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Determining the role of hyaluronan as an environmental cue for macrophages and dendritic cells Dong, Yifei (Jeff) 2018

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  DETERMINING THE ROLE OF HYALURONAN AS AN ENVIRONMENTAL CUE FOR MACROPHAGES AND DENDRITIC CELLS  by YIFEI (JEFF) DONG  B.Sc., The University of British Columbia, 2012  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)  February 2018  © Yifei (Jeff) Dong, 2018 ii  Abstract CD44 is a cell surface glycoprotein that binds to hyaluronan (HA), an extracellular matrix glycosaminoglycan. Immune cells widely express CD44, but only a few types of cells such as alveolar macrophages (AMФ) in the lung alveoli bind fluorescein-conjugated HA constitutively. During inflammation and immune responses, other immune cells such as monocytes are activated and can gain the ability to bind HA. However, the functional significance of CD44 and HA interactions remains unclear. Therefore, the aim of this study was to investigate the function of CD44 and HA binding and how they regulate immune cells such as AMФ in the tissue environment. HA has been described as a regulator of tissue inflammation, with HA fragments reported to stimulate immune cells. To test if HA fragments can induce inflammation or are consequences of inflammation, I stimulated macrophages and dendritic cells with various sizes of HA from different sources. Pharmaceutical grade HA and endotoxin-free HA fragments failed to stimulate an inflammatory response in vitro and in vivo, demonstrating they were not pro-inflammatory. Since AMФ constitutively bind HA, I then compared these cells from CD44+/+ and CD44-/- mice to study the role of CD44 and HA binding as regulatory environmental cues. Using adoptive transfer experiments and a mouse model of inflammation, I found CD44 expression and HA binding were required for the survival of mature AMФ, but not for the recruitment and differentiation of monocytes into AMФ. CD44 expression by AMФ was required for a cell surface HA coat, which maintained AMФ survival and numbers in the lung. Since AMФ are essential for regulating pulmonary surfactant lipid homeostasis, CD44 deficiency and the partial loss of AMФ in CD44-/- mice disrupted lipid homeostasis in their lungs. CD44-/- mice had iii  elevated lung surfactant phosphatidylcholine levels and CD44-/- AMФ exhibited an abnormal phenotype and accumulated cellular lipid droplets. They also suffered greater inflammation caused by oxidized phosphatidylcholine. Thus, CD44 deficiency led to a reduction of AMФ numbers and caused intrinsic defects in AMФ surfactant lipid homeostasis. Together, these results demonstrate a role of CD44 and HA binding in maintaining AMФ and lung homeostasis.   iv  Lay Summary Cells of the immune system such as macrophages and dendritic cells are essential for our health and ability to fight infections. Therefore, it is important to understand the environmental factors that control their function and survival. Hyaluronan is a large molecule present throughout the body and integral for tissue structure. However, it is not clear why some immune cells such as lung alveolar macrophages bind hyaluronan, and if this serves any function. In this study, I found that while different forms and sizes of hyaluronan do not cause inflammation in macrophages and dendritic cells as previously reported, binding to hyaluronan through its receptor CD44 was needed to maintain healthy alveolar macrophages. Without CD44 on their cells, mice had less alveolar macrophages and when challenged, suffered greater lung inflammation. Together, these results showed a new role of CD44 and hyaluronan in maintaining lung and alveolar macrophage health.   v  Preface A version of Chapter 2 has been published.  Dong, Y., Arif, A., Olsson, M., Cali, V., Hardman, B., Dosanjh, M., Lauer., Midura, R.J., Hascall, V.C., Brown, K.L., Johnson, P. (2016). Endotoxin free hyaluronan and hyaluronan fragments do not stimulate TNF-alpha, interleukin-12 or upregulate co-stimulatory molecules in dendritic cells or macrophages. Sci. Rep. 6, 36928.  I conducted all the experimental research for this publication with the following exceptions:  A.A. acquired the data for Figure 2.3e, 2.5a, and 2.8a, b, e. M.O. acquired the data for Figure 2.1, and Figure 2.2a, b. V.C., M.L., R.J.M., and V.C.H. the acquired data for Figure 2.6a and provided intellectual input. B.H. and M.D. provided technical assistance.  I wrote the manuscript with input from P.J. and edited the manuscript with all authors. These figures were reproduced with the permission from Scientific Reports.  A version of Chapter 3 has been published.  Dong, Y., Poon, G.F., Arif, A., Lee-Sayer, S.S.M., Dosanjh, M., and Johnson, P. (2017). Fetal and bone marrow monocyte derived alveolar macrophage survival is promoted by CD44 and its interaction with hyaluronan. Mucosal Immunol. advance online publication 25 October 2017. I conducted all the experimental research for this publication with the following exceptions:  vi  G.F.P acquired data and made Figure 3.1c-j, and Figure 3.6b-d. A.A. acquired the data for Figure 3.7d-e. For Figure 3.2, G.P. acquired the data, S.L. irradiated the mice and did the bone marrow reconstitution, I reanalyzed the data and made the figure. M.D. provided technical support. I wrote the manuscript with input from P.J. and edited the manuscript with all authors.  These figures were reproduced with the permission from Mucosal Immunology.  A version of Chapter 4 will be submitted for peer review.  Dong, Y., Arif, A., Poon, G., Lee-Sayer, S., Dosanjh, M., Roskelley, C.P., and Johnson, P. Lung surfactant lipid homeostasis is regulated by CD44 in alveolar macrophages. Manuscript in review. I conducted all the experimental research for this publication with the following exceptions:  A.A. helped to acquire the data for Figure 4.3a, c-d, e, g. and Figure 4.7a-b. M.D. provided technical support.  I wrote the manuscript with input from P.J. and edited the manuscript with all authors.   I have also contributed and authored the following work not included in this thesis: Dong, Y., Arif, A., Poon, G.F., Hardman, B., Dosanjh, M., Johnson, P. (2016). Generation and identification of GM-CSF derived Alveolar-like macrophages and dendritic cells from mouse bone marrow. J. Vis. Exp. Jun 25;(112). doi: 10.3791/54194. vii  Poon, G.F., Dong, Y., Marshall, K.C., Arif, A., Deeg, C.M., Dosanjh, M., Johnson, P. (2015). Hyaluronan binding identifies a functionally distinct alveolar macrophage-like population in bone marrow-derived dendritic cell cultures. J. Immunol. 195, 632-42. Lee-Sayer, S.S., Dong, Y., Arif, A., Olsson, M., Brown, K.L., Johnson, P. (2015). The where, when, how, and why of hyaluronan binding by immune cells. Front. Immunol. 6:150. doi: 10.3389/fimmu.2015.00150. Ding, H., Caza, M., Dong, Y., Horianopoulos, L.C., Hua, G., Johnson, P., Kronstad, J.W. ATG genes influence the virulence of Cryptococcus neoformans virulence through contributions beyond core autophagy functions. Manuscript in review.  Animal experimentation was conducted in accordance with protocols approved by the University Animal Care Committee and Canadian Council of Animal Care guidelines. Project titles and the certificate numbers applicable to this project include: Animal Care Certificate for Breeding Programs: A13-0015 CD44 and CD45 Breeding A16-0320 CD44 and CD45 Breeding Animal Care Certificate: A11-0307 Function of hyaluronan binding in the immune system A11-0316 Molecular Analysis of CD44 in Cell Migration and Adhesion A15-0213 Molecular analysis of hyaluronan and CD44 -innate immunity Biohazard Approval Certificate: B10-0110 Biohazard certificate for CD44 and CD45 projects B14-0155 Biohazard certificate for CS, CD44 and CD45 projects  viii  Table of Contents Abstract .......................................................................................................................... ii Lay Summary ................................................................................................................ iv Preface ........................................................................................................................... v Table of Contents ....................................................................................................... viii List of Figures ............................................................................................................ xiii List of Abbreviations ................................................................................................... xv List of Symbols .......................................................................................................... xix Acknowledgements ..................................................................................................... xx Dedication ................................................................................................................... xxi Chapter 1: Introduction ................................................................................................. 1 1.1 Mononuclear phagocytes in health and disease .................................................... 1 1.1.1 The immune system in health and disease ...................................................... 1 1.1.2 Introduction to the mononuclear phagocyte system ......................................... 3 1.1.3 Origin and development of monocytes ............................................................ 5 1.1.4 Monocytes at homeostasis and inflammation .................................................. 7 1.1.5 Monocyte-derived macrophages and DCs ....................................................... 9 1.1.6 Tissue resident macrophages ........................................................................ 12 1.2 Overview of the lung and its resident mononuclear phagocytes .......................... 16 1.2.1 Lung structure and function ........................................................................... 16 1.2.2 Pulmonary surfactant proteins and lipids ....................................................... 17 1.2.3 Mononuclear phagocytes in the lung ............................................................. 20 1.3 Alveolar macrophages ......................................................................................... 23 1.3.1 Alveolar macrophage development ............................................................... 23 1.3.2 Role of alveolar macrophage in lung homeostasis ........................................ 24 1.3.4 Role of alveolar macrophage in lung immunity .............................................. 26 1.4 CD44 and hyaluronan in the lung ......................................................................... 29 1.4.1 Hyaluronan .................................................................................................... 29 1.4.2 CD44 and hyaluronan binding ....................................................................... 32 1.4.3 CD44 and hyaluronan in dendritic cell function .............................................. 35 1.4.4 CD44 and hyaluronan in macrophage function .............................................. 36 ix  1.4.5 CD44 and hyaluronan in lung development ................................................... 37 1.4.6 CD44 and hyaluronan in lung inflammation ................................................... 38 1.5 Research aims and hypotheses ........................................................................... 40 Chapter 2: Endotoxin free hyaluronan and hyaluronan fragments do not stimulate TNF-α, interleukin-12 or upregulate co-stimulatory molecules in dendritic cells or macrophages. .............................................................................................................. 42 2.1 Introduction .......................................................................................................... 42 2.2 Material and methods .......................................................................................... 44 2.2.1 Animals .......................................................................................................... 44 2.2.2 Reagents ....................................................................................................... 45 2.2.3 Antibodies ...................................................................................................... 45 2.2.4 Cell isolation and culture ................................................................................ 46 2.2.5 Cell staining ................................................................................................... 47 2.2.6 Intratracheal instillation and bronchoalveolar lavage (BAL) ........................... 47 2.2.7 Cell stimulation and analysis of inflammation ................................................ 48 2.2.8 Fragmentation of HA ...................................................................................... 48 2.2.9 Inactivation of HA’ses .................................................................................... 49 2.2.10 Polymyxin B removal of LPS ........................................................................ 49 2.2.11 Triton X-114 removal of LPS ........................................................................ 49 2.2.12 Visualization of HA on agarose gel .............................................................. 50 2.2.13 Generation of HA-HC complexes in vitro ..................................................... 50 2.2.14 Detection of HA-HC complexes with Western blot ....................................... 50 2.2.15 Data analysis ............................................................................................... 51 2.3 Results ................................................................................................................. 52 2.3.1 Different sizes of pharmaceutical grade HA do not stimulate CSF-1 derived bone marrow macrophages but human umbilical cord HA does ............................. 52 2.3.2 CSF-2 but not CSF-1 derived BMDMs and BMDCs constitutively bind FL-HA ................................................................................................................................ 55 2.3.3 Different sizes of pharmaceutical grade HA do not simulate CSF-2 derived BMDMs and BMDCs, but human umbilical cord HA does ...................................... 57 x  2.3.4 Different sizes of pharmaceutical grade HA do not stimulate ex vivo splenic macrophages or DCs while human umbilical cord HA does ................................... 59 2.3.5 Administration of 20K and 200K HA in vivo does not induce lung inflammation ................................................................................................................................ 60 2.3.6 Fragmented HA is not inflammatory to CSF-2 derived BMDMs and BMDCs. 62 2.3.7 HC-HA complexes do not induce an inflammatory response in CSF-2 BMDMs and BMDCs ............................................................................................................ 64 2.3.8 Huc HA and HA’se are contaminated with endotoxin .................................... 66 2.4 Discussion ............................................................................................................ 69 Chapter 3: The survival of fetal and bone marrow monocyte-derived alveolar macrophages is promoted by CD44 and its interaction with hyaluronan .............. 75 3.1 Introduction .......................................................................................................... 75 3.2 Material and methods .......................................................................................... 77 3.2.1 Mice ............................................................................................................... 77 3.2.2 Reagents ....................................................................................................... 77 3.2.3 Flow cytometry (FC) ...................................................................................... 78 3.2.4 Cell isolation .................................................................................................. 79 3.2.5 Induction of lung inflammation and adoptive monocyte transfer .................... 80 3.2.6 Competitive BM reconstitution ....................................................................... 80 3.2.7 Intratracheal instillation of AMФ and PMФ ..................................................... 80 3.2.8 Intratracheal instillation of HA blocking CD44 antibody ................................. 81 3.2.9 AMФ Ki67 labeling ......................................................................................... 81 3.2.10 Generation of CSF-1 derived BMDM ........................................................... 81 3.2.11 Macrophage stimulation with CSF-2 and ROZ ............................................. 81 3.2.12 Measuring AMФ cell death and surface HA labeling .................................... 82 3.2.13 Confocal microscopy ................................................................................... 82 3.2.14 Data analysis ............................................................................................... 83 3.3 Results ................................................................................................................. 84 3.3.1 CD44-/- mice have reduced numbers of AMФ and CD44-/- AMФ are at a competitive disadvantage under both homeostatic and inflammatory conditions ... 84 3.3.2 CD44 provides an advantage to monocyte-derived AMФ .............................. 87 xi  3.3.3 LPS induced lung inflammation causes a transient loss of AMФ and differentiated monocytes gain HA binding .............................................................. 89 3.3.4 GFP+ monocytes mature into AMФ 30 days after the inflammatory stimulus . 92 3.3.5 CD44 exerts an advantage in monocyte-derived AMФ after inflammation ..... 95 3.3.6 The alveolar environment induces HA binding............................................... 97 3.3.7 CD44 and HA binding promote AMФ survival .............................................. 100 3.4 Discussion .......................................................................................................... 103 Chapter 4: CD44 deficiency disrupts lipid homeostasis in the lung ..................... 109 4.1 Introduction ........................................................................................................ 109 4.2. Material and methods........................................................................................ 111 4.2.1 Mice ............................................................................................................. 111 4.2.2 Reagents ..................................................................................................... 111 4.2.3 Flow cytometry (FC) .................................................................................... 112 4.2.4 Cell isolation ................................................................................................ 113 4.2.5 Intratracheal instillation of POVPC and CSF-2 ............................................ 113 4.2.6 Competitive BM reconstitution ..................................................................... 113 4.2.7 Adoptive transfer of AMФ in the alveolar space ........................................... 114 4.2.8 H2DCFDA labeling ....................................................................................... 114 4.2.9 NBD-PE and NBD-PC uptake ...................................................................... 114 4.2.10 AMФ treatment with HA’se and PC in culture ............................................ 114 4.2.11 AMФ survival in vitro with PC and OXPC ................................................... 115 4.2.12 Analysis of the BAL fluid ............................................................................ 115 4.2.13 Detection of AMФ cell surface HA.............................................................. 115 4.2.14 Labeling of AMФ with BODIPY 493/505, E06 antibody, PPARγ antibody, or anti-phosphoStat5 (pY694) antibody .................................................................... 116 4.2.15 Confocal Microscopy ................................................................................. 116 4.2.16 Data analysis ............................................................................................. 117 4.3 Results ............................................................................................................... 118 4.3.1 CD44-/-  mice with less AMФ have an enhanced alternatively activated phenotype ............................................................................................................. 118 xii  4.3.2 The cellular HA coat does not interfere with the detection of cell surface receptors ............................................................................................................... 120 4.3.3 Pulmonary PC surfactant levels are elevated in the lungs of CD44-/- mice .. 121 4.3.4 AMФ from CD44-/- mice have abnormal lipid accumulation and are more susceptible to OxPC toxicity ................................................................................. 122 4.3.5 The inflammatory response to OxPC is exacerbated in CD44-/- mice .......... 124 4.3.6 CSF-2 partially rescues lung lipid homeostasis disruption in CD44-/- mice ... 126 4.3.7 The alveolar environment regulates AMФ lipid droplet accumulation and CD36 expression while lipid droplet accumulation and CD11c expression are intrinsically affected by the loss of CD44. ................................................................................ 128 4.3.8 AMФ lipid droplet accumulation and CD36 expression can be modulated by extracellular PC .................................................................................................... 130 4.3.9 PPARγ expression is downregulated independently of CSF-2 and Stat-5 signaling in CD44-/- AMФ ...................................................................................... 133 4.4 Discussion .......................................................................................................... 135 Chapter 5: Concluding discussion and remarks .................................................... 140 5.1 Ability of HA to stimulate macrophages and DCs ............................................... 140 5.2 Contribution of monocyte-derived macrophages to AMФ repopulation .............. 142 5.3 Role of HA binding via CD44 for AMФ survival .................................................. 144 5.4 Effect of AMФ loss and CD44 deficiency in the lung .......................................... 145 Bibliography .............................................................................................................. 151    xiii  List of Figures Chapter 1 Figure 1.1: Schematic for the development of mononuclear phagocytes ........................ 3  Chapter 2 Figure 2.1: Stimulation of CSF-1 BMDMs by various HA preparations ......................... 52 Figure 2.2: FL-HA binding by CSF-1 BMDMs, peritoneal macrophages, and CSF-2 BMDMs and BMDCs ..................................................................................................... 55 Figure 2.3: Stimulation of CSF-2 BMDMs and BMDCs by various HA preparations ..... 57 Figure 2.4: In vitro stimulation of splenic macrophage and DCs by various HA preparations and the intratracheal instillation of HA into the lung .................................. 59 Figure 2.5: Sizes of various HA preparations and their effect on CSF-2 BMDMs and BMDCs .......................................................................................................................... 62 Figure 2.6:  Western blot demonstrating the generation of HA-HC complexes and their effect on stimulating CSF-2 BMDMs and BMDCs ......................................................... 64 Figure 2.7: The effect of Bov and Strp HA’se on the activation of CSF-2 BMDMs and BMDCs .......................................................................................................................... 66 Figure 2.8: Endotoxin contamination and removal from Huc HA and its effect on the production of pro-inflammatory cytokines by CSF-2 BMDMs and BMDCs .................... 67  Chapter 3 Figure 3.1: CD44 deficiency reduces AMФ numbers .................................................... 84 Figure 3.2: CD44 deficiency impairs AMФ repopulation following lethal irradiation ....... 87 Figure 3.3: Characterization of inflammatory cells in the BAL during LPS induced lung inflammation .................................................................................................................. 89 Figure 3.4: Adoptively transferred BM monocytes give rise to AMФ following LPS induced lung inflammation ............................................................................................. 92 Figure 3.5: CD44 deficiency impairs the number of monocyte-derived AMФ after LPS induced lung inflammation. ............................................................................................ 95 Figure 3.6: The alveolar environment and CSF-2 promote HA binding by macrophages ...................................................................................................................................... 97 xiv  Figure 3.7: CD44-/- AMФ have increased cell death and do not bind HA on the cell surface ........................................................................................................................ 100 Figure 3.8: HA binding to CD44 promotes AMФ survival ............................................ 102  Chapter 4 Figure 4.1: CD44-/- mice have abnormal AMФ ............................................................. 119 Figure 4.2: CD44 deficiency cause aberrant pulmonary surfactant PC and AMФ lipid accumulation ............................................................................................................... 121 Figure 4.3: POVPC induced pulmonary inflammation is exacerbated in CD44-/- mice . 124 Figure 4.4: CSF-2 increases the number of AMФ and rescues aberrant pulmonary surfactant PC accumulation ........................................................................................ 126 Figure 4.5: AMФ lipid homeostasis is controlled by the extracellular milieu and by the expression of CD44 ..................................................................................................... 128 Figure 4.6: Cell surface HA and extracellular PC can modulate in vitro AMФ lipid droplet level and CD36 expression ......................................................................................... 130 Figure 4.7: PPARγ expression is defective in CD44-/- AMФ......................................... 133  Chapter 5 Figure 5.1: Schematic showing the impact of AMФ loss and CD44 deficiency have on lipid homeostasis in the lung ....................................................................................... 145    xv  List of Abbreviations AEC  alveolar epithelial cell AMФ  alveolar macrophage(s) Bach2  B lymphoid transcription repressor BTB and CNC homology 2 BAL  bronchoalveolar lavage BM  bone marrow BMDC bone marrow derived dendritic cell BMDM bone marrow derived macrophage Bov  bovine testes BODIPY 4,4-difluoro-1,3,5,7,8-pentamethyl-4-bora-3a,4a-diaza-s-indacene FSC-A forward scatter area CCL  chemokine ligand CD  cluster of differentiation cDC  classical dendritic cell CDP  common dendritic cell progenitor C/EBP CCAAT/enhancer binding proteins CLP  common lymphoid progenitor cMoP  common monocyte progenitor CMP  common myeloid progenitor COPD  chronic obstructive pulmonary disease CSF-1  colony stimulating factor 1 (also known as M-CSF) CSF-2  colony stimulating factor 2 (also known as GM-CSF) DAMP  danger associated molecular pattern xvi  DMEM Dulbecco's Modified Eagle's medium DC  dendritic cell E. coli  Escherichia coli ECM  extracellular matrix EDTA  ethylenediaminetetraacetic acid EMP  erythro-myeloid precursors ELISA  enzyme-linked immunosorbent assay  EtOH  ethanol FC  flow cytometry FBS  fetal bovine serum  Fl-HA  fluorescein-conjugated HA  GFP  green fluorescent protein GMP  granulocyte-macrophage progenitor H2O2  hydrogen peroxide H2DCFDA 2',7'-dichlorodihydrofluorescein diacetate HA  hyaluronan HABP  hyaluronan binding protein HA-A647 Alexa Fluor 647 conjugated hyaluronan HA’se  hyaluronidase HAS  hyaluronan synthase HC  heavy chains HMW  high molecular weight HSC  hematopoietic stem cells xvii  Huc  human umbilical cord II  inter-alpha-inhibitor IL  interleukin IMФ  interstitial macrophage IPF  idiopathic pulmonary fibrosis i.t.  intratracheal instillation i.v.  intravenous LAL  Limulus Amebocyte Lysate LMW   low molecular weight LPS  lipopolysaccharide Lyve-1 lymphatic vessel endothelial hyaluronan receptor 1 mAb  monoclonal antibody MCP-1 monocyte chemoattractant protein-1 MDP  macrophage-DC progenitor MFI  mean fluorescence intensity moDC  monocyte-derived dendritic cell MPP  multi-potent progenitor MPS  mononuclear phagocyte system NBD-PC 1-palmitoyl-2-(6-[(7-nitro-2-1,3-benzoxadiazol-4-yl)amino]hexanoyl)-sn-glycero-3-phosphocholine NBD-PE n-(7-nitrobenz-2-oxa-1,3-diazol-4-yl)-1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine  NEAA  non-essential amino acids  xviii  NO  nitric oxide NR4A1 nuclear receptor subfamily 4 group A member 1 OxPC  oxidized phosphatidylcholine PAP  pulmonary alveolar proteinosis PAMP  pathogen-associated molecular pattern PC  L-α-phosphatidylcholine  pDC  plasmacytoid DC PFA  paraformaldehyde POVPC 1-palmitoyl-2-(5'-oxo-valeroyl)-sn-glycero-3-phosphocholine PPARγ peroxisome proliferator-activated receptor gamma Rc  rooster comb RPMI  Roswell Park Memorial Institute medium RT  room temperature ROZ  rosiglitazone ROS  reactive oxygen species SSC-A side scatter area Strp  Streptomyces hyalurolyticus S. equi Streptococcus equi SP  surfactant protein TCS  tissue culture supernatant TLR  toll like receptor TNF-α  tumor necrosis factor-alpha TSG-6 tumor necrosis factor-inducible gene 6 protein  xix  List of Symbols α alpha β beta γ gamma  kappa µ micro Ф phi  xx  Acknowledgements I give my sincerest thank you to Dr. Pauline Johnson. As my supervisor, she has gifted me great patience and provided me with continuous support and wise guidance. Her passion for science and drive for scientific rigor is always inspiring. I also express deepest gratitude to my family and friends, especially my parents who supported me throughout my years of education. Thank you for the unconditional love and support.  Thank you to the past and present members of the Johnson laboratory for their assistance and friendship. I am especially appreciative of Arif A. Arif and Dr. Grace F.T. Poon. I would also like to thank my committee members: Dr. Ninan Abraham, Dr. Kenneth Harder, Dr. Michael Murphy, and Dr. Georgia Perona-Wright for their intellectual expertise, valuable comments, and excellent support.  Thank you UBC flow cytometry core, CDM and MBF staff, and the Department of Microbiology and Immunology for their support over the years. I am grateful for the financial support from the University of British Columbia and the Robert Emmanuel and Mary Day endowment.  xxi  Dedication Two things awe me most, the starry sky above me and the moral law within me.  - Immanuel Kant 1  Chapter 1: Introduction 1.1 Mononuclear phagocytes in health and disease  1.1.1 The immune system in health and disease The immune system maintains homeostasis, mediates protection against pathogenic microorganisms, and helps to repair tissue damage (Medzhitov and Janeway, 1998). The skin and mucosal surfaces are the physical barriers which form the first line of defense against external insults (Iwasaki and Medzhitov, 2015). Typically, when these barriers are breached from an injury or acute infection, innate immune cells such as neutrophils, monocytes, macrophages, and DCs are recruited and help initiate the pro-inflammatory response to clear any foreign organisms (Janeway and Medzhitov, 2002). If the infection is not quickly resolved, antigen presenting cells such as activated DCs migrate to the draining lymph node and present pathogen specific antigens to the T and B cells of the adaptive immune system. These adaptive immune cells subsequently undergo exponential expansion and mount an antigen specific response against the offending pathogen. Finally, when the pathogen is neutralized, smaller subsets of T and B cells survive to become long-lived cells that establish immunological memory and will quickly recognize and respond to any secondary infection by the same pathogen (Iwasaki and Medzhitov, 2015; Kurtz, 2004). A strong immune system is essential for health, as various types of immunodeficiencies lead to opportunistic infections that severely lower the quality of life, or are life-threatening (Al-Herz et al., 2011).   The effectiveness of the immune system to distinguish between self and non-self-antigens and to respond appropriately are critical for the maintenance of health 2  and restoration of homeostasis. In cases when immune cells fail to recognize non-self-antigens, bacterial or viral pathogens will escape detection and cause life-threatening infections (Finlay and McFadden, 2006). Cancer may also develop or worsen when immune cells do not recognize malignant tumor cells as harmful (de Visser et al., 2006). Alternatively, over exuberant immune responses are also problematic. For instance, chronic respiratory diseases such as asthma are associated with the type of inflammation seen in allergic reactions, where lung immune cells activate in response to innocuous antigens or pollutants (Barnes, 2008). Likewise, food allergies may develop when gut immune cells become hypersensitive to food antigens (Tordesillas et al., 2017). In addition, more severe auto-immune diseases such as multiple sclerosis, Type 1 diabetes, or rheumatoid arthritis can develop with inappropriate inflammation and immune response to self-antigens (Davidson and Diamond, 2001; Medzhitov, 2008). Thus, the proper balance of immune cells and immune responses is necessary to maintain health, and to prevent disease.   3  1.1.2 Introduction to the mononuclear phagocyte system  Figure 1.1: Schematic for the development of mononuclear phagocytes  The MPS is a family of innate immune cells including monocytes, macrophages, and DCs that all have important roles in tissue homeostasis and immunity. Cells of the MPS are highly phagocytic and are found throughout the body with varying cell surface phenotype, function, ability to interact with T cells, and gene expression, indicating their plasticity and heterogeneity (Geissmann et al., 2010; Hume, 2008). Cells of the MPS are historically classified as cells that express the 4  CSF-1 receptor (CD115), proliferate and arise from BM precursors, circulate systemically as blood monocytes, and differentiate into DCs or macrophages in tissues (Hume, 2006; Sasmono et al., 2003; van Furth and Cohn, 1968). Although granulocytes such as neutrophils develop from the same common myeloid progenitors as cells of the MPS, they not included in the MPS definition because they are polymorphonuclear, and do not express CD115 at the protein level.  More recent ontological studies demonstrate tissue resident macrophages of the MPS are seeded at different stages of embryonic/fetal development (Ensan et al., 2016; Epelman et al., 2014; Goldmann et al., 2016; Gomez Perdiguero et al., 2015; Guilliams et al., 2013; Hashimoto et al., 2013; Hoeffel et al., 2015; Janssen et al., 2011; Schulz et al., 2012; Sheng et al., 2015; Yona et al., 2013; Zhu et al., 2017), and most are maintained independently from BM derived monocytes at the steady-state via self-renewal (Soucie et al., 2016). In addition, various studies show subsets of resident cDCs and pDCs in the spleen and lymph node originate from hematopoietic lineages that are distinct from monocytes and macrophages and require Flt3 signaling for their development (Liu et al., 2009; Liu et al., 2007; Miller et al., 2012; Naik et al., 2006; Onai et al., 2007). Thus, monocytes, tissue resident macrophages, moDCs, and cDCs and pDCs all develop from separate independent lineages at the steady-state (Figure 1.1). As a result of these new findings, there is an undergoing discussion to modify the definition of the MPS (Guilliams et al., 2014). For example, the MPS can be divided into DCs including cDCs and pDCs, monocytes and monocyte-derived cells, and self-maintaining tissue resident macrophages (Guilliams and Scott, 2017). Moreover, understanding the function 5  and contribution of various cells of the MPS to immune responses, such as monocytes and monocyte-derived macrophages versus embryonic/fetal-derived tissue resident macrophages during tissue inflammation, is an active area of research (Jenkins and Hume, 2014). 1.1.3 Origin and development of monocytes  Monocytes are a population of innate immune cells found in all vertebrates including mice and humans. They are well defined by their phenotype, morphology, and gene expression (Boyette et al., 2017; Cros et al., 2010; Etzrodt et al., 2012; Ginhoux and Jung, 2014; Ingersoll et al., 2010; Mildner et al., 2013; Schlueter and Glasgow, 2006; Sunderkotter et al., 2004; van Furth and Cohn, 1968). Monocytes are initially identified as CD11b+ CD115+ cells and are found in the BM, blood, and spleen (Auffray et al., 2009b; Swirski et al., 2009). In both mice and humans, there are two general subsets of monocytes (Geissmann et al., 2003; Ingersoll et al., 2010; Passlick et al., 1989): 1) classical monocytes that are Ly6Chi CCR2+ CD62L+ CX3CR1mid in mice or CD14+ CD16- in humans (Geissmann et al., 2003; Ginhoux and Jung, 2014; Jakubzick et al., 2013), and 2) non-classical (also known as patrolling) monocytes that are Ly6Clo CCR2- CD43+ CX3CR1hi in mice or CD14lo CD16+ in humans (Ginhoux and Jung, 2014; Hanna et al., 2011; Landsman et al., 2009; Yona et al., 2013). In addition, an intermediate transitional population between classical and non-classical monocytes may exist, particularly in humans where they are CD14+ CD16+ (Patel et al., 2017; Wong et al., 2011).  In adults, BM HSCs are stem cells that can self-renew and differentiate to maintain almost all immune cells in the body, although many tissue resident 6  macrophages do not require BM contribution at the steady-state (Ensan et al., 2016; Epelman et al., 2014; Goldmann et al., 2016; Gomez Perdiguero et al., 2015; Guilliams et al., 2013; Hashimoto et al., 2013; Hoeffel et al., 2015; Janssen et al., 2011; Schulz et al., 2012; Sheng et al., 2015; Yona et al., 2013; Zhu et al., 2017). Long term and short-term HSCs differentiate into MPPs that can either become CLPs which give rise to lymphoid cells, or CMPs which give rise to myeloid cells (Seita and Weissman, 2010). CMPs will differentiate into lineage negative CD117+ CD135+ CD115+ MDPs that become monocytes but could also generate DCs (Auffray et al., 2009a; Fogg et al., 2006). Recent studies then identified further restricted CDPs downstream of MDPs that give rise to only cDCs and pDCs (Liu et al., 2009; Naik et al., 2007; Onai et al., 2007), as well as CD135-  cMoPs that only generate monocytes (Hettinger et al., 2013; Kawamura et al., 2017). Interestingly, Yáñez et al. found while CMP derived MDPs give rise to monocytes and DCs, CMP derived GMPs give rise to monocytes and neutrophils, but not DCs, independently of MDPs (see Figure 1.1 for schematics). Moreover, MDP derived monocytes from CpG stimulation are transcriptionally different to GMP derived monocytes from LPS stimulation, suggesting additional complexity and function in monocyte populations can be regulated by inflammatory signals (Yanez et al., 2017).  CSF-1 is critical for the development Ly6Chi  monocytes (Cecchini et al., 1994), and CSF-1 deficiency causes monocytopenia (Dai et al., 2002). Transcription factors such as PU.1 (Laslo et al., 2006), GATA2 (Dickinson et al., 2011), as well as IRF8 and KLF4 are also critically important for the differentiation of mature Ly6Chi monocytes from BM progenitors (Kurotaki et al., 2013; Sichien et al., 2016). At the 7  steady-state, Ly6Chi monocytes egress from the BM via CCR2 chemokine signaling and enter peripheral circulation (Serbina and Pamer, 2006). In the blood, they can differentiate into Ly6Clo non-classical monocytes; this process requires NR4A1 (Hanna et al., 2011; Thomas et al., 2016) and Notch signaling (Gamrekelashvili et al., 2016). Interestingly, while the number of Ly6Chi monocytes is significantly impaired in IRF8 and KLF4 deficient mice, the number of Ly6Clo monocytes is less affected (Alder et al., 2008; Kurotaki et al., 2013), suggesting either non-classical monocyte can differentiate independently from classical monocytes (Carlin et al., 2013), or an enhanced differentiation and survival of Ly6Clo monocytes in the absence of IRF8 or KLF4.   1.1.4 Monocytes at homeostasis and inflammation At the steady-state, monocytes develop in the BM, circulate in the blood, and are present in the spleen which can be mobilized when needed (Swirski et al., 2009; Vanfurth and Sluiter, 1986). In humans and mice, more than 80% of the circulating monocytes are classical monocytes, with the remainder being intermediate and non-classical monocytes (Geissmann et al., 2003; Passlick et al., 1989; Patel et al., 2017; Wong et al., 2011). Typically, the half-life of classical monocytes in the blood is about 1 day and 5-7 days after differentiation into intermediate and non-classical monocytes (Gamrekelashvili et al., 2016; Patel et al., 2017; Yona et al., 2013). In mice, the maintenance and survival of monocytes are in part dependent on TNF-α signaling as mice lacking TNF or TNF receptor have increased monocyte cell death and impaired monocyte development (Wolf et al., 2017). In addition, the increased 8  expression of C/EBP-β (Tamura et al., 2016) and CX3CR1 (Landsman et al., 2009) in Ly6Clo monocytes are important for their survival in circulation. In naïve mice, Ly6Chi monocytes may traffic into tissues for surveillance, gain MHCII upon interaction with the tissue endothelium, and transport antigen from the tissues to the lymph node without differentiating into moDCs (Jakubzick et al., 2013). In contrast, Ly6Clo monocytes survey the blood vessels via integrin interactions with the endothelium and help remove damaged cells and debris in the vasculature (Auffray et al., 2007; Carlin et al., 2013; Thomas et al., 2015).  Upon inflammation or injury, classical monocytes are quickly recruited to the tissues (Dal-Secco et al., 2015; Liao et al., 2017; Shi and Pamer, 2011; Zigmond et al., 2014), and this is highly dependent on CCR2 binding to CCL2 and CCL7 (Serbina and Pamer, 2006; Tsou et al., 2007). There are also other chemokine receptors that can mediate monocyte recruitment during inflammation such as CCR1 and CCR5 (Kaufmann et al., 2001; Mack et al., 2001), or CCR7 and CCR8 (Qu et al., 2004). Additional production and egress of new monocytes from the BM help replace the loss of circulating monocytes during inflammation (Patel et al., 2017). The early accumulation of Ly6Chi monocytes in the tissues during an immune response contributes to inflammation, especially as they produce inflammatory cytokines such as TNF-α and NO upon stimulation (Serbina et al., 2003). The infiltration of inflammatory monocytes can have negative effects in some disease settings. For example, Ly6Chi monocytes accumulate during experimental autoimmune encephalomyelitis (Ajami et al., 2011; Mildner et al., 2009) and atherosclerosis (Swirski et al., 2006) to promote inflammation. Furthermore, ablating 9  classical monocytes via CCR2 deletion or CCL2 blocking reduces tumor burden in mice, suggesting their involvement by promoting metastatic seeding or by producing tumor promoting factors such as vascular endothelial growth factor (Headley et al., 2016; Qian et al., 2011). In contrast, depleting non-classical monocytes via NR4A1 or CX3CR1 deletion increases tumor metastasis, implicating their involvement in cancer immunosurveillance (Hanna et al., 2015). Non-classical monocytes normally crawl in the blood vessels and help remove cellular debris strictly in the vasculature (Carlin et al., 2013), although it has been reported that they can extravasate into tissues upon inflammation and gain repair functions such as arginase production (Auffray et al., 2007). For inflammatory monocytes that arrive at the site of inflammation or injury, they can differentiate into macrophages or moDCs depending on the signals in the tissue environment (Italiani and Boraschi, 2014; Martinez et al., 2006). When tissue homeostasis is re-established, they can be cleared by apoptosis (Bosurgi et al., 2017; Gautier et al., 2013; Janssen et al., 2011; Kiener et al., 1997; Serhan and Savill, 2005; Vandivier et al., 2006), or can contribute to the repopulation of tissue resident macrophages (Bleriot et al., 2015; Gundra et al., 2017; Scott et al., 2016).  1.1.5 Monocyte-derived macrophages and DCs  Although many tissue resident macrophages do not require monocytes for replenishment at the steady state (Ensan et al., 2016; Epelman et al., 2014; Goldmann et al., 2016; Gomez Perdiguero et al., 2015; Guilliams et al., 2013; Hashimoto et al., 2013; Hoeffel et al., 2015; Janssen et al., 2011; Schulz et al., 10  2012; Sheng et al., 2015; Yona et al., 2013; Zhu et al., 2017), monocytes give rise to some resident macrophages and moDCs in adult tissues such as the intestines (Bain et al., 2014; Varol et al., 2007), peritoneal cavity (Bain et al., 2016), skin (Ginhoux et al., 2006; Tamoutounour et al., 2013; Wu et al., 2016), and the lung (Gibbings et al., 2017; Plantinga et al., 2013). They may also serve as reservoirs of inflammatory cells that can quickly migrate to places where tissue homeostasis becomes disrupted. For instances, a recent study found that after the injection of senescent erythrocytes, monocytes are quickly recruited to the liver close to Kupffer cells and differentiate into macrophages to help clear dying erythrocytes and to recycle iron (Theurl et al., 2016).  Monocyte-derived macrophages in inflamed or injured tissue environments are heterogenous and express a variety of markers depending on their maturation state (Dal-Secco et al., 2015; Jakubzick et al., 2017; Kratofil et al., 2017). For example, Ly6Chi monocytes migrate to the inflamed lung during house dust mite antigen stimulation, lose Ly6C expression, become MafB+ CD45+ MHCII+ CD11c+ CD11b+ CD64+ cells, and can traffic to the mediastinal lymph node (Wu et al., 2016). Interestingly, these monocytes derived cells in the inflamed lung only express the macrophage linage specific transcription factor MafB (Soucie et al., 2016; Wu et al., 2016), but not the DC lineage transcription factor Zbtb46 (Meredith et al., 2012; Satpathy et al., 2012), suggesting that monocyte-derived inflammatory cells are distinct from the cDC lineage and adopt a macrophage identity. Thus, the nomenclature of monocyte-derived cells needs further clarification, especially since the identity and function of moDCs (Langlet et al., 2012; Plantinga et al., 2013), 11  TNF-α and inducible NO synthase producing DCs (Serbina et al., 2003), and monocyte-derived macrophages are overlapping.  The specific roles of inflammatory monocytes and monocyte-derived macrophages during lung inflammation are not well understood; although in a bleomycin induced lung fibrosis model, monocyte-derived macrophages are partially involved in driving tissue fibrosis (Misharin et al., 2017). In contrast, the functions of monocytes and monocyte-derived macrophages are better characterized in the intestines. Here, there is a constant replenishment of intestinal resident macrophages by monocytes due to the presence of the intestinal microbiota (Bain et al., 2014). Intestinal macrophages at the steady-state are highly phagocytic and bactericidal, but they are less responsive to inflammatory stimulation due to the production of TGF-β from intestinal stroma cells (Smythies et al., 2005). In contrast, monocytes and monocyte-derived macrophages are the main producers of inflammatory mediators such as IL-1, IL-6, TNF-α, NO, and ROS (Joeris et al., 2017). Moreover, monocyte-derived cells produce IL-1β and IL-23 to drive Th17 differentiation and IL-17 production (Arnold et al., 2016; Aychek et al., 2015; Lasiglie et al., 2011), as well as IL-12 to drive Th1 polarization and IFN-γ secretion (Schreiber et al., 2013), to protect against bacterial infection. Studies also demonstrate the ability of CD11b+ moDCs migrating to the lymph node and act as antigen presenting cells (Chakarov and Fazilleau, 2014; Randolph et al., 2000). However, the antigen presentation ability of moDCs is not very efficient compared to cDCs (Kamphorst et al., 2010; Schreiber et al., 2013), and thus they may be more important for the transportation of antigen from the tissue to the lymph node (Ersland 12  et al., 2010). In latter stages of inflammation or when danger signals are absent, monocytes and monocyte-derived macrophages differentiate into a more healing/reparative alternatively-activated phenotype (Italiani and Boraschi, 2014) in response to factors such as IL-4, IL-13, IL-10, or TGF-β (Casella et al., 2016; Gordon and Martinez, 2010; Makita et al., 2015; Sinha et al., 2005; Zhang et al., 2016). Various studies show the importance of these alternatively-activated macrophages in helping to resolve inflammation and directing wound repair in the liver, skin, and muscles (Duffield et al., 2005; Goren et al., 2009; Lemos et al., 2015). Finally, monocyte-derived macrophages can also contribute to the repopulation and renewal of tissue resident macrophages, where they fill any empty niche caused by inflammation and death of embryonic/fetal-derived tissue resident macrophages. Recent studies demonstrate this occurs in the liver and the lung (Bleriot et al., 2015; Dong et al., 2017; Guilliams and Scott, 2017; Gundra et al., 2017; Scott et al., 2016). Together, these all show the plasticity of monocytes during inflammation and disease, as well as the continuum of roles they participate in with the changing tissue environment.   1.1.6 Tissue resident macrophages  Macrophages are phagocytic immune cells found across all tissues and organisms. By expressing various PPRs (Brubaker et al., 2015) and receptors for recognizing apoptotic cells (Poon et al., 2014), macrophages can quickly sense any change in tissue homeostasis and are critical for initiating immune responses and for clearing dead cells. Recent findings highlight most tissue resident macrophages in 13  rodents develop from embryonic or fetal liver derived progenitors and are shaped transcriptionally and epigenetically by their local tissue environment (Ginhoux and Jung, 2014; Gosselin et al., 2014; Lavin et al., 2014). Likewise, the function of tissue resident macrophages varies depending on the tissue (Amit et al., 2017). Splenic red pulp macrophages for example are important for the clearance of erythrocytes and iron metabolism (Kohyama et al., 2009), whereas splenic marginal zone macrophages help clear apoptotic cells (Miyake et al., 2007). Microglial cells in the brain continuously survey the microenvironment (Nimmerjahn et al., 2005) and aid in brain development such as synaptic pruning (Paolicelli et al., 2011; Schafer et al., 2012). Bone osteoclasts aid in bone remodeling and resorption (Grigoriadis et al., 1994; Teitelbaum, 2000). Lung AMФ are required for pulmonary surfactant metabolism (Suzuki et al., 2008). Tissue resident macrophages fill the tissues during embryonic/fetal development, are long-lived, and self-renew at the steady-state; therefore most do not need significant contribution from BM progenitors such as monocytes in adulthood for maintenance (Ginhoux and Guilliams, 2016; Perdiguero and Geissmann, 2016). During embryogenesis, the first wave of primitive hematopoiesis develops from the extra-embryonic yolk sac around E7.0 to give rise to erythroblasts, megakaryocytes, and macrophages (Palis et al., 1999; Tober et al., 2007). A transient wave of RUNX1+ yolk sac hematopoietic progenitors around E7.25 differentiate into yolk sac macrophages that are the precursors to microglia, but not other tissue resident macrophages (Ginhoux et al., 2010). Following this, a second wave of EMPs develop from the hemogenic endothelium of yolk sac around 14  E8.0 to E8.5. After E8.5 when blood circulation is established, C-Myb+ EMPs colonize the fetal liver and differentiate into multiple hematopoietic lineages such as fetal liver monocytes (Bertrand et al., 2005; Palis and Yoder, 2001). Fetal liver monocytes replace yolk sac macrophages over time, seed multiple embryonic tissues, and differentiate into tissue resident macrophages in the lung, spleen, kidney, and liver during organogenesis (Hoeffel et al., 2015; Mass et al., 2016). However, other studies propose C-Myb- yolk sac macrophages that do not require a monocyte intermediate are the main precursors to tissue resident macrophages (Schulz et al., 2012) such as microglia, Langerhan cells, AMФ, and Kupffer cells (Gomez Perdiguero et al., 2015). Definitive HSCs only begin to emerge from the hemogenic endothelium of the aorta-gonado-mesonephros regions by E10.5 (Bertrand et al., 2010; Boisset et al., 2010; Kissa and Herbomel, 2010), and they are distinct from EMPs (Chen et al., 2011). Interestingly, a study by Sheng et al. showed the fate mapping of the stem-cell-factor receptor CD117 at E7.5 only labeled microglia and some Langerhan cells, whereas labeling at E8.5 labeled all tissue resident macrophages. Thus, the authors suggest all adult macrophages except microglia and some epidermal Langerhan cells are derived from either fetal HSCs or late EMPs (Sheng et al., 2015). The further development and maturation of resident macrophages require specific programming depending on the tissue. So far, studies show the transcription factor Id3 is required for liver Kupffer cells (Mass et al., 2016), Runx3 is required for Langerhan cells (Fainaru et al., 2004), Nr1h3 is required for splenic marginal zone macrophages (A-Gonzalez et al., 2013), SpiC is required for splenic red pulp macrophages (Kohyama et al., 2009), Gata6 is required for the 15  large peritoneal macrophages (Gautier et al., 2014; Rosas et al., 2014), and PPARγ (Schneider et al., 2014b) and TGF-β (Yu et al., 2017) are required for lung AMФ. Together, these fate mapping studies demonstrate the existence of three waves of progenitors during embryogenesis and fetal development that could give rise to long-living tissue resident macrophages with unique transcriptional programming (Mass et al., 2016) in various tissue organs.   Although embryonic/fetal-derived macrophages populate all tissues, they may be replaced to some extent by HSC derived progenitor cells over the course of a lifetime (Ginhoux and Guilliams, 2016; Perdiguero and Geissmann, 2016). Changes in tissue homeostasis such as aging, infections, or inflammation that lead to the loss of tissue resident macrophages may promote replenishment by infiltrating progenitors like Ly6Chi monocytes that fill the emptied niche (Guilliams and Scott, 2017). The continuous replacement of tissue resident macrophages by adult BM derived progenitors in the intestines (Bain et al., 2014), skin (Tamoutounour et al., 2013), heart (Epelman et al., 2014; Molawi et al., 2014), and pancreas (Calderon et al., 2015) supports this hypothesis. Thus, there are increasing complexity and heterogeneity of macrophages in tissues, where multiple populations may exist and include embryonic/fetal macrophages, adult monocyte-derived macrophages at the stead-state, as well as monocyte-derived cells with macrophage or DC functions during inflammation. Further investigation into the factors that regulate the maintenance of tissue resident macrophages and monocyte differentiation in homeostasis versus inflammation could lead to new strategies for treating diseases 16  where macrophages are critically involved, such as inflammatory diseases in the lung.   1.2 Overview of the lung and its resident mononuclear phagocytes 1.2.1 Lung structure and function  In mammals, the lung is the vital organ for respiration and thus evolved to help maximize the exposure of blood to oxygen. The respiratory organ begins from the conducting airways of the upper and lower respiratory tract, which then branches and divide up to 20 to 23 times into series of cartilaginous bronchi and bronchioles that terminate at the alveoli, providing a total surface area of around 90 m2 in humans (Colebatch and Ng, 1992). The alveoli are highly vascularized airways formed from a thin layer of epithelium and endothelium, separating blood from air. Capillaries surrounding all the alveoli in humans have a total surface area of up to 140 m2 and can filter more than 8000 liters of air each day (Effros, 2006; Kopf et al., 2014). Here, differences in gas partial pressure between the atmospheric air and blood facilitate the diffusion of carbon dioxide out of blood and oxygen in. In large airways, ciliated cells, goblet cells, club cells, and basal cells form the physical barrier; they also secrete mucus and antimicrobial peptides to protect against the outside environment (Iwasaki et al., 2017).  The alveolar epithelium is composed of Type 1 AECs (AEC1) and Type 2 AECs (AEC2). AEC2 are large round cells that can proliferate (Khalil et al., 1994; Nabeyrat et al., 1998) and produce surfactants which reduce the surface tension at the air-liquid interface in the lung epithelium, preventing lung collapse (Lopez-17  Rodriguez and Perez-Gil, 2014; Wright, 2003). AEC2 also differentiate into AEC1, which are long and thin to minimize the distance between the air and capillary blood (Castranova et al., 1988; Fehrenbach, 2001). The pulmonary interstitium contains fibroblasts, IMФ, and ECM components such as collagen and elastin which form the supportive connective tissue that separates the alveolar epithelium from the lung endothelium and vasculature, as well as adjacent alveoli (Davidson, 1990; Effros, 2006). In addition, hyaluronan is also found in the pleura space beneath the epithelium of the bronchioles (Allen et al., 1991). The pores of Kohn bridge adjacent alveoli and facilitate interalveolar movement of liquid, surfactants, and cells (Bastacky and Goerke, 1992). Since the alveolar epithelium is exposed to a variety of particles, allergens, and airborne microbes during respiration, its integrity is critical for lung health and homeostasis. Interestingly, megakaryocytes from the BM migrate to the lung where they generate up to 50% of the total platelets produced in the body (Lefrancais et al., 2017). This study also showed the lung contains hematopoietic progenitors that has the potential to seed other tissues. Thus, in addition to respiration, the lung is potentially an important organ for hematopoiesis.   1.2.2 Pulmonary surfactant proteins and lipids   Pulmonary surfactant is a mixture of lipids (90%) and proteins (5-10%). There are four major SPs associated with pulmonary surfactant and lung homeostasis: SP-A, SP-B, SP-C, and SP-D (Whitsett et al., 2015). Surfactant lipids mostly contain PC (70-85%), phosphatidylglycerol and phosphatidylinositol (10-15%), and cholesterol (3-8%). There are also minor amounts of phosphatidylethanolamine, sphingomyelin, 18  lysophosphatidylcholine, cholesterol esters, triglycerides, diglycerides and free fatty acids (Goerke, 1998; Lopez-Rodriguez and Perez-Gil, 2014; Vangolde et al., 1988). Surfactant lipid synthesis occurs in the endoplasmic reticulum of AEC2. Synthesized surfactants are transported by adenosine triphosphate-binding cassette transporter A3 (Ban et al., 2007) into lamellar bodies for storage with SP-B and SP-C (Whitsett et al., 2010). During respiration, the exposure to air (Ramsingh et al., 2011) and mechanical stretching (Frick et al., 2004; Nicholas et al., 1982; Patel et al., 2005) of the alveoli induce the fusion of lamellar bodies with the AEC2 cell membrane in a calcium ion dependent manner to secrete the surfactants (Ashino et al., 2000; Dietl et al., 2012; Frick et al., 2004). Once in the alveolar space, SP-B and SP-C help lamellar bodies unpack and remodel into different forms of surfactants (Hobi et al., 2016), including lamellar body-like particles, multi-layered packages, and tubular myelin (Goerke, 1998). Since SP-B and SP-C are hydrophobic, it is possible they help the adsorption of surfactant lipids into the air-liquid film (Parra et al., 2013). While SP-A and SP-D are known as collectins that have antimicrobial properties and help regulate lung immune responses (Clark et al., 2002; Hartshorn et al., 1997; Madan et al., 1997; Schagat et al., 2001; Wright, 2005; Wu et al., 2003), they are also integral to surfactant function. For example, the activity of the surfactant film is optimized by SP-A (Lopez-Rodriguez et al., 2016; Sanchez-Barbero et al., 2005), whereas SP-D regulates the turnover of surfactants by converting surfactant lipids into smaller forms for uptake (Ikegami et al., 2005; Ikegami et al., 2000; Korfhagen et al., 1998). The recycling of pulmonary surfactants is mostly mediated by AEC2 and AMФ. A previous study shows in the rabbit lung, up to 65% of the radiolabeled 19  surfactant phospholipids are internalized by AEC2 and approximately 20% by AMФ (Rider et al., 1992). Some surfactants may also be cleared via the upper respiratory airways (Pettenazzo et al., 1988). Furthermore, SP-A and SP-C potentially aid in PC (Bates et al., 2008) and cholesterol (Roldan et al., 2016) metabolism by AEC2, respectively.  Changes to the surfactant lipid and protein composition occur in different lung diseases. For example, acute respiratory distress syndrome patients have altered surfactant PC, phosphatidylglycerol, and cholesterol composition (Dushianthan et al., 2014; Gregory et al., 1991; Markart et al., 2007). In addition, IPF patients have increased SP-A and SP-D levels, which are important biomarkers (Greene et al., 2002) for the severity of the disease (Barlo et al., 2009; Kinder et al., 2009). Cigarette smokers and COPD patients also have dysregulated levels of surfactant proteins and phospholipids (Ilumets et al., 2011; Ishikawa et al., 2011; Lusuardi et al., 1992; More et al., 2010; Winkler et al., 2011), possibly due altered production from AEC2 (Zhao et al., 2010). Furthermore, disruptions in surfactant turnover and abnormal surfactant accumulation cause PAP, which could lead to the failure of lung immune responses and respiration (Carey and Trapnell, 2010). Thus, pulmonary surfactant regulation is critical to lung homeostasis, function, and normal respiration (Avery and Mead, 1959; Bour et al., 2014; Gupta and Zheng, 2017; Wert et al., 2009; Willson et al., 2005).  20  1.2.3 Mononuclear phagocytes in the lung  Like the intestines, resident immune cells such as macrophages and DCs in the lung are in close proximity with the mucosal epithelial membrane, the surrounding blood capillaries, and the outside environment. They work closely with AECs to monitor changes in the environment and will quickly respond when tissue homeostasis is disrupted. The activation of macrophages, DCs, and AECs require PAMPs or DAMPs to signal through pattern-recognition receptors such as TLRs, RIG-I-like receptors cytosolic DNA sensors, and nucleotide oligomerization domain-like receptors (Iwasaki et al., 2017). While monocytes are isolated from lung tissue preparations, their role and function as tissue resident cells in the lung at homeostasis are less well known (Kopf et al., 2014). Using the MacBlue x Cx3cr1gfp/+ reporter mouse, Rodero et al. showed lung resident monocytes patrol the vasculature and alveoli airways, whereas lung DCs only survey the airways (Rodero et al., 2015). In addition, given their plasticity, monocytes can also differentiate into DCs (Jakubzick et al., 2008; Plantinga et al., 2013) and IMФ in the lung (Landsman and Jung, 2007; Plantinga et al., 2013; Tan and Krasnow, 2016).   In the healthy lung, there are cDCs, pDCs, and moDCs. Lung pDCs are recruited from the BM, where they mature from Flt3, CSF-1, and IL-7 signaling (Gilliet et al., 2002; Vogt et al., 2009). CD103+ and CD11b+ cDCs can arise from CDP derived CD45+ MHCII+ CD11c- Flt3+ SIRPα- pre-DCs present in the lung tissue and require Flt3 signaling for differentiation and proliferation (Ginhoux et al., 2009; Plantinga et al., 2013). Both CSF-1 and CSF-2 appear to be dispensable for cDC development in the lung (Edelson et al., 2011; Ginhoux et al., 2009; Greter et al., 21  2012). Fate mapping suggest some cDCs, especially CD11b+ cDCs derive from monocytes (Jakubzick et al., 2008; Schraml et al., 2013), though it is unclear whether this is a significant phenomenon at the steady-state. In addition, it is difficult to isolate lung moDCs as it is hard to distinguish them from CD11b+ cDCs, due to the few differences in cell surface markers (Schlitzer et al., 2013). Functionally, DCs in the lung are critical for sampling antigen and initiating adaptive immune responses (Jahnsen et al., 2006). Their location in the lung is important for their ability to sample antigen. DCs situated closely to the alveoli sample antigen much more often and efficiently compared to DCs located by the bronchiolar airways (Thornton et al., 2012). Studies also show different subsets of lung DCs have unique and overlapping functions. CD103+ DCs can promote Th1 and Th2 responses (Furuhashi et al., 2012; Nakano et al., 2012), and are superior in antigen cross presentation for directing the anti-viral response (Helft et al., 2012; Ho et al., 2011). CD11b+ DCs can stimulate Th2 and Th17 responses (Plantinga et al., 2013; Schlitzer et al., 2013). In addition, for CD8+ T cell priming during influenza virus infection, CD103+ DCs support CD8+ T effector cell differentiation whereas CD11b+ DCs promote more CD8+ T central memory cell generation (Kim et al., 2014). TNF-α and NO producing moDCs are also important for the anti-viral response in the lung, by helping local CD8+ T cell proliferation (Aldridge et al., 2009) and stimulating the reactivation of Th1 cells (Iijima et al., 2011). Interestingly, while pDCs are potent producers of Type I interferons, they are not as critical for the anti-viral response (GeurtsvanKessel et al., 2008). Instead, pDCs in the lung help maintain tolerance in the lung against innocuous inhaled antigens (de Heer et al., 2004; Lombardi et al., 2012). In contrast, 22  CD11b+ cDCs and inflammatory moDCs may be involved in the induction and maintenance of allergic asthma (Hammad et al., 2010; Plantinga et al., 2013; van Helden and Lambrecht, 2013). In summary, these data demonstrate the importance of DCs for lung homeostasis and immunity.  There are also multiple types of macrophages in the lungs. Bronchial macrophages are found in the sputum from humans (Moniuszko et al., 2007). Humans also have intravascular macrophages in the lung capillaries, which are not present in rodents (Dehring and Wismar, 1989). While these first two types of macrophages are less well understood in humans because they are not found in mice, IMФ and AMФ have been extensively characterized in mouse models. Lung IMФ are phagocytic CD11bhi CD11clo CD64+ Ly6C- F4/80+ Siglec F- MHCII+ and CX3CR1+ cells that reside in the lung interstitium between adjacent alveoli. Given the difficulties and variabilities of isolating IMФ, their reported phenotype from different studies may differ (Becher et al., 2014; Gibbings et al., 2017; Misharin et al., 2013). IMФ are initially derived from fetal macrophages, but a second population of IMФ appear 4 days after birth, suggesting postnatal progenitors can also give rise to some IMФ (Tan and Krasnow, 2016). Recent studies show IMФ can repopulate from BM progenitors after irradiation where lungs are shielded (Gibbings et al., 2017) and the recruitment of CCR2 dependent splenic progenitors contribute to IMФ expansion during inflammation (Sabatel et al., 2017). In rhesus macaques, IMФ are under constant turnover like blood monocytes (Cai et al., 2014). In contrast, IMФ identified by microscopy were not derived from donor cells during parabiosis experiments using MacBlue x Cx3cr1gfp/+ and C57BL/6J mice (Rodero et al., 2015), 23  suggesting their independence from BM and blood progenitors. Thus, how much IMФ replenish from CCR2+ Ly6Chi monocyte differentiation or from self-renewal remain debated due to the lack of a definitive lineage tracing mice. Functionally, IMФ could help control immune responses by regulating T cells, likely through the production of IL-10 (Bouabe et al., 2011). In a model of low dose LPS stimulation followed by OVA peptide challenge induced lung allergy, IMФ are found to produce high levels of IL-10 to help suppress DCs and the Th2 response (Bedoret et al., 2009). Similarly, the expansion of IL-10 producing IMФ in the lung from bacterial CpG DNA stimulation is able to provide protection against house dust mite allergy (Sabatel et al., 2017). Unlike IMФ, AMФ reside in the alveolar space, on top of the AECs and within the pulmonary surfactant. The development and function of AMФ will be discussed in detail in the next section.   1.3 Alveolar macrophages  1.3.1 Alveolar macrophage development  In mice, AMФ make up more than 95% of the total cells in the alveolar air space at the steady-state (Becher et al., 2014; Guilliams et al., 2013; Kopf et al., 2014). During fetal development, monocytes migrate from the fetal liver to the developing lung and seed the tissue with immature CD11bhi CD11clo Siglec F- AMФ, which mature into long-lived CD11blo CD11chi Siglec Fhi MHCIIlo AMФ after birth (Gomez Perdiguero et al., 2015; Guilliams et al., 2013; Hashimoto et al., 2013). The proper maturation of AMФ require CSF-2 dependent PPARγ expression (Mass et al., 2016; Schneider et al., 2014b) and TGF-β signaling (Yu et al., 2017), whereas 24  the localization and engraftment of neonatal AMФ to the alveoli require the actin-bundling protein L-plastin (Todd et al., 2016). At the steady-state, AMФ will self-renew and do not require replenishment from blood or BM derived progenitors (Guilliams et al., 2013; Hashimoto et al., 2013). The alveolar environment may be rich in factors and signals such as CSF-2, PPARγ lipid ligands, and TGF-β to promote and maintain AMФ identity (Guth et al., 2009; Yu et al., 2017). In addition, AMФ can produce TGF-β which promotes their maintenance and gene expression in an autocrine manner (Yu et al., 2017). Because of this supportive environment, it may be possible for progenitor cells such as monocytes and immature macrophages to differentiate into fully functional AMФ in the alveolar space, especially if the niche is partially or completely empty. For instance, although AMФ self-renew independently from monocytes at the steady-state (Guilliams et al., 2013), and can rapidly proliferate after inflammation (Hashimoto et al., 2013), it is not clear whether monocytes recruited to the alveolar space during inflammation are capable of differentiating into mature AMФ and contribute to AMФ repopulation. 1.3.2 Role of alveolar macrophage in lung homeostasis In the healthy lung, AMФ remove particles, apoptotic cells, and debris from the alveolar space in order to maintain homeostatic tissue function (Hussell and Bell, 2014; Iwasaki et al., 2017; Kopf et al., 2014). In mice, the lung contains millions of alveoli, but there are only approximately a million AMФ, suggesting not every alveolus contain AMФ (Westphalen et al., 2014). They may move between alveoli through the pores of Kohn (Peao et al., 1993), although two-photon microscopy show AMФ move very slowly both in homeostasis and after stimulation (Thornton et 25  al., 2012). AMФ have a unique phenotype and can be identified as CD11blo CD11chi CD64+ CD200R+ CD205+ CD206+ Ly6C- F4/80+ Sirpα+ Siglec Fhi and MHCIIlo cells from the BAL. The expression of these cell surface markers may have functional importance in AMФ. For instance, CD11c can interact with CD18 to mediate the phagocytosis of inactivated complement component C3b-opsonized pathogens (Taborda and Casadevall, 2002), while CD205 is a receptor for the uptake of TLR9 agonists such as CpG (Lahoud et al., 2012). In addition, AMФ interact with AECs and receive negative regulatory signals via CD200 (Snelgrove et al., 2008), TGFβ (Morris et al., 2003), and IL-10 (Fernandez et al., 2004), which suppress their activation (Hussell and Bell, 2014). Yu et al. also showed autocrine TGF-β production and signaling by AMФ are critical for their development and maintenance by promoting PPARγ expression (Yu et al., 2017). Furthermore, Sirpα expressed by AMФ binds to surfactant proteins which reduces their cytokine production and NF-B activation in response to stimulation (Fournier et al., 2012; Gardai et al., 2003). In vitro engagement of Sirpα with surfactant proteins also suppresses the phagocytosis of apoptotic cells (Janssen et al., 2008), suggesting these interactions may be anti-inflammatory. Mature AMФ are critical for maintaining pulmonary surfactant homeostasis. PAP is a disease caused by excessive buildup of surfactant proteins and lipids in the lung airways. It impairs breathing, increases susceptibility to infections, and can lead to death. In humans, PAP develops due to auto-antibodies against the CSF-2 protein (Kitamura et al., 1999; Uchida et al., 2007), or because of mutations in the CSF-2 receptor (Martinez-Moczygemba et al., 2008; Tanaka et al., 2011). In mice 26  lacking the CSF-2 protein or the receptor, AMФ present in the alveolar space are foamy and immature (Shibata et al., 2001b), and cannot effectively degrade phospholipid surfactants (Yoshida et al., 2001). These changes lead to the accumulation of pulmonary surfactants and proteins (Dranoff et al., 1994), as well as pathology in the lung that resembles PAP in humans. In addition to CSF-2 and CSF-2 signaling, the Bach2 transcriptional repressor is important for proper AMФ function as Bach2-/- mice are defective in lipid processing and accumulate pulmonary surfactants similar to PAP (Nakamura et al., 2013). These studies demonstrate the importance of AMФ for the turnover of surfactant lipids and proteins, and for maintaining a healthy functioning lung. Since the alveolar environment can support the differentiation of monocytes and progenitor cells into AMФ phenotypically and transcriptionally in csf2rb-/- mice and rescue their PAP (Dong et al., 2017; Machiels et al., 2017; Misharin et al., 2017; van de Laar et al., 2016), the transplantation of macrophages or progenitor cells that are functionally similar to AMФ or can differentiate into AMФ may be used for treating PAP in humans (Happle et al., 2014; Suzuki et al., 2014). The observation that donor AMФ are maintained in the lung for at least 2 years after lung transplantation in humans (Eguiluz-Gracia et al., 2016; Nayak et al., 2016) suggests this is a promising therapy.  1.3.4 Role of alveolar macrophage in lung immunity As the predominant immune cell in the alveoli at steady-state, AMФ are important at initiating immune responses, by producing alarmins such IL-33 (Chang et al., 2011; Li et al., 2014) and IL-1α (Dagvadorj et al., 2015), as well as pro-inflammatory cytokines such as TNF-α (Herold et al., 2008; Poon et al., 2015) and 27  Type I interferons (Goritzka et al., 2015; Hogner et al., 2013; Pribul et al., 2008). Being macrophages, another important function of AMФ is the uptake and clearance of extracellular pathogens (Dockrell et al., 2003). In a model of intratracheal pneumococcal infection, mice with insufficient number of AMФ due to L-plastin deficiency could not control early bacterial burden and they rapidly succumb to the infection (Deady et al., 2014), which suggests the number of AMФ present in the lung is important for the primary immune response. This could explain why elderly individuals are more susceptible to pulmonary infections, as aging can reduce the number and function of AMФ (Li et al., 2017; Wong et al., 2017). But AMФ are not always successful at killing ingested bacteria. During Mycobacterium tuberculosis infection, AMФ may be hijacked to form foam cells that facilitate intracellular M. tuberculosis survival and granuloma formation (Peyron et al., 2008). In addition, M. tuberculosis surviving inside resident macrophages produce membrane phenolic glycolipids that induce CCL2 production and CCR2+ monocytes recruitment, leading to M. tuberculosis dissemination (Cambier et al., 2017). The response of AMФ to helminth infection is less well understood. While studies show the importance of alternatively activated macrophages in lung immunity against helminth infections (Bouchery et al., 2015; Hallowell et al., 2017), the relative importance and roles of AMФ (Reece et al., 2006) versus monocyte-derived alternatively activated macrophages (Borthwick et al., 2016) need additional investigation. During inflammatory resolution, cytokines such as IL-4, IL-13, and IL-10 in the tissue environment can induce alternatively activated macrophages to secrete factors such as matrix metalloproteinases, insulin-like growth factor 1, platelet-28  derived growth factor, TGF-β, and vascular endothelial growth factor-α to help in angiogenesis, cell proliferation, fibroblast activation, and tissue repair (Wynn and Vannella, 2016). When this process becomes dysregulated, pathological fibrosis can develop. The role of AMФ in lung fibrosis is not well understood. Although macrophages in the alveolar space have been identified to produce IL-33 that activates Type 2 innate lymphoid cells to drive bleomycin induced lung fibrosis (Li et al., 2014), it is not clear if these are true tissue resident AMФ or monocyte-derived macrophages that become similar to AMФ phenotypically but are more profibrotic in function (Misharin et al., 2017). Experiments showing that the absence of Ly6Chi monocyte recruitment to the lung reduces lung fibrosis (Gibbons et al., 2011; Okuma et al., 2004; Osterholzer et al., 2013) support the idea that monocytes and/or monocyte-derived macrophages promote fibrosis.  The observation that AMФ depletion also promotes greater pulmonary inflammatory responses (Knapp et al., 2003; Thepen et al., 1989) suggests that they are important for controlling excess inflammation, perhaps by inhibitory receptors such as CD200R (Snelgrove et al., 2008) and the scavenger receptor MARCO (Ghosh et al., 2011). The ability of a subset of AMФ to form connexin 43-mediated gap junctions with AECs which relayed immunosuppressive signaling to surrounding cells is another possible immunomodulatory mechanism (Westphalen et al., 2014). They also produce TGF-β (Khalil et al., 1993; Yu et al., 2017), which can generate regulatory T cells (Soroosh et al., 2013) and inhibit DC mediated T cell activation (Holt et al., 1993). The importance of AMФ for controlling lung immunity is also demonstrated by models where they are depleted. Clodronate ablation of AMФ 29  exacerbates allergic lung inflammation in response to OVA sensitization (Bang et al., 2011; Zaslona et al., 2014), whereas monocyte ablation attenuates the inflammation (Zaslona et al., 2014). AMФ loss induced by influenza virus infection also significantly increased bacterial burden and mortality from secondary Streptococcus pneumoniae infection (Ghoneim et al., 2013). The environmental signals present in the tissue milieu could also influence the function of AMФ. Gammaherpesvirus infection in mice induces a Type 1 inflammatory response in the lung which conditioned AMФ to protect against subsequent house dust mice antigen induced allergic asthma (Machiels et al., 2017). However, the authors did not distinguish whether this is due to fetal-derived resident AMФ that proliferated or monocyte-derived AMФ that formed from the gammaherpesvirus infection. In contrast, AMФ from mice challenged with OVA antigen induced allergic Type 2 inflammation are sensitized and became less immunosuppressive (Bang et al., 2011; Careau et al., 2006; Lauzon-Joset et al., 2014). Together, these studies highlight the importance of AMФ to lung immunity, where healthy AMФ initiate inflammation and prevent unnecessary responses, whereas significant AMФ loss or alteration of AMФ function are detrimental and can exacerbate diseases.  1.4 CD44 and hyaluronan in the lung 1.4.1 Hyaluronan   HA is a large glycosaminoglycan composed of repeating disaccharides of D-N-acetyl glucosamine and D-glucuronic acid. As a major component of the ECM, HA is distributed throughout the body and normally in high molecular mass forms of over 30  1000 kDa (Lee-Sayer et al., 2015). Because of its hygroscopic nature and viscoelastic properties, HA is important for maintaining tissue structural integrity, hydration, and lubrication (Fraser et al., 1997; Jiang et al., 2007, 2011; Monslow et al., 2015; Singh et al., 2014; Toole, 2009). HA is synthesized by HAS1-3 at the plasma membrane and extruded in to extracellular space for incorporation into the ECM and pericellular matrices (Tammi et al., 2011; Toole, 2004; Weigel, 2015; Weigel and DeAngelis, 2007). In mammalian tissues, three HA’ses (HYAL1-3) mediate HA degradation (Harada and Takahashi, 2007; Itano, 2008; Stern, 2005). HA is catabolized in the tissues after being taken up via CD44 (Harada and Takahashi, 2007) by phagocytic cells such as AMФ (Culty et al., 1992). Otherwise, HA drains into the lymphatics for uptake and degradation by sinusoid cells in the lymph node, spleen, and liver (Jadin et al., 2012). In addition to being an integral part of the ECM, HA is important for the development and homeostasis of the tissue microenvironment. This is demonstrated in mice where HAS2 deletion is embryonically lethal due to incomplete heart formation (Camenisch et al., 2000). In addition, tissue specific deletion of all three HAS and inhibition of HA synthesis reduce the survival of HSCs and their interaction with BM endothelial cells (Goncharova et al., 2012), suggesting a HA-rich BM microenvironment helps to support HSCs (Qu et al., 2014). HA is also a main component of the glycocalyx surrounding cells, which could help exclude other cells or molecules and protect against cell damage caused by oxidation (Presti and Scott, 1994; Sato et al., 1988).  In the tissues, HMW HA of over 1000 kDa is associated with homeostasis and may have anti-inflammatory roles. For example, inhaled HMW HA reduces 31  inflammatory cytokine production in a mouse model of lung cystic fibrosis (Gavina et al., 2013), and is used clinically to help treat human lung cystic fibrosis patients (Furnari et al., 2012; Ros et al., 2014). In addition, HMW HA is reported to induce regulatory T cell functions (Bollyky et al., 2009; Bollyky et al., 2007), and a modified form of HMW HA helps to promote tolerance to airway allergens (Gebe et al., 2017). Interestingly, contrary to the idea that HA is an important modulator in the tumour microenvironment and is associated with poor prognosis in patients (Chanmee et al., 2016), the accumulation of extra-large HMW HA in the ECM of naked mole rats is protective against cancer (Tian et al., 2013). Together, these studies suggest that HMW HA has roles in maintaining tissue health.  When homeostasis is disrupted by injury or inflammation, smaller fragments of LMW HA ranging from a few disaccharides to over 700 KDa can accumulate in the tissues (Cyphert et al., 2015; Lee-Sayer et al., 2015; Petrey and de la Motte, 2014). In humans, this is associated with chronic inflammatory diseases in the lung such as COPD (Papakonstantinou et al., 2015), asthma (Lauer et al., 2015), and IPF (Bjermer et al., 1989). HA fragments also form HA-HC complexes with the HC of the serum proteoglycan IαI (de la Motte et al., 2003), via covalent bonds catalyzed by the enzyme TSG-6 that is upregulated during inflammation (Dyer et al., 2016; Rugg et al., 2005). Interestingly, the HA-HC complexes enhance binding to CD44 (Lesley et al., 2004) and other receptors such as Lyve-1 (Lawrance et al., 2016), perhaps to mediate cell recruitment, migration, or retention at sites of inflammation. Previous studies implicate HA fragments as pro-inflammatory mediators that signal through TLR2 (Scheibner et al., 2006), TLR4 (Black et al., 2013; Taylor et al., 2004), TLR2 32  and TLR4 (Jiang et al., 2005), CD44 and TLR4 complexes (Taylor et al., 2007), or these in combination with NLRP3-mediated inflammasome activation (Yamasaki et al., 2009). However, direct interactions between HA and TLRs are not clearly demonstrated, and the downstream signaling mechanisms from HA that initiate inflammation are well understood. In addition, while LMW HA is largely recognized as a DAMP in the literature (Jiang et al., 2005; Jiang et al., 2011; Liang et al., 2011), a few studies show there can be endotoxin, nucleic acid, or protein contamination in LMW HA preparations (Ebid et al., 2014; Filion and Phillips, 2001; Huang et al., 2014b; Krejcova et al., 2009; Shiedlin et al., 2004). Moreover, since studies report inflammatory activities from LMW HA of various sizes or forms in vitro (Lee-Sayer et al., 2015), it is difficult to recapitulate and understand the effects of LMW HA fragments or complexes generated in vivo during inflammation. Thus, although the paradigm during the completion of this thesis is that HMW HA are present in healthy tissues and could help maintain homeostasis whereas LMW HA fragments are generated during tissue damage and considered to be inflammatory, further investigation into the mechanism of how LMW HA induces inflammation, as well as the characterization of which sizes or forms of LMW HA can induce inflammation are needed. 1.4.2 CD44 and hyaluronan binding CD44 is a type 1 transmembrane glycoprotein expressed on the surface of almost all cells of the body and is conserved in many organisms. It is the major cell surface receptor to extracellular HA (Aruffo et al., 1990; Lesley et al., 2000; Underhill, 1992a) and potentially has many other biological functions (Ponta et al., 33  2003). CD44 consists of an extracellular domain where the amino-terminal globular ‘link’ domain interacts with HA, a transmembrane domain that interacts with lipid rafts to mediate HA endocytosis and CD44 recycling (Thankamony and Knudson, 2006), and an intracellular domain that can bind ezrin, radixin, and moesin proteins (Yonemura et al., 1998). CD44 is also cleaved by the proteolytic processing activity of presenilin-mediated γ-secretase in the cytoplasm (Murakami et al., 2003; Wolfe et al., 1999). The released CD44 intracellular domain then acts as a signal transducer in the nucleus for the expression of CD44 (Okamoto et al., 2001) and matrix metalloproteinase 9 (Miletti-Gonzalez et al., 2012), induces in vitro macrophage fusion via NF-B (Cui et al., 2006), and contributes to in vitro fibroblast transformation (Pelletier et al., 2006). Although it is encoded by a single gene in humans and mice on chromosome locus 11p13 (Spring et al., 1988; Underhill, 1992b), CD44 proteins in cells are heterogenous in size because the gene transcript contains multiple exons that can be alternatively spliced, which produces different CD44 isoforms containing variant exons (Borland et al., 1998; Mackay et al., 1994; Screaton et al., 1992). CD44 can also be post-translationally modified (Camp et al., 1991). CD44 isoform expression and post-translational modification varies depending on cell types and their activation/differentiation states (Camp et al., 1991; Cichy and Pure, 2000; Ruffell et al., 2011). Interestingly, while CD44 is expressed at various levels on immune cells, only a few cells such as AMФ (Poon et al., 2015), bind HA through CD44 at the steady-state (Ruffell and Johnson, 2009). However, CD44 mediated HA binding is inducible by CD44 upregulation or post-translational modifications, particularly after cell 34  activation/proliferation (Levesque and Haynes, 1996, 1997; Maeshima et al., 2011). Since p53 can bind to the CD44 promoter and repress CD44 expression, the downregulation of p53 after cell activation may also upregulate CD44 (Watanabe et al., 2014). In addition, CD44 is post-translationally modified by glycosylation (Bartolazzi et al., 1996; Skelton et al., 1998), sulfation (Brown et al., 2001; Maiti et al., 1998), sialylation (Faller and Guvench, 2014), and glycosaminoglycan addition (Ruffell et al., 2011) to change its HA binding ability. Thus, the interaction between CD44 and HA is regulated on immune cells, and could have important functions depending on the context of cell homeostasis or activation, such as in AMФ which constitutively bind HA. CD44 is implicated in multiple immune processes (Jiang et al., 2011), such as the resolution of lung inflammation (Liang et al., 2007; Teder et al., 2002), the recruitment of immune cells such as lymphocytes and macrophages during inflammation (Cuff et al., 2001; DeGrendele et al., 1997; Egan et al., 2013), as well as T cell activation (Huet et al., 1989; Mempel et al., 2004) and differentiation (Baaten et al., 2010; Guan et al., 2009; Wu et al., 2014). Additional studies demonstrate the involvement of HA binding to CD44 in lymphocyte rolling and adhesion to the endothelium (Bonder et al., 2006), leukocyte extravasation during inflammation (Winkler et al., 2012), neutrophil recruitment (Khan et al., 2004; McDonald et al., 2008), and in inducing activated T cell death (Ruffell and Johnson, 2008). Furthermore, CD44 signaling may promote cell survival and proliferation by activating PI3K-Akt (Bates et al., 2001; Ghatak et al., 2002), STAT3 (Khurana et al., 2013), and vascular endothelial cadherin/CD31 (Tsuneki and Madri, 2014). CD44 35  also protects cells from apoptosis in vitro by forming a receptor complex with CD74 for macrophage migration inhibitory factor signaling (Shi et al., 2006). These results together with the observation that standard CD44 and CD44 isoforms are associated with cancer initiating cells and cancer progression (Lin and Ding, 2017; Ricardo et al., 2011; Roudi et al., 2014; Senbanjo and Chellaiah, 2017), suggest a role for CD44 in tumorigenesis. There is some knowledge on the function of CD44 on tumors, such as in promoting the survival and resistance of cancer cells to drug-induced apoptosis (Allouche et al., 2000; Bates et al., 2001; Herishanu et al., 2011), and assisting in antioxidant functions (Ishimoto et al., 2011; Tamada et al., 2012). Taken together, the role of CD44 and HA binding is multi-faceted and they are implicated in various cell processes and diseases. Surprisingly though, CD44-/- mice that do not express CD44 on any cells in their body and therefore are unable to bind HA are relatively normal in development (Protin et al., 1999), suggesting there are compensatory mechanisms or its functions are redundant (Nedvetzki et al., 2004), making it difficult to determine the function of CD44 and CD44 mediated HA binding in vivo.   1.4.3 CD44 and hyaluronan in dendritic cell function  Although DCs express CD44, they do not normally bind detectable levels of FL-HA in ex vivo or in vitro experiments. It is not known if this due to the level of their CD44 expression, the type of CD44 they are expressing, or the kind of HA used experimentally. HA fragments have been reported to activate DCs in vitro through TLR signaling (Termeer et al., 2002; Termeer et al., 2000a) and CD44 (Do et al., 36  2004). However, since direct interaction of HA with TLRs have not been proven, whether HA fragments can directly activate TLRs remains uncertain because of possible PAMP contamination in HA reagents. Interestingly, the HA binding peptide Pep-1 reduced DC clustering and antigen induced T cell activation in lymph nodes (Mummert et al., 2002), suggesting HA interactions are involved in antigen presentation between DCs and T cells. Overexpression of HA’se in the skin increases DC migration from the skin in a model of contact hypersensitivity (Muto et al., 2014). Similarly, LMW HA priming of DCs ex vivo increases their in vivo migration to the lymph node (Rizzo et al., 2014). These suggest the degradation of tissue matrix HA or exposure to LMW HA promotes DC migration. A recent study also found monocytes cultured in CSF-2 generated CD11c+ MHCII+ cells with endogenous HA on the cell surface that could interact with Lyve-1 in vivo, which promotes their migration into the lymphatics (Johnson et al., 2017). Given current limitations in differentiating cDCs from Langerhans cells and monocyte-derived cells, it is not definitive from this study whether in vivo cDCs also use cell surface HA and Lyve-1 for lymphatic trafficking. There is also the question whether moDCs are different from cDCs in their putative interactions with HA.   1.4.4 CD44 and hyaluronan in macrophage function  Similar to DCs, studies report the activation of macrophages by HA fragments (Horton et al., 1998a; Horton et al., 1998b; Horton et al., 1999a; Horton et al., 1999b), via NF-B signaling (Noble et al., 1996). Alternatively, CD44 and HA binding by macrophages facilitate the uptake and catabolism of HA (Culty et al., 1992), 37  promote their migration to damaged tissues (Wang and Kubes, 2016), and help protect against apoptosis in vitro by forming a complex with CD74 to signal for migration inhibitory factor activation (Shi et al., 2006). However, while CD44 is ubiquitously expressed at various levels in macrophages, AMФ are the only tissue resident macrophage that are reported to bind HA (Culty et al., 1994). Currently, it is not fully understood why these macrophages bind HA while others do not. One possible mechanism that upregulates HA binding by macrophages is by changing the expression and/or the post-translational modifications of CD44. Inflammatory cytokine signaling from TNFα or LPS and IFNγ upregulates CD44 expression and downregulates CD44 chondroitin sulfation, which promote HA binding by BM derived macrophages in vitro (Ruffell et al., 2011). Nevertheless, the in vivo function of HA binding by specific cells such as AMФ requires further investigation. Since AMФ constitutively bind HA, comparing ex vivo and in vivo AMФ from CD44+/+ and CD44-/- mice can lead to new insights on the roles and importance of CD44 and HA binding.  1.4.5 CD44 and hyaluronan in lung development  During early fetal lung development, HA accumulates in the interstitial and alveolar space. After birth, it is cleared in these areas by maturing AMФ and confined to regions around the blood vessels and bronchiolar airways (Allen et al., 1991; Johnsson et al., 2003; Underhill et al., 1993a). Since it is hygroscopic, the accumulation of HA during neonatal lung development is associated with greater lung water content (Sedin et al., 2000). The decrease in lung HA levels coincides with the increase and maturation of CD44 expressing pulmonary macrophages that 38  can take up HA, at least in vitro (Culty et al., 1992). When the antibody KM201 is used to block or neutralize these macrophages, HA clearance in the neonatal lung is significantly reduced (Underhill et al., 1993b). Thus, maturing AMФ are required for the clearance of HA during lung development. Interestingly, CD44-/- mice have normal lung development, suggesting other mechanisms of HA uptake compensate for the absence of CD44 during lung development. However, the implication of excess HA during lung development is unclear, although one study found it correlates with intrauterine infection in newborn infants (Johnsson et al., 2003).    1.4.6 CD44 and hyaluronan in lung inflammation   During lung inflammation, fibrosis, and asthma, there is the generation of LMW HA fragments and HA deposition in the tissues (Ayars et al., 2013; Bracke et al., 2010; Cheng et al., 2011; Garantziotis et al., 2009; Hallgren et al., 1989; Liang et al., 2011; Teder et al., 2002). It is not clear how HA fragments are generated in vivo, but it may be due to degradation by ROS and HA’ses in the inflammatory milieu (Monzon et al., 2010; Monzon et al., 2008). Studies report the accumulation of HA fragments and HA deposition contribute to lung inflammation by initiating TLR signaling and cytokine production (Jiang et al., 2005; Liang et al., 2011; McKee et al., 1996), impairing neutrophil clearance and TGF-β production (Teder et al., 2002), downregulating anti-inflammatory genes (Collins et al., 2011), promoting airway eosinophilia (Katoh et al., 2003; Swaidani et al., 2013), and supporting collagen deposition (Cheng et al., 2013). CD44 deficiency or blocking also exacerbates lung inflammation and enhances HA fragment accumulation in mice (Kumar et al., 2016; Liang et al., 2007; Teder et al., 2002; van der Windt et al., 2010). Therefore, CD44 39  mediated HA clearance correlates with the resolution of inflammation. Despite these observations, the specific of roles of HA fragments and macrophage CD44 expression during lung inflammation are not clear. Unlike HA fragments, HMW HA can help attenuate inflammation and promotes the restoration of homeostasis in the lung (Gebe et al., 2017; Lamas et al., 2016; Liu et al., 2008; Ruppert et al., 2014; Singleton et al., 2010). This is consistent with the hypothesis that LMW HA is associated with inflammation and HMW HA is associated with resolution. Interestingly, mice treated with bleomycin have less fibrosis after HA’se administration (Bitencourt et al., 2011; Skurikhin et al., 2015). This is counterintuitive because HA’se should generate more HA fragments associated with inflammation. Perhaps the effect of HA on lung fibrosis could be independent from inflammation and immune cells. CD44 and HAS2 overexpression promote the formation of myofibroblasts that drive lung fibrosis in mice and increase the invasion of fibroblasts from human IPF patients (Li et al., 2011a). Therefore, it may be possible to ameliorate lung fibrosis by inhibiting HAS2 expression which causes fibroblast senescence in vivo (Li et al., 2016). Surprisingly though, after bleomycin induced lung damage, HAS2 deletion reduces the level of cell surface HA on AEC2, which impairs their survival, proliferation, and ability to repair the lung (Liang et al., 2016). Together, these studies show CD44 and the accumulation of HA fragments is closely linked with lung inflammation and fibrosis. However, the size and localization of HA as well as how it can regulate the function of lung immune cells such as AMФ remain to be elucidated.  40  1.5 Research aims and hypotheses  The ability to bind HA via CD44 is a unique function of a few selected immune cells such as resident AMФ in the lung and CSF-2 derived macrophages generated from the BM in vitro. Although numerous studies report the pleiotropic effects of CD44 and HA in regulating cell functions and inflammation, less is known about their specialized roles in immune cells such as macrophages and DCs. In addition, the pro-inflammatory ability of HA fragments is evolving to be a controversial issue due to potential PAMP contamination in HA reagents. Therefore, in this study, I aim to 1) evaluate the ability of HA to cause inflammation from macrophages and DCs, and 2) determine the function of HA binding and CD44 in AMФ at homeostasis and during inflammation.  For the first aim, I tested the hypothesis that HA of various sizes and forms can stimulate different inflammatory response from macrophages and DCs because of their differential ability to bind HA. I used cells from CD44+/+ and CD44-/- mice to determine if CD44 is involved in HA mediated inflammatory signaling. In addition, the amount of endotoxin contamination from each HA reagent is evaluated. By stimulating BM derived in vitro macrophages and DCs, ex vivo splenic macrophages and DCs, and the lung in vivo with different types of HA, I can make a comprehensive conclusion on the ability of HA to stimulate inflammation in macrophage and DCs.   HA binding requires CD44, so I compared AMФ from CD44+/+ and CD44-/- mice for the second aim. Because of the observation that CD44-/- mice have less AMФ, I tested the hypothesis that CD44 and HA binding are needed for normal fetal-41  derived AMФ maintenance and/or the differentiation of monocyte-derived AMФ. In addition, since AMФ are partially depleted following treatment of i.t. LPS, I used this model to investigate the contribution of inflammatory monocytes and monocyte-derived cells to AMФ repopulation. CD44 may aid the recruitment and adhesion of leukocytes and comparing the numbers and fate of CD44+/+ and CD44-/- monocytes or monocyte-derived cells in the lung after LPS inflammation tested if CD44 and HA binding assist these processes. Together, these experiments help to understand the function of CD44 and HA binding on AMФ, and on monocyte-derived AMФ repopulation.  AMФ are critical for maintaining pulmonary surfactant homeostasis and they have immunoregulatory functions. CD44-/- mice have exacerbated lung immune responses and this is often attributed to excess HA accumulation. However, I hypothesize that the deficient number of AMФ in CD44-/- mice also disrupts lung homeostasis to sensitize the lung towards inflammation. Therefore, I performed detailed characterization comparing CD44+/+ and CD44-/- AMФ, and compared the pulmonary surfactant lipid homeostasis between CD44+/+ and CD44-/- mice to understand how CD44 deficiency impacts lung homeostasis and inflammation.   42  Chapter 2: Endotoxin free hyaluronan and hyaluronan fragments do not stimulate TNF-α, interleukin-12 or upregulate co-stimulatory molecules in dendritic cells or macrophages. 2.1 Introduction  At homeostasis, extracellular HA is present in its high molecular mass form (>1000 kDa), and its steady-state turnover is hypothesized to be mediated at least in part by tissue macrophages, as in vitro studies show the uptake and degradation of HA by AMФ (Culty et al., 1994). In vivo studies show the uptake of HA by macrophages in the developing lung (Underhill et al., 1993a), in the red pulp of the spleen, and in the liver (Jadin et al., 2012). HMW HA (800 kDa and 2700 kDa) administered in vitro suppresses the pro-inflammatory response to bacterial LPS by U937 macrophages (Yasuda, 2007), while inhaled HA reduces inflammatory cytokine production in a mouse model of lung cystic fibrosis (Gavina et al., 2013). Upon tissue inflammation, the size of HA at the site of damage is perturbed, as seen in mouse models of bleomycin injury (Teder et al., 2002) and cigarette smoke exposure (Bracke et al., 2010); and the amount of HA changes, as seen in the mouse model of ovalbumin induced asthma (Cheng et al., 2011). Shorter HA fragments (<400 kDa) are reported to become DAMPs capable of inducing pro-inflammatory responses in vitro and in vivo (Jiang et al., 2007; Lee-Sayer et al., 2015). HA’se expression is upregulated in the inflamed lung (Monzon et al., 2010) suggesting that HA turnover is increased upon inflammation. Furthermore, induced HA degradation in the skin can result in dendritic cell migration and promote allergic contact hypersensitivity (Esser et al., 2012; Muto et al., 2014). These results and 43  others have led to the model where HMW HA supports homeostasis and is anti-inflammatory, whereas LMW HA and HA oligosaccharides are pro-inflammatory (Cyphert et al., 2015). However, increased HA matrices in the tissue are also associated with inflammation. HA is produced by smooth muscle cells in response to a viral mimetic, and is organized into cable-like HA structures (Baranova et al., 2011; de la Motte et al., 2003). These structures are modified by TSG-6, a protein induced during inflammation that transfers HCs from the IαI onto HA to form HA-HC matrices (Baranova et al., 2011; de la Motte et al., 2003). In ovalbumin induced lung inflammation, wild type mice have increased HA production (Cheng et al., 2011), and HA-HC deposition occurs in the lung tissue with extensive eosinophilia and airway hyper-responsiveness (Swaidani et al., 2013). In contrast, ovalbumin treated TSG-6-/- mice have reduced HA, no HA-HC deposition and decreased eosinophilia and airway hyper-responsiveness, suggesting that HA-HC complexes influence the allergic response in the lung (Swaidani et al., 2013).  However, concerns regarding the pro-inflammatory effect of LMW HA and HA fragments have been raised. One study stimulated macrophage cell lines with various sizes of HA but could not detect any production of the inflammatory agents, NO and TNF-α (Krejcova et al., 2009). Another study stimulated glomerular mesangial cells with HA’se to generate HA fragments but found that endotoxin contamination was the cause of cytokine production (Ebid et al., 2014). In addition, administration of LMW HA (200 kDa) or human recombinant HA’se with endotoxin free HA into a mouse skin air pouch did not stimulate an inflammatory response 44  (Huang et al., 2014b). These results are in conflict with the idea that LMW HA is pro-inflammatory and raises the possibility that HA effects may occur due to the contamination of other inflammatory molecules. Indeed, human umbilical cord HA, which has been used in the past, has both protein and nucleic acid contamination (Shiedlin et al., 2004), and DNA contamination in HA samples has been shown to activate human monocytic cells (Filion and Phillips, 2001). Given that miniscule amounts of endotoxin, for example as low as 5 pg/ml of LPS, can induce pro-inflammatory IL-6 production in mouse dendritic cells (DCs) (Tynan et al., 2012), a small amount of contamination in HA can give misleading results. Therefore, the aim of this study was to systematically test and thoroughly analyze the ability of HA preparations to stimulate a pro-inflammatory response by macrophage CSF-1 or CSF-2 derived BMDMs and BMDCs in vitro by measuring the pro-inflammatory cytokines TNF-α, IL-1β, and IL-12, and the upregulation of CD40 and CD86 co-stimulatory molecules.  2.2 Material and methods 2.2.1 Animals C57BL/6J mice were from Jackson Laboratories, CD44−/− mice (Schmits et al., 1997) were backcrossed onto the C57BL/6J background for nine generations, and maintained at the University of British Columbia (UBC). Six to ten-week-old mice were used for experiments. All animal experiments were conducted in accordance with the Canadian Council of Animal Care Guidelines using protocols approved by the University Animal Care Committee. 45  2.2.2 Reagents Pharmaceutical grade HA of various sizes (HA15M-1 (~1680 kDa), HA200K-1 (~234 kDa), HA20K-1 (~28 kDa), HA5K-1 (<10 kDa) were purchased from Lifecore Biomedical and are referred to as 1.5M, 200K and 20K HA. The HA 4-mer (tetrasaccharide) was from Seikagaku. Rc HA (H5388) and Huc HA (H1504) were from Sigma-Aldrich. Fluorescein-labeled HA (FL-HA) was made by the Antibody Laboratory at UBC or according to reference (de Belder and Wik, 1975). Ultra-pure LPS was from Escherichia coli strain 0111:B4 from InvivoGen. Bov HA’se was from Sigma-Aldrich (H4272), and HA’se isolated from Streptomyces hyalurolyticus was from EMD Millipore (389561). Proteinase K was from Fisher BioReagents and saponin from Calbiochem. J558L cell-conditioned medium was the source of CSF-2 (Stockinger et al., 1996), L929 cell-conditioned medium was the source of CSF-1. Mouse recombinant TSG-6 was from R&D Systems (2326-TS-050). All other reagents and chemicals were from Sigma-Aldrich.  2.2.3 Antibodies F4/80 (BM8) phycoerythrin-cyanin 7 (PE-Cy7), CD44 (IM7) Pacific Blue, CD11b (M1/70) Pacific Blue, CD11c (N418) PE-Cy7, Gr1 (RB6-8C5) Pacific Blue, CD86 (P03.1) phycoerythrin (PE), MHC II (M5/114.15.2) allophycocyanin (APC), Ly6C (HK1.4) peridinin chlorophyll protein-cyanin 5.5 (Percp-Cy5.5), Ly6G (1A8) PE/Cy7, Siglec-F (E50-2440) PE, CD40 (HM40-3) fluorescein isothiocyanate (FITC), CD86 (GL1) PE, TNF-α (MP6-XT22) PerCP-eFluor 710, and IL-12/23p40 (C17.8) PE antibodies and streptavidin PE were purchased from eBioscience, Biolegend, or 46  the Antibody Laboratory at UBC. The Dako anti-IαI antibody was from the Programs of Excellence in Glycosciences at the Cleveland Clinic.   2.2.4 Cell isolation and culture CSF-2 BMDM and BMDC and CSF-1 BMDM were prepared as described previously (Poon et al., 2015; Ruffell et al., 2011). Briefly, bone marrow cells were isolated from the femurs and tibias of mice and treated with RBC lysis buffer (0.84% ammonium chloride, 2 mM Tris-Cl pH 7.2) for 5 min at RT. Two million bone marrow cells were cultured in a 100 × 15-mm petri dish (Falcon) in 10 ml complete RPMI 1640 medium (10% fetal bovine serum (FBS) 20 mM Hepes, 1× nonessential amino acid, 55 μM 2-mercaptoethanol, 50 U/ml penicillin/streptomycin, 1 mM sodium pyruvate (all from Invitrogen), 2 mM L-glutamine, (Sigma-Aldrich) supplemented with J558L supernatant containing 20 ng/ml of CSF-2). Non-adherent cells were harvested on day 7. For CSF-1 derived BMDMs, 1.5 x 107 bone marrow cells were cultured in 10 ml of DMEM, 20% FBS, 1 mM sodium pyruvate, 2 mM L-glutamine, 50 U/ml penicillin/streptomycin, and 8% L929 cell-conditioned media (LCCM) as a source of CSF-1. BMDMs were harvested using Versene at day 5 and re-plated in a 96-well plate overnight, and cells stimulated on day 6. Peritoneal macrophages were obtained by peritoneal lavage using Ca2+ and Mg2+ free Hanks' balanced salt solution (Invitrogen). Recovered cells were plated in DMEM with 20% FBS in a flat bottom 96-well plate for 2 hr, non-adherent cells discarded, and the remaining adherent cells (enriched for peritoneal macrophages) used for analysis. Splenic cells 47  were isolated from spleen homogenized by collagenase digestion (1 mg/ml collagenase IV (Worthington) in PBS and 5% FBS) for 20 min at RT.   2.2.5 Cell staining CSF-2 BMDM and BMDC cell suspensions were incubated in 2.4G2 tissue culture supernatant to block Fc receptors. Cells were then incubated with fluorescently labeled antibodies for 20 minutes at 4°C, followed by two washes of 300 µl 2% bovine serum albumin (BSA), 2 mM EDTA, in PBS. For measurement of intracellular cytokines, cells were stimulated in complete medium for 8 hr at 37°C, with 10 µg/ml Brefeldin A (Sigma) added after 2 hr to block protein transport. Cells were then labeled with antibodies as above, fixed in 4% paraformaldehyde (ThermoFisher), permeabilized in saponin buffer (0.1% saponin, 1% BSA in PBS), and labeled with antibodies specific to TNF-α or IL-12, for 30 minutes at RT. Brefeldin A was not used for flow cytometry of co-stimulatory molecules, and the cells were stimulated for 24 hr. Antibody labeled cells were analyzed by flow cytometry (BD LSRII).   2.2.6 Intratracheal instillation and bronchoalveolar lavage (BAL) Mice were anesthetized by isoflurane and instilled with 50 µl of PBS containing LPS (500 µg/ml), 200K HA (2 mg/ml) or 20K HA (2 mg/ml). Mice were euthanized 24 hr later by isoflurane overdose, and alveolar cells harvested by BAL via catheterization of the trachea and washing three times, each with 1 ml 1x PBS, 48  2% BSA, and 2 mM EDTA.  Cells were pelleted and then treated with RBC lysis buffer.  2.2.7 Cell stimulation and analysis of inflammation  Splenocytes, peritoneal macrophages, BMDMs or BMDCs were stimulated in 96-well, non-tissue culture treated, round bottom plates (Falcon). BMDMs were left to adhere overnight prior to stimulation (and peritoneal macrophages for 2 hr). Cells were cultured at 2 x 105 cells per well in 200 µl complete medium lacking CSF-1 or -2 and stimulated with 20 ng LPS (InvivoGen) or 20 µg of various HA preparations, with the exception of the HA 4-mer, where 2 µg was added. Alternatively, cells were stimulated with 100 µl of the reaction mixture containing 10 µg of HA complexed to HC in a final volume of 200 µl media. Cells were stimulated for 8 hr for intracellular cytokine measurement or 24 hr for ELISA (TNF-α, IL-12, IL-1β and CCL3, eBioscience) and co-stimulatory molecule expression. To measure IL-1β secretion, cells were primed with 2 ng of LPS in 200 µl media, and then 3 hr later, either 5 mM ATP or 20 µg of various HA preparations were added.   2.2.8 Fragmentation of HA 1.5M HA at 2 mg/ml was digested with 2 U/ml Bov HA’se overnight at 37°C to produce HA fragments. One ml of 4 mg/ml 1.5M HA in a microcentrifuge tube was sonicated in an ultrasonic bath (VWR) operating at 35 kHz for 24 hr. Two mg/ml of 1.5M and 200K HA were treated with 100 µM H2O2 for 24 hr at 37°C to generate fragmented HA.  49   2.2.9 Inactivation of HA’ses Bov and Strp HA’ses at 200 U/ml were degraded with 100 µg/ml proteinase K in 50 mM Tris pH 7.5, 5 mM CaCl2 for 30 min at 50oC then the enzyme denatured at 95° C for 5 min.   2.2.10 Polymyxin B removal of LPS  One ml of Huc HA at 200 µg/ml was incubated with 200 µl of packed polymyxin B coupled agarose beads (Sigma) in microcentrifuge tubes, and mixed end-over-end for 1 hr at RT. The supernatant was removed and treated a second time with the beads. 100 µl of the treated Huc HA was added to 100 µl of cells in complete media for stimulation.   2.2.11 Triton X-114 removal of LPS As previously described (Liu et al., 1997): 5 µl (1%) of cold Triton X-114 was added to 500 µl of ice-cold 2 mg/ml HA preparation and vortexed gently for 5 min. The mixture was then transferred to 37° C for 5 min to allow micelles to form and aggregate. Samples were then centrifuged at 10,000 x g for 30 seconds and kept at 37°C until the Triton X-114 formed a clear, immiscible layer at the bottom of the tube. The top aqueous fraction was collected. This cycle was repeated once more, and then used to stimulate cells. Endotoxin levels were measured by Endpoint Chromogenic limulus amebocyte lysate assay (Lonza).   50  2.2.12 Visualization of HA on agarose gel HA (2-4 µg) was loaded on a 1% agarose (Fisher) gel in 40 mM Tris, 20 mM acetic acid, 1 mM EDTA (TAE buffer) and run at 100 V for approximately 1.5 hr. The gel was stained overnight in 0.005% Stains-All (Sigma) in 50% ethanol filtered twice, then de-stained in 10% ethanol and imaged on a gel documentation light cabinet (Alpha Innotech Corp).   2.2.13 Generation of HA-HC complexes in vitro HA-HC complexes were generated as previously described (Lauer et al., 2013). Briefly, 1.5M or 20K HA (100 µg) was incubated with mouse serum (100 µl), and mouse recombinant TSG-6 (12.5 µg) in 1 ml of RPMI 1640 for 4 hr at 37°C. The generation of HA-HC complexes was verified, and then 100 µl of the reaction mixture, containing the HA-HC complexes, recombinant TSG-6 and mouse serum, was used to stimulate cells.   2.2.14 Detection of HA-HC complexes with Western blot HA-HC complexes generated above (10 µl) and control samples were incubated or not with Strp HA’se (2 µl) in PBS to a total of 25 µl for 1 hr at 37°C. Each sample (10 µl) was heated to 95°C for 5 mins and loaded on to a polyacrylamide gel, transferred to a membrane, blotted with the primary anti-IαI antibody (Cleveland Clinic) followed by a LI-COR donkey anti-rabbit secondary antibody for HC detection using an Odyssey Scanner.   51  2.2.15 Data analysis Flow cytometry was analysed using Flowjo VX (Treestar). Graphs were generated using Graphpad Prism 6. Data shown are the average ± standard deviation (SD). Significance was determined by a Student two-tailed, unpaired t test with *p < 0.05, **p < 0.01, ***p < 0.001, unless otherwise specified.   52  2.3 Results 2.3.1 Different sizes of pharmaceutical grade HA do not stimulate CSF-1 derived bone marrow macrophages but human umbilical cord HA does  Figure 2.1: Stimulation of CSF-1 BMDMs by various HA preparations a) Representative flow cytometry plots and bar graphs summarizing the percent of cells producing intracellular TNF-α and IL-12 p40 after stimulation by various HA preparations (see Key for details). b) Representative flow cytometry plots and bar graphs summarizing CD40 and F4/80 expression measured by MFI in response to stimulation by various HA preparations. c) Summary of production of CCL3 and IL-1β secreted by BMDMs after stimulation, measured by ELISA. d) Bar graph summaries comparing TNF-α and IL-12 p40 production between CD44+/+ and CD44-/- BMDMs after stimulation. e) TNF-α and IL-12 p40 production from F4/80 gated 53  peritoneal macrophages after stimulation. Graphs show an average of three mice from one experiment ± SD, repeated twice. Numbers in the flow cytometry plots refers to the percent of cells in each quadrant or box. Significance compared to the unstimulated control indicated as * p < 0.05, ** p<0.01, *** p < 0.001, unpaired student’s t-test.  In past studies, HA preparations purified from human umbilical cord or rooster comb were used that have a range of molecular masses. LMW HA (ranging from 4 to 200 kDa) derived from these preparations induced inflammatory activation of CSF-1 induced BMDMs and peritoneal macrophages (reviewed inLee-Sayer et al., 2015; Monslow et al., 2015). Now, commercial HA is purified after bacterial fermentation (Lifecore Biomedical, HTL Biotechnology) or enzymatic synthesis (Hyalose) and can generate HA of specific molecular mass ranges that are also endotoxin free. To compare the effects of HA from these different sources, CSF-1 induced BMDMs were challenged with a panel of commercially available purified HA of different specific molecular sizes (HA 1.5M, 200K, 20K, <10K daltons (see Methods for more details) from Lifecore Biomedical and HA 4-mer composed of four monosaccharides from Seikagaku), as well as HA from Rc, and Huc from Sigma-Aldrich. Cells were stimulated with 100 µg/ml of HA, or 10 µg /ml of HA 4-mer, or with 100 ng/ml of ultra-pure lipopolysaccharide (LPS, from E. coli strain 0111:B4) as the positive control. To assess the stimulatory activity of HA, key propagators of the inflammatory response were measured: the pro-inflammatory cytokines TNF-α, IL-12, and IL-1β; the co-stimulatory molecule CD40, and F4/80; and the chemokine CCL3 (MIP-1α). Only LPS and Huc HA treated macrophages produced significant amounts of TNF-α, IL-12, CCL3, and upregulated CD40 and F4/80 expression (Figure 2.1a-c), measured by intracellular cytokine production and MFI using flow 54  cytometry. The level of stimulation induced by Huc HA was similar to that of LPS. To test if the inflammatory effect of Huc HA on macrophages was dependent on the HA surface receptor CD44, CSF-1 induced BMDMs lacking CD44 expression were generated from the bone marrow of CD44-/- mice and stimulated with Huc HA. CD44-/- macrophages were not deficient in their pro-inflammatory cytokine production to Huc HA, demonstrating that the effect of Huc HA is not dependent on CD44 (Figure 2.1d). IL-1β was not detected from cells stimulated with any of the HA after priming with LPS (Figure 2.1c), suggesting that HA does not act like extracellular ATP, to activate the NLRP3 inflammasome.  Mouse CSF-1 BMDMs have similarities to in vivo F4/80+ peritoneal macrophages. Peritoneal cells were collected by lavage and stimulated by the same panel of HAs. Although these cells did not produce much intracellular IL-12 after stimulation, LPS induced substantial amounts of TNF-α, and again, this was mirrored by Huc HA (Fig. 2.1e).     55  2.3.2 CSF-2 but not CSF-1 derived BMDMs and BMDCs constitutively bind FL-HA  Figure 2.2: FL-HA binding by CSF-1 BMDMs, peritoneal macrophages, and CSF-2 BMDMs and BMDCs a) Representative histogram and summary graph showing the MFI of FL-Rc HA binding by CSF-1 induced BMDM. CD44-/- cells (dotted line and white bar) were used as a negative control, along with a fluorescence minus one (FMO) control for FL-HA on CD44+/+ cells (hatched bar). b) Representative histogram and graph showing the MFI of FL-Rc-HA binding by peritoneal macrophages. FMO for FL-HA on CD44+/+ cells (dotted line and hatched bar) is the negative control. c) Representative histogram and summary graph showing the MFI of FL-HA binding by CSF-2 BMDMs and BMDCs. CD44-/- cells (dotted line and white bar) were used as a negative control, along with a FMO control for FL-HA on CD44+/+ cells (hatched bar). The number in the flow cytometry plot refers to percent of cells in the gate. Key: Rc = rooster comb HA, Huc = human umbilical cord HA, H = 1.5M, M = 200K, L = 20K. Graphs show an average of six mice pooled from two experiments ± SD. Significance indicated as * p < 0.05, ** p<0.01, *** p < 0.001, unpaired student’s t-test.  56  To determine if this lack of response could be due to the inability of CSF-1 BMDMs to bind HA, we evaluated HA binding using FL-HA. We had previously shown that prior to stimulation, BMDMs and peritoneal macrophages do not bind appreciable levels of HA (Ruffell et al., 2011). Here we confirmed that no appreciable level of binding was seen in CSF-1 BMDMs or peritoneal macrophages by flow cytometry with FL-HA from Huc, Rc, and specific sizes of HA (Figure 2.2a-b). In contrast, CSF-2 elicited bone marrow derived cultures contain both CD11c+ HA binding and non-binding populations, which have been attributed to HA binding BMDMs and non-HA binding BMDCs, respectively (Poon et al., 2015). Figure 2.2c shows that a significant percentage of the CSF-2 cultured cells bound FL-HA. CD44+/+ CSF-2 BMDMs bound significant levels of Rc FL-HA, Huc FL-HA, 1.5 M FL-HA, and 200K FL-HA compared to CD44-/- CSF-2 BMDMs. However, specific binding of 20K FL-HA to CD44 was not detectable, but this and the lower labeling with the 200K FL-HA may be attributed to their lower fluorescein conjugation ratio. Given the ability of a significant percentage of these cells to bind FL-HA, we next determined if the different sized HA samples could generate an inflammatory response.  57  2.3.3 Different sizes of pharmaceutical grade HA do not simulate CSF-2 derived BMDMs and BMDCs, but human umbilical cord HA does  Figure 2.3: Stimulation of CSF-2 BMDMs and BMDCs by various HA preparations  a) Representative flow cytometry plots showing the percent of CD44+/+ CSF-2 BMDMs and BMDCs positive for intracellular TNF-α and IL-12 p40 production after stimulation. b) Representative flow cytometry plot showing the percent of CD40 and CD86 expressing cells after stimulation with various HA preparations. c) Bar graphs summarizing TNF-α and IL-12 production from CD44+/+ and CD44-/- BMDMs and BMDCs after HA stimulation. d) Bar graphs summarizing the percent of CD40 and CD86 expressing cells in CD44+/+ and CD44-/- cells after HA stimulation. e) IL-1β production from CD44+/+ CSF-2 BMDMs and BMDCs after priming with 10 ng/ml LPS then stimulation with various HA. Numbers in the flow cytometry plots are the percent of cells in each quadrant. Key: U = unstimulated, -ve = media only, LPS = lipopolysaccharide, Rc = rooster comb HA, Huc = human umbilical cord HA, H = 1.5M, M = 200K, L = 20K. Graphs show an average of six mice pooled from two experiments ± SD. Significance compared to the unstimulated control indicated as * p < 0.05, ** p<0.01, *** p < 0.001, unpaired student’s t-test.  In vitro stimulation of CSF-2 BMDMs and BMDCs with the panel of HA samples (HA 1.5M, 200K, 20K, <10K, 4-mer, Rc and Huc HA) demonstrated that only Huc HA led to significant levels of TNF-α and IL-12 being produced (Figure 58  2.3a and c). Similarly, no responses were obtained with pharmaceutical grade HA of 40K, 200K and 1.5M molecular mass from HTL Biotechnology. Only Huc HA treatment significantly upregulated CD40 and CD86 co-stimulatory molecules, and these elevated levels were comparable to those of LPS stimulation (Figure 2.3b and d). CD44-/- cells also produced TNF-α and IL-12, and increased co-stimulatory molecule expression to the same degree as CD44+/+ cells demonstrating a CD44 independent effect (Figure 2.3c-d). Again, IL-1β production was not detected from CSF-2 BMDMs and BMDCs primed with LPS and then stimulated with any of the HA for 24 hr (Figure 2.3e).   59  2.3.4 Different sizes of pharmaceutical grade HA do not stimulate ex vivo splenic macrophages or DCs while human umbilical cord HA does  Figure 2.4: In vitro stimulation of splenic macrophage and DCs by various HA preparations and the intratracheal instillation of HA into the lung a) Gating strategy for analyzing splenic F4/80+ CD11clow macrophages (box 1) and CD11c+ MHCIIhi DCs (box 2). b) Representative histograms showing FL-Rc-HA binding by CD44+/+ and CD44-/- splenic DCs and macrophages. c) Bar gaphs summarizing percent of cells positive for intracellular TNF-α and IL-12 p40 production by splenic DCs and macrophages after stimulation with various HA. d) Representative flow cytometry plots showing gating strategy to identify leukocytes and alveolar macrophages from the BAL. Alveolar macrophages are Siglec F+ CD11c+ and HA binding, while infiltrated leukocytes are CD11b+. e) Total number of cells in the BAL and the percent of CD11b+ cells after 24 hr following intratracheal instillation with PBS, LPS, 20K (L), or 200K (M) HA. Numbers in the flow cytometry plots refer to the percent of cells in the box or gate. Key: U = unstimulated, LPS = lipopolysaccharide, Rc = rooster comb HA, Huc = human umbilical cord HA, H = 1.5M, M = 200K, L = 20K. Graphs show an average of six mice pooled from two experiments ± SD. Significance compared to the unstimulated control indicated as * p < 0.05, ** p<0.01, *** p < 0.001, unpaired student’s t-test.  60  To test if the effect of HA was recapitulated with analogous ex vivo cells, splenic macrophages and DCs were tested for their ability to produce pro-inflammatory cytokines. Splenic macrophages were identified as CD11clow F4/80+ cells, whereas splenic DCs were identified as CD11chigh MHCIIhigh cells (Figure 2.4a). The HA binding of splenic F4/80+ macrophages and DCs was measured by FL-HA and was similar to the CSF-2 BMDMs and BMDCs: the majority of F4/80+ macrophages bound HA whereas the majority of splenic DCs did not (Figure 2.4b). When the splenic cells were stimulated with the panel of HA molecules, again only Huc HA stimulated a significant percentage of cells to produce TNF-α and IL-12 (Figure 2.4c). Thus, pharmaceutical grade HA (1.5M, 200K, 40K, 20K, <10K, 4-mer) and Rc HA were not pro-inflammatory for in vitro derived or ex vivo macrophages and DCs. Huc HA was the only HA that stimulated cytokine production and co-stimulatory molecule upregulation to similar levels seen with LPS stimulation, and this occurred independently of CD44.   2.3.5 Administration of 20K and 200K HA in vivo does not induce lung inflammation  Previous studies show that HA is upregulated in the lung in response to oxidative damage, and intratracheal instillation of short chain HA increases lung airway hyper-responsiveness (Lazrak et al., 2015). LMW HA is increased in cigarette smoke sensitized lung (Bracke et al., 2010), and induces chemokine production in ex vivo alveolar macrophages (McKee et al., 1996). Similar to CSF-2 derived macrophages and splenic F4/80+ macrophages, CD11b- Siglec F+ AMФ 61  isolated by BAL bind high levels of FL-RcHA (Figure 2.4d). To test if smaller sized HA can cause inflammation in the lung, 100 µg of 200K HA or 20K HA, PBS, or 25 µg LPS were introduced separately into mouse lungs via intratracheal instillation. At steady-state, the alveolar airspace predominantly contains AMФ, but upon inflammatory stimulation such as with LPS, CD11b+ leukocytes are recruited to the alveolar space (Andonegui et al., 2003). Thus, the degree of lung inflammation was measured after 24 hr by comparing the percentage of CD11b+ leukocyte infiltration into the alveolar airspace and the total number of cells retrieved from the BAL. LPS treated mice had significantly increased numbers of total cells and percentage of CD11b+ leukocytes, whereas the cell numbers in both 200K and 20K HA treated mice were similar to PBS treated mice (Figure 2.4e-f). Thus, no evidence was found to support the ability of these sizes of pharmaceutical grade HA to stimulate an in vivo inflammatory response leading to leukocyte recruitment in the lung.   62  2.3.6 Fragmented HA is not inflammatory to CSF-2 derived BMDMs and BMDCs  Figure 2.5: Sizes of various HA preparations and their effect on CSF-2 BMDMs and BMDCs  a) Representative agarose electrophoresis gel showing the size range of various HA preparations from Lifecore or fragmented by overnight sonication (S), H2O2 incubation, or Bov HA’se digestion. H = 1.5M, M = 200K, L= 20K. The gel on the right shows Rc and Huc HA.  b) Summary of the percent of cells producing TNF-α and IL-12 p40 after stimulation with the above HA preparations. c) Summary of the percent of cells expressing CD40 and CD86 in response to HA. Graphs show an average of six mice pooled from two experiments ± SD. Significance compared to the unstimulated control indicated as * p < 0.05, ** p<0.01, *** p < 0.001, unpaired student’s t-test.  One could argue that in vivo generated HA fragments may differ from purified HA available from commercial sources both by how the fragments are generated 63  (enzymatic or chemical cleavage) and how HA is associated with other proteins. Tissue HA can be fragmented enzymatically (Bracke et al., 2010), or by ROS (Agren et al., 1997), potentially generating altered HA structures that may be detectable by sentinel immune cells. To test whether immune cells are activated by HA fragments generated by these methods, CSF-2 derived BMDMs and BMDCs were stimulated with HA degraded physically by sonication, chemically via H2O2, or enzymatically using Bov HA’se. These methods generated a range of HA sizes (Figure 2.5a). However, stimulating cells with these prepared HA fragments did not elicit any significant TNF-α or IL-12 production (Figure 2.5b) and did not upregulate co-stimulatory molecule expression (Figure 2.5c).    64  2.3.7 HC-HA complexes do not induce an inflammatory response in CSF-2 BMDMs and BMDCs  Figure 2.6:  Western blot demonstrating the generation of HA-HC complexes and their effect on stimulating CSF-2 BMDMs and BMDCs  a) Representative anti-IαI (HC) Western blot showing the HC of IαI in serum and when bound to HA by TSG-6 and when released by Strp HA’se. b) Summary graphs of the percent of cells expressing TNF-α and IL-12 p40 production after stimulation with in vitro generated HA-HC complexes or with control samples containing just serum and TSG-6. Graphs show an average of six mice pooled from two experiments ± SD. Significance compared to the unstimulated control indicated as * p < 0.05, ** p<0.01, *** p < 0.001, unpaired student’s t-test.  As the formation and deposition of TSG-6 mediated HA-HC complexes occurs in vivo, for example during ovalbumin induced lung asthma in mice (Swaidani et al., 2013), HA-HC may be another form of HA that initiates the inflammatory response in macrophages or DCs. HA-HC complexes were generated in vitro by incubating mouse serum as a source of IαI, mouse recombinant TSG-6, and 1.5M or 65  20K HA at 37°C. The presence of HA-HC complexes was confirmed after HA’se treatment by Western blot using an IαI specific antibody (Figure 2.6a). In the serum, the HC of IαI is covalently associated with bikunin generating higher molecular mass forms of ~100 (preII) and 200 (II) kDa (Lauer et al., 2013). When TSG-6 adds HC to HA, the size will depend on the size of the HA and how many HC are conjugated: lane 4 shows a range of molecular weight for HC conjugated to 20K HA; lane 6 shows a band that did not enter the gel when HC was conjugated to 1.5M HA. Treatment of these samples with HA’se releases the HC from the HA and results in an increase in the HC band at 75 kDa (see arrow, lanes 5 and 7). This demonstrates that TSG-6 covalently bound HC to both 20K and 1.5M HA. These unpurified reaction mixtures containing the HA-HC complexes as well as recombinant TSG-6, serum and any unconjugated HA, were then used to stimulate CSF-2 BMDMs and BMDCs at a final concentration of HA of 10 µg/ml. However, no TNF-α or IL-12 was detected (Figure 2.6b), showing that neither the in vitro derived 20K HA-HC nor the 1.5M HA-HC complexes initiated pro-inflammatory cytokine production in CSF-2 derived BMDMs or BMDCs.  66  2.3.8 Huc HA and HA’se are contaminated with endotoxin  Figure 2.7: The effect of Bov and Strp HA’se on the activation of CSF-2 BMDMs and BMDCs a) Bar graphs summarizing the percent of cells producing TNF-α and IL-12 p40 after exposure of the cells to Bov and Strp HA’se stimulation for 8 hr; proteinase K (PK) and heat was used to degrade HA’se. b) Concentration of endotoxin present in 20 U/ml of Bov and Strp HA’se before or after polymyxin B treatment as measured by the LAL assay. c) Bar graphs summarizing the percent of TNF-α and IL-12 p40 producing cells assessed by intracellular cytokine labeling after stimulation with HA’ses treated with or without polymyxin B. Graphs show an average of six mice pooled from two experiments ± SD. Significance compared to the unstimulated control indicated as * p < 0.05, ** p<0.01, *** p < 0.001, unpaired student’s t-test.  Previous studies have used HA’se to generate lower molecular mass HA to stimulate cells in vitro. Bov or Strp HA’ses were added to CSF-2 induced BMDMs and BMDCs, and these enzymes induced TNF-α and IL-12 production (Figure 2.7a). However, HA’ses degraded with proteinase K and then denatured by heat inactivation also stimulated cytokine production, demonstrating that the ability of 67  these reagents to stimulate macrophages and DCs was independent of the HA’se protein. The presence of endotoxin contamination was confirmed in both Bov and Strp HA’ses by the LAL assay (Duner, 1993) (Figure 2.7b). Treatment with agarose beads coupled to polymyxin B, a cationic neutralizer of LPS (Cooperstock, 1974), was able to significantly lower endotoxin levels (Figure 2.7b), and this caused significantly less TNF-α and IL-12 production (Figure 2.7c), suggesting that the explanation for the positive effect was endotoxin contamination.   Figure 2.8: Endotoxin contamination and removal from Huc HA and its effect on the production of pro-inflammatory cytokines by CSF-2 BMDMs and BMDCs a) Level of endotoxin contamination in 100 µg/ml of the HA preparations measured by LAL assay. b) Endotoxin concentration in Huc HA before and after one (+) or two (++) treatments with polymyxin B or Triton-X 114 (TX), measured using the LAL assay. c) Percent of Huc HA stimulated cells making TNF-α and IL-12 p40 as assessed by intracellular labeling after polymyxin B treatment. d) Graphs showing the percent of cells making TNF-α and IL-12 p40, assessed after intracellular labeling, after stimulating the cells with decreasing amounts of LPS, Huc HA or Huc HA treated with polymyxin B (PB). The amount of LPS present in the Huc HA sample was as indicated and compared to an equivalent amount of LPS. The 68  effectiveness of polymixin B on reducing the Huc HA response is shown. e) Representative agarose electrophoresis gel showing the amount of Huc HA present before and after two rounds of TX-114 treatment. f) Percent of cells making TNF-α and IL-12 p40 production after stimulation with Huc HA or Huc HA after two cycles of TX-114 treatment. Graphs show an average of six mice pooled from two experiments ± SD. Significance indicated as * p < 0.05, ** p<0.01, *** p < 0.001, unpaired t-test.   The presence of endotoxin in Bov and Strp HA’se raised the possibility that the inflammatory properties of Huc HA may also be caused by contamination. The same concentration of various sized HA and HA fragments used in this study was tested in the LAL assay, revealing large amounts of endotoxin contamination in Huc HA (approximately 2 ng/ml in the 100 µg/ml of Huc HA used to stimulate cells), but no detectable levels of endotoxin were found in all other purified forms of HA (Figure 2.8a). Although previous studies have used polymyxin B to remove endotoxin from HA samples (Oertli et al., 1998; Sokolowska et al., 2014), the efficacy of this method of endotoxin removal was not determined. One study showed polymyxin B was only effective when removing endotoxin from low Huc HA concentrations (Wallet et al., 2010). Here, we found that polymyxin B treatment was not effective in removing endotoxin contamination from Huc HA at 100 µg/ml (Figure 2.8b), nor was it effective at reducing the pro-inflammatory effect of Huc HA, used at 100 µg/ml (Figure 2.8c). However, a titration comparing Huc HA and LPS at equivalent concentrations as the contamination (3000 pg/ml, 300 pg/ml, 30 pg/ml, and 3 pg/ml LPS) showed that contaminated Huc HA and LPS have a similar ability to stimulate cytokine production upon dilution, whereas the titration of Huc HA treated with polymyxin B revealed a limited but significant reduction in amount of TNF-α and IL-12 production, especially at the lower pg/ml concentrations (Figure 69  2.8d). Polymyxin B was able to remove small amounts (pg/ml) of endotoxin from the Huc HA sample, making the more diluted samples incapable of activating cells. Interestingly, IL-12 was not produced in response to lower concentrations of LPS whereas the cells were still able to produce TNF-α. As an alternative method for endotoxin removal, Huc HA was treated with two cycles of Triton X-114. This method uses micellar phase separation to remove LPS endotoxin from the aqueous phase (Liu et al., 1997). Huc HA treated for two Triton X-114 cycles significantly removed most of the endotoxin, leaving levels just at the point of detection by the LAL assay at 10 pg/ml, over a 100 times reduction (Figure 2.8b). Importantly, two cycles of Triton X-114 treatment did not significantly change the amount of Huc HA recovered in the aqueous phase (Figure 2.8e), and Huc HA treated with two Triton X-114 cycles significantly reduced TNF-α and IL-12 production to background levels (Figure 2.8f) demonstrating that the pro-inflammatory effect of Huc HA is due to endotoxin (LPS) contamination.  Taken together, we conclude that 1.5M, 200K, 40K, 20K, <10K, 4-mer, Rc, in vitro fragmented HA, as well as in vitro generated HA-HC complexes do not stimulate macrophages or DCs to make the key pro-inflammatory cytokines, TNF-α and IL-12. The stimulatory effect of Huc HA was attributed to endotoxin contamination and occurred independently of CD44 on macrophages and DCs.  2.4 Discussion This work re-evaluates the ability of various HA fragments below 250 kDa to act as DAMPs and concludes that pharmaceutical grade HA and HA fragments by themselves do not stimulate macrophages or DCs to produce TNF-α, IL-1β, IL-12, or 70  CCL3, and do not upregulate CD40 or CD86 co-stimulatory molecules. Here, both in vitro derived and ex vivo macrophages and DCs were treated with pharmaceutical grade HA of different molecular mass, with chemically and enzymatically generated HA fragments, and with in vitro generated HA-HC complexes, and none of the above pro-inflammatory responses were observed. In the context of these results, this raises the possibility that observed increases in lower molecular mass HA in inflamed tissues may be a consequence, not a cause, of a type 1 inflammatory response where TNF-α, IL-1β, and IL-12 are produced and the co-stimulatory molecules, CD40 and CD86, are upregulated. In this study, we cannot exclude that some other proinflammatory cytokine or chemokine could be made in response to HA, but if this was the case, we would have expected to see leukocyte recruitment in response to HA instilled into the lungs.  However, a full transcriptional and proteomic profile will be required to definitely determine if HA or HA fragments can activate any aspect of a pro-inflammatory response in macrophages and DCs. In contrast to the purified HA and HA fragments from Lifecore, Huc HA and HA’se treated samples induced a CD44 independent pro-inflammatory response that was subsequently shown to be due to endotoxin contamination. Notably, polymyxin B, a cationic neutralizing agent of LPS (Cooperstock, 1974) was able to remove endotoxin from the HA’se protein, but was much less efficient at removing endotoxin from Huc HA preparations. Perhaps the negatively charged HA molecules (Thalberg and Lindman, 1989) can interfere with the binding of cationic polymyxin B to LPS, which is also negatively charged (Schromm et al., 1998). Alternatively, both HA and LPS contain N-acetylglucosamine, which is important for binding to polymyxin(Mares 71  et al., 2009), therefore excess HA may compete with LPS for binding to polymyxin B. It is thus imperative to monitor the removal of endotoxin, ideally to 1-3 pg/ml levels, to minimize macrophage and DC activation. We found Triton X-114 to be more effective in removing endotoxin from HA preparations. Because of this potential endotoxin contamination in certain HA preparations and HA’ses, researchers should review the sources and methods in which past studies utilized and generated HA fragments. Some information in this regard can be found in two recent reviews (Lee-Sayer et al., 2015; Monslow et al., 2015). Furthermore, researchers should always check that the endotoxin levels of HA reagents are 1-10 pg/ml or lower.  Past studies used HA to stimulate cells at various concentrations ranging from 10 µg/ml to 1 mg/ml (Krejcova et al., 2009; McKee et al., 1996; Scheibner et al., 2006; Termeer et al., 2000b). In this study, a concentration of HA (100 µg/ml) was used to stimulate cells, and no response was observed. Other experiments with 1.5M, 200K, 20K and Rc HA at concentrations as high as 500 µg/ml and HA 4-mer concentrations of 50 µg/ml also did not elicit an inflammatory response.  Several forms of fragmented HA were used to stimulate cells in vitro in an attempt to cover the possible forms of HA that might be encountered in vivo. Various sizes and differently generated fragments as well as complexed HA-HC were used, all with no effect. Pharmaceutical grade HA is purified from fermenting Streptococcus equi under specific conditions, such that the rate of HA polymerisation and availability of substrates allow for the generation of HA in a specific size range (Chong et al., 2005). Both S. equi and vertebrate HAS are Class 1 HAS (Chong et al., 2005), and bacterial HAS are not known to generate a HA 72  molecule that is different from mammalian HA. It is also unlikely that the purification procedure introduces subtle changes in this HA as it remains capable of binding avidly to CD44 even after conjugation with fluorescein (Poon et al., 2015; Ruffell et al., 2011). In vivo, the form of HA present at both homeostasis and during inflammation is not clear; it could be nascent, or complexed with other HA binding proteins or matrix components. It is also unclear whether HA generated in vivo is inflammatory or possesses a different conformational structure that can be recognized by cells. While the addition of 200K and 20K sizes of pharmaceutical grade HA did not induce inflammatory leukocyte infiltration into the alveolar space in vivo, we cannot exclude the possibility that complexed forms of HA generated in vivo in response to an infection or damage are differentially recognized by pattern recognition receptors. For example, it is not known how well the HA-HC complexes generated in vitro mimic HA-HC complexes generated in vivo in response to an inflammatory insult. However, the in vitro data presented here suggest that HMW HA is not inflammatory, and smaller HA fragments, on their own, are insufficient to trigger an inflammatory response.  The conflicting roles of HA being anti-inflammatory and pro-inflammatory depending on its size and form has always been difficult to understand mechanistically. However, as studies demonstrating a direct effect of HA on inflammation have relied on stimulating cells or animals with exogenous HA (Lazrak et al., 2015; Li et al., 2011b; McKee et al., 1996; Noble et al., 1996; Scheibner et al., 2006; Vistejnova et al., 2014; Wallet et al., 2010), caution is needed, and efforts should be made to ensure that the effects are not due to picogram levels of 73  endotoxin contamination, from either the HA source or from the HA’se used to generate the fragments. Endotoxin contaminated Huc HA tested in this study activated macrophages and DCs regardless of the HA binding ability of CD44 or CD44 expression, consistent with the ability of LPS to induce pro-inflammatory effects through TLR4, which was also reported to be necessary for HA induced inflammation (Martin et al., 2008; Scheibner et al., 2006).  In vivo mouse models where LMW or fragmented HA are generated without introducing exogenous HA such as from the overexpression of human HA’se 1 (Muto et al., 2014) may provide a better approach. In this case, expression of human HA’se 1 in the skin did not lead to spontaneous inflammation. However, its induction enhanced DC migration to the draining lymph node, and this enhanced contact hypersensitivity if induced concurrently with a sensitizing agent (Muto et al., 2014). Irrespective of whether HA is a cause or consequence of inflammation, HA is increased upon inflammation, and its abundance and/or size in the tissue, and possibly the serum, may provide a useful biomarker for the state of inflammation. This is an ongoing area of research, for example in arthritis (Sasaki et al., 2015), breast cancer metastasis (Peng et al., 2015), and liver fibrosis (Neuman et al., 2016).   In conclusion, purified, endotoxin-free, exogenously added HA of various molecular mass, fragmented HA, in vitro complexed HA-HC, and HA’ses were unable to induce pro-inflammatory responses in in vitro derived and ex vivo murine macrophages and DCs, as measured by the production of TNF-α, IL-1β, IL-12, CCL3, and the expression of the co-stimulatory molecules, CD40 and CD86. HA 74  fragments of 200K and 20K did not stimulate the in vivo pro-inflammatory response of leukocyte recruitment after direct lung instillation. Endotoxin contamination was present in Huc HA, Bov HA’se, and Strp HA’se, and was responsible for their pro-inflammatory ability as this was lost upon endotoxin removal. This demonstrates the need to ensure HA preparations and HA’se treated samples are free of pg amounts of endotoxin contamination.   75  Chapter 3: The survival of fetal and bone marrow monocyte-derived alveolar macrophages is promoted by CD44 and its interaction with hyaluronan 3.1 Introduction  The airways are primary entry sites for pathogens and airborne particles (Iwasaki et al., 2017; Kopf et al., 2014) where AMФ are a first line of defense to phagocytose pathogens, particulates, and cell debris (Hussell and Bell, 2014; Iwasaki et al., 2017; Kopf et al., 2014). Shortly after birth, CSF-2 drives the differentiation of fetal monocytes into long-lived AMФ capable of self-renewal (Guilliams et al., 2013; Hashimoto et al., 2013; Schneider et al., 2014b; Shibata et al., 2001b). Their self-renewal is sufficient to maintain the AMФ population at homeostasis and does not require replenishment from BM derived monocytes (Guilliams et al., 2013; Hashimoto et al., 2013; Hussell and Bell, 2014). However, BM derived cells are capable of developing into AMФ, as after AMФ ablation by irradiation, subsequent BM reconstitution leads to BM derived AMФ in adult mice (Lavin et al., 2014). Yolk sac macrophages, fetal liver and adult monocytes can colonize the lungs of csf2rb-/- mice and develop into functional AMФ (van de Laar et al., 2016), showing that progenitors of different origins can differentiate into AMФ, raising the possibility that it is the alveolar environment, not the origin of the progenitor, that determines their differentiation into AMФ (Guilliams and Scott, 2017). In acute inflammation, the numbers of AMФ are reduced and then restored after inflammation is resolved. In influenza infection, the recovery of AMФ was attributed to AMФ self-renewal (Hashimoto et al., 2013) whereas 2 months after 76  acute LPS induced inflammation, the majority of AMФ were shown to be BM derived (Maus et al., 2006). Thus, in adult mice, the extent to which monocytes contribute to AMФ renewal after inflammation (Guilliams and Scott, 2017) is not clear, nor are the factors that regulate monocyte differentiation and AMФ renewal after inflammation. Such information would considerably aid the understanding of the AMФ renewal process, which could be crucial for the resolution of inflammation and return to homeostasis.  At homeostasis, AMФ are thought to exist in a somewhat immunosuppressive environment where they phagocytose pathogens without alerting other innate immune cells (Hussell and Bell, 2014). If they cannot clear the infection, they initiate innate immune responses; for example, in response to Streptococcus pneumoniae (Dockrell et al., 2003) and respiratory syncytial virus (Pribul et al., 2008). However, AMФ depletion can also exacerbate pulmonary inflammatory responses (Thepen et al., 1989), (Knapp et al., 2003), suggesting AMФ have an anti-inflammatory role, but the loss of AMФ  could also lead to greater damage to other cells which further stimulates the immune response. Murine AMФ are characterized by the expression of CD11c, Siglec F (Hussell and Bell, 2014), as well as their expression of CD44 and the ability to constitutively bind HA (Poon et al., 2015). CD44 is responsible for the binding and uptake of HA in AMФ in vitro and during lung development after birth (Culty et al., 1994; Underhill et al., 1993a). In bleomycin induced lung inflammation, CD44-/- mice have increased HA accumulation, a buildup of apoptotic neutrophils and the mice die from unremitting inflammation (Teder et al., 2002). This phenotype is partially rescued if the mice are reconstituted with CD44+/+ BM (Liang et al., 2007; 77  Teder et al., 2002), implicating a key role for CD44 in HA clearance and resolution of lung inflammation. CD44 also contributes to neutrophil recruitment to inflammatory sites (McDonald and Kubes, 2015), raising the possibility that CD44 may also contribute to monocyte recruitment to the lungs. Given the constitutive HA binding ability of AMФ, we sought to evaluate the significance of CD44 and HA binding in AMФ homeostasis and renewal under normal and inflammatory conditions.  3.2 Material and methods 3.2.1 Mice C57BL/6J (CD45.2+) and B6.SJL-PtprcaPepcb/BoyJ (CD45.1+) mice were from Jackson Laboratory. These mice, and CD44-/- mice42 backcrossed with C57BL/6J mice for 9 generations, were housed and bred at the University of British Columbia (UBC). Mating C57BL/6J and BoyJ mice gave C57BL/6JxBoyJ heterozygotes (CD45.1+, CD45.2+). C57BL/6 mice expressing GFP from an α-actin-CMV hybrid promoter were provided by Dr. Fabio Rossi at (UBC) and cells from heterozygous GFP+ mice were used for the adoptive transfer experiments. All animal experiments were conducted with protocols approved by the University Animal Care Committee in accordance with the Canadian Council of Animal Care guidelines for ethical animal research.  3.2.2 Reagents L-cell conditioned media (LCCM) containing CSF-1 was generated from L929 fibroblast cells and TCS from J558L cells was the source of CSF-2, as described 78  previously (Dong et al., 2016; Poon et al., 2015). FL-HA was prepared using hyaluronic acid sodium salt from Rooster comb (Sigma-Aldrich), as described previously (de Belder and Wik, 1975). High molecular weight HA (Lifecore Biomedical) was conjugated with Alexa Fluor-647 by AbLab (UBC) for HA-A647. Purified low endotoxin, HA-blocking, rat anti-mouse CD44 mAb, KM81, and rat IgG2a isotype matched mAb were purchased from Cedarlane Laboratories. Recombinant mouse CSF-2 (carrier free) was purchased from BioLegend. ROZ was purchased from Cayman Chemical Company.  3.2.3 Flow cytometry (FC) Cells were incubated with 2.4G2 TCS for 20 min to block Fc receptors, washed with FC buffer (PBS, 2% BSA and 2 mM EDTA), labeled with mAbs and/or FL-HA for 20 min at 4° C, washed twice in FC buffer, and resuspended in FC buffer containing propidium iodide, DAPI, or Live/Dead Fixable Aqua Dead Cell Stain (ThermoFisher) to label non-viable cells. For intracellular labeling, cells were fixed in 4% paraformaldehyde (PFA) for 10 min at RT, permeabilized with PBS, 2 mM EDTA, 0.1% saponin, 1% BSA for 30 min at RT, then incubated with intracellular mAb. The following mAb against mouse antigens were used for flow cytometry, cell isolation, or confocal imaging: CD11c (N418), F4/80 (BM8), CD44 (IM7), CD11b (M1/70), MHC II (M5/114.15.2), CD44 (IM7), CD45.1 (A20), CD45.2 (104), CD206 (C068C2), Ki67 (SolA15), Ly6C (HK1.4), Ly6G (1A8), Siglec F (E50-2440 or 1RNM44N), SIRPα (P84). Antibodies were purchased from Affymetrix eBioscience, 79  R&D Systems, BD Biosciences, or AbLab (UBC). Cells were processed on an LSR II (BD Biosciences) or MACSQuant (Miltenyi Biotec) flow cytometer.   3.2.4 Cell isolation Mice were euthanized by isoflurane overdose. BAL cells were harvested by catheterization of the trachea and washing thrice with 1 ml FC buffer. BAL cells were centrifuged and resuspended in RBC lysis buffer (0.84% NH4Cl in 10 mM Tris buffer, pH7.2) for 5 min, then centrifuged again and resuspended in FC buffer. The whole lung was then perfused with PBS via cardiac puncture of the right ventricle and harvested, minced, incubated in RPMI (with 0.7 mg/ml Collagenase IV (Worthington), 50 µg/ml of DNAse I (Worthington) for 1 h at 37°C, and passed through a 70 µm cell strainer to generate the lung single cell suspension which was treated with RBC lysis buffer, passed through a 35 µm cell strainer, and resuspended in FC buffer.  The peritoneal lavage, obtained with 5 ml of PBS, was centrifuged and treated with RBC lysis buffer for 5 min, then centrifuged and resuspended in FC buffer, incubated with 2.4G2 for 20 min, and labeled with biotinylated F4/80 antibody for 20 min. Then, F4/80+ PMФ were isolated with anti-biotin MicroBeads and LS columns (Miltenyi Biotec) according to the manufacturer’s instructions. Monocytes were isolated from the BM using the EasySep Mouse Monocyte Isolation Kit (Stemcell Technologies) according to the manufacturer’s instructions, and their purity was assessed by FC.   80  3.2.5 Induction of lung inflammation and adoptive monocyte transfer Mice were treated with 25 µg of Escherichia coli 0111:B4 LPS (Sigma-Aldrich) in 50 µl PBS via i.t. instillation. Four hours later, 0.5-1 x 106 GFP+ BM monocytes resuspended in 200 µl PBS were injected i.v. into C57BL/6J host mice. Alternatively, 1-2 x 106 CD45.2+ CD44+/+ and CD44-/- monocytes (1:1 ratio) were injected i.v. into the LPS treated CD45.1 host. The lung and BAL were harvested at indicated times following LPS for FC analysis. The weight of LPS treated mice were recorded daily for 7 days, and then on days 14 and 30.   3.2.6 Competitive BM reconstitution  8 x 106 CD45.1+ CD45.2+ CD44+/+ and CD45.2+ CD44-/- BM cells (1:1 ratio) were injected i.v. into lethally irradiated (24 h after gamma radiation 6.5 Gy, twice, 4 h apart) CD45.1+ host mice. The perfused lung and BAL were harvested after 7 weeks for analysis.  3.2.7 Intratracheal instillation of AMФ and PMФ  GFP+ AMФ (3 x 105) in 50 µl PBS were instilled i.t. into CD45.2+ mice. BAL was collected at the indicated time points for analysis. Alternatively, 3 x 105 CD45.2+ CD44+/+ and CD44-/- AMФ (1:1 ratio) in 50 µl PBS were instilled i.t. into CD45.1+ mice. Some mice were challenged with 25 µg of LPS 24 h following the instillation of donor AMФ. The BAL was collected 24 h or 8 days after AMФ instillation for analysis. PMФ were isolated from GFP+ mice as described above. CD45.1+ or 81  CD45.1+ CD45.2+ mice received 8 x 105 GFP+ PMФ in 50 µl PBS by i.t. instillation. The BAL was collected 2 weeks later for analysis.  3.2.8 Intratracheal instillation of HA blocking CD44 antibody  Four separate doses of 50 μg of the HA blocking CD44 mAb, KM81 or an isotype matched rat IgG2a mAb were instilled i.t. into isoflurane anaesthetized CD45.2+ mice on days 0, 2, 4, and 6. The BAL was collected on day 7 for FC analysis.   3.2.9 AMФ Ki67 labeling  AMФ were treated with 2.4G2 to block Fc receptors, labeled with CD44 antibody, and incubated with LIVE/DEAD Fixable Aqua Dead Cell Stain for 30 min. Cells were then fixed and permeabilized, incubated with 2.4G2, and labeled with fluorescent Ki67 antibody for FC analysis.    3.2.10 Generation of CSF-1 derived BMDM  BM was isolated from murine femurs and tibia, and CSF-1 derived BMDM were generated as previously described (Dong et al., 2016).   3.2.11 Macrophage stimulation with CSF-2 and ROZ CSF-1 BMDM (2 x 105) collected on day 5 and cultured in 200 μl DMEM (Invitrogen) with 10% FBS in non-tissue culture treated 96-well plates in the presence or absence of 4% CSF-2 containing supernatant for 48 h. Cells were then 82  analyzed by FC. Purified F4/80+ PMФ (2 x 105) were cultured in 200 μl complete RPMI 1640 media (with 10% FBS, 20 mM HEPES, 1x non-essential amino acid, 55 μM 2-mercaptoethanol, 1 mM sodium pyruvate, 2 mM L-glutamine, 50 U/ml Penicillin/Streptomycin, all from Invitrogen) in non-tissue culture treated 96-well plates in the presence or absence of 20 ng/ml of mouse recombinant CSF-2, 1 μM ROZ, or 20 ng/ml of CSF-2 and 1 μM ROZ for 7 days, with a media change at days 3 and 6. Cells were analyzed by FC on day 7.   3.2.12 Measuring AMФ cell death and surface HA labeling AMФ were labeled with DAPI and fluorescent Annexin V in 10 mM HEPES (pH 7.4), 140 mM NaCl, 2.5 mM CaCl2 for 20 min for FC analysis. Alternatively, 1 x 105 cells were plated in 96-well non-tissue-culture treated plates in complete RPMI 1640 media and incubated at 37°C. After 24 h, the cells were collected with Versene and analyzed for Annexin V and DAPI. Cell surface HA for FC was determined by biotinylated HABP (Millipore Sigma) and streptavidin-PE-Cy7 (Affymetrix eBioscience) labeling. AMФ were incubated in 100 µl PBS containing 20 U of Bov HA’se (Sigma-Aldrich) or 100 µl of KM81 hybridoma (ATCC TIB-241) TCS containing the CD44 blocking antibody KM81 for 1 h at 37°C to remove surface HA or block CD44-HA interactions, respectively.   3.2.13 Confocal microscopy Lungs were harvested from isoflurane anesthetized mice and fixed by immersing into ice cold 4% PFA for 2 h. Fixed tissue was embedded in NEG-50 83  freezing matrix (ThermoFisher) and snap frozen in dry ice cooled isopentane. 12 µm cryosections were cut and thaw mounted onto Superfrost Plus microscope slides (ThermoFisher). Tissue sections were fixed again with acetone, blocked with 10% goat serum, and labeled with CD11c and SiglecF antibodies followed by goat-anti hamster Alexa 647 and goat anti-rat Alexa 488 (ThermoFisher) respectively and biotinylated HABP followed by streptavidin Alexa 568 (ThermoFisher). Images were acquired using the 20x dry objective of a FV1000 laser scanning confocal microscope (Olympus) equipped with 405, 488, 543, and 633nm laser lines.   3.2.14 Data analysis  FC was analysed using FlowJo VX (Treestar). Flow cytometry plots shown were first gated by size, singlets, and live/dead stain.  Graphs were generated using GraphPad Prism 6. Data shown are the average ± standard deviation (SD). Significance was determined by the Student’s two-tailed, unpaired or paired t test with *p < 0.05, **p < 0.01, ***p < 0.001. Confocal microscopy was analyzed using the Fiji distribution package of ImageJ 1.51 (NIH). The mean pixel intensity of HABP for CD11c+ Siglec F+ CD44+/+ or CD44-/- AMФ was determined, data shown are individual cell measurements and the average ± SD, statistical significance was tested by non-parametric unpaired Mann-Whitney test with ***p < 0.001.  84  3.3 Results 3.3.1 CD44-/- mice have reduced numbers of AMФ and CD44-/- AMФ are at a competitive disadvantage under both homeostatic and inflammatory conditions  Figure 3.1: CD44 deficiency reduces AMФ numbers a) Representative flow cytometry plots and graph of 40 mice comparing the phenotype and number respectively, of AMФ isolated from the BAL of CD44+/+ and CD44-/- mice. b) Representative flow cytometry plots and graph of 16 mice showing the phenotype and number respectively, of AMФ from the lungs of CD44+/+ and CD44-/- mice. c) Flow cytometric analysis of 3 x 105 CD45.2+ donor CD44+/+ and CD44-/- AMФ mixed at a 1:1 ratio prior to i.t. instillation into CD45.1+CD45.2- host 85  mice. d) Proportion of donor CD44+/+ and CD44-/- AMФ in the BAL 24 h after instillation. e) Proportion of donor CD44+/+ and CD44-/- AMФ in the BAL after 8 days. f) Proportion of donor CD44+/+ and CD44-/- AMФ in the BAL after 8 days, treated with LPS via i.t. instillation 24 h after AMФ transfer. g) Percent of donor CD44+/+ and CD44-/- AMФ in the BAL 24 h after instillation. h) Percent of donor CD44+/+ and CD44-/- AMФ after 7 days with or without LPS treatment. i) Total number of host CD45.2- CD11c+ cells in the BAL after 7 days of with or without LPS treatment. j) GFP+ CD44+ AMФ were instilled into the lungs for 24 h (day 0) then LPS was instilled into the lungs and the number GFP+ AMФ counted at day 1 and day 7 post LPS. Relative cell numbers compared to day 0 are shown. In c-j, data show an average of two experiments ± SD, each with three to five mice. Significance indicated as * p< 0.05, ** p< 0.01, *** p < 0.001, non-paired (a-b, i-j) and paired Student’s t-test (g and h).  To determine the significance of constitutive HA binding and CD44 expression by AMФ in the alveolar space, we compared bone fide fetal monocyte-derived AMФ from CD44+/+ and CD44-/- mice. The number of AMФ isolated from the BAL of CD44-/- mice was significantly lower compared to CD44+/+ mice (Figure 3.1a). A reduction was also observed in AMФ isolated from the lung where equivalent numbers of lung cells were isolated from CD44+/+ and CD44-/- mice, ruling out differences in location or total cell numbers in the lungs between the mice (Figure 3.1b). No gross differences were observed in the expression of Siglec F, CD11c or CD11b between the CD44+/+ and CD44-/- AMФ (Figures 1a and b). This suggested an intrinsic defect in the CD44-/- AMФ, although the alveolar environment can also have a strong influence on AM survival and maturation (Guilliams et al., 2013; Guth et al., 2009; Schneider et al., 2014b; Shibata et al., 2001b). To distinguish between these possibilities, AMФ were isolated from CD45.2+ CD44+/+ and CD44-/- mice and instilled into the alveolar space of CD44+/+ CD45.1+ mice at a 1:1 ratio (Figure 3.1c). After 24 h, there was still a 1:1 ratio of donor CD44+/+ and CD44-/- AMФ in the BAL demonstrating no advantage of CD44 on the initial engraftment (Figure 3.1d and g). 86  However, by day 8 post instillation, there was greater than a 2:1 ratio of donor CD44+/+ AMФ over CD44-/- (Figure 3.1e and h), indicating a competitive advantage of CD44+/+ AMФ. Since the reduced percentage of CD44-/- AMФ was observed in a CD44 sufficient lung environment, this supports an intrinsic defect in these cells. To determine if instillation of the donor AMФ into an inflammatory lung environment would affect the outcome, we instilled LPS 24 h after the instillation of the 1:1 ratio of donor AMФ. Analysis of the cells at day 7 post LPS (day 8 post AMФ instillation) revealed a similar 2:1 ratio (Figure 3.1f and h), indicating no further advantage. It was noted that there were more CD45.2- CD11c+ host cells at day 7 after LPS treatment compared to untreated (Figure 3.1i). In a different experiment, a comparison of the number of instilled donor GFP+ CD44+/+ AMФ before and after LPS induced inflammation showed a 3-fold increase of AMФ on day 7 post LPS relative to day 0 and day 1 (Figure 3.1j), indicating that AMФ had proliferated in response to LPS induced inflammation. Thus CD44-/- AMФ have an intrinsic defect that compromised their ability to generate normal numbers of AMФ in the alveolar space at homeostasis and decreased their ability to compete with CD44+/+ AMФ under both homeostatic and inflammatory conditions.   87  3.3.2 CD44 provides an advantage to monocyte-derived AMФ   Figure 3.2: CD44 deficiency impairs AMФ repopulation following lethal irradiation a) Schematic showing BM reconstitution of lethally irradiated CD45.1+ host mice with 8 million BM cells from CD45.1+ CD45.2+ CD44+/+ and CD45.1- CD45.2+ CD44-/- mice at 1:1 ratio, analyzed after 7 weeks. b) Representative flow cytometry plots showing the identification and proportions of CD44+/+ and CD44-/- neutrophils, eosinophils, Ly6Chigh cells, AMФ in the lung, and AMФ in the BAL 7 weeks post reconstitution. c) Graph showing the proportion of CD44+/+ and CD44-/- cells in the different myeloid populations in the lung and BAL after reconstitution. Data show an average of two experiments ± SD, each with three to five mice. Significance indicated as * p<0.05, ** p< 0.01, *** p < 0.001, paired Student’s t-test.  88  To determine if this CD44 advantage also extends to BM monocyte-derived AMФ, we performed a competitive BM reconstitution using CD44-/- and CD44+/+ BM cells into a lethally irradiated CD44 sufficient host. Unlike at homeostasis, this creates an empty niche in the alveolar space, allowing full reconstitution of AMФ from BM monocytes (Lavin et al., 2014). The donor BM cells were distinguishable by CD44 expression and by CD45.1 and CD45.2 allogeneic markers: CD44-/- BM was CD45.2+ and CD44+/+ BM was CD45.1+ CD45.2+. BM cells were isolated, mixed at a 1:1 ratio and 8 million cells were injected intravenously (i.v.) into lethally irradiated CD45.1+ mice (Figure 3.2a). Seven weeks elapsed to allow for BM reconstitution and then the percentage chimerism was determined in the lung. Monocytes (Ly6Chigh CD11c-), neutrophils (Ly6G+), and eosinophils (Ly6Clow SSChigh) all showed equal chimerism of approximately 1:1 (Figure 3.2b and c). However, AMФ (CD11c+ FSChigh) showed a 3:1 ratio of CD44+/+:CD44-/- cells indicating a significant advantage for CD44+/+ AMФ in the lung (Figure 3.2b and c). Isolation of CD11c+ AMФ from the BAL, where they constitute the majority of cells, further demonstrated a 3:1 ratio and a distinct advantage for CD44+/+ AMФ (Figure 3.2b and c).  Although CD44 is expressed on neutrophils, eosinophils, and Ly6Chigh monocytes, it does not confer a competitive advantage after BM reconstitution in these cells, whereas it does for AMФ.   89  3.3.3 LPS induced lung inflammation causes a transient loss of AMФ and differentiated monocytes gain HA binding   Figure 3.3: Characterization of inflammatory cells in the BAL during LPS induced lung inflammation  a) Change in body weight of mice after treatment with 25 µg of LPS via i.t. instillation. b) Number of total leukocytes in the BAL during LPS induced lung 90  inflammation. c) Representative flow cytometry plots showing cell populations and their phenotype in the BAL at steady state (day 0) and during LPS induced lung inflammation. Histograms show HA binding of the different populations. d) Number of AMФ, gated as in c, in the BAL at steady state and throughout the response to LPS. e) Number of infiltrated monocytes, gated as in c, in the BAL in response to LPS. f) Number of neutrophils, gated as in c, in the BAL during LPS induced inflammation. Flow cytometry plots were first gated by size, singlets, and live/dead stain. Data show an average of two experiments ± SD, each with three to five mice. Significance indicated as * p< 0.05, ** p< 0.01, *** p < 0.001, non-paired Student’s t-test.  We next wanted to investigate if the CD44 advantage of monocyte-derived AMФ occurred in a more physiological setting. Furthermore, it was of interest to determine to what extent self-renewal or monocyte recruitment contributes to the AMФ population after inflammation. In order to study this, we first established a suitable acute lung inflammation mouse model using LPS. The course of LPS induced inflammation was monitored over time by weight loss (Figure 3.3a) and the total number of leukocytes present in the BAL (Figure 3.3b), which indicated the peak of inflammation occurred around day 3 and was resolved by day 14. Flow cytometry of the BAL cells, identifying the AMФ as well as incoming monocytes and neutrophils, revealed a significant depletion of CD11c+ SiglecF+ AMФ occurring early in the response at day 1, together with an influx of CD11b+ Ly6Ghigh neutrophils, followed by an influx of CD11b+ Ly6Chigh monocytes that peaked at day 3 after stimulation (Figure 3.3c-f). Despite their initial loss, AMФ quickly repopulated and expanded beyond pre-inflammatory values by day 7 (Figure 3.3d). Also by day 7, CD11b+ Ly6Chigh monocytes were replaced with CD11b+ Ly6C- CD11c+/- cells (Figure 3.3c). Most of these became CD11c+, gained Siglec F and lost CD11b expression by day 14, suggesting further differentiation. Interestingly, as the 91  monocytes differentiated, they also gained HA binding, another hallmark of AMФ. By day 30, these cells were either absent or indistinguishable from resident AMФ (CD11c+, Siglec F+, HA binding) (Figure 3,3c and e). Overall, this raises the possibility that the transiently depleted AMФ are capable of rapid self-renewal, whereas the recruited monocytes may undergo a slower differentiation process to eventually become AMФ one month after recruitment.  92  3.3.4 GFP+ monocytes mature into AMФ 30 days after the inflammatory stimulus  Figure 3.4: Adoptively transferred BM monocytes give rise to AMФ following LPS induced lung inflammation 93  a) Representative flow cytometry plots showing the expression levels of Ly6C, CD11b, Ly6G, CD11c, Siglec F, and FL-HA binding by AMФ at steady state. b) Representative flow cytometry plots showing the phenotype of GFP+ monocytes isolated from the BM and the phenotypic changes in the BAL after they were adoptively transferred i.v. into mice treated with 25 g LPS i.t. 4 h earlier. c) Number of GFP+ donor cells in the host BAL and lung throughout the LPS response. d) Representative flow cytometry plots and graph comparing Ki67 expression in GFP+ donor cells and host AMФ. e) Histograms representative of 12 mice from two experiments comparing the phenotype of GFP+ donor cells and host AMФ 45 days after LPS treatment. Data show an average from two experiments ± SD, each with three to five mice. Significance indicated as * p< 0.05, ** p< 0.01, *** p < 0.001, non-paired student’s t-test.  To determine whether the recruited monocytes disappear over time or differentiate into AMФ, we followed the fate of adoptively transferred GFP+ monocytes. Steady state AMФ were CD11b- Ly6C- Ly6G- CD11c+ Siglec F+ and HA binding (Figure 3.4a). GFP+ monocytes (CD11b+ Ly6Chigh Ly6G- CD11c- Siglec F- and non-HA binding) were isolated from the BM of GFP+ mice with ~95% purity (Figure 3.4b) and 0.5-1 x 106 monocytes were injected i.v. 4 h after i.t. of LPS. Perfused lung tissue and the BAL were harvested over time to analyze the fate of the recruited GFP+ monocytes. GFP+ cells were recruited to the BAL of inflamed mice on day 1 and maintained their monocyte phenotype: CD11b+ Ly6Chigh Ly6G- CD11c- Siglec F- and non-HA binding (Figure 3.4b). By day 3, monocytes were losing Ly6C expression and by day 7 this had largely disappeared. Cells started to gain CD11c at day 7 and by day 14 were mostly CD11c+ and CD11b-, and many had gained the ability to bind HA and express Siglec F (Figure 3.4b). By day 30, the GFP+ cells were indistinguishable from the characteristic phenotype of AMФ: CD11b- CD11c+ Siglec F+ and HA binding. These phenotypic changes in the GFP monocytes mirrored those of the host monocytes observed in Figure 3. In addition, the number of GFP+ cells in the BAL peaked at day 3, as did host monocytes, then the number 94  of GFP+ cells in the BAL decreased to a minimum at day 14, and then increased at day 30 (Figure 3.4c), suggesting that either the differentiated monocytes proliferated or entered the alveolar space from the lung tissue. The reduction in GFP+ cells in the lung between day 14 and 30 (Figure 3.4c) supports the latter possibility at day 30; however, detection of a small percentage of Ki67+ GFP+ cells at day 45 also supports cell proliferation, which occurred at a comparable rate to the host AMФ (Figure 3.4d). Furthermore, these GFP+ cells at day 45 were phenotypically indistinguishable from host AMФ, by the expression levels of CD11c, Siglec F, CD11b, HA binding, SIRPα, CD206, and F4/80 (Figure 3.4e).   95  3.3.5 CD44 exerts an advantage in monocyte-derived AMФ after inflammation   Figure 3.5: CD44 deficiency impairs the number of monocyte-derived AMФ after LPS induced lung inflammation.  96  a) Schematic showing the i.v. transfer of 1-2 x 106 monocytes isolated from the BM of CD45.2+ CD44+/+ and CD44-/- mice at 1:1 ratio into host CD45.1+ mice that were treated i.t. with 25 µg of LPS 4 h earlier to induce acute lung inflammation. The BAL was analyzed at day 14 and 30 post LPS treatment. b) Representative flow cytometry plots showing the purity and proportion of CD44+/+ and CD44-/- monocytes used for adoptive transfer. c) Representative flow cytometry plots showing the proportion of CD44+/+ and CD44-/- donor cells in the BAL at day 14 and comparing their phenotype. d) Representative flow cytometry plots showing the proportion of CD44+/+ and CD44-/- donor cells in the BAL at day 30 and comparing their phenotype. e) & f) Graphs comparing the percent of CD44+/+ and CD44-/- donor cells in the BAL at day 14 and 30, respectively. Data show an average of two experiments ± SD, each with three to five mice. Significance indicated as * p< 0.05, ** p< 0.01, *** p < 0.001, paired Student’s t-test.  Having identified that AMФ can repopulate a few days after depletion by an inflammatory insult such as LPS, and that recruited monocytes can differentiate into a phenotypically identical AMФ by day 30, we can now examine the effect of CD44 on AMФ repopulation by self-renewal and from monocytes. To determine when CD44 influenced monocyte-derived AMФ repopulation after inflammation, we repeated the above experiment using a 1:1 ratio of CD45.2+ CD44+/+ and CD44-/- monocytes purified from the BM of the respective mice injected i.v. into CD45.1+ mice, 4 h after i.t. LPS injection (Figure 3.5a-b). Analysis of the BAL cells 14 days after induction of inflammation revealed the percent and phenotype of CD44+/+ or CD44-/- cells within the CD45.2+ donor cells population were similar (Figure 3.5c and e), indicating no selective advantage of CD44 at this stage of differentiation in the BAL when differentiating monocytes begin to gain CD11c, Siglec F, and HA binding. However, at day 30 when monocytes fully adopted the AMФ phenotype, there was a similar 3:1 advantage of CD44+/+ cells over CD44-/- cells (Figure 3.5d and f). This demonstrated that CD44 did not affect the recruitment of monocytes. 97  Instead, the competitive CD44 advantage materializes by day 30 when monocytes have differentiated into AMФ and gained HA binding.   3.3.6 The alveolar environment induces HA binding  Figure 3.6: The alveolar environment and CSF-2 promote HA binding by macrophages a) Representative flow cytometry plot showing the expression of MHCII and F4/80 by purified PMФ. b) Representative flow cytometry plots showing the expression of 98  CD11c, CD11b, and HA-A647 binding by GFP+ PMФ before or c) 2 weeks after adoptive transfer i.t. into the BAL. d) Representative histograms of HA-A647 binding and CD11c expression of CSF-1 derived BMDM treated with (black line) or without (shaded) CSF-2 for 48 h in vitro, and graphs of their relative mean fluorescent intensity (MFI). e) Representative histogram and graph comparing FL-HA binding by PMФ treated for 7 days in vitro with CSF-2 alone, ROZ alone, or CSF-2 and ROZ. a and b are representative plots of six mice examined over two experiments. Graphs show an average of two experiments ± SD, each with three to five mice; Significance indicated as * p<0.05, ** p< 0.01, *** p < 0.001, paired Student’s t-test.  The alveolar environment and CSF-2 are important for the development and maturation of fetal monocyte-derived AMФ (Guilliams et al., 2013; Schneider et al., 2014b; Shibata et al., 2001b). To determine if the alveolar environment can induce HA binding in macrophages, we purified F4/80+ macrophages from the peritoneal lavage. The majority of the purified cells were F4/80high MHCIIlow peritoneal macrophages (PMФ) (Figure 3.6a), consistent with resident PMФ described in (Bain et al., 2016) and were phenotypically distinct from AMФ, being CD11c- CD11b+ and non-HA binding (Figure 3.6b). Purified GFP+ F4/80+ PMФ were instilled into the lungs of wild-type, non-irradiated mice and isolated from the BAL 2 weeks later where they had gained similar levels of CD11c and HA binding as the resident AMФ, but remained CD11b+ (Figure 3.6c). This demonstrated that the alveolar environment induces HA binding. CSF-2 may induce CD44 to bind HA in the alveolar space, as CSF-2, but not CSF-1 derived BM derived macrophages (BMDM) bound HA (Poon et al., 2015; Ruffell et al., 2011). Furthermore, CSF-2 derived BMDM have several phenotypic and functional similarities to AMФ (Poon et al., 2015). To investigate this, CSF-1 derived BMDM were cultured with CSF-2 for 48 h and this partially upregulated their HA binding ability and CD11c expression (Figure 3.6d). To test if CSF-2 also induced PMФ to bind HA, purified PMФ were cultured 99  with CSF-2 for 7 days in vitro. This did not induce HA binding in PMФ, neither did the PPARγ ROZ (Camp et al., 2000), however, together they significantly induced HA binding in the PMФ (Figure 3.6e).   100  3.3.7 CD44 and HA binding promote AMФ survival  Figure 3.7: CD44-/- AMФ have increased cell death and do not bind HA on the cell surface a) Representative flow cytometry plots and graph comparing Ki67 expression on ex vivo CD44+/+ and CD44-/- AMФ. b) Representative flow cytometry plots comparing ex vivo CD44+/+ and CD44-/- AMФ survival by DAPI and Annexin V labeling. c) Percent 101  of CD44+/+ and CD44-/- AMФ that are alive, apoptotic (Annexin V+) or dead (DAPI+ or Annexin V+, DAPI+) from ex vivo cells and percent live cells after 24 h in culture. d) Confocal microscopy of CD44+/+ and CD44-/- lung tissue labeled with DAPI (blue), CD11c (red), Siglec F (white), and HABP (green); merged (yellow) image is of CD11c and HABP. Image is representative from six mice over two experiments. e) Comparing mean pixel intensity of labeled HABP from confocal microscopy of CD44+/+ and CD44-/- AMФ in the lung tissue. Individual cell measurements were from 3 mice, 2 fields each, and the average ± SD with *** p < 0.001, non-parametric unpaired Mann-Whitney test. f) Flow cytometry of CD44 and the binding of biotinylated HABP to the surface of ex vivo CD44+/+ and CD44-/- AMФ detected by streptavidin-PE-Cy7 and the graph showing the average MFI of HABP binding from 6 mice. Graphs in a, c and f show the average from two experiments ± SD, each with cells from three to five mice. Significance indicated as * p< 0.05, ** p< 0.01, *** p < 0.001 non-paired Student’s t-test.    To determine how CD44 was providing an advantage for AMФ, we isolated CD44-/- and CD44+/+ AMФ from the BAL, and labeled these ex vivo AMФ with Ki67 antibody, a marker of cell proliferation. The percentage of Ki67+ AMФ was not significantly different (Figure 3.7a), suggesting no difference in cell cycling or proliferation rate at the steady state.  However, when ex vivo AMФ were labeled with DAPI and Annexin V, CD44-/- AMФ showed significantly more apoptotic and dead cells than CD44+/+ AMФ (Figure 3.7b-c). Less viable ex vivo CD44-/- AMФ were also observed after equal numbers were cultured for 24 h (Figure 3.7c).  This showed that CD44 was providing a survival advantage for AMФ, possibly by providing a survival signal to AMФ. Given the distinct constitutive HA binding ability of CD44 on AMФ, we investigated if CD44 was binding to HA in the alveolar space. HA was detected by HA binding protein (HABP), which revealed intense staining around the bronchioles and identified CD11c+ Siglec F+ AMФ as the major cells in the BAL that bound HA in vivo. In contrast, CD44-/- AMФ were poorly labeled by HABP, indicating significantly less surface HA (Figure 3.7d-e). This suggested that CD44+/+ AMФ, but not CD44-/- AMФ, were decorated with a HA coat. Further analysis by flow cytometry 102  on ex vivo AMФ from the BAL confirmed that CD44+/+ AMФ were decorated with surface HA to a much greater extent than CD44-/- AMФ (Figure 3.7f), suggesting the cell surface HA was bound to CD44.   Figure 3.8: HA binding to CD44 promotes AMФ survival a-b) Histogram and graph showing the MFI of HABP labeling of surface HA on ex vivo AMФ before and after Bov HA’se treatment. c) Representative flow cytometry plots of CD44+/+ or CD44-/- AMФ labeled with DAPI and Annexin V after PBS, Bov HA’se or CD44 mAb (KM81) treatment for 1 h, and d) Percent live CD44+/+ or CD44-/- AMФ (Annexin V- DAPI-) after treatment.  e) Representative flow cytometry plots of BAL cells from mice after treatment with isotype matched or CD44 mAb delivered i.t. on days 0, 2, 4, 6, then isolated on day 7 and labeled with CD44, HA-A647, DAPI, and Annexin V. f) Percent AMФ in the BAL that are alive, apoptotic (Annexin V+) or dead (DAPI+ or Annexin V+) after 7 days of isotype or CD44 mAb antibody treatment. g) Number of live AMФ in the BAL after 7 days of isotype or CD44 mAb antibody treatment. Data show an average of two experiments ± SD, each with three to five mice. Significance indicated as * p< 0.05, ** p< 0.01, *** p < 0.001, paired (b and d) and non-paired (f-g) Student’s t-test.    Incubation of CD44+/+ AMФ with Bov HA’se significantly reduced HABP labeling indicating HABP binding was specific for surface HA (Figure 3.8a-b). To determine the significance of this HA binding and its possible role in AMФ survival, ex vivo CD44+/+ and CD44-/- AMФ were incubated in PBS, with or without Bov HA’se 103  or the HA blocking CD44 monoclonal antibody (mAb), KM81, for 1 h, then labeled with DAPI and Annexin V. Both HA removal and KM81 addition decreased the viability of CD44+/+ AMФ to the levels observed with the CD44-/- AMФ, and these treatments had no effect on the CD44-/- AMФ (Figure 3.8c-d), indicating that the interaction of HA on AMФ promoted their survival. To determine if this interaction also had similar consequences in vivo, KM81 was instilled into the lung on days 0, 2, 4, and 6 to disrupt the interaction. On day 7, AMФ were isolated from the BAL, and the numbers of live, apoptotic, and dead cells were determined.  KM81 treated mice had significantly more apoptotic and dead cells, and less live AMФ cells than mice treated with an isotype-matched antibody (Figure 3.8e-g), demonstrating that the interaction of CD44 with HA on AMФ in the alveolar space is required for their optimal survival and the maintenance of normal AMФ numbers at homeostasis.  3.4 Discussion AMФ develop from fetal monocytes and do not depend on adult monocytes for self-maintenance at homeostasis (Guilliams et al., 2013; Hashimoto et al., 2013). Here, we show that the self-renewal of AMФ contributes to the rapid AMФ repopulation 7 days after LPS induced inflammation and AMФ depletion. We also show that, in contrast to steady state AMФ maintenance, adult monocytes contribute to the AMФ population after inflammation, albeit to a lesser extent and over a slower time frame of approximately 30 days. The relative contribution from AMФ and monocytes may depend on the extent of lung damage and inflammation, which in turn, may affect the extent of AMФ depletion. This initial depletion of AMФ may alter 104  the alveolar environment and expose a niche to allow monocyte engraftment and their subsequent differentiation into AMФ. While the transfer of GFP+ monocytes provided evidence for their differentiation into cells with an AMФ phenotype (CD11c+, CD11b-, Siglec F+, HA binding, SIRP+, CD206+, F4/80low) and similar proliferative capacity 45 days after lung inflammation, further transcriptomic and functional analysis is necessary before we can conclude that the monocytes have fully differentiated into bone fide AMФ.  Recent work shows monocyte-derived cells present in the lung 10 months after bleomycin induced injury that express equal levels of Siglec F and have a very similar transcriptional signature to tissue resident AMФ (Misharin et al., 2017).   This report shows that the CD44 expressed on both fetal and BM derived AMФ is important for their survival. When CD44+/+ and CD44-/- AMФ were instilled into the lung, CD44+/+ AMФ outcompeted CD44-/- AMФ under both non-inflammatory and inflammatory conditions. In addition, BM derived CD44+/+ AMФ were more prevalent than CD44-/- AMФ after competitive BM reconstitution. This was a selective effect of CD44 on AMФ, as donor derived monocytes, neutrophils and eosinophils all had equal proportions of CD44+/+ and CD44-/- cells after BM reconstitution. Although all these cells express CD44, only AMФ bind HA constitutively. Furthermore, when equal numbers of GFP+ CD44+/+ and CD44-/- BM derived monocytes were adoptively transferred into the blood after LPS induced lung inflammation, equal numbers of monocyte-derived cells were present at day 14. It was only by day 30, when they had gained the full HA binding AMФ phenotype, that the CD44+/+ AMФ had an 105  advantage. Together, this shows that CD44 exerts its competitive advantage on AMФ, not monocytes, correlating with its ability to bind HA.  Previous studies show the alveolar environment induces phenotypic and transcriptional changes to macrophages from other tissues to become more like AMФ (Lavin et al., 2014; van de Laar et al., 2016). HA binding, which is a key characteristic of AMФ, was also induced on PMФ by the alveolar environment. In vitro, HA binding by PMФ required both CSF-2 and the PPARγ agonist, ROZ, suggesting that these two signals may also be responsible for the induction of HA binding by macrophages in the alveolar space. CSF-2 induces the expression of the nuclear receptor PPARγ (Schneider et al., 2014b), and both are required for the differentiation of fetal monocytes into AMФ in vivo (Guilliams et al., 2013; Shibata et al., 2001b). The induction of constitutive HA binding by the alveolar environment is a striking feature, as monocytes and PMФ do not normally bind detectable levels of exogenously added HA, unless they are exposed to inflammatory stimuli (Brown et al., 2001; Levesque and Haynes, 1997; Ruffell et al., 2011). However, AMФ are not unique in their constitutive ability to engage HA, as splenic F4/80+ macrophages can also bind HA (Dong et al., 2016), and resident PMФ are recruited to the site of liver injury in a CD44 and HA dependent manner(Wang and Kubes, 2016). Together, these highlight the importance of environmental imprinting on HA binding and macrophage plasticity. Results from in vivo competition in wild type mice show that CD44 deficient AMФ have an intrinsic defect, which may be due to their inability to bind HA. The presence of cell surface HA is a major difference between CD44+/+ and CD44-/- AMФ. 106  CD44+/+, but not CD44-/-, AMФ are decorated with HA in vivo. This was somewhat unexpected as others have shown that alveolar macrophages readily take up and degrade HA in vitro and in vivo during lung development (Culty et al., 1994; Underhill et al., 1993a). Nevertheless, surface bound HA was functionally important as prevention of HA binding by a CD44 blocking antibody both ex vivo and in vivo or by enzymatic removal of HA on ex vivo cells reduced the survival of CD44+/+AMФ. Thus, we conclude the interaction of CD44 with HA provides a survival advantage for AMФ. Supporting this, CD44-/- mice had decreased numbers of AMФ in the alveolar space, and CD44-/- AMФ were more susceptible to cell death. Together, this demonstrates the importance of the HA coat and its interaction with CD44 in AMФ survival. In the alveolar space, AMФ reside between the surfactant and the alveolar epithelial cells (AEC) (Hussell and Bell, 2014). Thus, one possible source of HA in the alveolar space is AEC. Type II AEC (AEC2) express hyaluronan synthase 2 and display HA on their surface (Liang et al., 2016) that may promote interactions with AMФ. Other molecular interactions already exist between AEC and AMФ that provide inhibitory signals to prevent AMФ activation (Hussell and Bell, 2014). AMФ binding to AEC may also facilitate AMФ proximity to AEC derived growth and survival factors such as CSF-2, to help sustain lung homeostasis (Huffman et al., 1996). Alternatively, HA may be scavenged from these cells and from the alveolar space, or AMФ themselves may synthesize HA which is then bound by surface CD44. Extracellular HA binding by CD44 on AMФ could be integral to forming a HA-rich glycocalyx which may promote cell survival by protecting against environmental 107  stress, for example by protecting against reactive oxygen species (Presti and Scott, 1994; Sato et al., 1988), or against sheer stress (Gouverneur et al., 2006). Alternatively, HA binding to CD44 on AMФ may provide pro-survival signals. High expression of CD44 has been linked with promoting effector T cell survival and protection from Fas mediated apoptosis in Th1 cells (Baaten et al., 2010) and antibody induced CD44 ligation improves survival and resistance of cancer cells to drug induced apoptosis (Allouche et al., 2000; Bates et al., 2001; Herishanu et al., 2011). However, it has been difficult to identify HA mediated signaling events in cells (Lee-Sayer et al., 2015).  AEC2 have stem cell-like properties in the adult lung and contribute to tissue repair. In a mouse model of bleomycin induced lung damage, AEC2 cells lacking HAS2 had lower levels of surface HA, showed increased apoptosis, and a reduced ability to proliferate and form colonies, implicating a role for surface HA in the survival and proliferation of AEC2 (Liang et al., 2016). Furthermore, AEC2 from IPF patients also showed reduced surface HA and were less efficient in generating colonies compared to AEC2 from healthy individuals (Liang et al., 2016). Similarly, CD44 and its interactions with HA have been implicated in promoting the self-renewal and survival of cancer initiating cells (Chanmee et al., 2015). Here, we add to this accumulating evidence for a role of HA binding by CD44 in promoting the survival of cells with stem cell-like renewal capacity by showing HA-CD44 interactions support the survival of long-lived, self-renewing AMФ. The specific effect of HA binding by CD44 on AMФ survival highlights both CD44 and HA as possible targets for therapeutic intervention. Given the importance of AMФ in maintaining 108  lung immunosurveillance and homeostasis, strategies that improve the survival or self-renewal potential of these cells, especially in pathological conditions of lung inflammation, may result in therapeutic benefits.  109  Chapter 4: CD44 deficiency disrupts lipid homeostasis in the lung 4.1 Introduction AMФ are fetal-derived tissue resident macrophages (Guilliams et al., 2013) in the alveolar space that require CSF-2, PPARγ, and TGF-β for normal development and maturation (Schneider et al., 2014b; Shibata et al., 2001a; Yu et al., 2017). AMФ maintain lung homeostasis by catabolizing pulmonary surfactants present at the air/liquid interface in the alveolar space (Miles et al., 1988; Wright and Youmans, 1995), and by clearing pathogens, particulate matter, and cell debris (Hussell and Bell, 2014; Iwasaki et al., 2017; Kopf et al., 2014). Pulmonary surfactants are composed of phospholipids mostly in the form of PC species, as well as some cholesterol and surfactant proteins (Vangolde et al., 1988). In CSF-2-/- mice, AMФ fail to develop normally (Shibata et al., 2001b), and are immature and foamy as a result of defective surfactant phospholipid degradation (Yoshida et al., 2001). These defects lead to the accumulation of surfactant lipids and proteins in the alveolar space (Dranoff et al., 1994), and life-threatening PAP (Happle et al., 2014; Suzuki et al., 2014). Lung damage and inflammation can also lead to the accumulation of surfactant lipids as well the oxidation of surfactant lipids. For example, ROS and oxidized phospholipids are generated in the lungs of mice treated with acid aspiration or inactivated influenza virus, which cause inflammation and damage (Imai et al., 2008). Bleomycin induced damage also results in lipid accumulation in the lungs of mice, coinciding with macrophage accumulation of oxidized phospholipids and foamy phenotype, and leading to greater TGF-β1 production and fibrosis (Romero et al., 2015). In contrast, mice lacking a NADPH-oxidase subunit 110  that cannot generate ROS were protected from bleomycin-induced pulmonary fibrosis (Manoury et al., 2005). These data suggest damage from ROS and oxidized surfactant lipids contributes to lung pathology. In humans, signs of oxidative damage is found in the BAL fluid from patients with acute respiratory distress syndrome (Bunnell and Pacht, 1993; Lamb et al., 1999), and lung phospholipid peroxidation is associated with asthma and COPD progression (Kirkham and Rahman, 2006). Furthermore, OxPC is found in AMФ from patients diagnosed with idiopathic interstitial pneumonia (Yoshimi et al., 2005). Together, these observations demonstrate the importance of removing excess surfactant lipids and oxidized lipids from the alveolar space by cells such as AMФ in order to maintain lung homeostasis.  Mature murine AMФ at the steady state express high levels of CD11c and Siglec F (Hussell and Bell, 2014), and constitutively bind HA via CD44 (Culty et al., 1994; Poon et al., 2015). CD44 expressed by AMФ allow them to bind HA on the cell surface, forming a pericellular HA coat that promotes cell survival (Dong et al., 2017). AMФ from CD44-/- mice lack the HA coat, leading to a decrease in their viability and cell numbers (Dong et al., 2017). The physiological consequences of a sub-optimal number of AMФ are not clear, as CD44-/- mice do not exhibit clear pathology in the lung at steady-state. However, CD44 expression is lowered on AMФ isolated from human patients with diffuse panbronchiolitis (Katoh et al., 2001) and COPD (Pons et al., 2005), suggesting the loss of AMФ CD44 expression can be associated with lung inflammation. Additionally, the depletion of AMФ exacerbates pulmonary inflammatory responses (Knapp et al., 2003; Thepen et al., 1989). 111  Together, these studies suggest the defective number of AMФ in the CD44-/- mouse, and their lack of HA binding through CD44 may alter the induction and progression of lung inflammation. Indeed, CD44-/- mice are less able to resolve lung inflammation (Liang et al., 2007; Teder et al., 2002). Since AMФ are critical for pulmonary surfactant lipid clearance (Dranoff et al., 1994; Schneider et al., 2014b; Yoshida et al., 2001), reduced AMФ in CD44-/- mice may also lead to abnormal pulmonary surfactant lipid levels, which can increase their susceptibility to OxPC toxicity and lung inflammation. Here we investigated the effect of reduced numbers of AMФ in CD44-/- mice on their ability to maintain lung homeostasis.  4.2. Material and methods 4.2.1 Mice C57BL/6J (CD45.2+) and B6.SJL-PtprcaPepcb/BoyJ (CD45.1+) mice were from Jackson Laboratory. CD44-/- mice (Schmits et al., 1997), backcrossed with C57BL/6J mice for 9 generations, and all mice were housed and bred at the University of British Columbia (UBC). Mating C57BL/6J and BoyJ mice gave C57BL/6JxBoyJ heterozygotes (CD45.1+, CD45.2+). All animal experiments were conducted with protocols approved by the University Animal Care Committee in accordance with the Canadian Council of Animal Care guidelines for ethical animal research. 4.2.2 Reagents FL-HA was prepared using hyaluronic acid sodium salt from Rooster comb (Sigma-Aldrich), as described previously (de Belder and Wik, 1975). HA’se isolated 112  from Streptomyces hyaluronlyticus and biotinylated HABP were from Millipore. Recombinant mouse CSF-2 (carrier free) was purchased from BioLegend. BODIPY was from Thermofisher. PC was from Sigma-Aldrich. POVPC and NBD-PC were from Avanti Polar Lipids. Fatty acid free bovine serum albumin fraction V (FAF BSA) was from Roche Diagnostics. NBD-PE and H2DCFDA were from Thermofisher. Annexin V was from BD Biosciences.  4.2.3 Flow cytometry (FC) Cells were incubated with 2.4G2 TCS for 20 min to block Fc receptors, washed with FC buffer (PBS, 2% BSA and 2 mM EDTA), labeled with antibodies (Abs) and/or FL-HA for 20 min at 4°C, washed twice in FC buffer, and resuspended in FC buffer containing propidium iodide, DAPI, or Live/Dead Fixable Aqua Dead Cell Stain (Thermofisher) to label non-viable cells. For intracellular labeling, cells were fixed and permeabilized using the eBioscience Intracellular Fixation & Permeabilization Buffer kit, then incubated with intracellular mAb. The following Ab against mouse antigens were used for flow cytometry, cell isolation, or confocal imaging: CD11c (N418), CD11b (M1/70), CD14 (rmC5-3), CD36 (NL07), CD44 (IM7), CD45.1 (A20), CD45.2 (104), CD116 (698423), CD200R (OX110), CD206 (C068C2), MHC II (M5/114.15.2), Ly6c (HK1.4), Ly6G (1A8), Siglec F (E50-2440 or 1RNM44N), SIRPα (P84), MerTK (polyclonal BAF591), PPARγ (81B8), anti-phosphoStat5 (pY694), mouse IgG1 isotype, and anti-OxPC (E06). Antibodies were purchased from Thermofisher, eBioscience, R&D Systems, BD Biosciences, 113  Cell Signal, Avanti Polar Lipids, or AbLab (UBC). FC samples were processed on an LSR II (BD Biosciences).  4.2.4 Cell isolation Mice were euthanized by isoflurane overdose. BAL cells were harvested by the catheterization of the trachea and washing thrice with 1 ml FC buffer. For instances where the BAL supernatant was needed for further analysis, BAL was collected using PBS with 2 mM EDTA, centrifuged and resuspended in RBC lysis buffer (0.84% NH4Cl in 10 mM Tris buffer) for 5 min, and then centrifuged again and resuspended in FC buffer.  4.2.5 Intratracheal instillation of POVPC and CSF-2 Mice were treated with i.t. of 200 µg of POVPC in 50 µl PBS and the BAL was isolated for analysis after 3 days. Mice were treated with i.t. of 2 µg of CSF-2 in 50 µl PBS daily for 7 days, and then the BAL was isolated for analysis.   4.2.6 Competitive BM reconstitution  8 x 106 CD45.1+ CD45.2+ CD44+/+ and CD45.2+ CD44-/- BM cells (1:1 ratio) were injected i.v. into lethally irradiated (24 h after gamma radiation 6.5 Gy, twice, 4 h apart) CD45.1+ host mice. The BAL was harvested after 7 weeks for analysis.  114  4.2.7 Adoptive transfer of AMФ in the alveolar space 2-3 x 105 CD44+/+ or CD44-/- AMФ isolated from the BAL were resuspended in 50 µl PBS and adoptively transferred by i.t. into CD45.1+ CD44+/+ or CD45.2+ CD44-/- mice. BAL was collected after 7 days for analysis.   4.2.8 H2DCFDA labeling  AMФ were washed once with PBS, and resuspended in 5 μM H2DCFDA in PBS for 30 min at 37°C before blocking Fc receptors and antibody labeling. Cell fluorescence was then analyzed by FC. For data analysis, the H2DCFDA MFI from CD44+/+ AMФ were averaged from each experiment, and the fold difference was calculated by dividing the MFI from each CD44+/+ and CD44-/- sample by the average CD44+/+ MFI in each experiment.   4.2.9 NBD-PE and NBD-PC uptake AMФ were washed with PBS and incubated in 10 μM NBD-PE in RPMI for 30 min at 37°C. Alternatively, AMФ were incubated with 10 μM NBD-PC in RPMI containing 2% fatty acid-free BSA for 30 min at 37°C. Cell fluorescence was then analyzed by FC.   4.2.10 AMФ treatment with HA’se and PC in culture CD44+/+ AMФ were cultured with 20 ng/ml of recombinant CSF-2 in RPMI containing 1% BSA or 1 % FAF BSA, in the presence or absence of 2 U/ml HA’se or 115  500 µM of PC for 48 h in a 96-well non-tissue culture treated plate. Cells were then harvested with Versene for analysis.   4.2.11 AMФ survival in vitro with PC and OXPC  Freshly isolated AMФ were incubated with 1% EtOH (vehicle solvent), 50 µM PC, or 50 µM POVPC, a type of OxPC in PBS for 1 h at 37° C, then labeled with Annexin V and DAPI for FC analysis.   4.2.12 Analysis of the BAL fluid  BAL isolated using PBS 2 mM EDTA was centrifuged and the supernatant was collected as BAL fluid. SP-D concentration was determined using the mouse SP-D Quantikine ELISA Kit (R&D Systems). PC concentration was determined using the Phospholipase C kit (Wako Diagnostics). Total protein level was determined using the Pierce BCA Protein Assay Kit (ThermoFisher). HA concentration was determined using the Hyaluronan Quantikine ELISA Kit (R&D Systems). OxPC was determined using the E06 biotinylated antibody according to the manufacturer’s instructions (Avanti Polar Lipids).   4.2.13 Detection of AMФ cell surface HA Cell surface HA for FC was determined by biotinylated HABP and streptavidin-PE-Cy7 (Affymetrix eBioscience) labeling. AMФ were incubated in 100 µl PBS containing 2 U/ml of HA’se 15 min at 4°C to remove surface HA.  116  4.2.14 Labeling of AMФ with BODIPY 493/505, E06 antibody, PPARγ antibody, or anti-phosphoStat5 (pY694) antibody  AMФ isolated with PBS and 2 mM EDTA were fixed with 2% paraformaldehyde (PFA) immediately for 10 min at room temperature, then washed twice with PBS and incubated with 2.5 µM of BODIPY 493/503 in PBS for 30 min at 37° C. PFA fixed AMФ were labeled with the E06 antibody or PPARγ antibody intracellularly using the eBioscience Intracellular Fixation & Permeabilization Buffer Set. Alternatively, PFA fixed AMФ were labeled with the anti-Stat5 (pY694) after permeabilization with 90% methanol for 3 h at 4° C.  4.2.15 Confocal Microscopy  AMФ labeled with BODIPY or E06 were mounted in mounting medium (90% glycerol, 2.5% DABCO, 1x PBS) onto Superfrost+ microscope slides (Fisher). Whole cell image stacks were acquired using a Leica Sp8 White Light Laser Scanning Confocal Microscope. Maximum stack projections were analyzed using imageJ 1.51 (NIH). BODIPY stained lipid droplets were identified using a trainable segmentation function in ImageJ, followed by particle analysis, and droplet masks were applied onto the raw images to measure number, pixel intensity, and droplet size. E06 labeled droplets were counted and data from both BODIPY and E06 droplets were analyzed by Student’s t-test. PPARγ was stained for as described above, followed by DAPI to visualize nuclei Images were acquired using the 63x objective of a Leica Sp5 scanning confocal microscope equipped with 405 and 633nm laser lines. Total 117  nuclear PPARγ was measured within DAPI+ nuclear area and quantified as mean pixel intensity.   4.2.16 Data analysis  FC was analyzed using Flowjo VX (Treestar). Flow cytometry plots shown were first gated by size, singlets, and live/dead stain.  Graphs were generated using Graphpad Prism 6. Data shown are the average ± standard deviation (SD). Significance was determined by a student’s two-tailed, unpaired or paired t-test, or with non-parametric unpaired Mann-Whitney test where * p < 0.05, ** p < 0.01, *** p < 0.001.  118  4.3 Results 4.3.1 CD44-/-  mice with less AMФ have an enhanced alternatively activated phenotype  119  Figure 4.1: CD44-/- mice have abnormal AMФ a) Representative flow cytometry plots showing AMФ as the predominant cells in the BAL of CD44+/+ and CD44-/- mice. Unlabeled cells were used as negative controls. b) Representative flow cytometry showing the CD44 expression, HA binding, and autofluorescence of CD44+/+ and CD44-/- AMФ. c) Comparison of the size of CD44+/+ and CD44-/- AMФ as measured by FSC-A MFI in flow cytometry. d) Comparison of the granularity of CD44+/+ and CD44-/- AMФ as measured by SSC-A MFI in flow cytometry. e) Comparison of the granularity of CD44+/+ and CD44-/- BM derived AMФ 7 weeks after competitive BM reconstitution in irradiated host. f) Comparison of the autofluorescence of CD44+/+ and CD44-/- AMФ. g) Representative flow cytometry plots comparing the MFI of cell surface phenotype of CD44+/+ and CD44-/- AMФ. h) Graphs summarizing the phenotypic difference by MFI between of CD44+/+ and CD44-/- AMФ. Data show an average of two experiments ± SD, each with three to five mice. i) Representative flow cytometry plots comparing the MFI of various cell surface receptors on CD44+/+ and CD44-/- AMФ, with or without the removal of cell surface HA using HA’se. j) Graphs summarizing the MFI of cell surface receptors in CD44+/+ and CD44-/- AMФ, with or without HA’se treatment. Significance indicated as * p< 0.05, ** p< 0.01, *** p < 0.001, paired (e, i-j) and non-paired (c-d, f, h) student’s t-test.    Comparison of CD44+/+ and CD44-/- AMФ showed that both were CD11c+ Siglec F+ and were the major population in the BAL (Figure 4.1a), as reported previously (Dong et al., 2017). AMФ from CD44-/- mice lack CD44, do not bind FL-HA, and were slightly more auto-fluorescent than CD44+ AMФ (Figure 4.1b and f). This was not due to a difference in size as measured by forward scatter (FSC-A) (Figure 4.1c). The granularity (side scatter (SSC-A)) of CD44-/- AMФ was also slightly greater (Figure 4.1d), and this difference was still observed when lethally irradiated CD44+/+ mice were reconstituted with CD44-/- and CD44+/+ BM derived AMФ, (Figure 4.1e), indicating an intrinsic increase in the granularity of CD44-/- AMФ. Increased autofluorescence and granularity suggested that the CD44-/- AMФ may be enriched with lipid droplets and have a more foamy phenotype. Examination of several phenotypic markers of AMФ also revealed significantly higher expression on CD44-/- AMФ (Figure 4.1g-h). In particular, the markers of alternative activation: 120  CD206 and CD200R (Koning et al., 2010), as well as CD36 were notably increased.  CD36 is a receptor for oxidized lipids (Huang et al., 2014a), and its increased expression on CD44-/- AMФ is consistent with the possibility that these AMФ may contain more lipids and therefore take on a more foamy appearance (Huh et al., 1996; Podrez et al., 2002a).    4.3.2 The cellular HA coat does not interfere with the detection of cell surface receptors CD44+/+, but not CD44-/-, AMФ are decorated with an HA coat (Dong et al., 2017). To determine if this pericellular HA coat reduces receptor expression or interferes with antibody binding, ex vivo CD44+/+ and CD44-/- AMФ were treated with HA’se for 30 minutes prior to flow cytometry analysis. HABP detection of cell surface HA was significantly reduced in CD44+/+ AMФ after HA’se treatment to levels comparable to CD44-/- AMФ. In the absence of cell surface HA, the detection of receptors on CD44+/+ AMФ did not change (Figure 4.1i-j). Thus, the differences in expression of receptors between CD44+/+ and CD44-/- AMФ was not due to the presence of surface HA on the CD44+/+ AMФ.  121  4.3.3 Pulmonary PC surfactant levels are elevated in the lungs of CD44-/- mice  Figure 4.2: CD44 deficiency cause aberrant pulmonary surfactant PC and AMФ lipid accumulation  a) SP-D and b) surfactant PC in the BAL supernatant of CD44+/+ and CD44-/- mice. c) Representative confocal microscopy (z-stack) of ex vivo CD44+/+ and CD44-/- AMФ labeled with BODIPY and DAPI. d) Flow cytometry analysis of CD44+/+ and CD44-/- AMФ labeled with BODIPY. e) Number of BODIPY+ droplets per CD44+/+ or CD44-/- AMФ. f) Average size (µm2) of BODIPY+ droplets in CD44+/+ or CD44-/- AMФ. g) Representative confocal microscopy (z-stack) of ex vivo CD44+/+ and CD44-/- AMФ labeled with the anti-OxPC antibody E06 and DAPI. h) Flow cytometry analysis of CD44+/+ and CD44-/- AMФ labeled anti-OxPC antibody E06. i) Number of E06+ 122  droplets per CD44+/+ or CD44-/- AMФ. j and l) Representative flow cytometry plot and graph comparing H2DCFDA labeling of ROS in CD44+/+ and CD44-/- AMФ. k and m) Representative flow cytometry plot and graph comparing NBD-PE labeling by CD44+/+ and CD44-/- AMФ. n) Representative flow cytometry plots of CD44+/+ and CD44-/- AMФ treated with EtOH, PC, or POVPC labeled with Annexin V and DAPI. o) Percent live CD44+/+ or CD44-/- AMФ (Annexin V- DAPI-) after EtOH, PC, or POVPC treatment for 1 h. p) Comparing fold change in percent live CD44+/+ or CD44-/-  AMФ with different treatments. Data show an average of three to five mice from each experiment ± SD, repeated twice or three times, or five times (h and j); for confocal imaging three to four cells were analyzed per mice. Significance indicated as * p< 0.05, ** p< 0.01, *** p < 0.001, non-paired student’s t-test.   To determine whether CD44-/- mice, which have less AMФ (Dong et al., 2017) that were more foamy and phenotypically abnormal (Figure 4.1), have disruptions in pulmonary surfactant homeostasis, we compared the levels of PC in the BAL of CD44+/+ and CD44-/- mice. The level of SP-D was also compared since it regulates surfactant PC levels in the lung (Korfhagen et al., 1998). Although SP-D level was equal (Figure 4.2a), PC concentration was significantly elevated in the BAL supernatant from CD44-/- mice (Figure 4.2b), suggesting a dysregulation in surfactant lipid processing in CD44-/- lungs and/or AMФ.   4.3.4 AMФ from CD44-/- mice have abnormal lipid accumulation and are more susceptible to OxPC toxicity  Excess cytoplasmic lipid droplet accumulation is correlated with increasing intracellular granularity as measured by SSC-A in flow cytometry (Lee et al., 2004). Since CD44-/- AMФ were more granular (Figure 4.1d-e) and are exposed to greater concentration of extracellular PC, we labeled CD44+/+ and CD44-/- AMФ with the neutral lipid dye BODIPY to compare their cytoplasmic accumulation of lipid droplets. CD44-/- AMФ had greater BODIPY labeling by flow cytometry and confocal 123  microscopy, more BODIPY+ lipid droplets per cell, and larger lipid droplets compared to CD44+/+ AMФ (Figure 4.2c-f). Pulmonary surfactant PC can undergo oxidation, especially during inflammation when there are increased ROS levels and become OxPC (Podrez et al., 2002b; Romero et al., 2015; Yoshimi et al., 2005). The amount of OxPC in CD44+/+ and CD44-/- AMФ was also measured by intracellular labeling with the anti-OxPC antibody (E06) (Horkko et al., 1999). CD44-/- AMФ had higher E06 labeling and greater number of E06 labeled OxPC+ droplets per cell (Figure 4.2g-i). Thus, CD44-/- AMФ were more foamy, and had increased levels of lipid droplets and OxPC. Since OxPC may increase the oxidation stress in cells, we also detected a significant relative increase of ROS levels in CD44-/- AMФ compared to CD44+/+ AMФ using H2DCFDA (Figure 4.2j and l), a ROS reactive fluorogenic dye (Eruslanov and Kusmartsev, 2010). This suggested CD44-/- cells may be under greater oxidative stress. Excess accumulation of lipids and OxPC in the cell are also signs of phospholipidosis, which can be detected by measuring the uptake of fluorescently tagged phospholipid NBD-PE (Morelli et al., 2006). Consistent with the result that CD44-/- AMФ had greater lipid droplet and OxPC accumulation, CD44-/- AMФ had higher NBD-PE MFI, suggesting greater uptake (Figure 4.2k and m). Together, these data led us to test if CD44-/- AMФ were more susceptible to exogenous oxidized lipid induced toxicity. Ex vivo CD44+/+ and CD44-/- AMФ were incubated with the vehicle control 1% ethanol (EtOH), PC, or the OxPC POVPC for 1 h to test their susceptibility to OxPC induced cell death. Loss in cell viability caused by POVPC relative to EtOH or PC treatment was significantly greater in 124  CD44-/- AMФ compared to CD44+/+ AMФ (Figure 4.2n-p), indicating they are more sensitive to OxPC toxicity.   4.3.5 The inflammatory response to OxPC is exacerbated in CD44-/- mice  Figure 4.3: POVPC induced pulmonary inflammation is exacerbated in CD44-/- mice a) Percent weight change over 3 days from CD44+/+ and CD44-/- mice treated with POVPC intratracheally. b) Representative flow cytometry plots showing the gating 125  and proportion of AMФ, Ly6Chigh monocytes, neutrophils, eosinophils, and CD11b- cells in the BAL of CD44+/+ and CD44-/- mice 3 days after POVPC instillation. c-h) Graphs comparing the number of total leukocytes, AMФ, CD11b- cells, Ly6chigh monocytes, neutrophils, and eosinophils in the BAL of CD44+/+ and CD44-/- mice 3 days after POVPC instillation. i) Level of total protein in the BAL of CD44+/+ and CD44-/- treated with or without POVPC measured by BCA. i) Level of HA in the BAL of CD44+/+ and CD44-/- treated with or without POVPC measured by ELISA. j) Relative level of OxPC in the BAL of CD44+/+ and CD44-/-  treated with or without POVPC measured by E06 ELISA. Data show an average of two experiments ± SD, each with three to five mice. Significance indicated as * p< 0.05, ** p< 0.01, *** p < 0.001, non-paired student’s t-test and non-parametric unpaired Mann-Whitney test (i).    Because oxidized phospholipids are produced in the lung during inflammation (Imai et al., 2008) and sterile injury (Romero et al., 2015), we used a model of OxPC driven lung damage where CD44+/+ and CD44-/- mice were treated with i.t. of POVPC to determine if the increased susceptibility to OxPC toxicity in CD44-/- AMФ is physiologically important. CD44+/+ and CD44-/- experienced equal and maximum weight loss two days after POVPC stimulation (Figure 4.3a). Prior to POVPC instillation, AMФ make up over 95% of the cells in the BAL in both CD44+/+ and CD44-/- mice (Figure 4.1a), although CD44-/- mice have less total AMФ (Dong et al., 2017). Three days after POVPC instillation, the BAL of CD44+/+ and CD44-/- mice contained infiltrated monocytes, neutrophils, eosinophils, and CD11b- cells, in addition to AMФ (Figure 4.3b). Notably, equal numbers of AMФ were now present in the CD44+/+ and CD44-/- mice, and significantly greater numbers of infiltrated leukocytes were present in the BAL from CD44-/- mice (Figure 4.3c-h), indicating enhanced recruitment of inflammatory cells. Consistent with this, the amount of total protein (Figure 4.3i) and HA (Figure 4.3j) present in the BAL of CD44-/- mice were also significantly increased, indicating more lung damage. Although there was a 126  trend for increased OxPC in the CD44-/- BAL, this was not significant (Figure 4.3k). Thus, CD44-/- mice generated a greater inflammatory response to POVPC in vivo.  4.3.6 CSF-2 partially rescues lung lipid homeostasis disruption in CD44-/- mice  Figure 4.4: CSF-2 increases the number of AMФ and rescues aberrant pulmonary surfactant PC accumulation a) Representative flow cytometry plot showing the proportion of BAL AMФ from CD44-/- mice after 7 days of PBS or C-SF2 treatment. b) Number of AMФ in the BAL from CD44-/- mice after PBS or C-SF2 treatment. c) BAL PC concentration from CD44-/- mice after PBS or C-SF2 treatment. d) Representative flow cytometry plots comparing the phenotype of AMФ from CD44-/- mice after 7 days of PBS or C-SF2 treatment. e-f) Graphs comparing the MFI of SSC-A, BODIPY, CD36, Siglec F, CD11b, CD11c, CD116, and MerTK between AMФ from CD44-/- mice after 7 days of 127  PBS or C-SF2 treatment. Data show an average of two experiments ± SD, each with three to five mice. Significance indicated as * p< 0.05, ** p< 0.01, *** p < 0.001, non-paired student’s t-test.    CSF-2 can reverse lipid accumulation in CSF-2 deficient AMФ (Malur et al., 2011), by promoting AMФ maturation (Schneider et al., 2014b). It can also induce AMФ proliferation (Worgall et al., 1999) and helps to attenuate bleomycin induced lipid accumulation and fibrosis (Romero et al., 2015). To test if CSF-2 can rescue defects in lipid homeostasis caused by CD44 deficiency, we treated CD44-/- mice with i.t. of CSF-2 daily for 7 days. This did not change the composition of the BAL, which remained largely (>90%) AMФ (Figure 4.4a), but it did cause a significant increase in AMФ numbers, compared to the PBS treated controls (Figure 4.4b).  Furthermore, there was a significant decrease in the amount of PC in the BAL (Figure 4.4c). Analysis of the AMФ from the CSF-2 treated CD44-/- mice showed a significant reduction in CD36 expression (Figure 4.4d-e), but their granularity (SSC), neutral lipid content (measured by BODIPY), and the expression of other cell surface receptors were unaltered (Figure 4.4d-f). These results showed that CSF-2 increased the number of AMФ in CD44-/- mice, which reduced the levels of PC surfactant and decreased CD36 expression on CD44-/- AMФ. However, the accumulation of lipid droplets in CD44-/- AMФ did not change, indicating that CSF-2 alone did not rescue this defect.  128  4.3.7 The alveolar environment regulates AMФ lipid droplet accumulation and CD36 expression while lipid droplet accumulation and CD11c expression are intrinsically affected by the loss of CD44.     Figure 4.5: AMФ lipid homeostasis is controlled by the extracellular milieu and by the expression of CD44 a) Schematic diagram showing donor AMФ isolated from CD45.2+ CD44-/- mice are transferred by i.t. into CD45.1+ CD44+/+ host mice and analyzed 7 days later. b) 129  Representative flow cytometry plots comparing the SSC-A, BODIPY, CD36, and CD11c labeling by CD45.2+ CD44-/- donor AMФ and CD45.1+ CD44+/+ host AMФ 7 days after adoptive transfer. c) Graphs comparing SSC-A, BODIPY, CD36, and CD11c MFI of CD45.2+ CD44-/- donor AMФ, CD45.1+ CD44+/+ host AMФ, and ex vivo CD44-/- AMФ. d) Schematic diagram showing donor AMФ isolated from CD45.1+ CD44+/+ mice are transferred by i.t. into CD45.2+ CD44-/- host mice and analyzed 7 days later. e) Representative flow cytometry plots comparing the SSC-A, BODIPY, CD36, and CD11c labeling by CD45.1+ CD44+/+ donor AMФ and CD45.2+ CD44-/- host AMФ 7 days after adoptive transfer. f) Graphs comparing SSC-A, BODIPY, CD36, and CD11c MFI of CD45.1+ CD44+/+ donor AMФ, CD45.2+ CD44-/- host AMФ, and ex vivo CD44+/+ AMФ. Data show an average of two experiments ± SD, each with three to five mice. Significance indicated as * p< 0.05, ** p< 0.01, *** p < 0.001, paired (donor AMФ vs host AMФ) and non-paired (CD44-/- donor AMФ vs ex vivo CD44-/- AMФ in b, CD44-/- host AMФ vs ex vivo CD44+/+ AMФ in d) student’s t-test.    To distinguish between the environmental and cell intrinsic effects of CD44 loss on AMФ, CD45.2+ CD44-/- AMФ were isolated and adoptively transferred by i.t. into the alveolar space of CD45.1+ CD44+/+ mice (Figure 4.5a), with normal PC levels (Figure 4.2b). Donor CD44-/- AMФ retrieved from the BAL 7 days post adoptive transfer maintained a significantly higher granularity by SSC-A, BODIPY, and CD11c expression compared to host CD44+/+ AMФ, and they were not significantly different from the freshly isolated CD44-/- AMФ used as controls. However, donor CD44-/- AMФ significantly downregulated their CD36 expression to a level even lower than host CD44+/+ AMФ (Figure 4.5b-c). This indicated that the level of CD36 expression can change in CD44-/- AMФ in response to changes in the alveolar environment.  In the converse situation, donor CD45.1+ CD44+/+ AMФ were isolated and adoptively transferred by i.t. into the alveolar space of host CD45.2+ CD44-/- mice (Figure 4.5d), which had higher PC levels (Figure 4.2b). After 7 days, the donor CD44+/+ AMФ increased their granularity, BODIPY and CD36 levels to become like the host CD44-/- AMФ: with equal granularity (SSC-A), BODIPY labeling, and CD36 130  expression that were significantly higher than even the host CD44-/- AMФ (Figure 4.5e-f). These donor CD44+/+ AMФ were significantly different from freshly isolated naïve CD44+/+ AMФ, indicating that the extrinsic alveolar environment of the CD44-/- mice can influence these 3 parameters in CD44+/+ AMФ. Together, these data showed AMФ CD36 expression responded to the surfactant PC levels in both the CD44-/- and CD44+/+ AMФ, and the CD44-/- alveolar environment with greater PC levels also led to greater lipid droplet formation in CD44+/+ AMФ. However, when CD44-/- AM were exposed to lower levels of PC in the CD44+/+ alveolar space, concomitant drop in lipid droplets did not occur, suggesting an intrinsic defect in the regulation of lipid homeostasis by CD44-/-AMФ. The lack of change in CD11c expression in both the CD44+/+ and CD44-/- AMФ further supports that there are intrinsic defects in CD44-/- AMФ.  4.3.8 AMФ lipid droplet accumulation and CD36 expression can be modulated by extracellular PC  Figure 4.6: Cell surface HA and extracellular PC can modulate in vitro AMФ lipid droplet level and CD36 expression   a) Representative flow cytometry plots comparing the BODIPY and CD36 MFI of AMФ after 48 h culture in CSF- 2 or CSF-2 and PC. b and c) Graphs comparing the 131  BODIPY labeling and CD36 expression by AMФ cultured CSF- 2 or CSF-2 and PC for 48 h. d) Representative flow cytometry plots comparing the BODIPY and CD36 MFI of AMФ after 48 h culture in CSF- 2 or CSF-2 and HA’se. e and f) Graphs comparing the level of BODIPY labeling and CD36 expression by AMФ after 48 h culture in CSF- 2 or CSF-2 and HA’se for 48 h. g) Graph comparing NBD-PC uptake by CD44+/+ and CD44-/- AMФ. Data show an average of two experiments ± SD, each with three to five mice. Significance indicated as * p< 0.05, ** p< 0.01, *** p < 0.001, paired student’s t-test.    Since CD44-/- mice had greater accumulation of surfactant PC in the alveolar space (Figure 4.2b), and AMФ CD36 expression and BODIPY levels changed in response to alveolar environments with different PC levels (Figure 4.5), we tested if excess PC alone could increase AMФ lipid accumulation and CD36 expression by culturing ex vivo CD44+/+ AMФ in 500 μM of PC. After 48 h they had significantly increased BODIPY labeling indicating PC incorporation and upregulated CD36 expression (Figure 4.6a-c), showing increased extracellular PC drives both lipid droplet accumulation and CD36 expression. These changes mimicked the characteristics of the CD44-/- AMФ, suggesting that the increased PC levels in these mice can drive their increase in lipid droplets and CD36 expression.  However, because the transplantation of CD44-/- AMФ into CD44+/+ mice with normal PC environment did not rescue their granularity and lipid droplet accumulation (Figure 4.5a), CD44 and/or cell surface HA may intrinsically regulate AMФ lipid accumulation. To test this, we treated CD44+/+ AMФ in vitro with HA’se in the culture media. HA’se treatment removed cell surface HA to the level observed on CD44-/- AMФ (Figure 4.1i-j). CD44+/+ AMФ were maintained in 20 ng/ml of CSF-2 and treated with HA’se for 48 h and showed a significant increase in BODIPY labeling but unchanged CD36 expression compared to untreated cells (Figure 4.6d-f). This recapitulated the observation where CD44-/- AMФ maintained an increased 132  lipid droplet levels after transplantation into a normal CD44+/+ alveolar environment. Thus, intrinsically CD44-/- AMФ may have accumulated more lipid droplets because they have greater PC uptake without cell surface HA, or they have deficient lipid metabolism. To determine if cell surface CD44 and/or HA limited PC uptake, we cultured CD44+/+ and CD44-/- AMФ with NBD-PC, a fluorescently tagged PC, to compare their ability to uptake PC in vitro. Interestingly, NBD-PC MFI was not significantly different between the two (Figure 4.6g), suggesting PC uptake is normal in CD44-/- AMФ.     133  4.3.9 PPARγ expression is downregulated independently of CSF-2 and Stat-5 signaling in CD44-/- AMФ  Figure 4.7: PPARγ expression is defective in CD44-/- AMФ  a) Representative confocal microscopy showing CD44+/+ and CD44-/- AMФ labeled with intracellular PPARγ antibody and DAPI. b) Comparison of nuclear PPARγ MFI between CD44+/+ and CD44-/- AMФ as determined by confocal microscopy. c and d) Representative flow cytometry plot and graph comparing intracellular PPARγ expression between CD44+/+ and CD44-/- AMФ. e and f) Representative flow cytometry plot and graph comparing CD116 expression by MFI between CD44+/+ and CD44-/- AMФ. g and h) Representative flow cytometry plot and graph comparing p-STAT5 expression by MFI between CD44+/+ and CD44-/- AMФ. Data show an 134  average of three to five mice from each experiment ± SD, repeated twice or three times; for confocal imaging three to four fields containing cells were analyzed per mice. Significance indicated as * p< 0.05, ** p< 0.01, *** p < 0.001, non-paired student’s t-test.   CD44-/- AMФ were not fully rescued after transfer into a normal alveolar environment (Figure 4.5a-c), indicating there were cell intrinsic defects leading to lipid droplet accumulation and phenotypic abnormalities, and this was not due to altered PC uptake (Figure 4.6g). Similar to CD44-/- AMФ,  PPARγ deficiency also causes foamy AMФ and surfactant build up (Baker et al., 2010a; Schneider et al., 2014b; Shibata et al., 2001b). To determine whether CD44-/- AMФ had altered PPARγ expression that can contribute to their defective phenotype, PPARγ expression and localization from ex vivo CD44+/+ and CD44-/- AMФ were measured. Confocal microscopy showed almost all PPARγ detected in ex vivo AMФ were localized to the nucleus (Figure 4.7a) and there was significantly less PPARγ detected in CD44-/- AMФ compared to CD44+/+ AMФ (Figure 4.7a-d). Since PPARγ expression is regulated by CSF-2, and CSF-2 induction of PPARγ expression is critical for the development and maintenance of mature AMФ in the lung (Schneider et al., 2014b), we posited that CD44-/- AMФ may have reduced CSF-2 signaling. However, while there was significantly higher CSF-2 receptor expression in CD44-/- AMФ (Figure 4.7e-f), the phosphorylation of Stat-5, a downstream CSF-2 receptor signaling molecule was equal (Figure 4.7g-h). In addition, there was no significant difference in CSF-2 concentration detected from CD44+/+ and CD44-/- BAL (data not shown). Together, these results suggest CD44-/- AMФ have PPARγ deficiency was independent of CSF-2 signaling. 135  4.4 Discussion AMФ in the healthy lung self-renew and maintain tissue homeostasis (Hussell and Bell, 2014; Iwasaki et al., 2017; Kopf et al., 2014). Numerous studies have demonstrated improper development and absence of mature AMФ cause PAP (Dranoff et al., 1994; Happle et al., 2014; Shibata et al., 2001b; Suzuki et al., 2014; Yoshida et al., 2001). The loss of mature AMФ can arise from the lack of CSF-2 protein or receptor expression. In humans, PAP is rare and occurs due to autoimmunity against the CSF-2 protein (Kitamura et al., 1999) or hereditary mutations in the CSF-2 receptor (Suzuki et al., 2010). However, how a partial loss or change of AMФ due to aging, chronic diseases, or infections affect lung homeostasis and immune responses, as well as what threshold of AMФ loss could lead to increased pathology, are less clear. Here, we show that in CD44-/- mice that have a 50% loss of AMФ (Dong et al., 2017), have increased pulmonary surfactant lipid levels and altered AMФ lipid homeostasis, leading to increased susceptibility to OxPC toxicity and inflammation. These data also suggest that AMФ numbers regulate surfactant lipid levels in the alveolar space, which has physiological consequences during challenge with OxPC, which can be generated in different types of inflammation (Imai et al., 2008). We identified both environmental and cell intrinsic defects caused by the loss of CD44. The environmental defects can be attributed to the reduced numbers of AMФ being unable to turnover surfactant PC sufficiently, leading to increased PC levels in the alveolar space and a concomitant upregulation of CD36 expression.  136  CD44-/- AMФ expressed higher levels CD36, a scavenger receptor for various ligands including phospholipids and OxPC (Febbraio et al., 2001; Podrez et al., 2002b; Rahaman et al., 2006). They were also more granular, accumulated more lipid droplets and OxPC, and were in an alveolar environment with an abnormally high PC concentration. CSF-2 instillation partially restored the number of AMФ in CD44-/- mice, ameliorated the excess BAL PC levels, and downregulated CD44-/- AMФ CD36 expression. This result suggested CD36 expression was regulated by extracellular PC concentration, and maintaining a normal number of AMФ in the alveolar space helped to ensure the sufficient catabolism of pulmonary surfactants. Adoptively transferring CD44+/+ AMФ into CD44-/- lungs, where PC levels was increased, led to increased BODIPY labeling and CD36 expression, as did the addition of PC to CD44+/+ AMФ in vitro, demonstrating that elevated levels of PC promoted lipid droplet accumulation and upregulated CD36 expression. Conversely, CD36 expression was downregulated in CD44-/- AMФ when they were adoptively transferred into CD44+/+ lungs or when CSF-2 was instilled into the lungs of CD44-/- mice. These observations showed that PC concentration in the BAL was an environmental factor that regulated CD36 expression and this correlated with lipid droplet accumulation in CD44+/+ AMФ. However, PC levels in the alveolar space can also be controlled by the number of AMФ present, implicating that the decreased number of AMФ in CD44-/- mice disrupts surfactant lipid homeostasis.  Neither CSF-2 treatment nor the adoptive transfer of CD44-/- AMФ into CD44+/+ lungs rescued their accumulation of BODIPY+ lipid droplets or increased cell granularity. These abnormal phenotypes in CD44-/- AMФ persisted despite a rescue 137  of the PC levels in the alveolar space, suggesting an intrinsic defect in lipid metabolism in the CD44-/- AMФ. This may be related to their lower PPARγ expression, as PPARγ is a master regulator of lipid metabolism (Varga et al., 2011).  AMФ from PPARγ deficient mice were also foamy in appearance and had altered expression of cholesterol efflux and metabolism genes such as increased CD36 mRNA expression (Baker et al., 2010b).   In vitro experiments showed that the removal of the HA coat in CD44+/+ AMФ with HA’se increased BODIPY labeling, implicating the engagement of HA in regulating lipid droplet accumulation. Since NBD-PC uptake was not different between CD44+/+ and CD44-/- AMФ, the metabolism of lipid droplet formation or degradation may be regulated by HA interactions. For example, HA and CD44 may contribute to PPARγ expression and lipid droplet accumulation through TGF-β. Autocrine TGF-β production and signaling are integral for AMФ and lung homeostasis as TGF-β-/- mice have deficient AMФ numbers, PPARγ expression, and abnormal surfactant accumulation (Yu et al., 2017). Previous studies also showed HA and CD44 are involved in TGF-β signaling (Acharya et al., 2008; Ito et al., 2004b; Meran et al., 2011). Thus, CD44-/- AMФ may have deficient TGF-β production or signaling, which can downregulate their PPARγ expression and increase their lipid droplet accumulation. In addition, the upregulation of many cell surface receptors, including alternative activation markers such as CD206, CD200R, SIRPα, and CD36, as well as MHCII suggests that CD44-/- AMФ have received or are more responsive to factors that promote alternative activation. Since PPARγ is associated with polarization to alternative activation, changes in cell surface receptor 138  expression in CD44-/- AMФ may also relate to their deficient PPARγ expression. Thus, exactly how HA and CD44 regulate AMФ lipid homeostasis and cell surface receptor expression requires further investigation.  The increased inflammatory response to OxPC from CD44-/- mice suggests the reduced number of CD44-/- AMФ and their increased sequestration of OxPC negatively affected their inflammatory response to OxPC. During lung damage and inflammation, surfactant lipids such as PC can become oxidized (Imai et al., 2008; MacNee, 2001; Romero et al., 2015; Yoshimi et al., 2005). While these can be taken up by AMФ and metabolized, high concentrations cause cell death (Halasiddappa et al., 2013; Ramprecht et al., 2015), NLRP3 inflammasome activation (Yeon et al., 2017), and TLR activation (Imai et al., 2008; Kadl et al., 2011), which all can exacerbate inflammation and tissue damage. As OxPC is produced in multiple types of lung inflammation (Imai et al., 2008; Romero et al., 2015; Yoshimi et al., 2005), and oxidative stress has been considered as an important component of lung inflammation (MacNee, 2001; Rahman and Adcock, 2006), how lung immune cells respond to and clear OxPC are of broad interest. Thus, OxPC uptake by tissue resident cells such as AMФ may be essential to the resolution of inflammation, and any deficiencies in this process in humans, could contribute to their susceptibility to lung diseases.  Studies show the removal of AMФ prior to infection or inflammatory challenge exacerbates inflammation from i.t. of trinitrophenyl-keyhole limpet hemocyanin (Thepen et al., 1989), Streptococcus pneumoniae (Knapp et al., 2003), and influenza (Schneider et al., 2014a), indicating they have anti-inflammatory roles or 139  that the damage is much greater in the absence of AMФ. Since CD44-/- mice have less AMФ (Dong et al., 2017), they may be predisposed to greater damage, such as from OxPC. Since CSF-2 increased AMФ numbers and decreased surfactant PC levels, this is another potential mechanism to limit damage and inflammation from lung diseases. CSF-2 instillation has been shown to reduce lung fibrosis caused by bleomycin (Romero et al., 2015), and it is currently pursued as a therapy for treating PAP (Ohashi et al., 2012).  Overall, this study showed that CD44 deficiency leads to increased pulmonary PC levels in the alveolar space, which acted extrinsically to increase AMФ CD36 expression and contributed to lipid droplet accumulation. CD44 deficient AMФ also downregulated their PPARγ expression and their intrinsic dysregulation in lipid homeostasis was not rescued by environmental factors such as CSF-2 or normalized PC levels. The cumulative extrinsic and intrinsic defects sensitized CD44-/- mice to greater OxPC induced inflammation in the lung.  140  Chapter 5: Concluding discussion and remarks 5.1 Ability of HA to stimulate macrophages and DCs  This study demonstrated HA of various sizes and forms, including the HA-HC complexes generated in vitro by TSG-6 mediated covalent linking of HA with the HC of IαI, do not stimulate macrophages and DCs to produce Type 1 inflammatory cytokines such as TNF-α, IL-12, and IL-1β (Dong et al., 2016). The lack of co-stimulatory molecule upregulation from HA stimulation further confirmed the cells were not activated. These results were consistent in experiments using in vitro BMDM and BMDCs, ex vivo splenic macrophages and DCs, and in vivo in the lung. In addition, the levels of endotoxin contamination present in various HA reagents were evaluated and only Huc HA was contaminated with significant amounts of endotoxin. While polymyxin B beads helped to remove endotoxin from HA’se proteins, they were ineffective at cleaning high concentrations of Huc HA. Alternatively, Triton X-114 phase separation removed the contaminating endotoxin from Huc HA without significantly impacting the amount of Huc HA retrieved. These findings demonstrated the need to review previous conclusions from the literature that suggest LMW HA stimulate inflammation via TLR signaling, where studies use polymyxin B to rule out endotoxin contamination as the cause of HA induced inflammation. The possibility of PAMP contamination may also explain the various observations where HA induced inflammation requires TLR signaling. These new insights are now being recognized more (Safrankova et al., 2017; Weigel, 2017), and future studies should ensure HA reagents are free of contamination or use FDA 141  approved endotoxin free HA produced by biotechnology companies such as Lifecore Biomedical and Hyalose. Although the data from this study showed Type 1 inflammatory responses were not initiated by HA, it may also be possible that HA induces other types of inflammatory responses that was not tested in this study. For instance, HA may promote the production of Type 2 inflammatory cytokines such as IL-4 and IL-13, or alarmins such as IL-33 from macrophages and DCs. In addition, the types of HA tested in this study may not fully recapitulate the exact forms and sizes of HA generated during tissue injury, thus the in vivo consequences of increased LMW HA during inflammation require additional investigation. Future experiments could use RNA sequencing to help determine what types of transcriptional changes occur to immune cells following HA stimulation.   If HA is not an initiator of inflammation, or a DAMP, it may still be a useful biomarker that is associated with tissue damage and inflammation. In addition, there is the possibility that while HA does not activate macrophages and DCs to produce inflammatory cytokines and upregulate the co-stimulatory molecules tested in this study, it can cause other transcriptional changes in immune cells and tissues that are not yet discovered, such as the alternative activation of macrophages or fibrosis. The lack of inflammation from HA fragments is encouraging for PEGylated human recombinant HA’se therapies currently in clinical trials for treating certain types of tumors containing high amounts of HA (Hingorani et al., 2016; Infante et al., 2017). Regardless, the biological functions of HA fragments and HA-HC complexes in vivo during inflammation are poorly understood. Future research could test the effects of 142  increased HA production and deposition on the physical structure of damaged tissues such as an inflamed lung as a potential function of HA accumulation. Whether the degradation of HA and ECM in the lung can contribute to inflammation by promoting immune cell migration to the site of tissue damage is also worth investigating. Furthermore, whether HA interactions cause downstream signaling in cells and what those signaling pathway are, require additional elucidation. Because HA is integral to embryonic development, it is difficult to isolate the role of HA in tissues through lost-of-function studies. Future investigations could take advantage of CRISPR/cas9 targeted genome editing approach in vivo (de Solis et al., 2016; Dow et al., 2015; Manguso et al., 2017) and delete HAS or HA’se enzyme in situ to help understand the role of HA in vivo during homeostasis and in diseases.  5.2 Contribution of monocyte-derived macrophages to AMФ repopulation In the LPS model of acute lung inflammation, AMФ were initially depleted, but they repopulated in the alveolar space as inflammation resolved. Inflammatory monocytes were recruited to the alveolar space during this time and monocyte-derived cells were present in the alveolar space up to 45 days after the initial stimulation. By day 45, the monocyte-derived cells were phenotypically identical to AMФ and self-renewed at the same rate. These data suggest inflammatory monocytes contributed to AMФ repopulation after inflammation (Dong et al., 2017). Although this study did not compare the transcriptional programming of monocyte-derived AMФ versus the original fetal-derived AMФ and therefore cannot definitively conclude that the monocyte derived cell became identical to the resident AMФ both 143  phenotypically and functionally, another recent study shows they have similar transcriptional signature by RNA sequencing with only 330 differentially expressed genes (Misharin et al., 2017). In addition, since only adoptively transferred bone marrow derived macrophages were used to demonstrate their ability to differentiate and become AMФ in the BAL, it will be of broad interest to use a fate-mapping model where infiltrating monocyte are fluorescently marked in situ to determine the degree of monocyte contribution to AMФ repopulation, and if this is regulated by the intensity or type of inflammation. Nevertheless, observations from this study is consistent with the hypothesis that monocytes can differentiate into tissue resident macrophages when there is a niche made available from the depletion or death of the original resident macrophages (Guilliams and Scott, 2017; van de Laar et al., 2016). Thus, during inflammation, both the surviving fetal-derived macrophages and the infiltrating monocytes compete for available niche, growth factors, or survival factors to regenerate the tissue macrophage population (Guilliams and Scott, 2017). Whether monocyte-derived AMФ are functionally different to fetal-derived AMФ, particularly in the context of different diseases, requires further investigation. Although Misharin et al. suggest differentiating monocyte-derived macrophages contribute to lung fibrosis whereas fetal-derived AMФ do not (Misharin et al., 2017), it is unclear whether this functional difference is maintained when the intermediate monocyte-derived macrophages terminally differentiate into long term mature AMФ, especially upon re-stimulation. Perhaps as individuals age and become afflicted with various respiratory illnesses over time, their fetal-derived AMФ are replaced by monocyte-derived AMФ. By comparing the ontogeny and functional potential of 144  young AMФ and old AMФ, we will acquire new knowledge that can help answer questions such as why elderly individuals become more susceptible to pulmonary infections.   5.3 Role of HA binding via CD44 for AMФ survival  Although AMФ are characterized to bind HA ex vivo and uptake HA via CD44 in vitro, it is unclear if this interaction participates in any other cell function in vivo. This study identified a novel role for HA binding and CD44 in promoting AMФ survival (Dong et al., 2017). AMФ bound a layer of cell surface HA in vivo. Disrupting this HA binding by deleting CD44 or with a HA blocking CD44 mAb increased AMФ death, demonstrating the importance of the HA interaction with CD44 for maintaining the normal number of AMФ in the lung. However, the mechanism of how HA regulates AMФ survival needs further research. Does HA act as a physical barrier to protect against extracellular stresses such as ROS, or does its interaction with CD44 promote cell survival or anti-apoptotic signaling? One way to test if HA binding to CD44 signals to promote cell survival is to generate AMФ that express a mutant version of CD44 without the cytoplasmic domain, which will not prorogate intracellular interactions and/or signaling. However, the HA independent effects of deleting the cytoplasmic domain of CD44 will also need to tested in this case. In addition, the source of HA present on AMФ was not determined by experiments in this study. Thus, there is a limitation in the ability to conclude on the nature of the HA interaction by AMФ, such as whether it is pericellular HA binding, binding with HA on AECs, or both. Moreover, since HA can be degraded by factors such as ROS 145  or HA’se in the environment, it will be interesting to determine if the size and amount of HA bound by AMФ change during inflammation, and if so, how do these changes affect AMФ function and survival? Answering these questions may help the development of new treatments for patients afflicted with lung diseases, such as using HMW HA to promote cell survival and to restore homeostasis in diseases such as cystic fibrosis (Gavina et al., 2013; Lamas et al., 2016), asthma (Gebe et al., 2017), and IPF (Liang et al., 2016).   5.4 Effect of AMФ loss and CD44 deficiency in the lung  Figure 5.1: Schematic showing the impact of AMФ loss and CD44 deficiency have on lipid homeostasis in the alveolar space   By comparing CD44+/+ and CD44-/- mice, multiple deficiencies in CD44-/- AMФ were identified (Figure 5.1). In addition to the lower survival of CD44-/- AMФ due the lack of HA binding, they were phenotypically different and accumulated abnormal 146  amounts of intracellular lipids. Notably, CD44-/- AMФ accumulated excess intracellular OxPC and were more susceptible to OxPC toxicity induced cell death in vitro. This, together with the observation that CD44-/- mice had exacerbated inflammation from i.t of OxPC stimulation compared to CD44+/+ mice, suggested alterations in their lipid homeostasis and reduced AMФ numbers may sensitize the lung to inflammation. This may be another explanation of why CD44-/- mice have increased inflammation after bleomycin treatment (Teder et al., 2002), as bleomycin induces OxPC accumulation in the alveolar space (Romero et al., 2015). Thus, the production and clearance of OxPC are likely important processes that dictate the severity and outcome of lung diseases (Bochkov et al., 2017). However, because HA levels in the lung was also increased in CD44-/- mice compared to CD44+/+ mice after OxPC stimulation, the alternative hypothesis that greater HA deposition from OxPC also exacerbates inflammation in CD44-/- lungs needs to be tested. In addition to increased neutral lipid accumulation in CD44-/- AMФ, CD44-/- mice also had elevated levels of the PC surfactant in their lungs. Given PC is the major lipid component of pulmonary surfactant, this observation indicated that CD44 deficiency disrupted pulmonary lipid surfactant homeostasis. There are two major defects in CD44-/- mice that may have caused this: 1) the decreased number of AMФ also reduced the total PC turnover, or 2) the CD44-/- AMФ changed their lipid metabolism which led to lipid accumulation. The number of AMФ, CD36 expression, and pulmonary PC levels were rescued after CD44-/- mice were treated with i.t. of CSF-2. Although we did not determine how lung CSF-2 instillation helped to restore AMФ numbers and PC levels, these observations suggested the accumulation of 147  extracellular PC in CD44-/- lungs and the upregulation of CD36 in CD44-/- AMФ were associated with insufficient AMФ numbers. However, CD44-/- AMФ still accumulated greater amounts of intracellular lipid droplets despite CSF-2 treatment, and after they were transferred into CD44+/+ lungs, which were normal alveolar environments. Thus, there must be cell-intrinsic defects due to CD44 deletion, possibly through the downregulation of PPARγ. The changes in PPARγ regulation was not well tested in experiments from this study. For instance, the effect of CSF-2 treatment on CD44-/- AMФ PPARγ expression is of interest because it may help determine whether changing surfactant PC levels directly affect PPARγ expression. Similarly, knowing the expression of PPARγ after CD44-/- AMФ are transplanted into the CD44+/+ lung environment could clarify whether CD44 deficiency is linked directly to PPARγ expression. These additional observations could help understand the relationship between CD44 and PPARγ expression by AMФ. PPARγ is a member of the ligand activated PPAR nuclear hormone receptor family of transcription factors (Janani and Ranjitha Kumari, 2015). Upon interaction with natural ligands such the OxPC hexadecyl azelaoyl phosphatidylcholine (Davies et al., 2001), PPARγ is translocated to the nucleus where it can regulate gene expression for important processes such as lipid metabolism and tolerance to lipotoxicity (Medina-Gomez et al., 2007; Umemoto and Fujiki, 2012). PPARγ is also important for the differentiation of alternatively activated macrophages and the loss of PPARγ in tissue resident macrophages causes insulin resistance, glucose intolerance, and diet-induced obesity in mice (Odegaard et al., 2007). In addition, oxidized low-density lipoprotein degradation and cholesterol removal are defective in 148  macrophages lacking PPARγ (Chinetti et al., 2001; Moore et al., 2001). In the lung, PPARγ is essential for the proper differentiation of fetal monocytes into AMФ. AMФ from PPARγ deficient mice are foamy and lipid-laden (Schneider et al., 2014b), and excess surfactant lipids accumulate (Baker et al., 2010a; Malur et al., 2011; Schneider et al., 2014b). Together, these data implicate PPARγ as not only important for the development and maintenance of mature AMФ identity, but suggest that it may also be needed for the proper metabolism of surfactant lipids by AMФ. Thus, the defects found in CD44-/- AMФ lipid homeostasis may be due to their defective PPARγ expression. Interestingly, Yu et al. recently demonstrated that TGF-β is secreted by AMФ and it is required for normal PPARγ expression, as well as their development and self-maintenance at the steady-state (Yu et al., 2017). HA and CD44 interaction can regulate TGF-β receptor trafficking to lipid rafts and TGF-β signaling in epithelial cells (Ito et al., 2004a; Ito et al., 2004b), TGF-β activation in fibroblasts (Acharya et al., 2008), and can promote TGF-β mediated proliferation in fibroblasts (Meran et al., 2011; Meran et al., 2008). However, it is unclear from these studies whether potential endotoxin contamination may contribute to HA mediated TGF-β signaling during in vitro stimulations. Therefore, defective TGF-β activation or signaling in CD44-/- AMФ may downregulate PPARγ expression, which in turn alters their cell phenotype and lipid accumulation. Treating CD44-/- mice with i.t. of active TGF-β could test this hypothesis by determining if CD44-/- AMФ numbers, phenotype, and lipid homeostasis are restored after treatment.  Alternatively, CD44 may have regulatory functions that are independent from its interaction with HA. CD44 can be cleaved by presenilin-mediated γ-secretase 149  activity (Murakami et al., 2003). The release of the CD44 intracellular domain could signal in the nucleus by binding to DNA regulatory regions such as the 12-O-tetradecanoylphorbol 13-acetate–responsive element (Okamoto et al., 2001), as well as CCTGCG consensus sequences to modulate gene expression (Miletti-Gonzalez et al., 2012). It will be of interest to determine if the CD44 intracellular domain can also bind to the promoter or the regulating elements that control PPARγ expression through experiments such as chromatin immunoprecipitation. CD44 may also indirectly control PPARγ expression by regulating any genes upstream of PPARγ transcription, such as the C/EBP-α and C/EBP-β (Farmer, 2005; Wu et al., 1999). Interestingly, a product of HA catabolism is uridine diphosphate N-acetylglucosamine (Hascall et al., 2014), which is a substrate in pathways that control macrophage metabolism and subsequent polarization (Jha et al., 2015). In the absence of CD44 expression, there may be a change in HA catabolism or cell metabolism which could change the transcriptional programming of CD44-/- AMФ and lead to phenotypic and functional defects. The metabolic state of the cell is important for the function and polarization of macrophages. For example, lipolysis and fatty acid oxidation are critical for macrophage alternative activation (Huang et al., 2014a). Thus, it is of interest to compare the transcriptome of CD44+/+ and CD44-/- AMФ by RNA sequencing in order to reveal potential differences in gene expression that may regulate cellular HA or lipid metabolism. In summary, how CD44 deletion and/or the lack of HA binding cause these cell intrinsic defects in AMФ is of interest for future studies, and has implications in understanding how 150  macrophage function and identity are regulated during homeostasis and inflammation.   The observation that AMФ deficiencies and lipid accumulation lead to exacerbated lung inflammation has important implications for human diseases. For example, COPD patients have increased oxidized stress from ROS or lipid peroxidation products (Rahman and Adcock, 2006), impaired numbers of lung macrophages which are also more sensitive to inflammation (Barnes, 2004; Berenson et al., 2014), and increased susceptibility to opportunistic bacterial lung infections (Monso et al., 1995). Since AMФ are important for the control of pathogens at the start of infection, one hypothesis could be that the accumulation of oxidized phospholipids during COPD cause foamy AMФ and AMФ depletion, which could increase the susceptibility to secondary infections. Therefore, it is important to investigate whether AMФ loss and pulmonary lipid surfactant dysregulation during chronic lung inflammation promote opportunistic infections, such as by impairing AMФ bacterial uptake (Thimmulappa et al., 2012). Finally, understanding the factors that control lipid accumulation in macrophages has wider clinical implications in other human diseases such as atherosclerosis and diabetes.    151  Bibliography  A-Gonzalez, N., Guillen, J.A., Gallardo, G., Diaz, M., de la Rosa, J.V., Hernandez, I.H., Casanova-Acebes, M., Lopez, F., Tabraue, C., Beceiro, S., et al. (2013). The nuclear receptor LXRalpha controls the functional specialization of splenic macrophages. Nat Immunol 14, 831-839. 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