bcc possess a melanin-like pigment that quenches extracellular superoxide and protects it from oxidative killing in the macrophage phagosome (199, 200). Furthermore, Bcc produces at least four types of exopolysaccharides with roles in adhesion, evasion of host defenses, and resistance to antimicrobials (201). Cepacian is the predominate exopolysaraccharide produced by Bcc and cepacian produced by B. cenocepacia both inhibits neutrophil chemotaxis and scavenges ROS (202). Some isolates of B. cenocepacia express heme-binding proteins in their outer membrane that may enable the bacteria to withstand oxidative stress (203). B. cenocepacia biofilms also contain persister cells that survive antibiotic treatment by downregulating the TCA cycle to avoid the formation of toxic ROS byproducts (204). 1.5.2 Resistance to non-oxidative killing Phagocytes engage a number of non-oxidative killing strategies upon pathogen encounter that are especially important in the absence of ROS. CGD pathogens such as the Bcc posses a number of tools to combat non-oxidative killing that contribute to their pathogenesis in the disease. All species of the Bcc possess intrinsic resistance to aminoglycosides and a number of other antibiotics. Aminoglycosides are broad-spectrum antibiotics composed of polar aminated sugars that pass through the membrane of Gram-negative bacteria and bind to and impair the 30s subunit of prokaryotic ribosomes (205). Bcc species achieve aminoglycoside resistance primarily by altering their membrane permeability and utilizing efflux pumps to remove aminoglycosides from the cytosol. LPS on the outer membrane of Gram-negative bacteria is composed of 3 components: lipid A, oligosaccharides, and a polymer of glycan repeats called O-antigen (206). Lipid A contains negatively charged phosphates that bind electrostatically to positively charged aminoglycosides and antimicrobial (cationic) peptides produced by the innate immune system    27 that kill the target cell through a number of mechanisms (207). The Bcc modifies its LPS by attaching a 4-amino-4-deoxy-L-arabinose (Ara4N) residue to phosphate groups in lipid A and an inner core oligosaccharide residue to dramatically decrease membrane permeability (208). Bcc bacteria also integrate pentacyclic triterpenoids (surrogates for eukaryotic sterols) called hopanoids in their membrane that regulate the stablility, fluidity, and outer membrane permeability and contribute to resistance to various antibiotics, including polymixin (209, 210). Bcc bacteria also have a number of efflux pumps in their outer membrane that efficiently remove aminoglycosides and other antibiotics from the cell (211, 212).  Resistance of the Bcc to other antibiotics is achieved using a number of mechanisms. Multiple strains of Bcc have inducible β-lactamases that degrade penicillin and expanded-spectrum cephalosporins and aztreonam at lower levels (213). Some strains of Bcc obtain trimethoprim resistance by producing a dyhydrofolate reductase, the target of trimethoprim, that is not susceptible to the antibiotic (214). Bcc can form biofilms in vitro and produce acyl-homoserine lactones involved in cell-to-cell communication (215). Bcc biofilms exhibit increased resistance to ceftazidime and ciprofloxacin compared to planktonic Bcc (216), reduced drug penetration, and lower growth rate of sessile cells (217).  Several species of the Bcc produce extracellular proteases that proteolytically degrade host antimicrobials. A type II secretion system secretes the zinc metalloproteases ZmpA and ZmpB into the macrophage cytosol, where they cleave the potent antimicrobial peptides LL-37 and definsin-1, respectively (218, 219). Both metalloproteases can cleave host elafin and abolish its anti-protease activity (218). ZmpB also has proteolytic activity against alpha-1 proteinase inhibitor, α2-macroglobulin, type IV collagen, fibronectin, lactoferrin, transferrin, and immunoglobulins, indicating that it is a broad-specificity protease capable of degrading    28 substrates pertinent to host defense and host tissue integrity (220). Mutants in either ZmpA or ZmpB resulted in less virulence in a rat agar bead model of infection (220, 221). In addition to the abovementioned mechanisms of evading the host immune system, the Bcc also produce a variety of other virulence factors including cable pili and adhesins, flagella required for cellular invasion, one type II, one type III, two type IV, and one type VI secretion system, four types of iron-chelating siderophores, and a number of degradative proteins such as lipases and haemolysins (181, 222, 223). Many of these factors are not produced by all species of Bcc and their roles in human disease are not fully elucidated. Despite this robust arsenal of host evasion strategies available to the Bcc, it is still non-pathogenic in healthy humans. Nevertheless, a combination of these virulence factors clearly enables the Bcc to cause substantial morbidity and mortality in immunocompromised patients.   1.6 Interactions of the Bcc with phagocytes 1.6.1 Survival and trafficking within macrophages The ability of the Bcc to survive intracellularly and subvert macrophage killing has been investigated using multiple strains and macrophage cell types. Earlier studies with modified gentamicin-protection assays suggested that survival occurred with little to no replication depending on the bacterial isolate and macrophage type used (224-227). However, another study showed that the Bcc could replicate up to 3 log-fold within 24 hours in various types of macrophages (228). The gentamicin protection assay, which relies on the poor ability of gentamicin to enter eukaryotic cells, is the standard for assessing intracellular replication by bacteria (229), and the intrinsic resistance of B. cenocepacia to gentamicin has likely contributed to these confounding results. Recent studies have examined the intramacrophage growth of B.    29 cenocepacia using aminoglycoside-sensitive strains in the standard gentamicin-protection assay (230). These have shown that B. cenocepacia replicates within two macrophage cell lines but not in macrophages derived from the commonly used THP-1 human monocyte cell line (230, 231). Thus the ability of the Bcc to replicate intracellularly within primary human macrophages remains unclear. Trafficking of Bcc within macrophages has been thoroughly investigated. Engulfed bacteria reside within Bcc-containing vacuoles (BcCVs) that co-localize with EEA-1, a marker for early endosomes (224). BcCVs containing live bacteria delay phagosomal maturation and acidification and inhibit fusion with lysosomes for up to 6 hours (224). The delay in phagosomal maturation is required for survival of B. cenocepacia and mutants unable to mediate maturation delays are cleared from macrophages and are avirulent in a rat agar bead model of infection (232-235). BcCVs also delay NADPH oxidase formation on the phagosomal membrane and inhibit associated superoxide production (236). Intracellular growth in U937 cells was associated with escape from autophagosomes in the endocytic pathway to the cytoplasm, where the bacteria co-localized with the ER marker calnexin and replicated (227). Other groups however have not been able to demonstrate co-localization of calnexin and BcCVs in infected RAW264.7 macrophages (224). Intramacrophage survival of B. cenocepacia is dependent on a plasmid-encoded type IV secretion system, though the secreted effector molecules remain unknown (227).  1.6.2 Neutrophils and the Bcc The majority of literature involving the interactions of phagocytes and the Bcc is focused on macrophages and little is known about the interactions between Bcc and human neutrophils. The Bcc exhibits a number of mechanisms to evade oxidative killing, as described above.    30 Exopolysaccharides produced by B. cenocepacia are able to inhibit neutrophil chemotaxis in addition to their role in scavenging ROS (202). Exposure to live B. cenocepacia induces cell death in primary neutrophils and necrosis in a significant portion of CGD neutrophils (237). This effect was also seen when ROS were inhibited with diphenyleneiodonium, a NADPH oxidase inhibitor (237). However, the specific mechanism underlying B. cenocepacia-induced cell death in CGD neutrophils remains unknown. 1.7 Hypothesis and overarching aims Primary immunodeficiencies can cause aberrant inflammation and chronic or recurrent infections with uncommon pathogens that normally do not cause infections in healthy individuals. I hypothesize that defects in phagocyte antibacterial mechanisms and programmed cell death lead to inflammation and pathogen survival. The overall goal of this research is to gain a better understanding of the human immune mechanisms utilized to prevent infection with opportunistic pathogens. This research will also provide insight into the dysregulated systems in immunocompromised patients that cause disease and inflammation. Novel insight into the molecular pathways involved in controlling opportunistic pathogens may lead to the design of better therapeutics to treat life-threatening infections in patients.     31 Chapter 2: Burkholderia cenocepacia induces rapid neutrophil apoptosis in the absence of reactive oxygen species that promotes a pro-inflammatory macrophage response.  2.1 Introduction  Chronic granulomatous disease (CGD) is a primary immunodeficiency that affects approximately 1 in 200,000 people and results from genetic defects in components of the phagocyte nicotinamide adenine dinucleotide phosphate (NAPDH) oxidase (34, 40). This renders patients’ phagocytes unable to produce toxic reactive oxygen species (ROS) crucial for potent antimicrobial activity. CGD is characterized by multiple recurrent infections and various immune complications including granuloma formation. The Burkholderia cepacia complex (Bcc) comprises a group of opportunistic bacterial pathogens that are particularly problematic in the respiratory tract of patients with CGD and cystic fibrosis (CF) (178). Species of the Bcc possess intrinsic resistance to various antibiotics and cationic antimicrobial peptides and can therefore persist in the absence of oxidative killing, causing infiltration of polymorphonuclear neutrophils (PMN) and excessive inflammation (178).  Neutrophils are responsible for destroying invading microorganisms using a combination of cationic peptides, ROS, and the release of antimicrobial neutrophil extracellular traps (NETs) composed of nucleic acids bound to histones and granule proteases (80, 173, 238). After phagocytosing pathogens, neutrophils ideally undergo a form of programmed cell death called apoptosis (239). Apoptosis is a non-inflammatory form of programmed cell death that involves activation of caspases, cytoplasmic shrinkage, nuclear condensation and fragmentation, surface exposure of phosphatidylserine (PS) on the plasma membrane, membrane blebbing, and the formation of apoptotic bodies (115). Autophagy-induced cell death has emerged as an alternative    32 programmed cell death mechanism that lacks the classical features of apoptosis, particularly caspase activation (147). Autophagy is described as a catabolic process whereby damaged cytosolic proteins or intracellular microorganisms either in phagosomes or that have escaped to the cytosol are degraded by engulfment in double-membraned autophagosomes that proceed to fuse with lysosomes (149, 155). Autophagy-induced cell death occurs when the autophagic capacity is overwhelmed.  Non-inflammatory programmed cell death provides a method of controlled recycling of dying cells, however alternative inflammatory forms of cell death can occur upon pathogen interaction. Primary necrosis is a cell death mechanism that involves cell swelling and lysis. Some microorganisms can induce necrosis by escaping from phagosomes and/or secreting toxins into the cytosol that disrupt membrane integrity (140, 240). This allows pathogens to escape phagocyte killing and disseminate. Other pathogen-induced forms of cell death associated with the loss of membrane integrity include NETosis, characterized by release of antimicrobial NETs, and pyroptosis, characterized by caspase-1 activation and the production of inflammatory mediators interleukin (IL)-1β and IL-18 (170, 172). While NETosis is well described in neutrophils, the relevance of pyroptosis remains unclear. Both inflammatory forms of cell death are in part defined by the absence of the hallmarks of apoptosis, particularly caspase activation and exposure of PS on the outer membrane. When neutrophils undergo apoptosis, they expose “eat me” signals such as PS on their membrane surface that are recognized by resident tissue macrophages (112). Upon recognition, macrophages engulf apoptotic neutrophils in an anti-inflammatory process called efferocytosis, which provides the clearance of apoptotic cells before they reach secondary necrosis and release their potentially noxious intracellular contents (123, 124). Uptake of apoptotic cells by    33 macrophages also suppresses inflammation through the production of anti-inflammatory IL-10, TGF-β, and prostaglandin E2 (PGE2) and by inhibiting the production of pro-inflammatory mediators, including GM-CSF, IL-1β, IL-8, and TNF-α (123, 125, 126). In contrast, necrotic cells release their toxic contents before being recognized by phagocytes. While the anti-inflammatory macrophage response to spontaneous or sterilely induced apoptosis is well described, several reports suggest that macrophages can promote inflammation in response to neutrophils undergoing bacteria-induced apoptosis (241, 242).  ROS are instrumental in directing cell death and in their absence immune cells can undergo alternative cell death pathways upon stimulation with diverse inflammatory outcomes (159, 175). ROS are involved in initiating apoptosis and neutrophils from CGD patients reportedly display both delayed spontaneous apoptosis and decreased apoptotic rates following ingestion of heat-killed Staphylococcus aureus (243). In contrast, live but not UV-treated B. cenocepacia induces necrosis in CGD but not healthy control neutrophils (237). The different fate of CGD neutrophils upon challenge with B. cenocepacia may be caused by an inability of host cells to sufficiently kill the intracellular bacteria. Survival of bacteria may trigger host cell signaling pathways that lead to cell death. Overall, the cell death mechanisms underlying Burkholderia-induced neutrophil death in CGD remain unknown.  Dysregulated cell death may play a vital role in both the inflammatory complications and severity of infections in CGD. We hypothesized that B. cenocepacia induces pathological cell death in the absence of ROS with resulting hyper-inflammation. To investigate this hypothesis, we compared mechanisms of neutrophil cell death with and without ROS in response to challenge with B. cenocepacia clinical isolates and investigated the resulting inflammatory response by human macrophages.    34  2.2 Materials and methods 2.2.1 Isolation of cells Human venous blood was collected from adult volunteers according to the University of British Columbia Clinical Research Ethics Board protocol C04-0193. Immune cells were separated using centrifugation with Ficoll-Paque Plus (GE Healthcare). Peripheral blood mononuclear cells (PBMCs) were washed and resuspended in R10 consisting of: RPMI 1640 media (Gibco) supplemented with 10% (v/v) fetal calf serum (FCS) and 2 mM glutamax. Monocytes were isolated from PBMCs by plastic adherence in serum-free Dulbecco’s modified Eagle medium (DMEM, Gibco). Monocytes were cultured in plastic flasks at 37°C under 5% CO2 with DMEM containing 4.5 g/l glucose, 10% autologous serum, 2 mM glutamax, 10 ng/ml GM-CSF (PeproTech), 100 units/ml penicillin, 100 µg/ml streptomycin, and 250 ng/ml amphotericin B (Gibco) for maturation into human monocyte-derived macrophages. After 6 days macrophages were harvested using trypsin/EDTA, washed in media containing 20% (v/v) FCS to neutralize trypsin, and resuspended in DMEM supplemented with 4.5 g/l glucose, 10% (v/v) pooled normal human serum (PNHS), and 2 mM glutamax. Enumeration and confirmation of viability was performed using Trypan blue dye exclusion as assessed with a haemocytometer. Neutrophils were isolated by Ficoll-Paque centrifugation and dextran sedimentation and resuspended in R10. Human serum was collected from normal healthy donors in unhepranized serum tubes and stored at -80°C. To prepare PNHS, serum from at least 5 donors was thawed, pooled, and refrozen in aliquots. PNHS was used within 3 months of preparation.      35 2.2.2 Bacterial strains and growth conditions The bacterial strains used in this study are listed in Table 2.1. Isolates were selected from the Canadian Burkholderia cepacia Complex Research and Referral Repository (CBCCRRR) (University of British Columbia, Vancouver, BC) or the Bcc experimental strain panel (244). Bacteria were stored at -80°C in Mueller Hinton (MH) II Broth with 8% (v/v) dimethyl sulfoxide (DMSO) or maintained on Columbia agar containing 5% sheep blood (PML Microbiologicals) for a maximum of 2 passages. Bacteria were grown overnight in 3 ml cultures of Luria broth (LB) (10 g/l tryptone, 5 g/l yeast extract, 10 g/l sodium chloride) at 37°C and 250 rpm shaking. K56-2 expressing eGFP from the plasmid pDA42 (232) was grown on LB agar plates containing 100 µg/ml of tetracycline and overnight cultures were grown in LB containing 100 µg/ml of tetracycline. Overnight cultures were washed in phosphate-buffered saline (PBS), diluted in R10, and cultured for approximately 2 h until reaching the desired OD600 corresponding to 108 CFU/ml. Bacteria were opsonized with 10% (v/v) PNHS for 30 min tumbling at 37°C and diluted in R10 to appropriate concentrations. All strains of bacteria were opsonized for neutrophil infections with the exception of Serratia marcescens, which was highly susceptible to killing by PNHS. LB media components were purchased from BD (Franklin Lakes, NJ).            36 Table 2.1 Bacterial strains used in this study  Abbreviations: CF, cystic fibrosis; CGD, chronic granulomatous disease; CBCCRRR, Canadian Burkholderia cepacia complex Research and Referral Repository.   2.2.3 B. cenocepacia growth curves Overnight cultures of B. cenocepacia isolates were adjusted to an OD600 nm of approximately 0.1 in LB. Diluted overnight culture was added to LB with or without 10% (v/v) PNHS (final concentration) at an approximate OD600 nm of 0.01 in a Bioscreen 100 well honeycomb plate. Three technical replicates were performed and appropriate controls, with and without PNHS, were used as blanks. Growth at 37°C was monitored in a Bioscreen C machine (TYPE FP-1100-C, Oy Growth Curves Ab Ltd.) with measurement intervals of 15min and continuous shaking, controlled by EZ experiment software as previously described (73).  Species Strain name Characteristics Source/Reference K56-2  CF isolate, representative of ET12 transmissible lineage  (244) eGFP-expressing K56-2  K56-2 carrying pDA42-eGFP construct Provided by Dr. MA Valvano (232) MH1K Gentamicin-sensitive K56-2 Provided by Dr. MA Valvano (230) C8963 CF isolate, previously shown to induce necrosis in CGD neutrophils  CBCCRRR (237) CEP1067 CGD blood isolate, Canada CBCCRRR  (245) Burkholderia cenocepacia CEP0931 CGD endotracheal tube isolate, U.S.A CBCCRRR Staphylococcus aureus ATCC 29213  Pathogenic in CGD ATCC Serratia Marcescens ATCC 23862 Pathogenic in CGD ATCC PAO1 Low pathogenicity in CGD (246) Pseudomonas aeruginosa ATCC 27853 Low pathogenicity in CGD (247) Escherichia coli ATCC 25922 Low pathogenicity in CGD ATCC    37 2.2.4 Neutrophil challenge with bacteria Freshly isolated neutrophils were diluted in R10. Diphenyleneiodonium (DPI) is a potent inhibitor of the NADPH oxidase (248) that was used to inhibit ROS and mimic CGD. Neutrophils were incubated with 10 µM DPI (Sigma) to inhibit the NADPH oxidase for 30 minutes at 37°C. Opsonized bacteria diluted in R10 were added to neutrophils in 5 ml polypropylene tubes at a multiplicity of infection (MOI) of 5 bacteria per neutrophil. Tubes were tumbled at 37°C for 30 minutes then washed twice in PBS (250xg for 5 min). Neutrophils were resuspended in R10 containing 50 µg/ml meropenum and 1 µM DPI when required for cell death analysis or in DMEM containing 4.5 g/l glucose, 10% (v/v) PNHS, and 2 mM glutamax for co-culture with macrophages. 2.2.5 Association and intracellular killing assay Neutrophils were challenged with B. cenocepacia isolate K56-2 at a MOI of 5 for 30 minutes in R10. Cells were subsequently washed twice in PBS (250xg for 5 min) after challenge with B. cenocepacia and aliquots were removed for cytospinning. Slides were stained with Diff-Quik stain set (Dade Behring, IL) and bacterial association was measured by counting the number of bacteria associated with untreated or DPI-treated neutrophils. Greater than 200 neutrophils were counted for each condition using an Olympus BX50F upright microscope. Intracellular killing of B. cenocepacia was examined after challenging neutrophils with K56-2 at a MOI of 1 in R10 for 30 minutes. Cells were washed 4 times in PBS, which resulted in a 99.9% reduction in extracellular bacteria, resuspended in R10 with or without 1 µM DPI, and incubated at 37°C under 5% CO2. An aliquot was removed at each indicated time point, washed in PBS, and neutrophils were lysed in 1% Triton X-100 in distilled water. Lysates were serially diluted and CFU plated on LB agar.    38 2.2.6 Flow cytometric analysis of neutrophil cell death  Neutrophils challenged with bacterial isolates were washed twice and resuspended in R10 containing 10 µg/ml meropenum and 1 µM DPI where indicated, and incubated at 37°C under 5% CO2. For apoptosis and necrosis analysis, neutrophils were washed once with PBS and once with annexin V-binding buffer, and resuspended in binding buffer containing cells were stained with annexin V, which binds to exposed PS on the surface of apoptotic cells and 7-AAD, a cell impermeable dye that is a marker for leaky membranes indicative of necrosis. Cells were incubated for 15 minutes and 250 µl of binding buffer was subsequently added to each tube. Cells were transferred to 5 ml polystyrene tubes and analysis was performed on >20,000 events using a FACSCalibur system (BC Biosciences) and FlowJo software. Neutrophils were gated to exclude cellular debris and unbound bacteria and assessed as either viable (annexin Vlow/7-AADlow), apoptotic (annexin Vhigh/7-ADDlow), or necrotic (annexin Vhigh/7-AADhigh).  For flow cytometry studies of apoptotic cells containing B. cenocepacia, neutrophils were challenged with K56-2 carrying pDA42, which expresses enhanced green fluorescent protein gene (eGFP) (232). Previous studies have shown that eGFP is stably expressed in B. cenocepacia and has excitation/emission wavelengths of 488/509 (249, 250). EGFP (FL1) and annexin V-PE (FL2) were measured using a FACSCalibur system as above. The detection of eGFP-expressing K56-2 in neutrophils was comparable to the detection of K56-2 in neutrophils fixed in 4% paraformaldehyde (PFA) using a rabbit polyclonal antibody to B. cepacia (Speert laboratory) with a PE-conjugated secondary antibody. 2.2.7 Caspase activity by FLICA staining Neutrophils challenged with K56-2 were incubated at 37°C under 5% CO2. Caspase-3 activation was measured using a carboxyfluorescein (FAM)-labeled FLICA (fluorochrome-   39 labeled inhibitor of caspase) peptide (Immunochemistry Technologies, MN) specific for caspase-3 and -7 (FAM-DEVD-FMK). One hour before each indicated time point, neutrophils were washed with PBS and resuspended in 300 µl of R10 containing 10 µl FLICA for 45 min at 37°C under 5% CO2. Cells were washed twice in supplied apoptosis wash buffer and caspase-3/7 activation was measured by flow cytometry using a FACSCalibur system. Greater than 20,000 events were collected and neutrophils were gated to exclude cellular debris and unbound bacteria. The level of active caspase (FL1) was measured and represented as the percentage of cells with caspase activation above basal unstimulated levels and as the median fluorescent intensity (MFI) of caspase-positive cells. 2.2.8 Immunoblots K56-2-challenged neutrophils were cultured at a density of 2x106 cells/ml in R10 and maintained at 37°C under 5% CO2. At 2 h, 4 h, and 8 h post-infection, cells were washed twice in ice cold PBS and lysed at 2x107 cells/ml in 200 µl of RIPA buffer supplemented with 3x EDTA and 3x Halt protease inhibitor (Thermo Scientific) for 10 min on ice. Lysates were sonicated at 80% for 6 cycles of 10s on and 10s off and frozen at -20°C. Prior to loading on SDS PAGE, 0.5 µl benzonase was mixed with samples to reduce viscosity of nucleic acid and samples were boiled in Laemmli buffer for 5 min. Lysates from 5x105 cells per sample were resolved on 12% SDS-polyacrylamide gels and transferred onto PVDF membranes (Millipore). Blots were blocked for 1 h at room temperature and probed overnight at 4°C for caspase-3 (Cell Signaling) and GAPDH (Fitzgerald). Blots were subsequently probed for 1 h with fluorescent-labeled secondary antibodies, IRDye® 680 or 800CW (LI-COR Biosciences), and imaged on a LI-COR Odyssey infrared imaging system (LI-COR Biosciences) and quantified using the included analysis software.    40 2.2.9 Lactate dehydrogenase assay Neutrophils were challenged with K56-2 as described and washed twice in PBS. Where indicated, neutrophils were pretreated with 20 µM z-vad-fmk (R&D Systems), a cell-permeable irreversible pan caspase inhibitor. Cells were resuspended in R10 containing 10 µg/ml meropenum and 1 µM DPI and/or 10 µM z-vad-fmk where indicated. Supernatants from cell stimulations were transferred to 96-well v-bottom plates and spun at 4000xg for 5 min to pellet residual cells. Cell-free supernatants were diluted 1:10 in PBS and lactate dehydrogenase (LDH) release was measured using a cytotoxicity detection kit (Roche Applied Science) according to manufacturer’s instructions. Control samples included medium alone (blank) and cells treated with 1% Triton X-100 overnight. Data are represented as a percentage of maximum LDH release (Triton X-100) after subtraction of blank. 2.2.10 Cytokine release by macrophages Neutrophils were challenged with a gentamicin-sensitive strain of K56-2 (MH1K, (230)) at an MOI of 5 for 30 minutes as described. Cells were washed in PBS (250xg for 5 min) and resuspended in DMEM containing 10% FCS and 50 µg/ml of gentamicin for 30 minutes to kill extracellular bacteria. Cells were washed again in PBS and resuspended in DMEM supplemented with 4.5 g/l glucose, 10% (v/v) PNHS, 2 mM glutamax, 1 µg/ml of gentamicin to prevent the extracellular survival of bacteria from lysed cells, and 1 µM DPI where indicated for co-culture with macrophages. Macrophages were seeded at 6x104 cells/well in 96-well plates and B. cenocepacia-challenged neutrophils were added at a 1:1 neutrophil to macrophage ratio. Macrophages or neutrophils alone, with and without DPI, were included as controls. After 24 hours supernatants from co-cultures were transferred to 96-well v-bottom plates and spun at 3000xg for 5 min to pellet residual cells. Cell-free supernatants were frozen at -80°C for cytokine    41 analysis. Cytokines released into supernatants were quantified using sandwich ELISAs for TNF-α, IL-1β, IL-6, IL-8, IL-10, IL-12, and TGF-β (eBioscience). 2.2.11 Statistics Graphs displaying mean ± SEM and were generated with GraphPad Prism 5 (GraphPad Software Inc., San Diego, USA). Statistical significance was determined by performing a Student’s t test and one-way variances of analysis with the Bonferroni post-test where applicable.  2.3 Results 2.3.1 Impact of ROS on neutrophil death following challenge with B. cenocepacia clinical isolates  Neutrophils defective for ROS exhibited enhanced necrosis following challenge with the B. cenocepacia isolate C8963 (237).  However, the Bcc displays a high level of both intra-lineage and intra-species genetic diversity and causes variable levels of pathogenicity and clinical outcomes in different infection models and in humans (251-254). To determine whether enhanced neutrophil death in the absence of ROS following challenge with B. cenocepacia was a common effect within the species, neutrophils were challenged with a panel of B. cenocepacia clinical isolates at a MOI of 5. A MOI of 5 allowed most neutrophils to phagocytose at least one bacterium and resulted in longer neutrophil survival compared to higher MOIs assayed. B. cenocepacia isolates K56-2 and C8963 were isolated from patients with CF and CEP1067 and CEP0931 were isolated from patients with CGD (Table 2.1). Neutrophils were challenged with opsonized isolates for 30 minutes and cell death was measured by flow cytometry with staining for annexin V-PE, which binds to exposed PS on the surface of apoptotic cells and 7-AAD, a cell impearable dye that indicates leaky membranes or necrosis.     42 As seen in Figure 2.1 a-b, by 6 hours post-infection neutrophils challenged with C8963 and K56-2 exhibited significantly enhanced apoptosis in the absence of ROS. A significant increase in necrosis was also seen in DPI-treated neutrophils challenged with K56-2 compared to normal neutrophils. By 16 hours post infection, neutrophils challenged with C8963 and K56-2 were significantly more apoptotic in the absence of ROS (Figure 2.1c). A large portion of cells by 16 hours had proceeded to necrosis (Figure 2.1d) in the absence of efferocytosing phagocytes in our in vitro system. Significantly enhanced necrosis in neutrophils treated with DPI was seen following challenge with K56-2 compared to normal neutrophils. ROS did not have an apparent effect on apoptosis or necrosis at either time point following challenge with CEP1067. This isolate may behave as an anomaly as it has shown similar differences in cell death induction in dendritic cells compared to other B. cenocepacia strains (245). Although DPI-treated neutrophils challenged with C0931 had a trend towards higher cell death compared to normal cells, this trend failed to reach significance. DPI had no effect on spontaneous apoptosis or necrosis.             43  Figure 2.1 Neutrophil apoptosis and necrosis following challenge with clinical B. cenocepacia isolates Neutrophils were challenged with bacteria for 30 minutes. At each time point, neutrophils were washed and stained with annexin V-PE and 7-AAD prior to analysis by flow cytometry. Cells that stained with only annexin V were classified as apoptotic and those that stained with both annexin V and 7-AAD were classified as necrotic. A. apoptotic cells at 6 h p.i, B. necrotic cells at 6 h p.i., C. apoptotic cells at 16 h p.i., D. necrotic cells at 16 h p.i. N ≥ 5 for each condition. *P ≤ 0.05 ; **P ≤ 0.01 ; ***P ≤ 0.001 as measured using Student’s t tests.   2.3.2 Impact of ROS on neutrophil death following challenge with a panel of pathogenic and non-pathogenic bacteria in CGD To determine whether enhanced neutrophil death in the absence of ROS was specific for the Bcc, we challenged neutrophils with a number of different bacteria that are or are not common pathogens in CGD. Information related to the panel of isolates used in this study are   SpontaneousC8963K56-2CEP1067CEP0931020406080***NormalDPI % Apoptotic neutrophilsSpontaneousC8963K56-2CEP1067CEP09310510152025*% Necrotic neutrophilsSpontaneousC8963K56-2CEP1067CEP0931020406080* *NormalDPI % Apoptotic neutrophilsSpontaneousC8963K56-2CEP1067CEP09310510152025**% Necrotic neutrophils6h p.i. 16h p.i.ABC D   44 found in Table 2.1. Neutrophils were challenged with S. aureus, S. marcescens, and B. cenocepacia, which are isolated from 12%, 5%, and 8% of patients with pneumonia and 9%, 6%, and 12% of patients with bacteremia, respectively (34). The clinical isolate K56"2 was chosen as a representative B. cenocepacia isolate for this and for future experiments because it is well-characterized and amenable to genetic manipulation. E. coli and P. aeruginosa rarely cause infections in CGD and common labortatory strains of these bacteria were chosen for comparison controls. Pseudomonas is recorded to cause some infections in CGD, however since B. cepacia was formerly known as Pseudomonas cepacia, it is acknowledged that some of the Pseudomonas isolates recorded in the literature were actually Bcc (34). The prevalence of the Bcc is therefore likely underestimated and that of Pseudomonas overestimated (34). All bacteria were opsonized with 10% PNHS prior to incubation with neutrophils except for S. marcescens, which was extremely susceptible to killing by PNHS. Neutrophils were challenged with bacteria at MOI 5 for 30 minutes tumbling to allow for sufficient interaction. After 16 hours resting, cell death was examined by flow cytometry using annexin V-PE as a marker for apoptosis and 7-AAD as a marker for membrane integrity, or necrosis. As seen in Figure 2.2, challenge with S. marcescens, S. aureus, P. aeruginosa, and E. coli did not induce apparent apoptosis in normal neutrophils after 16 hours, but did result in necrosis. In the absence of ROS, challenge with these bacteria was in fact anti-apoptotic compared to the spontaneous level of neutrophil apoptosis. A significant protective effect against apoptosis in the absence of ROS was observed following challenge of S. marcescens and P. aeruginosa isolate PAO1. This trend failed to reach significance with S. aureus, P. aeruginosa isolate ATCC 27853, and E. coli. ROS also contributed to necrosis following challenge with S. marcescens and P. aeruginosa. A strong trend towards protection against necrosis in the absence of ROS was seen with S. aureus    45 but again failed to reach significance. B. cenocepacia was the only strain observed where ROS was protective against neutrophil death. Unlike in Figure 2.1, B. cenocepacia isolate K56-2 did not induce significant differences in apoptosis in normal and DPI-treated neutrophils by 16 hours post-infection. However, the data suggests that significantly more cells had gone necrotic at this time point, which may explain this discrepancy and reflects the variability of using primary human cells. This data suggests that neutrophils defective in ROS are not simply prone to cell death following exposure to bacteria. It also provides evidence that enhanced neutrophil death in the absence of ROS does not occur following exposure to all CGD pathogens and that the effect may be specific for the Bcc.     46                    Figure 2.2 Neutrophil apoptosis and necrosis following challenge with a panel of pathogenic and non-pathogenic CGD bacteria Neutrophils were challenged with bacteria for 30 minutes. At 16 h p.i., cells were washed and stained with annexin V-PE and 7-AAD prior to analysis by flow cytometry. Cells that stained with only annexin V-PE were classified as apoptotic (A) and those that stained with both annexin V-PE and 7-AAD were classified as necrotic (B). N ≥ 3 for each condition. *P ≤ 0.05 as measured using Student’s t tests.    SpontaneousS. marcescens ATCC 23862S. aureus ATCC 29213P. aeruginoa PAO1P. aeruginosa ATCC 27853E. coli ATCC 25922B. cenocepacia K56-2020406080NormalDPI**% Apoptotic PMNSpontaneousS. marcescens ATCC 23862S. aureus ATCC 29213P. aeruginoa PAO1P. aeruginosa ATCC 27853E. coli ATCC 25922B. cenocepacia K56-201020304050****% Necrotic PMNAB   47 2.3.3 Intracellular killing of B. cenocepacia by neutrophils The Bcc is killed by normal but not CGD neutrophils in vitro, presumably due to their resistance to non-oxidative killing (255). We aimed to confirm that phagocytosed B. cenocepacia survive intracellularly within neutrophils in the absence of ROS. Normal and DPI"treated neutrophils were challenged for 30 minutes with live opsonized K56-2 at an MOI of 1. To examine association of B. cenocepacia, neutrophils were stained using Diff"Quik and the number of intracellular bacteria counted microscopically. As seen in Figure 2.3 a-b, opsonized B. cenocepacia was readily taken up by both normal and DPI-treated neutrophils within 30 minutes. Normal and DPI-treated neutrophils were associated with 65.1% and 73.89% of the inoculating bacteria, respectively, which was not significantly different by a Student’s t test. To measure the rate of intracellular killing, cells were washed extensively following infection to remove 99.9% of extracellular bacteria. Neutrophils were then lysed and CFU plated. As seen in Figure 2.3 c, the survival of K56"2 within DPI"treated neutrophils within 165 minutes was nearly two log-fold greater than its survival in normal neutrophils.      48                 Figure 2.3 Intracellular survival of B. cenocepacia in neutrophils A, B. Diff-Quik stains of human neutrophils following 30 min infection with B. cenocepacia isolate K56-2. Arrows indicate intracellular bacteria. A. Normal neutrophils, B. DPI-treated neutrophils. C. Intracellular survival of K56-2 within normal and DPI-treated neutrophils. Bacteria were tumbled with neutrophils at an MOI of 1 for 30 min. Following 4 washes, neutrophils were lysed and CFU plated. The experimental limit of detection is 103. N = 3. P ≤ 0.05 as measured using a two-way analysis of variance.   2.3.4 Composition of live and apoptotic neutrophils containing B. cenocepacia Removal of bacteria by neutrophil phagocytosis ideally leads to phagocytosis-induced apoptosis. This allows for the resolution of inflammation and tissue healing through efferocytosis by scavenger macrophages (256). Many bacterial components can modulate neutrophil apoptosis. Previous studies have shown that some species of the Bcc release a hemolysin that     Number of intracellular bacteria following 30 min challenge with B. cenocepacia (1x10^6 CFU/ml )45105165100102103104105106NormalDPIp=0.035CTime p.i. (min)CFU/ml   49 induces apoptosis in human neutrophils (257). In contrast, LPS and bacteria-free supernatants from B. cenocepacia were both shown to be anti-apoptotic to neutrophils in a dose-dependent manner (237). We aimed to determine whether the enhanced apoptosis seen in DPI-treated neutrophils after challenge with B. cenocepacia was associated with phagocytosis of the bacteria or simply exposure to the bacteria.  Neutrophils were challenged with K56-2 expressing constitutive eGFP for 30 minutes as described above. Previous studies have shown that eGFP is expressed and remains stable in B. cenocepacia and emits strong fluorescence (249). At 8 hours post-infection, cells were stained with annexin V-PE, which binds to exposed PS on the surface of apoptotic cells and is therefore a marker for apoptosis. As in earlier experiments, positive staining for annexin V can represent either apoptotic cells, or necrotic cells. Since less than 5% of K56-2-challenged neutrophils undergo necrosis by 6 hours (Figure 2.1), it is reasonable to assume that the majority of annexin V positive cells detected by 8 hours are apoptotic. Flow cytometry was used to distinguish cell death in cells that have ingested eGFP-expressing B. cenocepacia and those that were exposed to, but did not phagocytose, the bacteria.  Figure 2.4ab shows that normal and DPI-treated uninfected neutrophils exhibited similar levels of annexin V staining and no signal for eGFP-expressing bacteria. As seen in Figure 2.4c, B. cenocepacia did not induce apoptosis in normal neutrophils. Furthermore, eGFP was only detected in 12% of normal cells, indicating that most cells do not contain detectable bacteria at 8 hours post-infection. Only 9% of normal neutrophils were positive for both eGFP and annexin V. In contrast, 69% of DPI-treated neutrophils contained detectable eGFP-expressing B. cenocepacia at 8 hours post-infection (Figure 2.4d). Of the total DPI-treated neutrophils, 60% were annexin V-positive and contained B. cenocepacia. Overall the majority of normal    50 neutrophils were negative for annexin V and B cenocepacia, whereas the majority of DPI-treated cells were positive for annexin V and contained B. cenocepacia. Preliminary data using a polyclonal antibody specific for the Bcc on fixed and permeabilized cells showed similar results.  It is important to note that of the neutrophils that contained detectable bacteria, approximately 75% were apoptotic in both normal and DPI-treated cells. This suggests that the induction of apoptosis may be associated with the phagocytosis and the presence of intracellular B. cenocepacia within neutrophils rather than a direct effect of ROS.              Figure 2.4 Composition of live and apoptotic neutrophils with detectable B. cenocepacia.  Cells were challenged with eGFP-expressing K56-2 and stained with annexin V/PE after 8 hours. A. Normal, unstimulated neutrophils, B. DPI-treated, unstimulated neutrophils, C. K56-2-challenged normal neutrophils, D. K56-2-challenged, DPI-treated neutrophils. Values represent averages of 4 donors ± SEM.      51 2.3.5 B. cenocepacia-induced caspase activation  Caspase activation is a hallmark of apoptosis and caspase-3 and -7 are well-described effector caspases that cleave cellular substrates and lead to PS exposure on the cell surface (258). Caspase-3 exhibits much more efficient cleavage activity than activity caspase-7 (259). To assess early apoptotic processes, we evaluated the level of caspase-3 activation in neutrophils following challenge with B. cenocepacia. Caspase-3 activation was measured by flow cytometry using a FLICA probe specific for active caspase-3/7 at 1, 3, and 6 hours post-infection. In normal neutrophils, a portion of neutrophils activated caspase-3/7 within 1 hour and the signal began to decrease by 3 hours post"infection. In contrast, a greater proportion of DPI-treated neutrophils activated caspase-3/7 and its activation persisted over time (Figure 2.5a,b). Most striking was the difference in MFI of FLICA positive cells. As seen in Figure 2.5a,c, DPI-treated neutrophils exhibited a much higher level of FLICA binging, suggesting that more caspase-3/7 molecules were activated within a given cell. Caspase-3 activation was confirmed by Western blotting for the unprocessed form of the protein at 4, 8, and 20 hours post"infection. DPI-treated cells stimulated with B. cenocepacia undergo profound cell death by 18 hours post-infection (Figure 2.1c-d) and we found that neutrophils at 20 hours were difficult to fully lyse and to extract appreciable levels of protein. Therefore quantification was measured on Western blots for 4 and 8 hours post-infection. As early as 4 hours post"infection, significantly less full"length caspase-3 was observed in B. cenocepacia!challenged neutrophils pre"treated with DPI compared to untreated cells (Figure 2.5d-e), indicating that full-length caspase-3 was cleaved to its active form. The difference in caspase activation between normal and DPI-treated cells continued to increase at 8 hours post-infection.     52 Figure 2.5 B. cenocepacia-induced caspase-3 activation in neutrophils   A. At each time point cells were incubated with a FLICA probe specific for caspase-3/7 and fluorescence was measured by flow cytometry. Grey shadowed: unstimulated. Blue line: K56-2 stimulated. Representative graph of N = 4 B. Cells with fluorescence above the level of unstimulated controls were classified as FLICA positive. N = 4. C. Mean fluorescent intensity (MFI) of FLICA positive cells. N=3 D. Representative Western blot for uncleaved caspase-3. E. Densitometry of Western blot normalized to GAPDH. N = 6. *P ≤ 0.05 ; **P ≤ 0.01 ; ***P ≤ 0.001 as calculated by two-way analyses of variance using a Bonferroni posttest.                                       1 3 6020406080NormalDPIp=0.0312*Time p.i. (hours)% FLICA positive PMN1 3 6050100150200NormalDPI*****Time p.i. (hours)MFIB CWestern 4-8h4h8h0.00.51.01.5UnstimulatedDPI UnstimulatedK56-2DPI K56-2****Time p.i.Uncleaved caspase-3 (Normalized fluorescence)E   53 To examine the necessity of caspase activation on B. cenocepacia-induced neutrophil death, neutrophils were pre-treated with the pan-caspase inhibitor z-vad"fmk prior to B. cenocepacia challenge. At 20 hours post-infection, the LDH in cell supernatants was measured as an indicator of cytotoxicity. As seen in Figure 2.6, caspase inhibition had no effect on the cytotoxicity of normal B. cenocepacia!treated neutrophils. In contrast, pre"treatment with z"vad"fmk significantly reduced the level of B. cenocepacia-induced cytotoxicity in DPI"treated neutrophils to that of normal neutrophils. This data shows that caspase activation is necessary for enhanced cell death in ROS-deficient neutrophils following challenge with B. cenocepacia, which is indicative of apoptotic cell death.        Figure 2.6 Caspase activation is required for enhanced cell death in DPI-treated neutrophils  Normal and DPI-treated cells were pre-treated with the pan-caspase inhibitor z-vad-fmk prior to challenge with B. cenocepacia isolate K56-2. Cytotoxicity was determined by measuring LDH in cell supernatants 20 hours post-infection. N = 5. *P ≤ 0.05 ; **P ≤ 0.01 as measured using Student’s t tests.   Normal DPI0204060UnstimulatedK56-2K56-2 + z-vad-fmk***Effect of caspase inhibitor zvad-fmk on cell death% of Max (LDH)   54 2.3.6 ROS-deficient neutrophils challenged with B. cenocepacia induce a pro-inflammatory response in macrophages Apoptotic cells in the body are quickly efferocytosed by phagocytes, most commonly tissue-resident macrophages. Phagocytosing-macrophages promote resolution of inflammation by releasing anti-inflammatory mediators and down regulating the production of pro-inflammatory cytokines (123). In particular, secretion of TGF-β and IL-10 is well documented following either phagocytosis of apoptotic neutrophils expressing PS on the surface or upon interaction with apoptotic membrane or PS vesicles (127, 128, 260). Since DPI-treated neutrophils exhibited a high level of PS surface exposure consistent with apoptosis in response to B. cenocepacia (Figure 2.1a,c), we examined their ability to impart an anti-inflammatory phenotype on primary human monocyte-derived macrophages (MDM). Macrophages were differentiated in autologous serum supplemented with GM-CSF, which is associated with differentiation to an alveolar macrophage-like phenotype (261). A gentamicin-sensitive strain of B. cenocepacia K56-2 (MH1K) was used in order to kill extracellular bacteria following phagocytosis by neutrophils and to ensure that any effect on macrophages was due to neutrophils alone and not due to the differential survival of B. cenocepacia in normal versus DPI-treated cells. The growth rate of MH1K did not differ from that of K56-2 (Appendix A, Figure A.1) and MH1K had a minimum inhibitory concentration (MIC) of ≤ 1 µg/ml of gentamicin. As described above, normal and DPI-treated neutrophils were tumbled with opsonized MH1K for 30 minutes. Cells were washed and incubated in gentamicin to kill extracellular bacteria, washed again, and resuspended in a low level of gentamicin (1 µg/ml) to ensure killing of any bacteria released from lysed neutrophils. Neutrophils were added to macrophages at a 1:1 ratio and the levels of IL-10, TGF-β, IL-1β, TNF-α, IL-6, IL-12p70, and    55 IL-8 in cell-free supernatants were measured by ELISA after 24 hours. No CFU were detected in culture supernatants at this time.  As shown in Figure 2.7, the only inflammatory mediator produced by neutrophils was IL-8 following challenge with B. cenocepacia. The levels of IL-8 produced by normal and DPI-treated neutrophils were not significantly different. DPI-treated neutrophils challenged with MH1K induced high levels of TNF-α, IL-6, and IL-8 from macrophages compared to normal neutrophils challenged with MH1K. Neither normal nor DPI-treated unstimulated neutrophils induced the secretion of inflammatory mediators from macrophages. We were not able to detect IL-10, TGF-β, IL-1β, or IL-12p70 in any of the conditions examined. These findings suggest that macrophages are skewed towards a pro-inflammatory phenotype following exposure to neutrophils challenged with B. cenocepacia. DPI-treated B. cenocepacia-challenged neutrophils induced much higher levels of pro-inflammatory mediators from macrophages compared to normal neutrophils. The chemokine IL-8 is potent at recruiting neutrophils to sites of infection and TNF-α and IL-6 are involved in promoting a potent inflammatory response. This data shows that although B. cenocepacia induces apoptosis in ROS-deficient neutrophils, the apoptotic neutrophils do not promote an anti-inflammatory response in macrophages consistent with resolution but rather induce the secretion of the pro-inflammatory mediators TNF-α, IL-6, and IL-8. The resulting combination of high levels of surviving bacteria and inflammation induced by apoptotic neutrophils may contribute to morbidity in patients with CGD.        56                  Figure 2.7 Cytokines produced by macrophages cultured with B. cenocepacia-challenged neutrophils. Normal and DPI-treated neutrophils (PMN) were challenged with MH1K for 30 minutes. PMN were washed and resuspended in gentamicin to kill extracellular bacteria, then added to macrophages (MDM) at a ratio of 1:1. Cell-free 24 h supernatants were assayed for IL-10, TGF-β, IL-1β, TNF-α, IL-6, IL-12p70, and IL-8 by ELISA as described by the manufacturer. IL-10, TGF-β, IL-1β, or IL-12p70 were not detected in any of the conditions examined. N = 6 – 8. *P ≤ 0.05 ; **P ≤ 0.01 as measured by a Student’s t test.    MDMMDM + PMNMDM + PMN/MH1KPMNPMN/MH1K05000100001500020000**DPINormalTNF-α (pg/ml)MDMMDM + PMNMDM + PMN/MH1KPMNPMN/MH1K02000400060008000*NormalDPIIL-6 (pg/ml)AMDMMDM + PMNMDM + PMN/MH1KPMNPMN/MH1K0100000200000300000400000500000NormalDPI**IL-8 (pg/ml)BC   57 2.4 Discussion  Bacteria of the Bcc cause severe lung infections that are associated with neutrophil infiltration and tissue inflammation in patients with CF and CGD. B. cenocepacia was previously shown to survive in the presence of ROS-deficient neutrophils and induce neutrophil necrosis within 20 hours in vitro (57, 255). This study was designed to investigate earlier events that lead to neutrophil necrosis. We show here that B. cenocepacia survives intracellularly within ROS-deficient neutrophils and induces signaling that leads to early apoptosis compared to healthy controls. Many apoptotic ROS-deficient neutrophils progressed to secondary necrosis by 16 hours post-infection in the absence of efferocytosing phagocytes. This effect was in contrast to other genera of bacteria, including other CGD pathogens, for which ROS was protective against cell death. ROS are involved in a number of biological pathways, including in initiating apoptosis (262), and the requirement of ROS for neutrophil apoptosis following exposure to a number of bacteria and their virulence factors, including E. coli (263), S. aureus (264), and pyocyanin from  P. aeruginosa (265) has been demonstrated. Like B. cenocepacia, B. multivorans and B. cepacia are significant Bcc pathogens in CGD. Whether they induce increased rates of apoptosis in ROS-deficient neutrophils remains to be established.  A number of bacteria and bacterial toxins exhibited a dose-dependent effect on neutrophils, where low MOIs induced apoptosis and high doses induced necrosis (266-268). We too observed a dose dependent effect where lower MOIs could in fact be anti-apoptotic and higher MOIs could induce very rapid cell death. Furthermore, we found that a high level of variability in working with primary neutrophils. Together, these factors contribute to the variability in this study, particularly in the difference in K56-2-induced apoptosis seen in Figure 2.1 and 2.2.    58 We investigated the role of apoptosis by evaluating caspase-3 activation and caspase dependency and early and late PS exposure. We also aimed to examine whether the extrinsic caspase-8 mediated or intrinsic caspase-9 mediated pathways of apoptosis were involved in this system. However a number of factors prevented us from examining these upstream factors. First, caspases have general cleavage motifs with overlapping specificity for several caspases. Most commercially available caspase activation assays use short peptide-based substrates and inhibitors that exhibit promiscuity for different cleavage motifs (259). Importantly, caspase-3 is not only able to cleave most substrates more efficiently than the caspases to which the substrates are specific but is also more abundant in neutrophils (259, 269, 270). Since standard substrate-based assays and FLICA assays to distinguish specific caspase activation are dependent on these motifs, we did not trust the sensitivity and validity of these experiments for initiator caspases. The use of in vitro transfections to reduce caspase expression was hindered by the fact that neutrophils are terminally differentiated and short lived. Furthermore, not only do neutrophils have a low level of de novo protein synthesis, but also caspases exist as pre-formed zymogens (inactive enzymes) in the cell, making RNA interference to reduce caspase levels impractical (271). Finally, we were not able to detect sufficient levels of either caspase-8 or -9 using monoclonal antibodies for Western blotting. Therefore, the upstream signaling of apoptosis induced by B. cenocepacia remains to be established and examining other upstream factors is a future direction of this study. A growing number of pathogens have been shown to induce neutrophil apoptosis to evade host defenses, including P, aeruginosa (272), Streptococcus spp (266, 273), Yersinia pestis (274), Francisella tularensis (275), Shigella dysenteriae (276), and S. aureus (267). Apoptosis is thought to provide a survival method for these pathogens to be transferred to less-antimicrobial    59 macrophages through the Trojan horse model of efferocytosis. Whether apoptosis is a desired outcome of the pathogen or a consequence of effective immunity likely depends on the specific microorganism and its ability to escape the apoptotic cell and survive within macrophages. B. cenocepacia-induced neutrophil death requires live bacteria (237), suggesting that apoptosis may be triggered by a virulence factor. A variety of bacterial virulence factors have been described that modulate specific pathways of cell death in leukocytes, including apoptosis (277, 278). These include effector proteins that are secreted or injected into target cells via type III or type IV secretion systems (268). Effector proteins in the host cell cytoplasm can interact with a number of pro-survival and apoptotic pathway regulators, including Bcl-2 family proteins (279) and other regulators of mitochondrial membrane permeability (280), and NF-κB and MAP kinase pro-survival pathways (281, 282). Several secretion systems have been implicated in the pathogenicity of the Bcc, including a type II, a type III, two type IV, a type V, and a type VI secretion system (178). Importantly, the type II secretion system of B. cenocepacia secretes a hemolysin (283, 284) and a type IV secretion system was required for intracellular survival in epithelial cells and macrophages (227). Molecules secreted into culture supernatants do not contribute to B. cenocepacia-induced neutrophil death (237), but the role of potential Bcc virulence factors such as zinc metalloproteases, hemolysins, and lipases activated intracellularly in immune cells remains to be determined. As previously mentioned, macrophages efferocytose apoptotic cells and transfer them to the phagolysosome where they are degraded, limiting the likelihood that they will progress to secondary necrosis (122-124). Efferocytosis is considered an anti-inflammatory process that has a profound influence on resolution of inflammation (123, 124). Macrophages produce IL-10, TGF-β, and PGE2 during the engulfment of apoptotic cells that inhibit inflammatory mediator    60 production (123, 125, 126). Even in the absence of entire apoptotic cells, apoptotic cell membranes alone could diminish the production of IL-12 by macrophages exposed to LPS in vitro (128) and PS vesicles can stimulate the production of TGF-β (127). Anti-inflammatory responses are critical to restore lung homeostasis following infection and to prevent inflammation. The role of apoptotic cells in this response is demonstrated by the fact that intratracheal administration of apoptotic cells enhances the resolution of acute pulmonary inflammation induced by lipopolysaccharide (127). However, an anti-inflammatory response has the often undesirable ability to dampen innate antimicrobial defenses. Apoptotic cells suppress phagocytosis and bacterial killing by alveolar macrophages in vitro and PGE(2) induced by efferocytosis impairs lung recruitment of neutrophils and clearance of Streptococcus pneumoniae (285). Furthermore, macrophage phagocytosis of uninfected apoptotic, but not necrotic cells fuels intramacrophage growth of Trypanosoma cruzi and Coxiella burnetii (286, 287). Thus the inflammation induced by infected apoptotic neutrophils may in fact be necessary to provide an efficient antimicrobial response. Most studies involving the inflammatory response of macrophages to apoptotic neutrophils have used apoptotic neutrophils induced by age, irradiation, or sterile stimuli. Thus anti-inflammatory macrophage programming may not always apply when neutrophils undergo pathogen-induced apoptosis. During the course of infectious disease, the immune system must balance the need to eliminate or control the infectious agent and the need to minimize the destruction of tissue architecture (288). Skewed inflammatory and anti-inflammatory responses correlate with resistance and susceptibility to infectious diseases, respectively. Several studies have reported differential effects between spontaneous and pathogen-induced apoptotic neutrophils on the inflammatory state of macrophages. Neutrophil apoptosis    61 triggered by M. tuberculosis, E. coli, or S. aureus induces a pro-inflammatory response in macrophages that is not triggered by uninfected apoptotic neutrophils or the bacteria alone (241, 289, 290). We found that although B. cenocepacia induced a fast rate of apoptosis in ROS-deficient neutrophils, these neutrophils induced high levels of the chemokines IL-8 and pro-inflammatory cytokines IL-6 and TNF-α from human macrophages and a distinct absence of the anti-inflammatory mediators IL-10 and TGF-β. This suggests that B. cenocepacia-induced apoptotic ROS-deficient neutrophils induce pro-inflammatory programming in macrophages. We found that ROS-deficient neutrophils were unable to effectively kill intracellular B. cenocepacia and that the bacteria was detected in most apoptotic neutrophils. Therefore, the pro-inflammatory macrophage programming induced by these apoptotic neutrophils may reflect the inability to kill the invading pathogen. It is important to note that while B. cenocepacia-induced apoptotic neutrophils induced high levels of TNF-α , IL-6, and IL-8, we did not detect the production of IL-1β or IL-12p70, two potent pro-inflammatory cytokines involved in immune activation that are produced by macrophages in response to B. cenocepacia (Chapter 3). Future directions are aimed at further characterizing the inflammatory profile of macrophages in response to B. cenocepacia-infected neutrophils. A limitation of this study is the use of DPI as a model of CGD neutrophils. Though most potent as an inhibitor of NOX enzymes such as the NADPH oxidase (291), DPI also inhibits other flavoenzymes, including the nitric oxide synthase, cyclooxygenase, and the mitochondrial electron transport chain. Some studies have observed discrepancies between DPI-treated and CGD cells, however this is most often described in macrophages and monocytes (50, 292, 293) rather than in neutrophils. We found that DPI had an inhibitory effect on the growth of K56-2, even at low concentrations. Therefore, all experiments with involving bacteria incubated with    62 DPI for longer than 1 hour were conducted using only 1 µM of DPI, which was not inhibitory to the growth of K56-2. The effect of this concentration of DPI on the growth of other bacteria in this study is a focus of future directions. The neutrophil-like cell lines PLB-985 and X-CGD PLB-985, in which the gp91phox locus of the NADPH oxidase has been disrupted by homologous recombination (294), were also assayed in this study as an alternative model but had little to no antibacterial capabilities towards B. cenocepacia. Apocyning is a naturally occurring methoxy-substituted catecol that inhibits the formation of superoxide, likely by blocking the sulfhydryl groups on the NADPH oxidase subunits and inhibiting enzyme assembly (295). Apocynin could therefore be used to confirm our results with DPI. Ideally, we would like to confirm our results in primary human CGD neutrophils when available.  In summary, we show that B. cenocepacia associates similarly with normal and DPI-treated cells, however survives intracellularly only in the absence of ROS. DPI-treated cells challenged with B. cenocepacia exhibited increased apoptosis compared to normal neutrophils that was associated with the presence of intracellular bacteria, caspase-3 activation, and PS exposure and depended on caspase activity. However, B. cenocepacia-induced apoptotic DPI-treated neutrophils were highly inflammatory to macrophages. Collectively, these studies suggest that B. cenocepacia infections in CGD are exacerbated by a high level of bacterial survival and inflammatory neutrophil apoptosis.     63 Chapter 3: Human macrophages provide a rich replication niche that allows B. cenocepacia to escape neutrophil killing and enhances replication  3.1 Introduction Pulmonary innate immunity protects against inhaled pathogens through a combination of bronchial/alveolar epithelial cells that clear foreign agents and phagocytes that engulf pathogens and activate various arms of host defense (296). A small number of phagocytes, mainly alveolar macrophages and dendritic cells, reside in the lung, however polymorphonuclear neutrophils (PMN) and monocytes are quickly recruited to sites of infectious inflammation (297). Alveolar macrophages are the most abundant phagocyte in the lung and play a key role in immunity against respiratory infections through their phagocytic, immunomodulatory, and antigen-presenting abilities (298). Macrophages express several pattern recognition receptors (PRRs) that recognize highly conserved pathogen-associated molecular patterns (PAMPs) and stimulate the production of chemokines and pro-inflammatory cytokines that recruit leukocytes and activate the inflammatory response (92). Macrophages can also phagocytose microbes and channel them through the endocytic pathway that eventually fuses with a lysosome containing proteolytic enzymes and low pH and leads to their destruction (93). Macrophages also exhibit antigen-processing and presentation capacity and are important in the development of an adaptive immune response.  Circulating neutrophils in the blood are mobilized in high numbers to infectious loci. Neutrophils possess a larger repertoire of antimicrobial agents compared to macrophages and are inherently better at killing most pathogens (299). Neutrophils produce reactive oxygen species    64 (ROS) on the plasma and phagosomal membranes, release antimicrobial peptides and proteases from intracellular granules, and release neutrophil extracellular traps (NETs) composed of DNA and histones that bind to and kill microbes (80, 238). Importantly, neutrophils contain high levels of catalytic proteases and potent mammalian antimicrobial peptides including various defensins and cathelicidins that are absent or scarce in macrophages (207, 300-302). Neutrophils can also produce hypochlorous acid, one of the most potent bacterial oxidants, using myeloperoxidase (MPO). MPO is found in monocytes but is lost during maturation to macrophages (303). While macrophages and neutrophils have distinct functions in immunity based on their specialized antimicrobial effector mechanisms, capacity to activate immune cells, and location in the body, they can also work collaboratively to combat disease (Reviewed in (304, 305)). Macrophages secrete cytokines and chemokines that activate and recruit neutrophils to infectious loci and can prolong their survival (305-308). Efferocytosis is a form of phagocyte cooperation whereby macrophages phagocytose apoptotic leukocytes and promote the resolution of inflammation (112, 122). Phagocytosis of intact apoptotic bodies prevents their progression to necrosis and the subsequent release of cytotoxic cellular constituents. Furthermore, efferocytosing macrophages produce anti-inflammatory cytokines, including interleukin (IL)-10, transforming growth factor (TGF)-β, and prostaglandin E2 (PGE2) (123, 125, 126).  Efferocytosis has also been described as an innate antibacterial mechanism (309). Phagocytosis of apoptotic neutrophils can enhance the limited antimicrobial methods available to macrophages through the acquisition of neutrophil granules and use of their contained antimicrobial molecules (310, 311). Furthermore, efferocytosis acts as an alternative antibacterial mechanism against Mycobacterium tuberculosis (Mtb). Mtb survives in macrophages by inhibiting apoptosis and persisting in arrested phagosomes; alternatively Mtb can induce necrosis    65 to evade immunity and disseminate (312, 313). Martin et al. showed that when Mtb-containing apoptotic cells were efferocytosed by uninfected macrophages in vivo, the highly compartmentalized bacteria are delivered to the lysosome and killed (309). This may explain the bacteria’s strategy of avoiding apoptosis for survival. It is important to note that macrophage-neutrophil interactions are not always beneficial to the host. The Trojan horse model is used by a number of pathogens, including Leishmania (314), Chlamydia (315), and Mycobacterium marnium (316), whereby neutrophil death is induced and efferocytosis is used to establish a productive infection in macrophages. Furthermore, while interaction of macrophages with dead neutrophils helped to reduce replication of Leishmania major in C57BL/6 mice, it exacerbated intramacrophage parasite growth in BALB/c mice (317).  The Burkholderia cepacia complex (Bcc) is a group of bacteria that are ubiquitous in nature, existing in the rhizosphere, ground water, and in association with amoebae, insects, plants, and animals (318). The Bcc are also opportunistic pathogens that cause respiratory infections in patients with cystic fibrosis (CF) and chronic granulomatous disease (CGD) (178). The Bcc can persist in immunocompromised lungs and survives intracellularly within host cells, including macrophages and epithelial cells, by subverting antimicrobial mechanisms (319). B. cenocepacia is intrinsically resistant to many host-derived antimicrobial peptides (208). It also delays phagosomal maturation and acidification (320), assembly of the NADPH oxidase (236), and inhibits phagosomal fusion with lysosomes (224). Survival in macrophages is associated with escape from the endocytic pathway to the cytoplasm (227). A number of studies using modified gentamicin-protection assays to evaluate intracellular growth have shown that B. cenocepacia survives with little to no replication depending on the    66 bacterial strain and macrophage type used (224-227). However, another study showed that the B. cenocepacia could replicate up to 3 log fold within 24 hours in various macrophage types (228). The intrinsic resistance of B. cenocepacia to gentamicin has likely contributed to these confounding results. Recent studies examining the intramacrophage growth of B. cenocepacia using aminoglycoside-sensitive strains in the standard gentamicin-protection assay (230) have shown that B. cenocepacia replicates within in U937 and RAW 264.7 macrophage cell lines but not in macrophages derived from the commonly used THP-1 human monocyte cell line (230, 231). Thus the ability of the Bcc to replicate intracellularly within primary human macrophages remains unclear. B. cenocepacia is killed primarily by ROS within human neutrophils and can induce neutrophil apoptosis at high multiplicities of infection (237, 255), (Chapter 2). The aim of this chapter was to study the collaborative effects of primary human monocyte-derived macrophages (MDM) and neutrophils on survival of B. cenocepacia. We also investigated the potential mechanisms that provide a growth benefit to B. cenocepacia in the presence of macrophages.  3.2 Materials and methods 3.2.1 Isolation of cells Human venous blood was collected from adult volunteers according to the University of British Columbia Clinical Research Ethics Board protocol C04-0193. Immune cells were separated using centrifugation with Ficoll-Paque Plus (GE Healthcare). Peripheral blood mononuclear cells (PBMCs) were washed and monocytes were isolated by plastic adherence in serum-free DMEM. Monocytes were cultured in plastic flasks at 37°C under 5% CO2 with DMEM containing 4.5 g/l glucose, 10% autologous serum, 10 ng/ml GM-CSF (PreproTech),    67 2mM glutamax, 100 units/ml penicillin, 100 µg/ml streptomycin, and 250 ng/ml amphotericin B (Gibco) for differentiation into macrophages. GM-CSF was included to increase cell survival and to differentiate towards an alveolar macrophage-like phenotype (261). After 6 days macrophages were harvested using trypsin, washed in DMEM containing 20% (v/v) FCS to neutralize residual trypsin activity, and resuspended in DMEM supplemented with 4.5 g/l glucose, 10% (v/v) pooled normal human serum (PNHS) and 2 mM glutamax. Enumeration and confirmation of viability was performed using Trypan blue dye exclusion as assessed using a haemocytometer. Neutrophils were isolated by Ficoll-Paque centrifugation and dextran sedimentation and resuspended in R10 for bacterial infections or in DMEM containing 4.5 g/l glucose, 10% (v/v) PNHS and 2 mM glutamax for co-culture with macrophages. Human serum was collected from normal healthy donors in unhepranized serum tubes and stored at -80°C. To prepare PNHS, serum from at least 5 donors was thawed, pooled, and refrozen in aliquots. Human serum was used within 3 months at -80°C. 3.2.2 Bacterial strains and growth conditions B. cenocepacia strain K56-2 is a prototypic isolate of the transmissible ET12 clone and has been described previously (244, 321). B. cenocepacia strain MH1K (gentamicin-sensitive K56-2) was generously provided by Dr. Miguel Valvano (230). Bacteria were stored at -80°C in Mueller Hinton (MH) II Broth with 8% (v/v) dimethyl sulfoxide (DMSO) or maintained on Columbia agar containing 5% sheep blood (PML Microbiologicals) for a maximum of 2 passages. Bacteria were grown overnight in 3 ml cultures of Luria broth (LB) (10 g/l tryptone, 5 g/l yeast extract, 10 g/l sodium chloride) at 37°C and 250 rpm shaking. Overnight cultures were washed in phosphate-buffered saline (PBS), diluted in R10 for neutrophils infections or DMEM for all other experiments, and cultured for approximately 2 h until reaching the desired OD600    68 corresponding to 108 CFU/ml. Bacteria were opsonized with 10% (v/v) PNHS for 30 min tumbling at 37°C and diluted in culture media to appropriate concentration. LB media components were purchased from BD (Franklin Lakes, NJ). 3.2.3 B. cenocepacia growth curves Overnight cultures of B. cenocepacia isolates were adjusted to an OD600 nm of approximately 0.1 in LB. Diluted overnight culture was added to LB or LB with a final volume of 10% (v/v) PNHS at an approximate OD600 nm of 0.01 in a Bioscreen 100 well honeycomb plate. Three technical replicates were performed with appropriate controls with and without serum were used as blanks. Growth at 37°C was monitored in a Bioscreen C machine (TYPE FP-1100-C, Oy Growth Curves Ab Ltd.) with measurement intervals of 15min and continuous shaking, controlled by EZ experiment software as previously described (73). 3.2.4 Neutrophil challenge with B. cenocepacia and incubation with macrophages Freshly isolated neutrophils were diluted in R10. To inhibit ROS, neutrophils were incubated for 30 min with 10 µM diphenyleneiodonium (DPI, Sigma), a potent inhibitor of the NADPH oxidase. Opsonized K56-2 was diluted in R10 and added to neutrophils in 5 ml polypropylene tubes at a multiplicity of infection (MOI) of 5. Tubes were tumbled at 37°C for 30 min then washed thrice in PBS (250xg for 5 min). Neutrophils were resuspended in DMEM containing 4.5 g/l glucose, 10% (v/v) PNHS and 2 mM glutamax and 1 µM DPI where indicated for co-culture with macrophages. Macrophages were seeded at 6x104 cells/well in 96-well plates and B. cenocepacia-challenged neutrophils were added at a 1:1 neutrophil to macrophage ratio. Macrophages or neutrophils alone, with and without DPI, were included as controls. Co-cultures were incubated 37°C under 5% CO2 for 24 h and supernatants were subsequently serially diluted and CFU plated.    69 3.2.5 Co-culture of macrophages, neutrophils, and B. cenocepacia and cytokine analysis Macrophages were seeded in 96-well tissue culture plates at 6x104 cells/well in DMEM supplemented with 4.5 g/l glucose, 10% (v/v) PNHS and 2 mM glutamax. Freshly isolated neutrophils were added to macrophages at a 1:1 or 10:1 neutrophil to macrophage ratio and opsonized K56-2 was subsequently added at a MOI of 0.1 (6x103 per well).  Co-cultures were incubated at 37°C under 5% CO2 for 24 hours and supernatants were subsequently serially diluted and CFU plated. Remaining supernatants were transferred to v-bottom 96-well plates and spun at 4000xg to pellet residual cells. Cell-free supernatants were frozen at -80°C for cytokine analysis. Cytokines released into supernatants were quantified using sandwich ELISAs for IL-1β, IL-6, IL-10, IL-12p70,  (eBioscience) and IL-8 (BD Pharmaceuticals). 3.2.6 Phagocytosis of B. cenocepacia in co-culture with macrophages and neutrophils Macrophages were seeded on acid-washed coverslips in 24-well tissue culture plates at 7.5x104 cells/well in DMEM containing 4.5 g/l glucose, 10% (v/v) PNHS and 2 mM glutamax. Neutrophils were added at a 1:1 ratio with macrophages and plates were spun at 250xg for 5 min to bring neutrophils onto coverslips. Opsonized K56-2 was added at a MOI of 10. At 30 min, 1 h, 2 h, and 4 h post-infection, coverslips were washed twice in PBS and stained with Diff-Quik (Dade Behring, IL). Cells were visualized using an upright Olympus BX50F microscope. Greater than 200 macrophages and 200 neutrophils were scored at each time point as containing zero, one to five, or five or more bacteria per cell. 3.2.7 Gentamicin protection assay A gentamicin protection assay was used to evaluate intramacrophage growth of B. cenocepacia as previously described (230). Macrophages were seeded at 4x104 cells per well in 96-well plates as described above. Opsonized bacteria diluted in DMEM containing 4.5 g/l    70 glucose and 10% (v/v) PNHS were added to macrophages at an MOI of 1 (time = 0 h p.i.) and cells were incubated for 2 hours at 37°C under 5% CO2. Infected macrophages were washed thrice in PBS and incubated in DMEM with 10% (v/v) FBS containing 50 µg/ml of gentamicin for 30 min to kill extracellular bacteria. Media was then removed and replaced with fresh medium containing 10 µg/ml of gentamicin for the remainder of the experiment. At 3 h, 7 h, and 12 h post-infection, macrophages were washed in PBS and lysed in 1% Triton X-100 in water. Lysates were serially diluted and recovered viable intracellular bacteria were quantified by plating on LB agar.  3.2.8 Bacterial culture with macrophages Macrophages were diluted in DMEM containing 4.5 g/l glucose, 10% (v/v) autologous serum and 2 mM glutamax and were seeded at 6x104 cells/well in a 96-well tissue culture plate. The next day media was removed and replaced with fresh media containing opsonized K56-2 at MOI 0.1. Where indicated, macrophages were pretreated with 10 µM cytochalasin D for 30 minutes and cultured with K56-2 in media containing 5 µM cytochalasin D. Conditioned media was collected from 24 hour macrophage culture supernatants and filtered through a 0.4-µm filter. Fresh conditioned media was diluted 1:2 or 1:4 with fresh media and used for incubation with K56-2. K56-2 was cultured with macrophages or macrophage-conditioned media for 24h at 37°C under 5% CO2. Culture supernatants were subsequently serially diluted and CFU plated on LB agar. Remaining supernatants were transferred to v-bottom 96-well plates and spun at 3000xg to pellet residual cells and cell-free supernatants were used to assess lactate dehydrogenase (LDH) release.    71 3.2.9 Bacterial culture with lysed macrophages Macrophages were detached with trypsin and counted on day 6 of differentiation. An aliquot was pelleted, the media discarded, and the cells were lysed by freezing on dry ice and immediately thawing at 37°C for 5 cycles with vortexing. Lysis was confirmed by visualization with Trypan blue exclusion and by examining LDH release in culture supernatants using a cytotoxicity detection kit (Roche Applied Science). Lysates were resuspended to their original concentration and added to wells of a 96-well plate at 6x104 freeze-thawed macrophages per ml. Intact macrophages and media alone conditions were included as controls. The next day opsonized K56-2 was added at 6x103 CFU/ml to each condition and plates were incubated at 37°C under 5% CO2. After 24 h, culture supernatants were serially diluted and CFU plated on LB agar.  3.2.10 Transwell assays Macrophages diluted in DMEM containing 4.5 g/l glucose, 10% (v/v) autologous serum and 2 mM glutamax and were seeded at 1.8x105 cells/well in a 24-well tissue culture plate. The next day media was removed and replaced with either fresh media or media containing opsonized K56-2 at 3x104 CFU/ml (MOI 0.1). Transwell inserts (0.4-µm, VWR) were added and either media or K56-2 in media was added to the top. Growth of bacteria above and below transwell inserts, in wells with and without macrophages was measured by serially diluting culture supernatants after 24 h incubation at 37°C under 5% CO2 and plating CFU on LB agar plates. Media from media-control compartments above and below transwells were cultured on LB agar plates to ensure lack of bacterial growth.     72 3.2.11 Lactate dehydrogenase assay Supernatants from cell stimulations were transferred to 96-well v-bottom plates and spun at 3000xg for 5 min to pellet any residual cells. Cell-free supernatants were diluted 1:10 in PBS and LDH release was measured using a cytotoxicity (LDH) kit (Roche Applied Science) according to manufacturer’s instructions. Control samples included medium alone (blank) and cells treated with 1% Triton X-100 overnight. Data are presented as a percentage of the maximum LDH release (Triton X-100) after subtraction of blank. 3.2.12 Statistics Graphs display mean ± SEM and were generated with Prism 5 (GraphPad Prism). Statistical significance was determined by performing a Student’s t test.  3.3 Results 3.3.1 Macrophages promote bacterial survival from B. cenocepacia-infected neutrophils In addition to its role in removing apoptotic cells and resolving inflammation, efferocytosis has recently been described as an innate antibacterial mechanism (309). As discussed above, Mtb survives intracellularly in macrophages within arrested, immature phagosomes and inhibits macrophage apoptosis, which is associated with reduced bacterial growth (313). A strategy for Mtb is to induce necrosis to evade host defenses and allow bacterial escape and dissemination (312). Martin et al. found that when apoptotic cells were efferocytosed by uninfected macrophages in vivo, the compartmentalized bacteria was delivered to the lysosome and killed (309).  To determine whether macrophages could contribute to killing of B. cenocepacia within neutrophils, especially in the absence of ROS, B. cenocepacia-infected neutrophils were co-   73 incubated with human monocyte-derived macrophages (MDM) and the number of CFU was determined after 24 hours. Surprisingly, co-incubation with macrophages resulted in a significant increase in the CFU of B. cenocepacia-infected neutrophil culture supernatants (Figure 3.1). This effect was independent of the ability of neutrophils to produce ROS to kill B. cenocepacia since culture of macrophages with either normal or DPI-treated neutrophils resulted in a 29-fold and 6-fold increase in supernatant CFU, respectively. The lower enhancement of growth seen in DPI-treated neutrophils may be due to the bacteria reaching stationary phase since the concentration of B. cenocepacia was much higher.         Figure 3.1  Macrophages enhance survival of B. cenocepacia following phagocytosis by neutrophils.  Neutrophils (PMN) were challenged with B. cenocepacia at MOI 5 for 30 minutes, washed thrice, and cultured with macrophages (MDM) for 24 hours in 96-well plates. N ≥ 7. *P ≤ 0.05.  3.3.2 Effect of macrophages and neutrophils on survival of B. cenocepacia The ability of macrophages to enhance growth of B. cenocepacia when in co-culture with B. cenocepacia-infected neutrophils could be caused by a number of mechanisms. For example, macrophages release various pro- and anti-inflammatory mediators that could affect the  Raw TurduckinNormalDPI100101102103104105106107108109PMNPMN + MDM**CFU/ml   74 antimicrobial effector activity of neutrophils. Alternatively, the ability of macrophages to enhance growth of B. cenocepacia in this system could be entirely neutrophil-independent.  To determine the ability of macrophages to enhance the growth of B. cenocepacia in the absence of neutrophils, we cultured B. cenocepacia in 96-well plates i) with and without neutrophils, ii) with and without adhered macrophages or iii) with both neutrophils and macrophages at a 1:1 or 10:1 ratio. Bacteria were added at a MOI of 0.1 bacteria:macrophage or neutrophil ratio with a low starting input of 3x104 CFU/ml.  In this system. B. cenocepacia grew approximately 2-log fold within 24 hours in DMEM supplemented with 10% (v/v) PNHS and 2mM glutamax. As seen in Figure 3.2, macrophages enhanced the growth of bacteria alone by an additional 17"fold. This was in sharp contrast to the bactericidal effects of neutrophils, which significantly decreased the growth of B. cenocepacia approximately 3-fold when incubated at MOI 0.1 or 10-fold when incubated with ten times the number of neutrophils. It should be noted that since co-cultures were plated in 96-well plates, the level of interaction between bacteria and immune cells is much lower compared to experiments where cells and bacteria are tumbled together and may contribute to the lower than expected level of bacterial killing by human neutrophils.  When macrophages and neutrophils were co-cultured at a 1:1 ratio with B. cenocepacia, macrophages had a dominant effect over neutrophils since bacterial growth was still enhanced by approximately 5-fold. We found that approximately ten times the number of neutrophils compared to macrophages was required to abolish the enhanced growth provided by macrophages. This data suggests that macrophages not only enhance the growth of B. cenocepacia but also have a dominant effect over neutrophils, requiring a large number of neutrophils to be recruited to contain the infection.    75                 Figure 3.2  Effect of macrophages and neutrophils on the survival of B. cenocepacia.  K56-2 was cultured in 96-well plates in DMEM + 10% (v/v) PNHS with and without adherent MDM, with and without PMN, or with MDM and PMN at a MOI of 0.1. PMN were added at a 1:1 or 10:1 ratio to MDM. After 24 hours culture supernatants were serially diluted and CFU plated. N ≥ 5. *P ≤ 0.05 ; **P ≤ 0.01. ; ***P ≤ 0.001; n.s. not significant.    3.3.3 Modulation of B. cenocepacia-induced inflammatory mediators by neutrophils  In addition to their well-defined role as potent antimicrobial phagocytes important in host defense, neutrophils also exhibit immunomodulatory effects that govern inflammation. Neutrophils are capable of inhibiting inflammatory responses by degrading pro-inflammatory but not anti-inflammatory cytokines with neutrophil elastase (322), scavenging IL-1β (323), and producing the anti-inflammatory cytokine IL-1 receptor antagonist even in the absence of IL-1 itself (324) Furthermore, phagocytosis of apoptotic neutrophils by macrophages elicits an anti-inflammatory response characterized by reduced secretion of pro-inflammatory cytokines and heightened production of anti-inflammatory cytokines (125, 126). Thus although macrophages exerted a dominant effect over neutrophils on the growth and survival of B. cenocepacia,  K56-2K56-2 + 1x PMNK56-2 + 10x PMNK56-2 + MDMK56-2 + MDM + 1x PMNK56-2 + MDM + 10x PMN103104105106107108*****n.s.*CFU/ml   76 neutrophils may function to modulate cytokine production of macrophages to dampen inflammation.  The release of pro- and anti-inflammatory cytokines and chemokines from macrophages after 24-hour culture with B. cenocepacia with and without neutrophils was examined. Both whole B. cenocepacia and its isolated LPS induce secretion of high levels of pro-inflammatory cytokines and chemokines from human and murine monocytes and macrophages (325-330). As seen in Figure 3.3, B. cenocepacia was a potent inducer of the pro-inflammatory cytokines IL-6, IL-1β, IL-12p70 and the anti-inflammatory cytokine IL-10 from human monocyte-derived macrophages. B. cenocepacia induced IL-8 from both macrophages and neutrophils. The addition of neutrophils to macrophages with B. cenocepacia at a ratio of 1:1 did not have a significant impact on the production of any of the cytokines or chemokines investigated. When neutrophils were added in excess at a ratio of 10 neutrophils : 1 macrophage, the release of IL-6, IL-1β, and IL-10 significantly decreased suggesting a dampening of inflammation. IL-12p70 did show a similar trend, however the effect was diluted by macrophages from one donor that exhibited abnormally high production of IL-12p70. Total IL-8 levels in co-cultures did not decrease with the addition of neutrophils, likely due to the production of IL-8 by neutrophils themselves. These findings indicate that neutrophils do not reduce the production of inflammatory mediators by macrophages when at an equal concentration. However, neutrophils may play a role in dampening macrophage-induced inflammation at higher concentrations, which many in fact be physiologically relevant based on the model of excessive neutrophil infiltration during B. cenocepacia infections.       77 Figure 3.3 B. cenocepacia-induced inflammatory mediators. MDM were incubated with B. cenocepacia isolate with and without the presence of PMN at a 1:1 or 10:1 PMN to MDM ratio. PMN with and without B. cenocepacia were included as controls. Cytokine levels in 24 hour cell-free supernatants were determined by ELISA. N = 4 for IL-1β and N = 6 for all other cytokines. *P ≤ 0.05 ; ***P ≤ 0.001; n.s. not significant.    MDMMDM + PMNMDM + 10x PMNMDM + K56-2MDM + PMN + K56-2MDM + 10x PMN + K56-2PMN10x PMNPMN + K56-210x PMN + K56-201000020000300004000050000***IL-6 (pg/ml)MDMMDM + PMNMDM + 10x PMNMDM + K56-2MDM + PMN + K56-2MDM + 10x PMN + K56-2PMN10x PMNPMN + K56-210x PMN + K56-20100000200000300000400000n.s.IL-8 (pg/ml)MDMMDM + PMNMDM + 10x PMNMDM + K56-2MDM + PMN + K56-2MDM + 10x PMN + K56-2PMN10x PMNPMN + K56-210x PMN + K56-2050010001500***IL-10 (pg/ml)MDMMDM + PMNMDM + 10x PMNMDM + K56-2MDM + PMN + K56-2MDM + 10x PMN + K56-2PMN10x PMNPMN + K56-210x PMN + K56-2K56-20200400600800*IL-1β (pg/ml)MDMMDM + PMNMDM + 10x PMNMDM + K56-2MDM + PMN + K56-2MDM + 10x PMN + K56-2PMN10x PMNPMN + K56-210x PMN + K56-202004006008001000n.s.IL-12p70 (pg/ml)   78 3.3.4 Phagocytosis of B. cenocepacia by macrophages and neutrophils As shown in Figure 3.2, macrophages had a dominant effect over neutrophils in their ability to affect survival and growth of B. cenocepacia. We hypothesized that this may be due to a greater ability of human macrophages to phagocytose B. cenocepacia compared to neutrophils when in co-culture in our in vitro system. To assess the competition between phagocytes to engulf B. cenocepacia, we cultured macrophages and neutrophils together at a 1:1 ratio on acid-washed coverslips with bacteria. Neutrophils were spun down onto coverslips prior to infection to ensure that bacteria had the same opportunity to interact with the two cell types. After 30 minutes, 1 hour, 2 hours, and 4 hours, coverslips were washed with PBS and stained with Diff-Quik to visualize bacteria within macrophages and neutrophils. As seen Figure 3.4, by 1 hour of incubation with B. cenocepacia, significantly more macrophages remained uninfected compared to neutrophils. Most neutrophils that were infected contained less than 5 bacteria per cell. By 2 hours post-infection, the number of neutrophils and macrophages containing no bacteria or 1-5 bacteria per cell did not significantly differ. Approximately 50% of each cell type contained at least one bacterium. By 4 hours of incubation, the majority of neutrophils and macrophages contained at least one bacterium and slightly more neutrophils remained void of bacteria. Seventy percent of macrophages contained 5 or more bacteria per cell and many cells were filled with high levels of bacteria (10 or more per cell). Fifteen percent of macrophages contained no bacteria and 15% contained between 1 and 5 bacteria per cell. In contrast, only 41% of neutrophils contained more than 5 bacteria per cell and 38% contained between 1 and 5 bacteria per cell.  This data shows that the dominant effect of macrophages over neutrophils in impacting B. cenocepacia survival and growth is not due to increased phagocytosis of macrophages compared    79 to neutrophils. Although neutrophils were faster to phagocytose B. cenocepacia than macrophages, by 2 hours incubation with bacteria, neutrophils and macrophages contained the same profile of phagocytes containing no bacteria, 1 to 5 bacteria per cell, and greater than 5 bacteria per cell. By 4 hours, a far greater number of bacteria are contained in macrophages compared to neutrophils.  Figure 3.4 Relative phagocytosis of B. cenocepacia by neutrophils and macrophages Neutrophils and macrophages were co-incubated at a 1:1 ratio on glass coverslips with B. cenocepacia at MOI 10. At each timepoint, coverslips were washed and stained with Diff-Quik. Greater than 200 of each cell type was counted per time point at 40x magnification and each cell was classified as containing either 0 bacteria, 1-5 bacteria, or ≥ 5 bacteria. A. Microscopic cell counts. B. Representative images of macrophages (i) and neutrophils (ii) at each time point. N ≥ 4. *P ≤ 0.05 ; **P ≤ 0.01. ; ***P ≤ 0.001               01- 5> 5 01- 5> 5 01- 5> 5 01- 5> 5020406080100NeutrophilMacrophage*******ABacteria / cell% of cell type   80 Several factors may contribute to the observation that macrophages contain more bacteria at later time points compared to neutrophils. We suspected that neutrophils may lyse during our time course since their cytoplasms are filled with bacteria at earlier time points. However, preliminary experiments found no evidence of this when we incubated B. cenocepacia with macrophages and neutrophils alone and compared the level of LDH released as a percentage of the maximum LDH released from each cell type treated with Triton X-100 (Figure 3.5). In fact, macrophages were more susceptible to B. cenocepacia-induced LDH release. Since there is evidence that B. cenocepacia grow in macrophages (225, 228, 231) but are killed in neutrophils, some of the bacteria detected in macrophages at later time points may originate from intracellular replication. Macrophages were also observed to have a larger cytoplasm and may simply have a greater capacity to fill with more bacteria compared to neutrophils.                Figure 3.5 B. cenocepacia-induced phagocyte lysis. PMN and MDM were cultured with B. cenocepacia at a MOI of 0.1 or 1 for 24 hours. Cell-free culture supernatants were removed and assayed for LDH using a cytotoxicity detection kit. Data is represented as a percentage of the maximum LDH release of each cell type incubated with 1% Triton X-100 (% of Max). N=3. *P ≤ 0.05.   SpontaneousMOI 0.1MOI 1010203040MDMPMN *% of Max (LDH)   81 3.3.5 Intracellular growth of B. cenocepacia in human monocyte-derived macrophages As discussed above, the ability of B. cenocepacia to replicate within macrophages is controversial and several contradictory reports exist. Furthermore, B. cenocepacia survival and growth depends on the types of macrophage used (231). A recent study by Al-Khodor et al showed that B. cenocepacia isolate J2315 grew approximately 2 log-fold in 24 within human MDM (228). We therefore investigated the ability of opsonized B. cenocepacia to grow in human MDM in our system. Macrophages were challenged with a gentamicin-sensitive strain of K56-2, MH1K, in a gentamicin-protection assay as previously described (230) to evaluate intracellular growth over time. The growth rate of MH1K did not differ from that of K56-2 (Appendix A, Figure A.1) and MH1K had a minimum inhibitory concentration (MIC) of ≤ 1 µg/ml of gentamicin. Macrophages were incubated with bacteria at a MOI of 1 for 2 hours, washed, and incubated in media containing 50 µg/ml of gentamicin for 30 minutes. The first time point used therefore occured at 3 hours post-infection. For later time points, macrophages were cultured in media containing 10 µg/ml of gentamicin to prevent the growth of bacteria released from any lysed macrophages. No CFU were detected in culture supernatants at any of the time points observed.  As seen in Figure 3.6, B. cenocepacia isolate MH1K grew approximately 1 log-fold in 11 hours in human MDM cultured in DMEM containing 10% (v/v) PNHS. A significant amount of growth was seen as early as 7 hours post-infection. We also assayed time points at 15h and 24h post-infection. However, we observed that many macrophages appeared lysed by microscopy. Since any bacteria released from lysed macrophages would be rapidly killed in 10 µg/ml of gentamicin, the number of CFU detected once macrophages start to lyse is an underestimate of intracellular bacterial growth. Indeed, we saw a decrease in the number of CFU detected by 15    82 hours post-infection at an MOI of 1 and at earlier time points using higher MOIs (data not shown).                    Figure 3.6 Intracellular growth of B. cenocepacia within macrophages. Human MDM were incubated with gentamicin-sensitive K56-2 (MH1K) with a MOI of 1 for 2 hours. MDM were washed thrice and incubated in media containing 50 µg/ml of gentamicin for 30 minutes to kill extracellular bacteria. Media was then replaced with media containing 10 µg/ml of gentamicin to prevent intracellular growth. At indicated time points, MDM were washed and lysed in 1% Triton X-100. N ≥ 6. *P ≤ 0.05 ; **P ≤ 0.01.   3.3.6 Effect of macrophage-released factors on growth of B. cenocepacia Previous studies have shown that macrophage-released mediators can enhance the growth of bacteria in vitro (331, 332). A number of experiments were performed in order to investigate whether macrophages released a factor that contributed to the enhanced growth of B. cenocepacia in co-culture of macrophages.  The necessity of macrophage/B. cenocepacia contact for enhanced bacterial growth was examined using a transwell system. Macrophages were seeded in 24-well plates and cultured with either B. cenocepacia or separated from B. cenocepacia by 0.4 µm transwell inserts. The experimental set-up allowed for bacteria and macrophages to communicate via secreted    3h7h11h103104105106**Hours p.i.CFU/ml   83 molecules but not by physical interaction. Macrophages and B. cenocepacia were cultured together or separated by inserts for 24 hours and CFU in supernatants were determined. B. cenocepacia was also cultured below or above transwell inserts in the absence of macrophages as controls. No CFU were found to pass through the pores of the inserts after 24-hour culture. As seen in Figure 3.7a, B. cenocepacia cultured on the same side of inserts as macrophages exhibited enhanced growth, however bacteria separated from macrophages did not. Growth of B. cenocepacia in the absence of macrophages did not differ from its growth when physically separated form macrophages by transwell inserts.  The transwell inserts are separated from the bottom of the transwell by a distance of 1.0 mm. If macrophages did release a factor to enhance growth of B. cenocepacia, this distance may have been too great for the diffusion of the molecule to reach the top of the transwell insert and impact the bacteria. We therefore cultured B. cenocepacia in macrophage-conditioned media to ensure that any macrophage-released factors had adequate contact with bacteria. Briefly, media from macrophages cultured for 24 hours at 37°C under 5% CO2 was filtered and diluted either 1:2 or 1:4 with fresh media to ensure that media was not entirely spent of nutrients required for bacterial growth. B. cenocepacia was cultured with diluted conditioned media for 24 hours in 96-well plates. Bacterial culture in wells containing macrophages was included as positive controls. As seen in Figure 3.7b, growth of B. cenocepacia in regular media did not differ from that of bacteria in macrophage-conditioned media at either dilution. Only culture with intact macrophages enhanced the growth of B. cenocepacia after 24 hours.         84                               Figure 3.7 Effect of macrophage-released factors on B. cenocepacia growth. A. Macrophages were seeded in the bottom of 24-well tissue culture plate and B. cenocepacia isolate K56-2 was cultured above or below 0.4 µm transwell inserts in wells with or without seeded macrophages. After 24 hours, CFU in B. cenocepacia-containing compartments were counted. N = 6. B. Conditioned media from 24 hour macrophage culture was filtered and diluted and used for culture with K56-2. N = 4. *P ≤ 0.05 ; ***P ≤ 0.001.   3.3.7 Role of phagocytosis in macrophage-induced growth of B. cenocepacia  Conditioned media contains factors that are released from resting macrophages but is not representative of the repertoire of molecules secreted from macrophages stimulated with B.    K56-2K56-2 + MDM100101102103104105106107108K56-2 on top (separated)K56-2 on bottom (together)*ACFU/mlK56-2 K56-2 + MDM K56-2 + 1/4 cond. media K56-2 + 1/2 cond. media100101102103104105106107108***BCFU/ml   85 cenocepacia. Cytochalasin D (Cyt D) is a potent inhibitor of actin polymerization required for endocytosis and exocytosis (333). Macrophages were treated with Cyt D to inhibit the phagocytosis of B. cenocepacia and the release of possible bacterial growth factors during incubation the bacteria. As seen in Figure 3.8a, the ability of macrophages to induce growth of B. cenocepacia was lost when macrophages were pre-treated with Cyt D, suggesting that macrophages are actively involved in enhancing the growth of B. cenocepacia.  Together with the lack of enhanced growth seen in our assays using transwell inserts and macrophage-conditioned media, this finding indicates that the enhanced growth of B. cenocepacia that is induced by macrophages requires that the bacteria be taken up into the cell. We hypothesized that the ultimate goal of the bacteria might be to enter macrophages to induce lysis. Subsequent breakdown of cellular components may provide a rich nutrient source to enhance bacterial growth. The release of LDH from normal macrophages or those pre-treated with Cyt D cultured with B. cenocepacia at MOI 0.1 was measured after 24 hours (Figure 3.8b). Untreated macrophages cultured for 24 hours released approximately 7% of the maximum amount of LDH as determined by macrophages treated with 1% Triton X-100. B. cenocepacia induced approximately 24% of  maximum LDH release from macrophages. In contrast, when macrophages were pre-treated with Cyt D to inhibit phagocytosis of bacteria, the level of LDH released when cultured with B. cenocepacia was identical to that of macrophages alone. Uptake of B. cenocepacia therefore did induce low but significant macrophage death after 24 hours. To determine whether macrophage lysis contributed to the growth of B. cenocepacia, macrophages were lysed by 5 cycles of freeze-thawing and cultured with bacteria. Intact macrophages were included as a positive control. As seen in Figure 3.8c, the addition of macrophage lysates to B. cenocepacia had no observable effect on its growth.     86  Figure 3.8 Uptake of B. cenocepacia by intact macrophages is necessary for enhanced bacterial replication. A. Normal MDM or those pre-treated with 5 µM of Cyt D were incubated with B. cenocepacia isolate K56-2 for 24 hours and CFU counted. B. cenocepacia alone was included as a control. N = 8. B. LDH released from normal MDM or those pre-treated with 5 µM of Cyt D following challenge with K56-2 for 24 hours, represented as percentage of maximum release of macrophages treated with 1% Triton X-100. N = 6. C. Growth of K56-2 following 24-hour incubation with live MDM or MDM lysed by 5 cycles of freeze-thawing. N = 4. *P ≤ 0.05 ; **P ≤ 0.01. ; ***P ≤ 0.001.  Collectively, this data indicates that uptake of B. cenocepacia by human macrophages is required to enhance bacterial growth. Although B. cenocepacia induces cell death in a significant proportion of macrophages, this is likely a consequence of bacterial uptake and intracellular   K56-2 K56-2 + Cyt DK56-2 + MDMK56-2 + MDM + Cyt D100101102103104105106107108*******CFU/mlK56-2K56-2 + MDMK56-2 + lysed MDM100101102103104105106107108***CFU/mlMDMMDM + Cyt DK56-2 + MDMK56-2 + MDM + Cyt D K56-2K56-2 + Cyt D0102030******% of Max (LDH)A BC   87 replication rather than a primary goal for growth, since bacterial growth was not enhanced when cultured in cell lysates. This suggests that B. cenocepacia establishes a replication niche in human macrophages that allows it to not only escape from neutrophil killing but also greatly enhance its growth compared to in media alone in vitro. Whether growth of B. cenocepacia is enhanced in macrophages in vivo compared to its growth extracellularly on the surface of the human lung remains unknown.  3.4 Discussion The purpose of this chapter was to examine the ability of human macrophages to contribute to the capacity of neutrophils to kill B. cenocepacia, especially in the absence of ROS. Since B. cenocepacia is able to survive intracellularly within neutrophils in the absence of ROS, we hypothesized that macrophages may utilize efferocytosis to channel B. cenocepacia within neutrophils to the lysosome. As previously described for Mtb, delivery of bacteria to the lysosome via engulfed apoptotic cells can prevent the bacteria from subverting macrophage antibacterial mechanisms, presumably due to the extra layer of compartmentalization (334).  Surprisingly, we found that macrophages significantly enhanced the growth of B. cenocepacia when co-cultured with B. cenocepacia-infected neutrophils (Figure 3.1). This effect occurred for both normal and ROS-deficient (DPI-treated) neutrophils. Upon further investigation, this effect of macrophages on the growth of B. cenocepacia was found to be independent of neutrophils, since macrophages alone enhanced the growth of B. cenocepacia when cultured in the absence of neutrophils (Figure 4.2). The enhanced growth seen in Figure 4.1 with B. cenocepacia-infected neutrophils therefore may be an effect of macrophages on bacteria that have escaped from neutrophils. While the MOI of 5 used in this experiment allowed    88 most neutrophils to phagocytose a low number of bacteria per cell in this system, there was a high variability in phagocytosis and some cells contained higher numbers of bacteria per cell (Chapter 2,(237)). Remnants of neutrophils filled with bacteria following infections were observed in cytospins by microscopy during these studies, indicating that neutrophils may lyse when they phagocytose too many B. cenocepacia. Therefore, the growth enhancing effect of macrophages may be specifically on B. cenocepacia that have escaped through neutrophils via cell lysis.  As described above, the ability of B. cenocepacia to replicate within human macrophages remains controversial. The gentamicin protection assay, which relies on the poor ability of gentamicin to enter eukaryotic cells, is the standard for assessing intracellular replication by bacteria (229). Infected macrophages are washed and incubated in gentamicin to kill off extracellular bacteria, then at desired time points cells are lysed and CFU are measured. Due to the Bcc’s intrinsic resistance to aminoglycosides and other relevant antibiotics, gentamicin protection assays were traditionally modified to include extremely high levels of gentamicin (≥ 250 µg/ml) in combination with other antibiotics such as ceftazidime (≥ 500 µg/ml), which can have adverse effects on both the bacteria and eukaryotic cells. Even with these high levels of antibiotics, extracellular B. cenocepacia is often still able to survive (Speert laboratory, data not shown). Recently, aminoglycoside-sensitive strains of B. cenocepacia and B. multivorans were constructed that carry a deletion in the operon encoding an AmrAB-OprA-like efflux pump (230, 231). Studies using gentamicin-sensitive strains have revealed that both B. cenocepacia and B. multivorans can replicate in U937 and RAW 264.7 macrophage cell lines but not in macrophages derived from the commonly used THP-1 human monocyte cell line (230, 231). We investigated the ability of a gentamicin-sensitive strain of K56-2 (MH1K, (230)) to replicate intracellularly    89 within human macrophages derived from primary monocytes in autologous serum, which we think is the best in vitro model of physiological human macrophages. Our data indicate that B. cencocepacia is able to replicate approximately 1 log-fold in 11 hours in primary human macrophages.  The intracellular growth of B. cenocepacia has been previously reported (225, 228, 230, 231, 335), however studies have shown that Bcc growth is primarily extracellular (336) and the Bcc is considered an extracellular or facultative intracellular pathogen (337). We found that primary macrophages provide an intracellular niche with a better growth environment compared to media alone. B. cenocepacia exhibited approximately 2 log-fold growth in culture media (DMEM containing 4.5 g/L glucose supplemented with 10% PNHS and 2 mM glutamax) and great than 3 log-fold growth when macrophages were included. Thus, although growth was less than is seen in nutrient-rich media such as Luria Broth, B. cenocepacia grew efficiently in culture media. Therefore we suspect that the macrophage replication niche is rich in nutrient sources in order to provide such enhanced growth. We included PNHS in culture media because isolates K56-2 and MHI1K were found to have limited uptake by human neutrophils without opsonization compared to other isolates of B. cenocepacia and are serum-resistant (Appendix A, Figure A.1, (73)). Growth of B. cenocepacia is however slightly impeded in serum (Appendix A, Figure A.1) and may contribute to the discrepancies in the rate of intracellular growth observed in other studies (230, 231).    We aimed to determine whether macrophages could promote the growth of B. cenocepacia through alternative means in addition to providing a replication niche. A series of experiments were conducted to examine the ability of macrophages to release a growth-promoting factor. Macrophages did not enhance bacterial growth when physically separated by    90 transwell inserts or when uptake of B. cenocepacia was blocked with cytochalasin D. Furthermore, 24 hour macrophage-conditioned media had no impact on bacterial growth, suggesting that macrophages do not secrete a growth-promoting factor. We found that B. cenocepacia did induce a low level of macrophage lysis in our system, however incubation with macrophage lysates did not enhance growth of B. cenocepacia, suggesting that the bacteria do not simply invade macrophages to induce lysis and the release of potential nutrients. As previously mentioned, macrophages may upregulate the expression of potential nutrients upon internalization of bacteria and therefore the macrophage lysates used here are not a perfect representation of bacteria-induced macrophage lysates. However, the absence of any growth benefit of macrophage lysates combined with the relatively low level of macrophage lysis induced by B. cenocepacia suggests that macrophage lysis does not play a prominent role in bacterial growth promotion compared to the observed intracellular growth. Overall, we were not able to identify any additional growth-promoting capabilities of macrophages in addition to providing an intracellular replication niche. The intramacrophage location of B. cenocepacia proliferation remains unknown. Previous reports have shown that engulfed Bcc resides within Bcc-containing vacuoles (BcCVs) that co-localize with early endosomes but delay phagosomal maturation and inhibit fusion with lysosomes (224). Growth in U937 cells was associated with bacterial escape from vacuoles to the cytosol (227, 338). Although bacteria are tagged with ubiquitin and autophagic machinery is recruited, ultimately selective autophagy fails and B. cenocepacia co-localizes with markers of the endoplasmic reticulum (ER) and proliferates (227, 228). Another study was not able to demonstrate co-localization of calnexin in the ER and BcCVs in RAW264.7 macrophages (224). Intracellular survival of B. cenocepacia was dependent on a plasmid-encoded type IV secretion    91 system, though the secreted effector molecules remain unknown (227). Similarly, a role for type III and type IV secretion systems in intracellular replication has been reported for Legionella (339), Brucella (340), Chlamydia (341), and Salmonella species (342).  The intracellular macrophage factors that provide a growth benefit for B. cenocepacia remain to be explored. A study of changes in B. cenocepacia gene expression upon internalization into murine macrophages demonstrated the upregulation of genes involved in motility, including regulators of flagellin and chemotaxis, and in decreasing membrane permeability (343). Changes were not seen in specific virulence factors other than a type VI secretion system, consistent with the opportunistic nature of the Bcc. Metabolic adaptation accounted for approximately a third of differentially regulated genes, particularly in genes responsible for carbohydrate and amino acid transport and metabolism. However, nearly 40% of upregulated genes were poorly characterized and thus may represent undescribed survival mechanisms (318, 343).  The capacity of neutrophils to kill B. cenocepacia was undermined by the presence of macrophages in this system. Despite phagocytosing B. cenocepacia faster, approximately ten times the number of neutrophils compared to macrophages was required to over-ride the bacterial growth promotion conferred by macrophages. Similarly, we only observed that neutrophils exerted immunomodulatory effects on cytokine production by macrophages at high levels. Whether this discrepancy occurs physiologically is unknown. The stationary in vitro system in which phagocytes and bacteria were cultured does not provide optimal interaction between bacteria and neutrophils as a tumbling set-up would. Neutrophils are recruited at high levels to infectious loci so it is quite likely that an excess of neutrophils compared to    92 macrophages might be present during B. cenocepacia infection and have a profound impact on bacterial survival and the inflammatory response. Indeed, we found that B. cenocepacia induced a profound level of IL-8 from macrophages, which is responsible for neutrophil recruitment. An excess of neutrophils was able to decrease both the burden of bacteria and the level of inflammatory mediators that activate other immune cells. We show here that human macrophages provide a rich replication niche for B. cenocepacia that can allow the bacteria to escape killing by neutrophils and proliferate. The ability of B. cenocepacia to survive and replicate intracellularly may contribute to its ability to evade the host response and establish chronic lung infections in some patients.      93 Chapter 4: Recurrent subacute post-viral onset of ataxia associated with PRF1 mutation  4.1 Introduction Hemophagocytic lymphohistiocytosis (HLH) is a classification of immune disorders that lead to immune dysregulation with uncontrolled activation of T cells and macrophages and associated hypercytokinemia (10, 13). Common diagnostic symptoms of FLH include high-grade fever, hepatosplenomegaly, progressive cytopenia, hemophagocytosis in the bone marrow, elevated ferritin levels, low natural killer (NK) cell activity, and hypertriglyceridemia (14). HLH is classified as either primary or familial (FHL), when there is an identified underlying genetic cause or a family history, or secondary or acquired HLH, which tends to manifest at an older age and is generally associated with an underlying infection, rheumatic illness, or malignancy. Overall, a genetic diagnosis can be made in 40-80% of patients with HLH (10, 18, 19) FHL is an autosomal recessive disorder caused by mutations in genes involved in the granule-perforin cytotoxic pathway. The pathway is mediated by cytotoxic lymphocytes (CTL) and NK cells and is the primary means of eliminating abnormal cells arising from intracellular pathogens or malignant transformation (23). Secretory (cytotoxic) granules of CTLs and NK cells are composed primarily of perforin and serine proteases such as granzyme, which when delivered to a target cell can induce rapid apoptosis in as little as 20 minutes (Figure 1.1) (133). Perforin is responsible for mediating the delivery of granzymes released into the intracellular space into the cytosol of target cells, where granzymes cleave specific substrates involved in activating apoptosis, such as BID and effector caspases (134, 135). Genetic mutations involved    94 in FHL have been mapped to perforin1 (PRF1; FHL2) (21), UNC13D (FHL3)(24), STX11 (FHL4) (25) and STXBP2 (FHL5) (26, 344). The locus responsible for FHL-1 is mapped to chromosome 9q21.3, but the specific mutated gene has not been identified (20). While symptoms of neurodegeneration are rare, several reports have identified neurological presentation with eventual progression of classical symptoms of HLH. Central nervous system (CNS) involvement can present as seizures, ataxia, brain lesions, encephalitis, and demyelination consistent with multiple sclerosis (345-348). Childhood neurodegeneration is a diagnostic challenge because it can arise as a primary or secondary consequence of many genetic and acquired etiologies, including from various infectious causes (349). Genetic bases include disorders of metabolism, myelination, vascular integrity and inflammation (350). Dysfunction of the innate immune system is the most common primary inflammatory cause of neurodegeneration (351). The aberrant release of cytokines by activated microglia and circulating monocytes damage the myelin sheath and enhance blood-brain barrier permeability, allowing further recruitment of leukocytes to the CNS.  Inflammatory neurodegenerative disorders rarely have diagnostic clinical or radiographic features and are generally subclassified as affecting primarily the white matter, the gray matter, or both. Even when neuroradiological signs suggest a particular classification, diagnosis remains challenging and requires additional investigations. Consequently, many neurodegenerative disorders do not have a precise diagnosis.  Pro- and anti-inflammatory cytokines can both cause and mitigate neurodegeneration (352, 353). Interleukin (IL)-1β is a potent pro-inflammatory cytokine associated with the induction and maintenance of the innate immune response. It has been implicated in both neurodegeneration and neuroprotection (reviewed by Fogal and Hewett (354)). IL-1β is over-   95 expressed within the CNS of patients with multiple sclerosis, in animal models of experimental autoimmune encephalomyelitis, and systemically in some patients with active HLH with CNS involvement (11, 355, 356). We present a family with two daughters who died of a previously undescribed neurodegenerative disorder triggered by infection. Both girls had biallelic mutations in PRF1 without diagnostic features of FHL2.   4.2 Materials and methods 4.2.1 Patient data and standard protocol approvals  Family members gave informed consent/assent for protocol H09-01228 (University of British Columbia, Vancouver, Canada) and for protocol 76-HG-0238 (National Institutes of Health, Bethesda, MD, USA). Clinical data on each affected individual were obtained through retrospective chart review and interviews of the parents.  4.2.2 Exome sequencing and analysis  Exome sequencing was performed on the nuclear family (both affected sisters, their three unaffected siblings, and both parents; Figure 4.1) as previously described. NextGENe software version 2.2.2 (SoftGenetics, LLC, State College, PA, USA) was used to align sequences to the human reference genome (NCBI, Build 37 v2) and call variants. Each sample alignment was performed allowing one mismatched base and ≥ 85% of the read matching the reference sequence. Variant calling required that the variant was observed in ≥ 20% and ≥ 3 short reads, with a minimum of 5x coverage.  Following generation of a variant list fitting autosomal recessive inheritance by NextGENe Viewer’s Variant Comparison Tool, we ranked the variants using VAR-MD as    96 described. To accept variants for ranking, we required a variant to meet the following criteria in at least four of the seven exomes: a cutoff for genotype confidence score of ≥ 16, or for variants that had scores between 7 and 16, a genotype confidence score of at least one-fourth the coverage value.  4.2.3 Cell stimulation and measurement of cytokines  Peripheral blood mononuclear cells (PBMCs) from patient IV-5, unaffected siblings, parents, and seven healthy controls were purified over a gradient of Ficoll-Paque Plus (GE Healthcare, Piscataway, NJ, USA). PBMCs were diluted in RPMI 1640 supplemented with 10% heat-inactivated fetal calf serum (FCS) and 2mM L-glutamine and plated in 96-well plates at 2x105 cells/well. PBMCs were stimulated in triplicate with 10 ng/ml of E. coli Ultra-Pure LPS (InvivoGen, San Diego, CA, USA) for 4 hours with or without addition of 5 mM ATP for the last hour of incubation. Culture supernatants were spun to remove residual cells and cell-free supernatants were stored at -80°C. IL-1β was measured in thawed supernatants using an ELISA Ready-SET-Go kit (eBioscience, San Diego, CA, USA).  A Luminex-based cytokine/chemokine assessment was performed on patient and healthy adult control whole blood. Whole blood was mixed 1:1 with RPMI 1640 (Invitrogen, Carlsbad, CA, USA) and 200 µl was added to wells of a 96-well tissue culture plate. Cells were stimulated with 10 ng/ml LPS, 1 µg/ml PAM, 100 µg/ml pIC, 10 µg/ml PGN, and 10 µM R848 in 96-well tissue culture plates in duplicate. After 24 hours incubation, cytokine levels (IL-1β, IL-6, IL-8, IL-10, IL-12p40, IL-12p70, IP-10, MIP-1α, MIP-1β, TNF-α, IFN-α2, and IFN-γ) were measured in the supernatants using a Luminex immunobead-based 12-plex assay (Masterplex software, MiraiBio, Alameda, CA, USA).     97 4.2.4 Statistics Data are represented as mean ± SEM. Statistical analysis was performed using SPSS v.19 (IBM, North Castle, NY, USA), applying independent Student t tests. 4.3 Results 4.3.1 Clinical features The proband and her sister were daughters of a con-sanguineous couple of Lebanese origin. The parents were first cousins but had additional loops of consanguinity with a resulting estimated coefficient of inbreeding for the proband of 0.13, the equivalent of the offspring of half-sibs.  The proband (IV-5, Figure 4.1) was born at term after an unremarkable pregnancy and delivery. She had normal early growth and development and good health aside from upper respiratory and middle ear infections. Beginning at 18 months, her weight gain declined, and at 22.5 months, a week after a bout of gastroenteritis with an eczematous rash, she developed ataxia, abnormal eye movements and dysarthria as well as brain white matter abnormalities. She had elevated cerebral spinal fluid (CSF) protein concentration (0.56 g/l, compared to reference: 0.10–0.35 g/l). Bone marrow biopsy showed iron depletion. Additional testing was otherwise unremarkable.  Soon after partial resolution of the ataxia and dysarthria, she developed progressive disease. At age 32.5 months, her brain MRI documented further white matter degeneration, and a skeletal muscle biopsy showed mild variation in fiber size and type II fiber atrophy on light microscopy. Fiber and mitochondrial ultrastructure as well as mitochondrial respiratory chain activity were normal. Her CSF had unremarkable chemistry and cytology and PCR showed no detectable nucleic acid indicative of infection with Cytomegalovirus, Herpes virus type 6,    98 Varicella-Zoster, Epstein–Barr virus, Adenovirus, West Nile virus, Enterovirus, Herpes simplex viruses 1 and 2, and toxoplasmosis. By age 42 months, she developed mild splenomegaly and lymphadenopathy without hepatomegaly or detectable acute viral infection. Two months later, she was admitted to hospital for vomiting, tachycardia, tachypnea and fever (39°C). She had then hypertrophic cardiomyopathy with mild diastolic dysfunction, mild pancytopenia, and elevated AST and ALT levels. Again her bone marrow biopsy was unremarkable except for iron depletion. Her immune studies showed normal numbers and distribution of T, B, and NK cells as well as unremarkable responses to mitogens and normal complement activity. No viral or bacterial pathogens were identified. During her recovery, she manifested persistent neck rigidity suggestive of meningeal inflammation.  Shortly after this hospitalization, she developed seizures. Over the next 17 months, she gradually became blind, lost cognitive and motor skills, and developed dysphagia. Throughout this period, she manifested neck rigidity following each of many infections but had no more recorded fevers until the week before her death when her temperature was 38.5°C. She died at 61 months of age with upper respiratory infection and cardiac failure secondary to left ventricular hypertrophy with outlet obstruction and mitral regurgitation. The family declined an autopsy.  The proband’s sister (IV-3; Figure 4.1) was the product of an uneventful pregnancy. She had mild developmental delay and hearing loss. At age 75.5 months, concurrent with varicella infection, she had a seizure in the context of fever, deterioration of mental status and was diagnosed with varicella meningitis. CT brain imaging showed dilated ventricles and cerebellar swelling. Following 2 weeks of treatment with acyclovir and steroids, she regained baseline neurological function. Subsequently, however, she became ataxic and was unable to walk. Three months after initial presentation her brain MRI showed diffuse white matter changes. She    99 gradually lost additional motor skills and strength; eight months after presentation the nerve conduction velocities of her sensory and motor evoked responses were at the lower limits of normal in the right upper and lower limbs (Right Tibial Motor at ankle: latency 3.22 ms, conduction 40.97 m/s). Her CSF showed normal protein, glucose and amino acid levels. CSF neurotransmitters were normal except for an elevated neopterin level; her skin fibroblasts had normal 6-pyruvoyltetrahy-dropterin synthase activity, excluding 6-pyruvoyltetrahydropterin synthase deficiency as the cause. All other testing was unremarkable. At age 88.5 months, her brain MRI showed progression of the white matter changes. She died at 91 months of age, 1 year before the birth of her younger sister, the proband. No autopsy was performed.                  100               Figure 4.1 Familial pedigree and photographs of the affected girls.  A. The pedigree shows two loops of consanguinity; however, molecular studies and family history analysis confirmed additional loops. The affected girls are noted by shaded shapes and the proband by an arrow. B. Photograph of the elder affected sister. C. Photograph of the proband.  4.3.2 Defective cytokine production by the proband  In the context of her normal B and T cell studies, the absence of fever with apparent infection and neurodegeneration triggered by minor infections, we suspected that the proband had a defect in pyrogen generation mediated by innate immunity. To assess this, we tested her PBMCs for the production of the pyrogen IL-1β in response to an LPS challenge in the presence of ATP (357). At 59 months (37 months after onset), the proband’s PBMCs failed to produce IL-      101 1β, in contrast to healthy controls, the asymptomatic siblings, and the parents (Figure 4.2). PBMCs were also stimulated with a panel of pathogen-associated molecular patterns (PAMPs) to stimulate cytokine production. As seen in Figure 4.3, the proband exhibited a broad decrease in inflammatory cytokines assessed by Luminex. The proband died before these abnormalities could be investigated further.         Figure 4.2 IL-1β production is defective in PBMCs from the proband. PBMCs were treated with 10 ng/ml LPS with or without 5 µM ATP for 4 hours. IL-1β in culture supernatants was measured by ELISA. All experiments were performed in technical triplicates. ‘Healthy controls’ represents the mean of 7 biological replicates ± SEM. Production of IL-1β by proband PBMCs following stimulation with LPS and LPS + ATP is profoundly decreased in comparison to other healthy controls and to all 5 unaffected family members. All heterozygous family members showed no difference from controls. Control experiments with media only, or media with ATP, yielded negligible concentration of IL-1β in all samples (data not shown). N = 7 healthy controls, N = 5 unaffected family members, N = 1 proband.  Healthy ControlsUnaffected siblingsProbandHealthy ControlsUnaffected siblingsProband01000200030004000IL-1β (pg/ml)Healthy controlsUnaffected family membersProbandLPS LPS + ATP   102 Figure 4.3 Cytokine and chemokine levels detected by Luminex. PBMCs were stimulated with a panel of PAMPs and cytokine/chemokine production was measured by Luminex. Graphs depict the concentration (pg/mL) of the indicated cytokine/chemokine in 5 healthy controls (average; one standard deviation indicated by the bars) and the proband. PAMPs used included: PAM (triacylated lipopeptide, 1 µg/ml); plC     103 (polyinosinic:polycytidylic acid, 100 µg/ml); LPS (lipopolysaccharide, 10 ng/ml); R484 (Imidazoquinoline compound, 10 µM); PGN (peptidoglycan, 10 µg/ml).  4.3.3 Exome sequencing To determine the molecular etiology for this unusual innate immune response, we performed exome sequencing on the DNA of the affected girls and immediate family members (parents and unaffected siblings; Figure 4.1). Analysis for homozygous variants unique to the affected siblings identified a mutation (NM_005041.4: c.673C4T, p.R225W) in exon 3 of the PRF1 gene. Both parents and all of the unaffected siblings were heterozygous for the same variant. The p.R225W mutation is in the transmembrane region of perforin and has been associated with FHL2 (12, 21, 358, 359) In an animal cell model, p.R225W causes decreased cytotoxicity against Jurkat target cells and impaired trafficking of secretory granules (360).  4.4 Discussion Two sisters are reported with fatal neurodegeneration showing similar patterns of white matter cerebellar degeneration. Both sisters had pathogenic biallelic mutations in PRF1 and the proband had a profoundly defective innate immune response that may have contributed to disease. The PRF1 mutation identified has been associated with FHL2 (12) but the sisters did not manifest diagnostic features of FHL2 (361). Specifically, they had minimal or no fevers, no hypertriglyceridemia, no hypofibrinogenemia, and no hyperferritinemia. The youngest had only mild, transient cytopenia. Because both sisters died before the PRF1 mutation was identified, they were not tested for reduced NK-cell activity or elevated soluble IL-2 receptor levels. Notably, two bone marrow biopsies in the proband did not demonstrate increased hemophagocytosis found in 82% of FHL2 patients (362).    104 Mutations in the gene encoding PRF1 account for 20-40% of global FHL cases and up to 50% of FHL cases in North America (22). Perforin co-localizes with granzyme B in granules of cytotoxic NK and T cells and is secreted upon conjugation between target and effector cells. Once secreted, perforin inserts into the membrane of target cells and mediates apoptosis (23). The mechanism of perforin has been the focus of much debate. It was previously thought that perforin formed pores in the target cell that allowed passage of granzyme into the target cell, however more recent studies suggest that granzymes are endocytosed by the target cell and that perforin aids in their escape to the cytosol (136). While the molecular basis for mutations in perforin leading to defective immunity are explicable, the role of perforin in neurodegeneration remains unclear. Neurodegeneration has been reported as a possible feature of FHL2 and sporadic HLH, with a frequency of 37–69% (362-365). A third of patients have neurological symptoms at diagnosis (365), and 36% of those with PRF1 mutations have some CNS involvement (365). Although there is an association of variations in PRF1 with susceptibility to multiple sclerosis (366), very few patients with HLH have had neurological symptoms as the sole presenting feature, and consistent with the expected ascertainment and reporting bias (363, 364, 367, 368), all developed the diagnostic features of FHL2-HLH, with the exception of one individual who was diagnosed by brain histopathology but without identification of a mutation (346, 347, 363, 368-372). The sisters reported herein are therefore the first individuals reported with neurodegeneration, biallelic pathogenic PRF1 mutations and no diagnostic features suggestive of FHL–HLH.  Potential explanations for this lack of features include the nature of the triggering pathogen, pathogen–host interaction, the immunological status defined by prior infectious    105 exposures, genetic background or another primary cause of neurodegeneration. However, the sisters did not appear to share a common triggering pathogen or common environmental exposures. Notwithstanding the limitations of exome sequencing (361), extensive evaluation of all exome variants in the siblings failed to identify an alternative primary causal mutation for neurodegeneration or candidate modifier variant even though the high degree of consanguinity suggests that the affected children may both have inherited such modifier variants for the phenotype associated with the PRF1 mutation. A genome-wide analysis of a larger cohort of patients harboring PRF1 mutations, presenting with and without neurodegeneration might, however, identify such modifiers.  In contrast to the febrile cytokine storms observed in FHL2, the proband stopped developing fevers with infection during the course of her disease. Consistent with this, her PBMCs did not produce IL-1β or other pro-inflammatory cytokines classically over-expressed in FHL2 (11). This observation, consistent with a defect in the innate immune response, suggests that the link between PRF1 mutations and cytokine over-expression may be indirect or complex. The discrepant cytokine phenotype between FHL2 patients and the sisters described can be explained by genetic modifiers or by a difference in the timing of cytokine testing. Studies of cytokine expression in patients with classical FHL have been performed early in the disease course, most typically before treatment (11), whereas our proband was tested long after onset of symptoms. Determining whether this cytokine profile is a distinguishing feature of the PRF1-related neurodegenerative disorder manifest by these girls, or whether loss of cytokine production or cytokine depletion is a feature common to the later stages of FHL2 will require testing of additional FHL2 patients.     106 The brain MRI findings in HLH are nonspecific. They include multiple focal or diffuse areas of signal abnormality within the cerebral and cerebellar white and gray matter (348, 363, 371). The lesions can have a nodular appearance and enhance after contrast, a finding suggestive of perivascular involvement (363, 367). The involvement of the cerebellum with development of cerebellar atrophy is a prominent feature observed in the majority of HLH patients. Cerebellar gray matter involvement usually appears later in the disease, and thus is not constant. Involvement of the cerebral gray matter is less common. Even when the patient cohort is limited to those with perforin mutations, the MRI findings are also not consistent, although cerebellar disease predominates (346, 347, 371, 373-375). In the absence of systemic findings of HLH, therefore, one is unable to make a clinical diagnosis of PRF1-related neurodegeneration, and molecular testing is required.  In summary, we extend the neurodegenerative phenotype associated with biallelic mutations of PRF1 and show that the neurodegenerative phenotype can occur in the absence of the hematological and immune signs of FHL2-HLH. We hypothesize that PRF1-related neurodegeneration is an under-recognized condition, and suggest that when suspicion of immune-mediated neurodegeneration arises, this diagnosis should be considered because it is potentially curable with hematopoietic stem cell transplantation (14, 375).        107 Chapter 5: Conclusion  Primary immunodeficiencies can be the underlying cause of severe and often chronic infections with unusual pathogens that are difficult to treat, cause major financial burden, and cause high morbidity and mortality. Studying the molecular pathways in normal and immunodeficient cells that participate in host defense against opportunistic infections can shed light on how the human body mediates health and disease. Such investigations are thus beneficial for understanding essential immunological mechanisms in both immunodeficient patients as well as the healthy general population. A major theme of this thesis was the human immune response to opportunistic pathogens. Chapters 2 and 3 involved the use of primary neutrophils and primary human monocyte-derived macrophages differentiated in autologous serum to be as physiologically relevant as possible. Nearly all of the literature reported on the interactions between B. cenocepacia and macrophages have used human or murine macrophage cell lines. The variability of these cell types is beautifully shown in a study by Schmerk et al, who found that isolates of the Burkholderia cepacia complex (Bcc) proliferate within the U937 macrophage cell line but were killed within macrophages derived from the THP-1 monocyte cell line (231). While in vivo animal models of infection are highly valuable, cells from such models often provide information that is quite different from data derived from human cells and in vitro studies using primary human cells are instrumental in understanding human immunity to human pathogens. In Chapter 4, we investigated the immune response of peripheral blood mononuclear cells (PBMCs) to bacterial ligands ex vivo. These experiments provided a link between aberrant innate immunity associated with a genetic mutation in perforin1 and the phenotype of neurodegeneration that was exhibited    108 by the sisters described. Ultimately, translational studies and bed-to-benchside research of this sort is the best method of deciphering the roles of individual molecules in human immunity. Although we aimed to use in vitro models that were as physiologically relevant as possible, a limitation of this study was the use of inhibitors, particularly diphyleneiodonium (DPI) to mimic chronic granulomatous disease (CGD) in Chapter 2. As previously discussed, DPI is not specific and therefore can have effects on other cellular systems. Since CGD only occurs in 1 in 200,000 live births and is associated with high levels of morbidity and mortality, we were not able to use primary CGD neutrophils for these studies at this time. This thesis explored the ability of innate immune cells with underlying defects to respond to live opportunistic bacteria and bacterial pathogens. Chapter 2 and 3 investigated the ability of Burkholderia cenocepacia to subvert phagocytic antimicrobial responses. CGD results from genetic defects in the NADPH oxidase, responsible for producing reactive oxygen species (ROS) during the oxidative burst, that are crucial for bacterial and fungal killing. The Bcc causes severe lung infections in CGD characterized by excessive neutrophil infiltration and inflammation and is the second leading infectious cause of death (34, 44). We found that B. cenocepacia survives within ROS-deficient neutrophils and induces rapid apoptosis. In contrast, healthy neutrophils were extremely efficient at killing B. cenocepacia and did not undergo significant apoptosis compared to untreated cells. Although previous literature shows that macrophages exposed to apoptotic cells or cell membranes exhibit an anti-inflammatory phenotype consistent with resolution (239), we show that B. cenocepacia-induced apoptotic ROS-deficient neutrophils were highly pro-inflammatory to macrophages. This suggests that in the presence of abundant live bacteria, inflammation persists to enhance pathogen killing, overriding the traditional anti-inflammatory response to apoptotic cells.     109 In contrast to the effective ability of normal neutrophils to kill B. cenocepacia, human primary monocyte-derived macrophages provided a rich replication niche that enhanced the growth of B. cenocepacia compared to its growth in tissue culture media alone. Furthermore, the intracellular niche allowed B. cenocepacia to escape neutrophil killing unless neutrophils were co-incubated in large excess to macrophages. Similarly, neutrophils were only able to dampen the pro-inflammatory response of macrophages to B. cenocepacia when co-cultured at high levels. This is consistent with our earlier findings that inflammation decreases only when bacteria are successfully killed. These findings extend our knowledge of the phagocyte and bacterial interactions that govern lung immunity. While many studies examine the interplay between bacteria and one type of immune cell, we recognize the diverse ecology of the lung and have attempted to show a broader picture of host defense. These studies fill gaps in the literature of innate immunity against B. cenocepacia including the limited knowledge of B. cenocepacia interactions with neutrophils. We provide a link between the survival of B. cenocepacia in neutrophils and the induction of cell death, specifically via apoptosis, with resulting inflammation rather than resolution. This extends our understanding of apoptosis in infected cells and the ability of innate immunity to respond in an appropriate manner. Furthermore, this finding is consistent with the symptoms of Bcc respiratory infections in patients with CGD.  The ability of B. cenocepacia to escape neutrophil killing by establishing a replication niche in macrophages may contribute to its persistence in the lung and ability to cause chronic infections such as are seen in patients with cystic fibrosis. Despite this ability to subvert host immunity, infections with the Bcc are very rarely seen in healthy individuals in the absence of an underlying defect in immunity. Furthermore, species of the Bcc are ubiquitous in the    110 environment and are found in diverse environments such as the soil, plant and animal surfaces, the plant rhizosphere and as contaminants in cosmetic and pharmaceutical solutions, sterile solutions, unpasteurized bovine milk, and gelatin (376). Thus, exposure to the Bcc is likely not uncommon. The lack of Bcc infections in healthy individuals despite its ability to subvert neutrophil killing via macrophages suggests that there are other factors at play that govern the balance between infection and immunity. We found that when neutrophils were present at ten times the levels of macrophages that the enhanced growth of B. cenocepacia provided by the macrophages was abolished and the induction of inflammatory mediators was dramatically reduced. Although this may seem like an excess of neutrophils, neutrophils are quickly mobilized to sites of infection and it may be quite likely that physiologically there is a similar, if not greater, ratio of neutrophils to macrophages during acute infection. Although the studies conducted for this thesis are entirely in vitro, they do shed light on the complex strategies that might come into play in preventing infections in immunocompetent individuals.  Chapter 4 described a novel presentation of a primary immunodeficiency caused by a previously described mutation in perforin associated with familial hemophagocytic lymphohistiocytosis (FHL) triggered by infection. Whereas opportunistic pathogens in CGD most often cause severe infections that lead to organ failure or septicemia, the infectious agent in the sisters with FHL triggered a rapid decline in health without overt infection. During the course of neurodegeneration, we found that the patient’s PBMCs exhibited a profound lack of cytokine production for which the role of perforin is unknown. The links between perforin, neurodegeneration, and aberrant cytokine production remain unknown and are a focus for future directions. However, this study does extend the phenotype associated with mutations in perforin    111 and suggests previously undescribed roles for perforin in cytokine production and neurological homeostasis. Overall, this thesis describes the diverse effects that opportunistic pathogens can exert on immunodeficient individuals and highlights the breadth of knowledge on host defense that remains unknown. The major findings were that B. cenocepacia induces rapid apoptosis in the absence of ROS and that the apoptotic neutrophils induce pro- rather than anti-inflammatory cytokines from macrophages. B. cenocepacia was able to escape neutrophil killing by establishing a rich replication niche in macrophages. Finally, we reported a novel presentation of a mutation in perforin1 that is associated with FHL in two sisters. The girls did not exhibit symptoms consistent with FHL but had fatal neurodegeneration and their PBMCs showed a marked inability to produce cytokines upon stimulation.  5.1 Future directions  This thesis addresses the complex roles of innate immunity that govern infection and inflammation. In Chapters 2 and 3, the roles of macrophages and neutrophils in controlling infection with B. cenocepacia were examined. We found that B. cenocepacia induces rapid apoptosis in neutrophils in the absence of ROS, however the majority of the signaling upstream of caspase-3 in this process remains unknown. Specifically, it would be interesting to evaluate the role of any bacterial factors, including secretion systems previously shown to regulate cell death and/or bacterial intracellular survival. We would also like to investigate whether extrinsic caspase-8 mediated or intrinsic caspase-9 mediated apoptosis is triggered following challenge with B. cenocepacia and which other cellular signaling molecules are specifically involved. Improving our understanding of the specific pathways that activate B. cenocepacia-induced    112 neutrophil cell death in CGD may provide novel targets for therapeutics aimed at decreasing airway inflammation. Indeed, a number of caspase inhibitors are currently undergoing clinical trials to prevent disease such as infection-induced tissue damage, septic shock, and ischemia (Reviewed in (377)), and they may prove advantageous in CGD to dampen inflammation in combination with antibiotic treatment. Another future direction for Chapter 2 is to further investigate the inflammatory phenotype induced in macrophages by B. cenocepacia-challenged neutrophils. This could include examining macrophages for traditional markers of M1 skewing, including CD86 and CD80 expression (378). We would like to extend our experiments in Chapter 2 to CGD patient cells to examine whether our findings of B. cenocepacia-induced apoptosis is a true effect in CGD. As discussed, we used DPI to inhibit the NADPH oxidase due to the low prevalence of CGD patients. However, a number of CGD patients participate in similar studies in North America and a future direction would be to contact some patients to participate in our study. Chapter 3 of this thesis assessed the interactions between macrophages and neutrophils in combating B. cenocepacia survival. We found that human monocyte-derived macrophages had a great capacity for allowing intracellular proliferation of B. cenocepacia and that the bacteria grew better intracellularly than extracellularly. In the future, it would be interesting to further characterize the location and the nutrient factors provided by this niche. An analysis of genes upregulated by B. cenocepacia when internalized by macrophages found that approximately 40% of genes were poorly characterized and may represent undescribed survival mechanisms. Therefore it is unknown whether B. cenocepacia upregulates specific metabolic pathways for intracellular growth. Characterizing the nutrient sources that allow B. cenocepacia to proliferate is of great interest for future directions.    113 Despite the ability of macrophages to provide a replication niche for B. cenocepacia that allows it to escape neutrophil killing, healthy people generally to not become infected. Thus a future direction would be to investigate this discrepancy, using either in vitro or in vivo systems. In vitro, it would be interesting to skew macrophages towards an inflammatory M1 phenotype using LPS, IFN-γ, or higher levels of GM-CSF prior to infection, to evaluate whether priming macrophages may induce greater antibacterial properties (379). Furthermore, in vivo studies with macrophage depletion may shed light on their role during B. cenocepacia infection.  Chapter 4 of this thesis described a novel presentation of disease with underlying perforin1 mutation associated with neurodegeneration and profoundly impaired cytokine production. This study raises many questions about the broad role of perforin in maintaining health. We are particularly interested in the causes and effects of such low cytokine production. It would be interesting to see if this cytokine phenomenon extends to other patients with mutations in perforin1 or with the specific biallelic perforin mutation identified here (PRF1; p.R225W). One defining symptom of FHL is elevated cytokine production by activated macrophages and cytotoxic T cells. Since cytokine production in this thesis was unfortunately limited to one biological replicate at one time point throughout the course of disease, we would aim to further characterize the cytokine production in other patients with similar mutations.    While this thesis extends the current understanding of the causes of B. cenocepacia infection and inflammation in CGD and also describes a novel form of disease associated with a perforin1 mutation, it generates a number of new questions. Future work on the topics described will aim to extend the understanding of the innate immune responses described in this thesis.   114 Bibliography 1. Medzhitov R. 2008. 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Fischer U, Schulze-Osthoff K. 2005. Apoptosis-based therapies and drug targets. Cell Death Differ 12: 942-61 378. Menzies FM, Henriquez FL, Alexander J, Roberts CW. 2010. Sequential expression of macrophage anti-microbial/inflammatory and wound healing markers following innate, alternative and classical activation. Clinical & Experimental Immunology 160: 369-79 379. Mosser DM. 2003. The many faces of macrophage activation. Journal of Leukocyte Biology 73: 209-12     164 Appendix Appendix A   Growth curves of B. cenocepacia isolates K56-2 and MH1K   Figure A.1 B. cenocepacia isolates K56-2 and MH1K display similar growth curves Growth of K56-2 and MH1K (gentamicin-sensitive K56-2) in LB with and without 10% pooled human serum. Growth at 37°C was monitored in a Bioscreen C machine (TYPE FP-1100-C, Oy Growth Curves Ab Ltd.) with measurement intervals of 15min and continuous shaking (73).  0.001$0.01$0.1$1$0$ 4$ 8$ 12$ 16$ 20$ 24$OD600%Time%(h)%K56+2$in$LB$MH1K$in$LB$K56+2$in$LB$+$10%$PNHS$MH1K$in$LB$+$10%$PNHS$

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