UBC Theses and Dissertations

UBC Theses Logo

UBC Theses and Dissertations

Regulation of mucosal T cell responses by intestinal helminths and retinoic acid metabolism Chenery, Alistair Lee 2016

Your browser doesn't seem to have a PDF viewer, please download the PDF to view this item.

Item Metadata

Download

Media
24-ubc_2016_may_chenery_alistair.pdf [ 4.31MB ]
Metadata
JSON: 24-1.0300054.json
JSON-LD: 24-1.0300054-ld.json
RDF/XML (Pretty): 24-1.0300054-rdf.xml
RDF/JSON: 24-1.0300054-rdf.json
Turtle: 24-1.0300054-turtle.txt
N-Triples: 24-1.0300054-rdf-ntriples.txt
Original Record: 24-1.0300054-source.json
Full Text
24-1.0300054-fulltext.txt
Citation
24-1.0300054.ris

Full Text

Regulation of Mucosal T Cell Responses by Intestinal Helminths and Retinoic Acid Metabolism  by  Alistair Lee Chenery B.Sc., The University of British Columbia, 2011  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Pathology and Laboratory Medicine)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  April 2016  © Alistair Lee Chenery, 2016 ii  Abstract  Mucosal immune diseases such as asthma and inflammatory bowel disease are associated with major environmental factors - diet, geography, hygiene, infections - that contribute to disease risk. The mucosal immune system is in direct contact with the external environment and must balance protective immune responses with tolerance to innocuous antigens. Since chronic inflammation at mucosal sites depends on a diverse set of T cell-driven responses, understanding the factors that regulate mucosal T cell differentiation and function is key to developing better treatments to a variety of inflammatory diseases. Further, immunological cross-talk can occur between mucosal organs such as the intestine and the lung, but the role of T cells in this cross-talk is poorly defined. The work herein investigates the effect of external factors, specifically intestinal infections and dietary immunomodulators, on mucosal T cell responses in the context of inflammation in the intestine and the lungs. Using a mouse model of infection with the intestinal helminth Trichuris muris, I show infection-mediated alterations in the lung microenvironment that can protect against murine models of allergic airway inflammation. I further show that intestinal T. muris infection has a systemic effect on hematopoiesis in the bone marrow. In other studies, I examine the role of the dietary vitamin A metabolite, retinoic acid, on T cell function during intestinal inflammation. Specifically, I investigate how metabolism of retinoic acid by the enzyme Cyp26b1 modulates T cell differentiation and function and show that Cyp26b1 controls regulatory T cell and T helper 17 differentiation in vitro. Further, I posit a role of Cyp26b1 in regulating effector T cell function in vivo using a murine model of T cell-driven inflammatory bowel disease. Thus, the results presented here provide further insight into helminth-mediated immune regulation, intestine-to-lung mucosal immune crosstalk, and dietary immunomodulation that regulate mucosal T cell responses.  iii  Preface  All work was conducted by me under the guidance of Dr. Colby Zaph. I was involved in all experimental planning, performed all of the experiments, and performed all data analyses.  A version of chapter 2 has been published: Chenery, A.L., Antignano, F., Burrows, K., Scheer, S., Perona-Wright, G., & Zaph, C. Low dose intestinal Trichuris muris infection alters the lung immune microenvironment and can suppress allergic airway inflammation. 2016. Infect. Immun. 84:491–501.  I contributed to the experimental design, performed all forms of data acquisition/analysis, and wrote the entire manuscript.  A version of chapter 3, at the time of writing, has been submitted for publication: Chenery, A.L., Antignano, F., Hughes, M.R., Burrows, K., McNagny, K.M., & Zaph, C. Chronic Trichuris muris infection alters hematopoiesis and causes IFN-gamma-expressing T cells to accumulate in the bone marrow. 2016. (Submitted).  I contributed to the experimental design, performed all forms of data acquisition/analysis, and wrote the entire manuscript.  A version of chapter 4 has been published: Chenery, A., Burrows, K., Antignano, F., Underhill, T.M., Petkovich, M., & Zaph, C. The retinoic acid metabolizing enzyme Cyp26b1 regulates CD4 T cell differentiation and function. 2013. PLoS One 8:e72308.  I contributed to the experimental design, performed all forms of data acquisition/analysis, and wrote the entire manuscript. iv   Ethics approval for studies presented in chapter 2 and 3 was obtained from the University of British Columbia Animal Care Committee: protocol number A13-0010. Ethics approval for studies presented in chapter 4 was obtained from the University of British Columbia Animal Care Committee: protocol number A11-0329. All animal studies performed were in accordance with the Canadian Guidelines for Animal Research. v  Table of Contents Abstract .......................................................................................................................................... ii Preface ........................................................................................................................................... iii Table of Contents ...........................................................................................................................v List of Tables ..................................................................................................................................x List of Figures ............................................................................................................................... xi List of Acronyms and Abbreviations ....................................................................................... xiii Acknowledgements ................................................................................................................... xvii Chapter 1: Introduction ................................................................................................................1 1.1 Overview ......................................................................................................................... 1 1.2 Organization of the mucosal immune system ................................................................. 2 1.3 T helper cell differentiation and function ....................................................................... 5 1.4 Mucosal immune crosstalk ............................................................................................. 8 1.5 Mucosal inflammatory diseases .................................................................................... 11 1.5.1 Allergic airway inflammation ................................................................................... 12 1.5.1.1 Experimental models of AAI ............................................................................ 15 1.5.2 Inflammatory bowel disease ..................................................................................... 17 1.5.2.1 Experimental models of IBD ............................................................................ 20 1.6 Modulation of mucosal immune responses................................................................... 22 1.6.1 Intestinal helminth infections .................................................................................... 22 1.6.1.1 The Trichuris muris model of intestinal helminth infection ............................. 25 1.6.2 Retinoic acid ............................................................................................................. 28 1.6.2.1 RA metabolism in T cells ................................................................................. 31 vi  1.7 Rationale, hypothesis and specific aims ....................................................................... 31 Chapter 2: Alteration of the lung immune microenvironment by the intestinal helminth Trichuris muris .............................................................................................................................34 2.1 Overview ....................................................................................................................... 34 2.2 Materials and methods .................................................................................................. 35 2.2.1 Mice .......................................................................................................................... 35 2.2.2 Propagation of T. muris eggs, worm antigen, and oral infections ............................ 35 2.2.3 Induction of acute lung inflammation with papain ................................................... 36 2.2.4 Antibodies and flow cytometry ................................................................................. 37 2.2.5 Gene expression analysis of lung T cells .................................................................. 38 2.2.6 Lung leukocyte ex vivo culture stimulation .............................................................. 38 2.2.7 RNA isolation and quantitative real-time PCR ......................................................... 39 2.2.8 Enzyme-linked immunosorbent assay (ELISA) ....................................................... 40 2.2.9 Histology ................................................................................................................... 40 2.2.10 Depletion of CD4+ T cells ..................................................................................... 41 2.2.11 In vivo neutralization of IL-12 .............................................................................. 41 2.2.12 In vivo blockade of IL-10 activity ......................................................................... 41 2.2.13 HDM model of AAI .............................................................................................. 42 2.2.14 Statistics ................................................................................................................ 42 2.3 Results ........................................................................................................................... 42 2.3.1 Low dose T. muris infection alters the lung immune microenvironment ................. 42 2.3.2 Lung CD4+ T cells increase IFN-γ production during low dose T. muris infection 44 2.3.3 Lung myeloid cells increase IL-10 expression during low dose T. muris infection . 46 vii  2.3.4 Low dose T. muris infection protects against innate-mediated allergic airway inflammation ......................................................................................................................... 49 2.3.5 Low dose T. muris infection protects against acute airway inflammation in the absence of CD4+ T cells ........................................................................................................ 51 2.3.6 Rag1−/− mice are partially protected from acute innate-mediated airway inflammation during low dose T. muris infection ................................................................. 53 2.3.7 Neutralization of IL-10 activity in T. muris-infected Rag1−/− mice during acute airway inflammation ............................................................................................................. 54 2.3.8 Low dose T. muris infection protects against the house dust mite model of asthma 56 2.4 Discussion ..................................................................................................................... 58 Chapter 3: Chronic Trichuris muris infection affects hematopoiesis in the bone marrow ...62 3.1 Overview ....................................................................................................................... 62 3.2 Materials and methods .................................................................................................. 63 3.2.1 Mice and intestinal worm infections ......................................................................... 63 3.2.2 Isolation of bone marrow cells .................................................................................. 63 3.2.3 In vivo neutralization of IFN-γ .................................................................................. 63 3.2.4 Colony forming unit (CFU) assay............................................................................. 64 3.2.5 Antibodies and flow cytometry ................................................................................. 64 3.2.6 Statistics .................................................................................................................... 65 3.3 Results ........................................................................................................................... 65 3.3.1 Chronic T. muris infection induces increased Sca-1 expression on hematopoietic cells and increases LSK cells in the BM ............................................................................... 65 viii  3.3.2 Chronic T. muris infection alters bone marrow HSC and MPP populations and is dependent on adaptive immunity .......................................................................................... 68 3.3.3 Changes in hematopoiesis during chronic T. muris infection is IFN-γ-dependent ... 70 3.3.4 A mixed population of IFN-γ-expressing T cells accumulate in the BM during chronic T. muris infection ..................................................................................................... 72 3.3.5 IFN-γ-expressing T cells in the BM display activated and central memory phenotypes during chronic T. muris infection ...................................................................... 73 3.4 Discussion ..................................................................................................................... 75 Chapter 4: The RA-metabolizing enzyme Cyp26b1 controls Th cell differentiation and function .........................................................................................................................................78 4.1 Overview ....................................................................................................................... 78 4.2 Materials and methods .................................................................................................. 79 4.2.1 Mice .......................................................................................................................... 79 4.2.2 In vitro T cell polarization assays ............................................................................. 79 4.2.3 Treg suppression assay ............................................................................................. 79 4.2.4 In vitro stimulation of CCR9 expression .................................................................. 80 4.2.5 T cell proliferation assay ........................................................................................... 80 4.2.6 Antibodies and flow cytometry ................................................................................. 80 4.2.7 RNA isolation and quantitative real-time PCR ......................................................... 81 4.2.8 T cell transfer model of colitis .................................................................................. 82 4.2.9 Statistics .................................................................................................................... 82 4.3 Results ........................................................................................................................... 83 4.3.1 Expression of Cyp26b1 is dispensable for T cell development ................................ 83 ix  4.3.2 Naturally-occurring Treg cells are normal in Cyp26b1−/− mice ................................ 84 4.3.3 Cyp26b1 modulates iTreg and Th17 cell polarization in vitro ................................. 85 4.3.4 Cyp26b1-deficient T cells fail to induce intestinal inflammation ............................ 88 4.3.5 Cyp26b1-deficient T cells do not have altered expression of intestinal homing molecules .............................................................................................................................. 90 4.4 Discussion ..................................................................................................................... 92 Chapter 5: Conclusions ...............................................................................................................95 5.1 Research summary and significance ............................................................................. 95 5.2 Study limitations ......................................................................................................... 100 5.3 Future directions ......................................................................................................... 102 5.4 Concluding remarks .................................................................................................... 107 Bibliography ...............................................................................................................................108 Appendices ..................................................................................................................................128 Appendix A chapter 2 supplemental material ......................................................................... 128 A.1 Supplemental figures .............................................................................................. 128    x  List of Tables  Table 1.1 Helminth-mediated protection against mouse models of inflammatory disease. ......... 24 Table 2.1 Primer sets used for qRT-PCR analysis. ....................................................................... 39 Table 4.1 Primer sets used for qRT-PCR analysis. ....................................................................... 81  xi  List of Figures Figure 1.1 Simplified schematic of mucosal immune regulation. .................................................. 3 Figure 1.2 Common Th cell lineage differentiation paradigm. ...................................................... 6 Figure 1.3 Th cell lineage fidelity during differentiation. .............................................................. 7 Figure 1.4 Immunological cross-talk between the intestine and the lungs. .................................. 10 Figure 1.5 Mechanisms of type 2 immune responses during AAI. .............................................. 14 Figure 1.6 Dysregulation of Th1-driven immune responses during IBD. .................................... 19 Figure 1.7 The T. muris model of intestinal helminth infection. .................................................. 27 Figure 1.8 Overview of RA synthesis and regulation of gene expression. ................................... 30 Figure 2.1 Low dose intestinal infection with T. muris alters the lung immune microenvironment........................................................................................................................................................ 44 Figure 2.2 Lung CD4+ T cells upregulate IFN-γ expression but not IL-10 during low dose T. muris infection. ............................................................................................................................. 46 Figure 2.3 Low dose T. muris infection induces expression of IL-10 in lung myeloid cells. ...... 48 Figure 2.4 Low dose T. muris infection suppresses papain-induced allergic airway inflammation........................................................................................................................................................ 50 Figure 2.5 Depletion of CD4+ T cells does not reverse T. muris-mediated protection from papain-induced allergic airway inflammation. ......................................................................................... 52 Figure 2.6 Low dose infection with T. muris in Rag1−/− mice partially protects from papain-induced lung inflammation. .......................................................................................................... 54 Figure 2.7 Blockade of IL-10R in T. muris-infected Rag1−/− mice following papain challenge. 55 Figure 2.8 Time dependence of low dose T. muris-mediated suppression of house dust mite antigen-induced allergic airway inflammation. ............................................................................ 57 xii  Figure 3.1 Induction of Sca-1 expression and LSK cells in the BM during low dose T. muris infection. ....................................................................................................................................... 67 Figure 3.2 Low dose T. muris infection alters hematopoietic progenitor populations in the BM. 69 Figure 3.3 Alterations in BM hematopoietic progenitor populations during low dose T. muris infection is IFN-γ-dependent. ....................................................................................................... 71 Figure 3.4 Low dose T. muris infection induces an accumulation of IFN-γ+ T cells in the BM. . 73 Figure 3.5 Activated and central memory-like phenotype of IFN-γ+ T cells in the BM of low dose T. muris-infected mice. ......................................................................................................... 75 Figure 4.1 Cyp26b1 is dispensable for normal lymphoid development. ...................................... 83 Figure 4.2 Cyp26b1 is not required for nTreg cell development and suppressive function. ........ 85 Figure 4.3 Cyp26b1 limits iTreg and Th17 cell differentiation in vitro. ...................................... 87 Figure 4.4 Cyp26b1-deficient T cells fail to promote intestinal inflammation following adoptive transfer into Rag1−/− mice. ............................................................................................................ 89 Figure 4.5 Deficiency in Cyp26b1 does not alter expression of intestinal homing molecules on T cells. .............................................................................................................................................. 91 Figure A.1 Neutralization of IL-12 does not reverse T. muris infection-mediated protection from papain-induced allergic airway inflammation. ........................................................................... 128 Figure A.2 T. muris infection does not affect lung Treg cells. ................................................... 129  xiii  List of Acronyms and Abbreviations AAI: Allergic airway inflammation ADH: Alcohol dehydrogenase Ag: Antigen AMP: Anti-microbial peptide ANOVA: Analysis of variance APC: Allophycocyanin APC: Antigen presenting cell ATRA: All-trans retinoic acid BAL: Bronchoalveolar lavage CCR: Chemokine (C-C motif) receptor CD: Cluster-of-differentiation CD: Crohn's disease cDNA: Complementary deoxyribonucleic acid CFU: Colony forming unit CRABP: Cellular retinoic acid binding protein CRBP: Cellular retinol binding protein CTCM: Complete tissue culture medium Ctrl: Control Cyp26b1: Cytochrome P450 family 26, subfamily b, polypeptide 1 DC: Dendritic cell DMEM: Dulbecco's Modified Eagle's Medium DSS: Dextran sulfate sodium xiv  ELISA: Enzyme-linked immunosorbent assay FACS: Fluorescence-activated cell sorting FITC: Fluorescein isothiocyanate Foxp3: Forkhead box p3 GREAT: IFN-γ Reporter with Endogenous PolyA Tail HD: High dose HDM: House dust mite H&E: Hematoxylin and eosin IBD: Inflammatory bowel disease IFN-γ: Interferon γ Ig: Immunoglobulin IL: Interleukin ILC: Innate lymphoid cell ILC2: Type 2 innate lymphoid cell i.n.: Intranasal i.p.: Intraperitoneal iTreg: Inducible regulatory T cell KO: Knock out LD: Low dose LN: Lymph node LSK: Lineage−Sca1+cKit+ MHC: Major histocompatibility complex mesLN: Mesenteric lymph node xv  MPP: Multipotent progenitor mRNA: Messenger ribonucleic acid nd: Not detected ns: Not significant nTreg: Natural regulatory T cell OVA: Ovalbumin PAS: Periodic Acid-Schiff PBS: Phosphate buffered saline PE: Phycoerythrin PMA: phorbol 12-myristate 13-acetate PRR: Pattern recognition  RA: Retinoic acid RALDH: Retinal dehydrogenase RAR: Retinoic acid receptor RARE: Retinoic acid response element  RBC: Red blood cell ROR-γt: RAR-related orphan receptor-γt RXR: Retinoid X receptor Sca-1: Stem cells antigen-1 SEM: Standard error of the mean SLAM: Signaling lymphocytic activation molecule  Tbx21: T-box transcription factor Tbx21/Tbet Tc: Conventional T cell xvi  Teff: Effector T cell TGF-β: Transforming growth factor β Th: T helper Th1: T helper type 1 Th2: T helper type 2 Th17: T helper type 17 Tm: Trichuris muris TNF-α: Tumor necrosis factor α Treg: Regulatory T cell TSLP: Thymic stromal lymphopoietin UC: Ulcerative colitis Vert-X: IL-10-eGFP reporter WT: Wild type xvii  Acknowledgements  I would like to formally acknowledge the great support and guidance provided by my supervisor Colby Zaph. I thank the helpful feedback from my supervisory committee throughout my program - Mark Scott, Megan Levings, Jacqueline Quandt, and Georgia Perona-Wright. I would also like to thank Haydn Pritchard, my graduate program advisor, for providing me with support and advice.   The work performed was largely supported by operating grants provided by the Canadian Institutes of Health Research (CIHR). I am grateful for support provided by the University of British Columbia four-year doctoral fellowship.  I owe a lot to my fellow lab colleagues - Frann Antignano, Kyle Burrows, Menno Oudhoff, and Sebastian Scheer - for providing technical training and maintaining a wonderfully interactive and friendly work environment. I further thank the services and facilities provided by staff in the Biomedical Research Centre - Andrew Johnson & Justin Wong (UBC Flow), Takahide Murakami (Genotyping), Ingrid Barta (Histology), Helen Merkens & Rupi Dhesi (BRC Core), Krista Ranta, Leslie Rollins, & Weidong Yuan (Animal Unit). Thank you to the rest of the members of the BRC for making up such a great collaborative and diverse training environment.  Finally, I would like to thank my family for their unconditional support over the years. I especially thank my close friends that I met during my time at UBC for their friendship and many fun experiences we've shared together.1  Chapter 1: Introduction  1.1 Overview  The mucosal immune system is a vast network of tissues and immune cells that play an important role in host defense at organs that have a direct interface with the external environment. This environmental exposure consequently translates into contact with a diverse and large number of microorganisms and potential antigens. This warrants the need for appropriate immune responses against potential threats balanced with proper immunoregulation to prevent host tissue damage 1. Immunity at these barrier organs thus depends on a highly organized arrangement of immune cells to efficiently detect antigens from the outside and to relay these antigens to lymphoid tissues in order to mount appropriate immune responses. Consequently, T cells play a central role during protective immune responses against pathogens as well in maintaining tolerance to harmless antigens at mucosal sites 2. The critical role of T cells in this context is evidenced by the fact that dysregulation in T cell responses is often associated with chronic mucosal inflammatory diseases. Therefore, there is a need to better understand the factors that can control T cell function prior to or during the development of mucosal disease which may provide the impetus to develop newer therapies. This dissertation will focus on the regulation of T cell function by intestinally-derived factors - infectious stimuli and dietary immunomodulators - in the context of two distinct mucosal inflammatory diseases that affect the lung and intestinal tissues.   2  1.2 Organization of the mucosal immune system Mucosal tissues include the nasal/oral cavities, airways, gastrointestinal tract, and urogenital tract. These mucosal tissues are dynamic sites of immune activity and regulation which are important for maintaining overall homeostasis (Figure 1.1). In particular, barrier organs such as the intestine and lungs, which are directly exposed to the external environment over very large surface areas, have the unique challenge of mounting protective immune responses against potential pathogens while maintaining tolerance to innocuous antigens and commensal organisms. The mucosal-associated immune system is immense given that the gastrointestinal tract alone harbors the largest number of immune cells in the body 3.    3  Figure 1.1 Simplified schematic of mucosal immune regulation. Epithelial cells form a barrier against lumenal microbiota and external antigens. Goblet cells secrete mucins into the lumen forming a protective mucus layer. The microbiota composition is controlled by epithelial cell secreted AMPs. DCs extend their processes to sample lumenal antigens and migrate into inductive sites such as draining lymph nodes where they present antigen to naive T cells. Activated Teff cells can then migrate from the lymph node into the lamina propria of the mucosal site. In the lamina propria, immune regulation is maintained by Treg cells that can suppress DC and Teff responses to innocuous antigens. Lamina propria plasma cells secrete IgA that gets transported across the epithelial layer of the mucosa.   A single layer of mucosal epithelial cells provides a physical boundary between the internal tissues and the outside environment as well as contributing to innate immune responses 4. Depending on the anatomical location, epithelial cells can differentiate into several cell types (e.g. Paneth cells, goblet cells, absorptive cells, ciliated cells) with precise functions such as absorption, secretion, and immune surveillance. Within the epithelial layer, goblet cells secrete and maintain a protective mucus layer that coats all mucosal lumenal surfaces 5. Mucins are heavily glycosylated proteins and are the major gel-forming component of the mucus layer. Since humans harbor a natural microbiota that outnumber host cells ten-to-one 6, regulation of these commensal populations and defense against pathogenic bacteria can be mediated by the secretion of antimicrobial peptides (AMPs) such as β-defensins from epithelial cells 7. Within the epithelium are populations of intraepithelial lymphocytes that have an immune surveillance role by being able to rapidly release cytokines after encounter with antigens.8 Amid the subepithelial regions are dendritic cells (DCs) which sample antigens and can migrate to the draining lymph nodes to induce tolerance or orchestrate adaptive T and B cell responses 9. Other mucosal inductive sites can include organized lymphoid structures, such as Peyer's patches in the small intestine which are composed of overlying M cells that can directly sample lumenal antigens and contain antigen presenting cells (APCs) with distinct T cell zones and B cell zones 10.  4   Underlying the epithelium, the lamina propria contains a vast network immune cells which includes effector and memory populations of T and B cells, innate immune cell types (neutrophils, macrophages, mast cells) as well as non-immune cells such as fibroblasts 11. Plasma cells secrete immunoglobulin A (IgA) that traverses the epithelium, important in toxin neutralization and controlling microbial populations 12. Mucosa-associated granulocytic populations include eosinophils, and mast cells which are all involved in type 2 immune responses against parasites and allergens by releasing cytotoxic mediators, cytokines, and chemokines 13. Neutrophils are another granulocyte that function via phagocytosis, secreting antimicrobial factors, and by releasing extracellular traps in response to invasive microbial pathogens 14. Monocytes and macrophages also have diverse functions during mucosal immune responses with roles in both pro-inflammatory responses (phagocytosis, antigen presentation, cytokine release) and tissue repair responses, during the resolution phases of inflammation by producing anti-inflammatory cytokines such as IL-10 and regulating epithelial cell renewal 15.   In addition to these common immune cells, several populations of innate lymphoid cells (ILCs), lymphoid cells lacking antigen-specific receptors, have been recently identified that play major roles early on during all types of mucosal immune responses that complement the associated adaptive immune responses; for example, type 2 ILCs (ILC2) are a major source of interleukin (IL)-4, IL-5, and IL-13 that precede and shape subsequent adaptive type 2 responses 16. However, given the vastness and complexity of the mucosal-associated immune system, numerous cellular and molecular mechanisms that regulate mucosal immune homeostasis are yet to be fully characterized. Another challenge will be translating data from mouse studies (predominately cited in this dissertation) to better understanding the human mucosal immune system. 5   1.3 T helper cell differentiation and function  Mucosal immune responses and chronic inflammation invariably involve the induction of a variety of cluster-of-differentiation 4 (CD4)+ T helper (Th) cell subsets, differentiating in a context-specific fashion depending on the inflammatory milieu (Figure 1.2). Consequently, the balance between these Th cell subsets plays an important role in health and disease. Classically, naïve Th cells, upon encountering APCs and associated polarizing cytokine stimuli, undergo activation and differentiation into many possible effector Th cell lineages. Th cell lineage choice is influenced by co-stimulatory signals, antigen dose, and is maintained by specific transcription factors induced by downstream cytokine signaling. The Th1 lineage, induced by IL-12, is responsible for initiating cell-mediated immune responses against intracellular pathogens and is characterized by expression of the transcription factor Tbet and production of the pro-inflammatory cytokine interferon-γ (IFN-γ) 17. Th2 cells induced by IL-4, are involved in antibody-mediated humoral immunity, defense against helminths, and allergic immune responses, and are characterized by expression of the transcription factor GATA3 and mainly produce IL-4, IL-5, IL-13 17. Th17 cells, induced by transforming growth factor β (TGF-β) and IL-6, are important in the maintaining barrier function and driving responses against extracellular pathogens at mucosal sites; Th17 cells are controlled by the transcription factor RAR-related orphan receptor (ROR)-γt and can produce IL-17A, IL-17F, IL-21, IL-22, and IL-26 18. On the other end of the spectrum, regulatory T (Treg) cells, which include natural thymus-derived Treg  and inducible Treg cells (by TGF-β), are involved in suppressing immune responses and preventing uncontrolled immunopathology; Tregs are maintained by the master transcription factor forkhead box p3 (Foxp3), and can upregulate inhibitory surface molecules such as CTLA-6  4, outcompete effector T cells (Teff) for IL-2, and can produce the immunosuppressive cytokines TGF-β and IL-10 19.   Figure 1.2 Common Th cell lineage differentiation paradigm. Naive CD4+ T cells differentiate into several lineages depending on context-dependent immune stimuli. Th1 cells differentiate during responses to intracellular pathogens such as viruses, are maintained by the transcription factor Tbet and characteristically express IFN-γ, required for cell-mediated immunity. Th2 cells differentiate during allergic and anti-helminth immune responses, are maintained by the transcription factor GATA3, and produce the type 2 cytokines IL-4, IL-5, and IL-13. Mucosal Th17 cells differentiate in response to extracellular pathogens such as fungi and bacteria, are maintained by the transcription factor ROR-γt, and can secrete IL-17A and IL-17F. Treg cells, naturally arising from the thymus or induced from naive T cells, promote tolerance to self or other harmless antigens, are identified by the master transcription factor Foxp3, and can produce anti-inflammatory cytokines such as TGF-β and IL-10.    In order to mount the appropriate adaptive immune response to a given pathological stimulus, Th cell lineages are antagonistic toward the differentiation of other lineages 17 (Figure 7  1.3). Based on in vitro studies, Th cell lineage commitment is relatively stable and T cell clones maintain heritable gene expression patterns, largely controlled by epigenetic mechanisms 20,21; DNA can be methylated on CpG motifs within gene promoters and regulatory elements, repressing gene expression. In Th cells, inappropriate lineage-specific cytokine expression can be repressed by this process; for instance, during Th2 cell differentiation the Ifng promoter is methylated to block Th1 programs 22. Lineage-specific gene expression profiles are also enforced epigenetically the by post-translational modifications of histones which affect the accessibility of chromatin to transcriptional machinery 23.  Figure 1.3 Th cell lineage fidelity during differentiation. Th cell lineages are antagonistic to the differentiation of opposing Th cell lineages in order to maintain lineage fidelity. One possible mechanism of Th cell lineage fidelity is that lineage-associated cytokines directly block the 8  transcriptional programs needed for other Th cell lineages; for example, IFN-γ produced during Th1 cell differentiation directly interferes with the differentiation of Th2 and Th17 cells.   It is important to note that in vivo, these epigenetic mechanisms of Th lineage commitment may not be absolute due to plasticity observed between Th cell lineages that do not fit the classical paradigms of Th cell differentiation completely 24. Thus, a better understanding of the mechanisms that regulate Th cell differentiation and function in the context of mucosal immunity is key to better defining the pathogenesis of a variety of inflammatory diseases as well as identifying novel treatment strategies that may target these immune mechanisms.  1.4 Mucosal immune crosstalk  The mucosal immune system is interconnected since mucosal immunization at one site can provide immunity at other mucosal tissues 25,26. For example, gastrointestinal immune responses can induce systemic B cell-derived Ig secretion that can be reactive at other mucosal tissues such as the lungs 27. Other adaptive immune responses are also implicated in this mucosal crosstalk, highlighted by the challenges of administering protective vaccines to populations that have heavy enteric pathogen burdens interfering with immunity, particularly in underdeveloped countries 28. However, the mechanisms involved in cross-mucosal immune responses, especially with regards to Th cell-dependent responses, are poorly defined. Thus, a better understanding of this immune crosstalk is critically important in defining better strategies for treating mucosal inflammatory diseases as well as for being informative to the design of more effective mucosal vaccines.   Several experimental studies in mice and clinical reports suggest that direct immunological cross-talk can occur between the lungs and the intestine 29 (Figure 1.4). For 9  example, the intestinal microbiota in early life has been shown to play a role in determining susceptibility to allergic asthma later in life 30,31. Antibiotic-mediated depletion of the intestinal microbiota in perinatal mice results in enhanced allergic responses in the lung during adulthood 32. This is consistent with the fact that certain intestinal bacteria and their associated metabolites can induce specific T cell responses, such as polysaccharide A from Bacteroides fragilis promoting Treg cell responses 33. As well, there is evidence that trafficking of immune cells can occur directly between the lung and the intestine; lung DCs have the ability to prime and license lung-residing T cells to home toward intestinal tissues via specific homing receptors 34. Further, it has been shown that upon oral ingestion of antigens, intestinal epithelial cells can distally regulate allergic immune responses in the airways by manipulating IgA responses 35. Additionally, interstitial pneumonia has been reported to occur in mouse models of chronic colitis 36 and a subset of inflammatory bowel disease (IBD) patients have been reported to exhibit extraintestinal inflammation in the lungs 37. Overall, these findings suggest that local immune responses originating at one mucosal site can strongly impinge on immune responses at distal mucosal tissues. However, whether this gut-lung immune cross-talk, especially with regards to T cell-mediated responses, plays a definitive physiological role in health and disease still remains unclear.   10    Figure 1.4 Immunological cross-talk between the intestine and the lungs. Studies in the literature suggest that immunological crosstalk occurs specifically between the intestine and the lungs. The composition of the intestinal microbiota early in life can influence susceptibility to asthma in adulthood. Antigen-specific immune responses in the lungs can lead to DC-mediated imprinting of lung T cells to upregulate the intestinal homing receptor CCR9 which allows T cells to directly home from the lung to the intestine. Subsets of IBD patients have been found to exhibit extraintestinal inflammation in the lungs.     11  1.5 Mucosal inflammatory diseases  If normal mucosal immune homeostasis is perturbed, dysregulated inflammatory processes can manifest into chronic disease under the right conditions. As such, a variety of factors can contribute to the development of mucosal inflammatory diseases. For instance, genetics are known to play a role in the etiology of chronic inflammatory diseases, including allergic disease and IBD 38,39. However, since mucosal inflammatory diseases are often multifactorial and have become prevalent in a relatively short period of time, genetic factors alone cannot be the sole driver of disease. The fact that chronic inflammatory and autoimmune diseases tend to occur specifically in developed countries, environmental factors appear to be strongly determinant of conferring disease susceptibility to an individual, based on epidemiological studies; specific environmental factors include diet, exposure to microorganisms/pathogens, urban environments, and exposure to airborne pollution 40,41. Despite the heterogeneity of these mucosal diseases, there are commonalities to be found in terms of pathological features. For example, disruption in epithelial barrier function is associated with both asthma and IBD 1,42. There is also a prominent role for T cells in mediating chronic immunopathology during disease (further elaborated below with respect to asthma and IBD). Therefore, therapies targeting T cell-mediated mechanisms of disease may bypass the difficulties associated with treating these heterogeneous diseases.     12  1.5.1 Allergic airway inflammation  Asthma is a broad term that encompasses a heterogeneous group of related diseases that affect the airways. One of the most common forms is allergic asthma which occurs as a result of hypersensitivity to environmental antigens encountered in the airways 43. The incidence of asthma has been increasing at an accelerated rate globally, and is especially prevalent in young children 44. Asthma tends to occur in westernized and affluent countries 43, again further implicating a strong environmental component that affects disease risk. Key clinical symptoms of asthma include wheezing and shortness of breath, although there is a great deal of variability in the definitions of symptoms globally 45. The typical treatment involves the use of nonspecific inhaled glucocorticoids which often only provide short-term relief from exacerbations and do not necessarily treat the underlying causes of the disease 46. Further, given the heterogeneity of asthma, efficacious and targeted treatment options are generally very limited 47.  Immunologically, type 2 allergic airway inflammation (AAI), a key process involved in the development of asthma, has a complex and multifaceted pathology (Figure 1.5). AAI is dependent on early-responding innate immune cells, such as DCs, orchestrating an eventual adaptive Th2 cell response following exposure to aeroallergens 48. Allergens that directly damage the airway epithelium, often by proteolytic activity, can cause epithelial cells to directly release cytokines such as IL-25, IL-33, and thymic stromal lymphopoietin (TSLP) 49 - cytokines that can activate ILC2s during type 2 immune responses and activate DCs 50–52. Upon activation, ILC2s produce high levels of type 2 cytokines such as IL-5 and IL-13 53 - IL-5 is involved in the recruitment and activation of eosinophils into the airways, a pathogenic cell type that can release damaging mediators and cytokines during AAI 54; in parallel, IL-13 induces goblet cell hyperplasia and mucus overproduction 55. ILC2s have been shown to directly interact with other 13  immune cells such as CD4+ T cells 56. Th2 cells, induced by APCs upon presentation of antigens loaded on major histocompatibility complex (MHC)-II, play a key role in further propagating the allergic immune response by producing IL-4, IL-5, IL-13, and by promoting class switching to IgE in B cells 57. Mast cells and basophils upon activation by IgE can degranulate and release chemical mediators such as histamine and can also secrete type 2 cytokines 13.  During chronic AAI, tissue remodeling, fibrosis, and airway hyperresponsiveness can occur 43. Together, this suggests that there are numerous immune cell types with distinct roles in the pathogenesis of diseases involving AAI.    14  Figure 1.5 Mechanisms of type 2 immune responses during AAI. Allergens encountered in the airways can cause epithelial cell damage, leading to the release of alarmin cytokines IL-25, IL-33, and TSLP. These alarmin cytokines activate ILC2s that provide an early source of type 2 cytokines such as IL-5 which activates and recruits eosinophils and IL-13 which can stimulate goblet cell hyperplasia and mucus production. Eosinophils can release damaging mediators and cytokines and can also be involved in tissue remodeling by airway smooth muscle cells. DCs in this type 2 inflammatory environment can direct the differentiation of CD4+ T cells toward Th2 cells which further mediate AAI by primarily producing the type 2 cytokines IL-4, IL-5, and IL-13. Th2 cells also promote allergen-specific IgE class-switching in B cells which can lead to the activation of IgE-responding granulocytes such as mast cells and basophils which degranulate and release harmful inflammatory mediators such as histamine, prostaglandins, and leukotrienes.   As mentioned before, the balance between Th cell subsets is a key aspect of determining the outcome of specific mucosal immune responses. Consequently, Th2 cell-dependent responses during AAI can be antagonized by generating a counteracting Th1 cell response or by the induction of suppressive Treg cell-driven immune responses 58–60. Mechanistically, this Th2 cell lineage antagonism can occur at the epigenetic level, demonstrated by the finding that the Il4 promoter is heavily marked with the repressive H3K27me3 mark in Th1 cells 61. Another strategy is to target Th2 cell-driven effector responses such as through antibody neutralization of IL-13 which clinically resulted in improved lung function in asthmatics 62. However, there is still much to be known mechanistically about how mucosal-associated Th cell-responses are controlled during homeostasis and AAI. Thus, a better understanding of the factors that modulate Th cell differentiation is critical in better understanding the mechanisms of AAI.  Treg cells have a significant role in AAI since they have the ability to suppress the function of effector T cells, including Th2 cells. This is supported by the fact that patients harboring mutations in the FOXP3 gene have enhanced susceptibility to allergic disease and asthma 63. Treg cells are also involved in the induction of airway tolerance, another mechanism critical in preventing AAI 64. In mouse models of AAI, the induction of allergen-specific Treg 15  cells accompanies Th2 cell responses and is necessary to limit the extent of airway disease 65. Additionally, treatments such as allergen-specific immunotherapy (involving progressively increased doses of allergens to induce tolerance) and corticosteroids involve the promotion of Treg cell responses 66. Treg cell suppressive functions during asthma can include secretion of IL-10 and TGF-β, direct cell contact-mediated suppression and outcompeting effector cells for growth factors 67; however, there have been known inconsistencies between in vivo and in vitro experimental data in determining the relevant suppressive mechanism in the context of AAI. Further, there is controversy in distinguishing between the functions of thymus-derived (nTreg) and inducible Treg (iTreg) cells in vivo 66.  1.5.1.1 Experimental models of AAI  A variety of well-characterized murine models of AAI have been established to dissect the cellular and molecular mechanisms of type 2 responses in the airway. All of these AAI models have their unique advantages and disadvantages.  One of the most well-used murine models of AAI is the chicken egg-derived ovalbumin (OVA)-alum model which involves systemic Th2 cell-driven immunization with OVA-alum and several subsequent airway challenges with OVA 68. Advantages of using this model include a robust Th2 cell inflammatory response in the lung and a variety of available tools such as transgenic mice with T cell receptors restricted to OVA and OVA-MHC tetramer reagents to track antigen-specific T cell responses. However, OVA is not a natural allergen for humans and the use of an adjuvant in the model diminishes its physiological relevance.  Other AAI models offer a more natural means of allergen sensitization such as the papain model of AAI. Papain is a cysteine protease derived from Papaya, which was commonly used as 16  a meat-tenderizer and was originally reported as a workplace respiratory allergen that caused asthma in production factory workers 69. When papain is intranasally administered to mice daily over three consecutive days, epithelial damage induces the activation of ILC2s and results in acute type 2 inflammation, characterized by an influx of eosinophils into the airways 70. The papain model of AAI is useful in studying the early stages of type 2 responses in the lung, is technically simpler than the OVA model, and provides a natural route of allergen exposure. However, papain is not a common allergen for humans and its use as a model is limited to the early stages of AAI, unless more chronic exposure models are employed.  In contrast to OVA and papain, house dust mite (HDM) antigens (from Dermatophagoides pteronyssinus) are very common allergens implicated in human asthma 71. HDM antigens also contain proteases similar to papain in stimulating ILC2-dependent type 2 immune responses in the lung 72. HDM antigen is delivered intranasally during a sensitization phase and subsequent challenge phase of the model which results in an adaptive Th2 cell-driven AAI. Therefore, out of all of three models of AAI, the HDM model provides a more complete model of asthma that is physiologically relevant in terms of exposure route and more translatable to human disease. However, disadvantages include the fact that HDM antigen is a relatively crude extract containing a complex mixture of antigens which complicates the interpretation of disease mechanisms. In addition, variability during the manufacturing of HDM antigen, such as differing endotoxin levels, can affect reproducibility at given doses.  In summary, while there is a variety of well-established experimental models of AAI that individually, have their unique set of advantages, a combination of these models is the best means to study disease mechanisms while providing the most relevance to human disease.  17  1.5.2 Inflammatory bowel disease  IBD is a broad term that includes several distinct inflammatory diseases affecting the gastrointestinal tract with the two major types being Crohn's disease (CD) and ulcerative colitis (UC). Generally, IBD involves specific genetic susceptibilities being precipitated by environmental factors leading to dysregulated inflammatory responses in the gut 73. Similar to allergic disease, IBD incidence is highest in westernized countries, with Canada particularly having one of the largest populations of affected individuals 74. Also similar to allergic diseases, the incidence of IBD is increasing at an accelerated rate globally 75. What distinguishes between the two major forms of IBD is that CD can occur at any part of the entire gastrointestinal tract (from the oral cavity to the anus) while UC is localized to only affect the colonic mucosa. CD is characterized by a fluctuation between relapsing disease and states of remission. Clinical features of CD include segmented areas of chronic inflammation that span the whole intestinal wall, granuloma formation, deep fissuring ulcers, and the potential for developing fistulas in severe cases 76. Other major gross features of CD are lumenal narrowing, intestinal adhesions, and creeping fat 77. Common symptoms of CD include abdominal pain, diarrhea, fever, and weight loss 76. In contrast to CD, UC is continuous along the length of the colon and affects only the mucosa layer. Clinical features of UC include ulceration, edema, and hemorrhaging along the colon 76. UC patients typically present with abdominal pain, recurrent bloody/mucoid diarrhea, increased frequency of bowel movements, and anemia if bleeding is severe 78. Distinguishing histological features of UC include, inflammatory cell infiltrate in the lamina propria, neutrophilic infiltration of the colonic crypt epithelium, and disruption of crypt architecture 79.   The immunology of IBD involves a disruption in barrier function and loss of tolerance to the intestinal microbiota with pro-inflammatory responses driven by both innate and adaptive 18  mucosal immune cells 73 (Figure 1.6). Similar to other mucosal inflammatory diseases, T cells play a central role in disease progression. Under conditions of immune homeostasis in the steady state, tolerance to commensal organisms is primarily maintained by Tregs, regulatory B cells, and various other innate cells with immunosuppressive functions such as production of the anti-inflammatory cytokines IL-10 and TGF-β 80–82. However, during IBD there is an expansion of myeloid cells such as monocytes, macrophages and DCs. Through the sensing of microbial antigens by pattern recognition receptors (PRRs), these myeloid cells produce chemokines as well as the pro-inflammatory cytokines IL-1β, IL-6, IL-12, and tumor necrosis factor-α (TNF-α) 2.   19  Figure 1.6 Dysregulation of Th1-driven immune responses during IBD. IBD is characterized by immune responses against the naturally-occurring intestinal microbiota. A breach in the barrier function of the intestine can lead to the activation of DCs and macrophages by the sensing of microbial stimuli through PRRs. This leads to the production of inflammatory cytokines such as IL-1β, IL-6, IL-12, and TNF-α which can stimulate ILC1 cells to produce IFN-γ. Th1 cells can also differentiate under these pro-inflammatory conditions to produce elevated levels of IFN-γ which can further lead to tissue damage, completing a viscous cycle of chronic inflammation. Treg cells which are important in preventing this Th1 cell response to the microbiota may be defective in the context of disease.  The central role of TNF-α in mediating IBD is illustrated by the widespread use of TNF-α neutralizing drugs as treatments 83. With regards to other innate cells, intestinal ILC populations are also expanded during IBD which can provide cytokines and direct lymphocyte recruitment 84. During chronic inflammation, this can lead to polarized Th cell responses, with Th1 cells being the predominant subset with enhanced IFN-γ production in the lamina propria during CD 85. Conversely, UC is Th2 cell-driven disease with levels of IL-13 being elevated in UC patients that is thought to cause epithelial cell barrier disruption 86. Th17 cells are also increased in both CD and UC patients compared to healthy controls, but whether or not they are a pathogenic cell population has been a controversial issue in the field. Th17 cells are considered a plastic lineage and studies in mice have shown that these cells can acquire Th1-associated features during the pathogenesis of colitis 87. However, antibody neutralization of IL-17a in patients has been reported to be ineffective 88, and other mouse studies have reported a protective function for Th17 cells 89. Thus, the development of IBD and specific clinical manifestation of disease subtypes hinges heavily on Th cell differentiation and function in the intestine.  As mentioned above, deficiencies in Treg cell number and function is also associated with increased susceptibility to IBD 90. In many mouse models of colitis, the depletion of Treg cells results in exacerbated disease 91. In mice, it is well established that Treg cell-intrinsic 20  mutations that affect suppressive function are known to contribute to increased inflammatory disease severity 92. However, since the number Treg cells tends to increase in the intestine of IBD patients and that Treg cells from these patients have been shown to have comparable suppressive function to healthy controls ex vivo, Treg cell-extrinsic factors must play an important role in the failure of Treg cell-mediated suppression during IBD 90. For example, the inflammatory cytokine environment in the intestine during IBD can destabilize Treg cells and can cause effector T cells to be resistant to the suppressive effects of Treg cells 93. However, despite strong evidence from mouse models, it is very difficult to determine the precise in vivo role of Treg cells during IBD, since human activated effector T cells also express Foxp3 94. Additionally, studies investigating Treg cell versus effector T cell function during IBD are often removed from the intestinal microenvironment 90.  1.5.2.1 Experimental models of IBD  A great deal of what is known about the immunology of IBD has derived from data generated using experimental animal models of colitis. These include chemically-based, genetic, and cell-mediated models of IBD.  One method to induce colitis in mice is by means of chemical insult of the intestinal mucosa, resulting in barrier disruption. This is commonly achieved by administering dextran sulfate sodium (DSS) in the drinking water of mice which damages the intestinal epithelium. Over one week of constant exposure to DSS, treated mice develop acute colitis that results in gross morbidity typically measured by loss of total body weight, diarrhea, bloody stool, and shortened colon length 95; histologically, the colonic epithelium becomes eroded with a substantial loss in crypt architecture, loss of goblet cells, and a prominent influx of inflammatory 21  cells after DSS treatment. The pro-inflammatory cytokines upregulated during DSS colitis are similar to those commonly increased during human IBD such as TNF-α and IFN-γ 96. While the DSS model offers a rapid means to induce colitis, it is still relatively complex in terms of deciphering the relative contribution of all immune cells involved in the disease model.  Targeted gene deletion is another method to induce a spontaneous form of colitis in mice. When genes involved in immune regulation are knocked out, immunopathology can develop. For example, mice deficient in genes encoding the anti-inflammatory cytokines TGF-β and IL-10 will go on to develop spontaneous colitis over time 97. These genetic models depend on the presence of intestinal microbiota as is the case for IL-10-deficient animals, with specific strains of bacteria being required to precipitate the development of colitis 98. Therefore, the use of genetically manipulated mice can be useful in assessing the role of genes potentially implicated in the development of IBD, but are generally limited due to the inherent variability and possibility for off-target effects of the mutations.  In contrast to chemically-based and genetic models, the T cell transfer model of colitis is a powerful mouse model of IBD used to study more specific, cell-intrinsic mechanisms of disease. In this system, naive CD4+ T cells (CD25−CD45RBhi) are isolated from donor mice by fluorescence-activated cell sorting (FACS) and transferred into immunodeficient recipient mice 99; after approximately five weeks following T cell transfer, recipient mice develop severe wasting disease that results in significant weight loss and morbidity associated with the development of colitis. In the absence of Treg cells, the transferred T cells react against the intestinal microbiota of the recipient mice that causes chronic intestinal inflammation 100. This is characterized by differentiation of the transferred cells into Th1 cells producing increased levels of IFN-γ, and blocking this Th1 response is sufficient to prevent disease 101. Further, this model 22  offers the powerful advantage of being able to use genetically manipulated donor mice, such as gene knock out (KO) strains, to test the T cell-intrinsic role of any gene on the development of colitis. However, the requirement for immunodeficient recipient mice limits the model's translatability to human IBD. Regardless, the T cell transfer model of colitis offers the best means to study specific factors that control Th cell differentiation in the context of chronic intestinal inflammation.  1.6 Modulation of mucosal immune responses  A major goal for the development of new therapies to treat mucosal inflammatory diseases is to identify immunomodulatory pathways that mediate protection against disease. Amidst the plethora of environmental factors that can affect the development of mucosal diseases, this dissertation will focus on two examples that can potently influence Th cell differentiation and offer further mechanistic insight into the regulation of mucosal immune responses.  1.6.1 Intestinal helminth infections  Parasitic organisms have had a long co-evolutionary history with their mammalian hosts. Notably, helminth parasites have been infecting human beings well before any known forms of  recorded history 102. Helminths are a diverse group of parasitic worms divided into two major phyla, the nematodes (round worms) and the platyhelminths (flat worms) 103. Intestinal worms, infect approximately 2 billion people world-wide, are mainly prevalent in underdeveloped countries, and are considered agents of the most neglected of tropical diseases 103. Infections with these intestinal worms in malnourished individuals, especially young children whom are more 23  immunologically vulnerable, can cause stunting of growth and impaired mental development 104. However, among affected adult populations, a large proportion of chronically infected individuals are asymptomatic which most likely represents a consequence of helminth-induced immune regulation in these hosts 105. One important aspect of this host-parasite interaction is the Th cell responses elicited by the helminth infection.  Host anti-helminth responses typically require the induction of immune responses to mount either direct killing of the helminth or physical expulsion of the helminth from the mucosa - primarily through increased epithelial cell turnover and enhanced mucus production 106. The latter strategy is conceptually more beneficial to the host as it limits the potential for off-target tissue damage. Many of these anti-helminth responses hinge heavily on the development of Th2 cell responses. In fact, the Th2 arm of the adaptive immune system has been shown to have been under intense evolutionary pressure induced by these helminth parasites 107. In the case of helminth parasites that are able to chronically persist in the host, parasite-induced immunoregulation is often evoked to prevent excessive host tissue damage as a means to tolerize the presence of the parasite when anti-helminth responses fail 108,109. This is due to parasitic helminths having evolved sophisticated strategies to evade host immune responses and promote tolerance as a survival mechanism. Interestingly, this helminth-induced immune regulation can lead to bystander effects on the host's susceptibility to autoimmune and allergic disease as posited by recent iterations of the "Hygiene Hypothesis" which suggests that westernized countries are more susceptible to inflammatory disease due to the elimination of helminth infections from increased hygiene 110. In other words, this suggests that normal immune homeostasis was shaped in the context of tolerating these intestinal parasites and their removal may perturb a dynamic immune equilibrium established through a long co-evolution with these 24  organisms. Epidemiological evidence supports these hypotheses in addition to a variety of animal models of intestinal helminth infection showing protection against several inflammatory diseases such as autoimmunity and allergic disease 111,112 (Table 1.1). This helminth-induced immunomodulation has attractive therapeutic potential as clinical trials using live intestinal helminth infection with the pig whipworm Trichuris suis has had some success in treating both IBD and multiple sclerosis 113,114. However, live infections need not be the primary means of therapy as isolated helminth-derived molecules or the discovery of novel helminth-induced immunoregulatory pathways could also serve this purpose 115.  Intestinal helminth species Disease modulation Heligmosomoides polygyrus asthma116, diabetes117, colitis118, multiple sclerosis119 Schistosoma mansoni asthma120, colitis121, diabetes122, multiple sclerosis123 Taenia crassiceps diabetes124 Trichinella spiralis asthma125, diabetes126, colitis127, multiple sclerosis128 Nippostrongylus brasiliensis asthma129  Table 1.1 Helminth-mediated protection against mouse models of inflammatory disease.    In terms of specific mechanisms, intestinal helminths employ a broad range of immunomodulatory capacities which often converge on influencing CD4+ Th cell differentiation and function 130. This modulation of Th cell responses have been shown to result in bystander suppression of various mouse models of inflammatory diseases; for example, mice infected with Heligmosomoides polygyrus are protected from experimental models of asthma and type 1 diabetes primarily by inducing Treg cell and IL-10-dependent suppressive immune responses 116,131. Another common immunomodulatory strategy that helminths use is directly interfering 25  with antigen presentation by secreting protease inhibitors such as cystatins which can affect antigen processing, ultimately impairing T cell priming 132. Taken together, the mechanisms by which helminth parasites modulate mucosal Th cell responses is an attractive research area that may inform not only on mucosal disease pathogenesis but also reveal potent immune pathways that could be exploited for therapeutics.  1.6.1.1 The Trichuris muris model of intestinal helminth infection  A large variety of intestinal helminths have been adopted as infection models in the laboratory. All species of intestinal helminths elicit their own unique host immune response and this translates into a vast selection of experimental tools to study particular immunological mechanisms. However, the biology of a parasitic organism must be taken into account since different stages in its life cycle and invasiveness into tissues outside of the intestine can dramatically influence the host immune response.  Intestinal whipworm parasites belonging to the genus Trichuris have served as an appropriate helminth model to study both anti-helminth Th2 responses and parasite-induced host immunoregulation during chronic infections. Trichuris spp. have very specific mammalian host species but are very similar in morphology and life cycle; for example, T. trichiura is the human-infecting whipworm that currently infects greater than 800 million people globally 133. In contrast, T. muris  is a naturally-occurring nematode parasite in wild rodent populations that has been isolated for infections in laboratory mice 134 (Figure 1.7). Trichuris spp. also have a very simple life cycle consisting of larval development in the cecum of a single primary host, molting into mature worms, shedding of immature eggs into the feces, embryonation of the eggs in the soil, and finally ingestion of mature eggs in a new host to complete the cycle 135. Trichuris spp. 26  are also intestinally-restricted which further justifies their use as an experimental model to study localized mucosal immune responses.  Immunity to T. muris in mice hinges heavily on mounting an anti-worm Th2 cell response that ultimately drives the coordinated intestinal epithelial cell turnover, goblet cell hyperplasia/mucus production, and smooth muscle contractility needed to physically expel the worms 136; however, if an inappropriate Th1 cell response occurs in response to T. muris infection, the worms will persist leading to chronic intestinal infection. Consequently, due to genetic differences, all strains of laboratory mice can be classified as being either resistant (Th2-cell mounting) or susceptible (Th1 cell-mounting) to T. muris infection; wild type C57BL/6 mice are resistant to T. muris infection while AKR mice are susceptible 137. This immune dichotomy is a powerful tool to assess the role of any gene on the balance between Th2 and Th1 cell responses in the intestine: for instance, mutant mice on the C57BL/6 background can be assessed for disruption in Th2 immune responses 138. Another added level of flexibility with the T. muris system is that modulating the infection dose will affect the immune outcome; for example, giving a high dose (>200) of T. muris eggs to C57BL/6 mice will result in worm clearance while giving the same mouse a low dose (<30) of eggs will result in a chronic Th1 cell-driven infection139. Chronic infection with T. muris also results in host immune suppression, such as the induction of IL-10, which mitigates immunopathology and morbidity in mice 140. This dose-dependent effect is hypothesized to be due a threshold of antigen load that is required for optimal Th2 responses 136. Thus, the T. muris system of intestinal helminth infection is a versatile and highly informative infection model to study factors that control mucosal Th2 vs. Th1 cell responses in the intestine.  27   Figure 1.7 The T. muris model of intestinal helminth infection. Wild type C57BL/6 (B6) mice can either be infected with a low dose or high dose of T. muris eggs by oral gavage which will result in two different immunological outcomes. B6 mice infected with a low dose (~30) of worms will mount a Th1 cell-driven anti-T. muris intestinal response which is non-productive in expelling the worms, resulting in a chronic intestinal infection. Conversely, mice infected with a high dose (~200) of worms mount a Th2 cell-driven anti-T. muris response that is completely effective at expelling all of the worms. Therefore, a Th1/Th2 cell dichotomy can be established for any given strain of mice on the B6 genetic background.      28  1.6.2 Retinoic acid  Vitamin A is an essential nutrient and is primarily found in the form of dietary retinol and carotenoids 141. Retinoids are critical in many physiological processes by regulating cellular proliferation, differentiation, and apoptosis 142. Retinoic acid (RA) is a vitamin A metabolite that plays a critical role during embryonic development 143 but also has important immunomodulatory functions after development 144. What highlights RA’s critical importance to the immune system is that vitamin A deficiency can lead to profound immunological impairments in children, causing increased susceptibility to infections, which occurs in countries where malnutrition is a problem 145. Vitamin A deficiency also impairs protective immunizations in children as a result of depressed immune responses to vaccines 146. As RA is highly enriched in the intestine, vitamin A deficiency has a dramatic effect on intestinal immune responses and physiology with affected children having a higher risk for gastroenteritis 147. These findings are confirmed by the fact that vitamin A supplementation can ameliorate this associated morbidity 148. In this context, RA has an important role in intestinal immune homeostasis by being a key factor involved in the induction of oral tolerance to innocuous antigens 149. Further, RA promotes epithelial barrier function, controls intestinal permeability 150 and is required for the secretion of IgA by intestinal plasma cells 151.  Before synthesis, RA precursors such as retinol are transported in the blood by binding to serum albumins and retinol binding proteins 152. Retinoids are generally stored in the form of retinyl esters in the liver 142. Upon entering circulation, these retinyl esters are hydrolyzed into retinol 153. Within cells, retinol binds to cellular retinol binding proteins (CRBPs) and is converted into retinal by alcohol dehydrogenase (ADH) 153. Mucosal biosynthesis of RA from retinal is a tightly controlled process, largely restricted to intestinal CD103+ DCs, intestinal 29  macrophages, intestinal epithelial cells and mucosal stromal cells in the mesenteric lymph node (mesLN) which selectively express the required enzymes, notably retinal dehydrogenase (RALDH) to synthesize RA from retinal 153,154. At other mucosal sites such as the lung, alveolar macrophages possess the ability to synthesize RA 155. The predominant stereoisomer synthesized by this process is all trans-RA (ATRA) 156. Transport of RA into the nucleus is facilitated by cellular RA binding proteins (CRABPs) where it performs its transcriptional regulatory functions 157.  Functionally, RA binds nuclear RA receptors (RARs) and retinoid X receptors (RXRs) that form heterodimers which transcriptionally regulate genes that contain specific RA response elements (RAREs) 158 (Figure 1.8). There are three isoforms of RARs, RARα, RARβ, and RARγ, and the three are differentially expressed in different cell types, with both CD4+ and CD8+ T cells primarily expressing RARα 159. One important immunological function of RA is the regulation of Th cell responses. For instance, through the paracrine secretion of RA, intestinal DCs can modulate activation and imprint tissue homing specificity on CD4+ T cells by upregulating the gut-specific receptor, chemokine (C-C motif) receptor 9 (CCR9) and integrin α4β7 on the T cells 160. In parallel to controlling intestinal T cell homing, RA has also been shown to be required for the migration of ILC1 and ILC3 into the intestinal tissues through very similar mechanisms 161. RA can induce both immune tolerance in the steady-state and effector responses during inflammation. RA produced by DCs has been shown to strongly promote the differentiation of iTreg cells in synergy with TGF-β 162 and limits the plasticity of Treg cells such as by blocking conversion to Th17 cells by upregulating the microRNA miR-10a 163. However, under pro-inflammatory conditions, RA can modulate the migration and function of Th17 cells in the intestine 164. Furthermore, it has been demonstrated that RA signaling occurs in T cells 30  during the early stages of an inflammatory response 165, suggesting that RA may be required for effector T cell function. Indeed, during infection with the intracellular protozoan parasite, Toxoplasma gondii, optimal Th1 effector cell responses absolutely require RA signaling 166. Thus, RA signaling is critical for both effector and regulatory T cell functions.   Figure 1.8 Overview of RA synthesis and regulation of gene expression. Retinol is converted to retinal by ADH which is then subsequently converted to RA via RALDH. RA may either be oxidized into inactive polar metabolites by Cyp26 enzymes, or transported into the nucleus via CRABPs. RXR/RAR heterodimer co-repressor complexes bind RARE target sequences of DNA. Upon RA binding, the complex becomes co-activating, allowing the transcription of RA-regulated genes.   31  1.6.2.1 RA metabolism in T cells  Despite the importance of RA signaling in T cells, very little is known about the molecular mechanisms that control RA bioavailability, signaling and metabolism in T cells and how these processes ultimately affect T cell differentiation and function. The cytochrome P450 family 26 (Cyp26) enzymes catalyze the oxidation of RA into inactive polar metabolites and are absolutely required in preventing the teratogenic effects of excessive RA during embryonic development 167.  Cyp26 enzymes are induced by RA and therefore act as negative regulators of RA signaling 168. Cyp26a1, Cyp26b1, and Cyp26c1 are the main enzymes involved in RA degradation though it has been shown from mouse studies that Cyp26b1 is the primary negative regulator of RA responsiveness in T cells 169; Cyp26b1 is highly induced in the presence of RA and is downregulated by the cytokine TGF-β1 in CD4+ T cells and modulates RA-dependent expression of the gut-homing receptor CCR9 on T cells 169. Thus, regulation of RA signaling by Cyp26b1 likely plays a central role in T cell function.  However, the specific role of Cyp26b1 in T cells has not been investigated in vivo in the context of mucosal inflammation.  1.7 Rationale, hypothesis and specific aims  The regulation of Th cell-dependent responses has thus far been shown to be critical for overall immune homeostasis at mucosal sites. A disruption in the balance of Th cell responses is clearly evident in the lung during AAI and in the intestine during IBD. Both of these disease archetypes have strong environmental factors that contribute to disease susceptibility. Infections and dietary supplements are two of many such external factors that can affect overall mucosal immune homeostasis. However, much is still yet to be elucidated about how these factors control Th cell differentiation in the context of these mucosal inflammatory diseases. 32   While intestinal helminth infections are correlated with immune hyporesponsiveness in populations within endemic regions 170, there is still little mechanistic insight into how these infections alter Th cell-dependent responses that could contribute to protection against allergic and autoimmune diseases. Further, since mucosal immune responses are not locally restricted to a particular organ, how intestinally-restricted helminth infections affect distal mucosal Th cell responses is still an important question that needs to be addressed. In addition, the systemic impact of these infections, such as on hematopoiesis in the bone marrow (BM), is also often neglected in these contexts. Thus, there is a need to better understand the effects of intestinal helminth infections systemically and on distal mucosal Th cell responses, such as Th2 cell-driven responses in the lung during AAI.  RA has a clear role in both regulatory and effector T cell responses, yet the factors that control RA-mediated signaling is not well-defined in vivo. There is also evidence indicating that helminth-infected individuals have associated nutritional impairments such as the reduced absorption of vitamin A 171; since these helminths often evade host immunity, it is conceivable that they can also alter RA-mediated processes as a means of immunomodulation. Likewise, the role of the RA-metabolizing enzyme Cyp26b1, while being strongly induced by RA as a negative feedback mechanism, has no previously defined role in Th cell differentiation and function during mucosal disease processes. Since these factors have such a potent effect on T cell biology, there is a need to better characterize the role of intestinal helminth infections and Cyp26b1 in the context of T cell-driven responses during mucosal disease.   33   Based on the above rationale, I hypothesize that Th1 cell-driven intestinal helminth infections can impinge on distal Th2 cell responses and that RA metabolism can modulate mucosal T cell responses during homeostasis and disease.   I have devised three specific aims to test this central hypothesis:  1) To assess the impact of intestinal helminth infections on the lung mucosal Th cell responses. The T. muris model of intestinal helminth infection will be used to determine the effect of infection on distal Th cell responses in the lung and its impact on the development of AAI.  2) To investigate the systemic effects of intestinal helminth infections. The T. muris infection model will be used to assess alterations in hematopoiesis in the bone marrow during infection.  3) To determine the role of the RA-metabolizing enzyme, Cyp26b1, on Th cell function during mucosal immune responses. The role of Cyp26b1 will be characterized in terms of T cell development, differentiation, and in vivo function using the T cell transfer model of colitis. 34  Chapter 2: Alteration of the lung immune microenvironment by the intestinal helminth Trichuris muris  2.1  Overview  The development of AAI in the lungs depends on type 2 immune responses after exposure to allergens. While it is known that mucosal immune crosstalk can occur between the intestine and the lungs 34, it is unclear how Th cell responses originating in the intestine can impinge on AAI-mediated pathology. Previous studies have largely attributed Treg responses as being responsible for gut-to-lung suppression of AAI 172. However, it is not known whether chronic Th1 cell-driven intestinal helminth infections, through this mucosal crosstalk, can directly affect the lung microenvironment. To address this, we employed the T. muris model of chronic intestinal helminth infection and characterized the distal immune effects on the lung. Using this model, we found that after the establishment of a chronic Th1-driven intestinal infection, a robust Th1 cell response is detectable in the lungs in parallel with the induction of myeloid-derived anti-inflammatory IL-10. Despite these immune effects, no overt lung pathology was found in mice chronically infected with T. muris. To assess the impact of these changes to the lung microenvironment, we exposed infected mice to protease allergens and found that chronic T. muris infection completely blocks the development of AAI. Investigating the immune mechanisms of this infection-induced protection from AAI, we identify non-redundant roles for innate and adaptive immune cells in our model.  35  2.2 Materials and methods 2.2.1 Mice  C57BL/6J, GREAT (IFN-γ Reporter with Endogenous PolyA Tail), and Rag1−/− mouse strains were originally purchased from the Jackson Laboratories (Bar Harbor, ME).  Vert-X (IL-10 eGFP reporter) mice, originally generated by Christopher Karp 173, were re-backcrossed on the C57BL/6 background and provided by Dr. Georgia Perona-Wright (UBC). All mice were bred and housed in specific pathogen-free animal facilities at the UBC Biomedical Research Centre.    2.2.2 Propagation of T. muris eggs, worm antigen, and oral infections  T. muris eggs were propagated by first infecting several immunodeficient Rag1−/− mice with 300 embryonated eggs per mouse, constituted in 300 μl sterile mouse drinking water, delivered by oral gavage using a ball-tipped feeding needle. After 35 days of infection, which allows for the development of fully mature worms, the chronically infected Rag1−/− mice were euthanized by CO2 administration and subsequent cervical dislocation. Intact adult worms were physically removed from the cecums and proximal colons of the mice and all worms were pooled in a tissue culture dish containing Dulbecco's Modified Eagle's Medium (DMEM) supplemented with penicillin and streptomycin. The pooled worms were incubated at 37°C to enable mating and release of eggs overnight. Worms were then physically removed and the remaining culture was centrifuged to pellet worm eggs. The supernatant of the worm culture was collected for protein dialysis with phosphate buffered saline (PBS) to yield T. muris antigen. Worm eggs were filtered and then washed several times with sterile mouse drinking water, sealed, and stored in the dark at room temperature for six weeks to allow for the eggs to fully 36  embryonate. The concentration of eggs was determined by counting embryonated eggs in a known volume using a dissecting microscope (Olympus, Tokyo, Japan).  Mice were infected with 30 hand-counted, embryonated T. muris eggs by oral gavage to induce chronic intestinal infection. For acute T. muris infections, mice were orally gavaged with 200-300 embryonated eggs. Most infections were carried out over a period of 21 days before sacrifice, as previously performed using this infection model 174. Sacrificed mice were assessed for worm burdens by manually counting worms in the cecums using a dissecting microscope.  2.2.3 Induction of acute lung inflammation with papain  The papain model has been previously described 70. Mice were anesthetized under aerosolized isoflurane and immediately instilled with 10 μg of papain from Papaya latex (Sigma, St. Louis, MO) intranasally (i.n.) in 40 μl of sterile PBS on days 18, 19, and 20 post-infection with T. muris. The day after the last instillation of papain, mice were injected intraperitoneally (i.p) with 2,2,2-tribromoethanol (avertin, Sigma), tracheas were cannulated, and bronchoalveolar lavages (BAL) were performed using 3 x 1 ml of sterile 10% fetal bovine serum in PBS. BAL fluid was then red cell-lysed using ammonium chloride buffer. Lung tissue was digested in 200 U/ml collagenase Type IV (Sigma) for 1 hour, red cell-lysed, and centrifuged in a 30% percoll solution to purify leukocytes. BAL fluid and lung cells were analyzed by flow cytometry.     37  2.2.4 Antibodies and flow cytometry  Absolute numbers of cells were determined via hemocytometer or with latex beads for BAL fluid samples. Intracellular cytokine staining was performed by stimulating cells with phorbol 12-myristate 13-acetate (PMA), ionomycin, and Brefeldin-A (Sigma) for 5 hours and fixing/permeabilizing cells using the eBioscience (San Diego, CA) IC buffer kit. All antibody dilutions and cell staining were done with PBS containing 2% FCS, 1 mM EDTA, and 0.05% sodium azide. Fixable Viability Dye eFluor 506 was purchased from eBioscience to exclude dead cells from analyses. Prior to staining, samples were Fc-blocked with buffer containing anti-CD16/32 (93, eBioscience) and 1% rat serum to prevent non-specific antibody binding. Fluorescein isothiocyanate (FITC)-conjugated anti-neutrophil Ly6B (7/4) was purchased from Abcam (Cambridge, MA). Phycoerythrin (PE)-conjugated anti-Siglec F (E50-2440) and allophycocyanin (APC)-conjugated anti-Ly6G (1A8) were purchased from BD Biosciences (San Jose, CA). FITC-conjugated anti-IL10 (JES5-16E3); PE-conjugated anti-CD4 (RM4-4), anti-IL-5 (TRFK4); PE-Cy7-conjugated anti-CD3e (2C11), anti-IFN-γ (XMG1.2); eFluor 450-conjugated anti-CD8a (53-6.7); APC-conjugated anti-CD4 (GK1.5); APC-eFluor 780-conjugated B220 (RA3-6B2), anti-Ly6C (HK1.4), and anti-F4/80 (BM8) were purchased from eBioscience. Pacific Blue-conjugated anti-CD45 (I3/2), anti-CD11b (ICRF44); and Alexa Fluor 647-conjugated anti-CD11c (N418) were produced in-house (AbLab Biologics, UBC). Data were acquired on an LSR II flow cytometer (BD Biosciences) and analyzed with FlowJo software (TreeStar, San Carlos, CA).  38  2.2.5 Gene expression analysis of lung T cells  Naive or infected T. muris GREAT mice were euthanized by avertin i.p. injection and cervical dislocation. Whole lungs were digested in 200 U/ml collagenase in CTCM for 1 hour at 37° C. Lung cells were RBC-lysed using ammonium chloride buffer and then leukocyte-purified using 30% percoll density centrifugation. Lung cells were Fc-blocked and surface stained for CD4 and then stained for viability using 7-AAD. Live CD4−IFN-γ+ and CD4+IFN-γ+ were FACS sorted into separate fractions for each individual mouse lung sample. Purified cell fractions were then subjected to RNA isolation using RLT buffer and RNeasy RNA extraction kits (QIAGEN) according to the manufacturer's instructions. RNA was converted into cDNA using High Capacity cDNA reverse transcription kits according to the manufacturer's instructions (Applied Biosystems). Quantitative PCR was then performed as described below.  2.2.6 Lung leukocyte ex vivo culture stimulation  In some experiments, ex vivo lung leukocytes, following digestion and percoll purification, were plated in complete tissue culture medium (CTCM) (DMEM supplemented with 10% FBS, penicillin, streptomycin, l-glutamine, β-mercaptoethanol, and HEPES) at 4x106 cell per well. Cells were either unstimulated, polyclonally stimulated with 1 μg/ml anti-CD3/CD28 antibodies, or stimulated with 50 μg/ml T. muris antigen for 3 days of culture. Cells were then stimulated for intracellular cytokine staining for flow cytometry as described above and the supernatants were harvested for ELISA of cytokine production.   39  2.2.7 RNA isolation and quantitative real-time PCR  Lung tissues were mechanically homogenized using a bead basher and RNA was extracted using the TRIzol method according to the manufacturer's instructions (Ambion, Austin, TX). Complementary DNA (cDNA) was generated using High Capacity cDNA reverse transcription kits according to the manufacturer's instructions (Applied Biosystems, Foster City, CA). Quantitative PCR was performed using the SYBR FAST master mix (Kapa Biosystems, Woburn, MA) and SYBR green-optimized primer sets run on an ABI 7900 real-time PCR machine (Applied Biosystems). Ct values of all genes measured in each sample were normalized relative to beta-actin (Actb) gene expression. Primers and sequences used are shown in Table 2.1.  Primer Forward (5’-3’) Reverse (5’-3’) Cd4 CTTCGCAGTTTGATCGTTTTGAT CCGGACTGAAGGTCACTTTGA Ifng GGATGCATTCATGAGTATTGCC CCTTTTCCGCTTCCTGAGG Il4 TCGGCATTTTGAACGAGGTC CAAGCATGGAGTTTTCCCATG Il5 GATGAGGCTTCCTGTCCCTACTC TCGCCACACTTCTCTTTTTGG Il13 GGTCTTGTGTGATGTTGCTCA CCTGGCTCTTGCTTGCCTT Il10 CTGAAGACCCTCAGGATGCG TGGCCTTGTAGACACCTTGGTC Tbx21 CAACAACCCCTTTGCCAAAG TCCCCCAAGCAGTTGACAGT  Table 2.1 Primer sets used for qRT-PCR analysis.     40  2.2.8 Enzyme-linked immunosorbent assay (ELISA)  Immulon plates were coated overnight at 4° C with either anti-IFN-γ (XMG1.2), anti-IL-5 (TRFK4), or anti-IL-10 (JES5-16E3) capture antibodies (eBioscience). Blocking and assay buffer used was PBS containing 10% fetal calf serum. Wash buffer consisted of PBS containing 0.05% Tween 20. Cytokine standards (eBioscience) and samples (in duplicate) were incubated on plates for 2 hours at room temperature. Biotin-conjugated detection antibodies (eBioscience) and streptavidin-conjugated horseradish peroxidase (Jackson ImmunoResearch, West Grove, PA) were incubated on plates for one hour at room temperature. Plates were developed using 3,3',5,5'-tetramethylbenzidine (TMB) substrate (Mandel Scientific, Guelph, ON), stopped with 1N HCl, and read at 450 nm on a Spectramax 384 plate reader (Molecular Devices, Sunnyvale, CA).  2.2.9 Histology  Lung lobes were fixed in buffered 4% paraformaldehyde solution for at least 48 hours and then embedded into paraffin blocks. 5 μm thick sections were stained with Periodic Acid-Schiff (PAS). Airways were assessed microscopically, using 4 representative fields from each animal at 20X magnification for PAS-staining as an indication of mucus hyperproduction by goblet cells. Images were captured for each field and were assigned randomized codes to blind the analysis. PAS+ cells in the airways in each lung section were quantified.    41  2.2.10 Depletion of CD4+ T cells  To deplete CD4+ T cells in vivo, mice were injected i.p. with either 300 μg (per injection) purified anti-mouse CD4 (GK1.5, made in-house) in PBS or PBS alone on days 15 and 17 post-infection, prior to airway challenge with papain. An isotype IgG control antibody was not used since CD4 depletion has been previously shown to be tolerizing to anti-rat Ig responses 175. Depletion efficiency was assessed by flow cytometry using an anti-CD4 antibody (RM4-4, eBioscience) that binds an epitope distinct from the depleting antibody's epitope.  2.2.11 In vivo neutralization of IL-12  T. muris-infected mice were injected i.p. with either 500 μg (per injection) purified anti-mouse IL-12p40 or control IgG1 (Bio X Cell, NH) starting on day 6 post-infection with T. muris and then treated every 3 days until day 18 post-infection (5 injections total) (as previously described 176), prior to airway challenge with papain.  2.2.12 In vivo blockade of IL-10 activity  T. muris-infected mice were injected i.p. with either 500 μg (per injection) purified anti-mouse IL-10R to block IL-10 receptor ligation or control IgG1 (Bio X Cell, NH) starting on day 6 post-infection with T. muris and then subsequently every 3 days until day 18 (5 injections total), prior to airway challenge with papain.    42  2.2.13 HDM model of AAI  The HDM model has been previously described 72. For the first 3 days of airway sensitization, mice were anesthetized under aerosolized isoflurane and instilled i.n. with 100 μg of HDM antigen (GREER, Lenoir, NC). On days 13-17 post-sensitization, mice were i.n. challenged with 25 μg of HDM antigen daily before sacrifice on day 18. BAL and lung tissues were processed as above to assess airway disease. Splenocytes were stimulated with 100 μg/ml HDM antigen for 3 days prior to ELISA of cytokines in the culture supernatants.  2.2.14 Statistics  Data are presented as means ± S.E.M. Statistical significance between two groups was determined by the Student’s t-test while comparisons between 3 or more groups were made by ANOVA with a Bonferroni post-hoc test using GraphPad Prism software.  Results were considered statistically significant with a *p < 0.05, **p < 0.01, or ***p < 0.001.  2.3 Results 2.3.1 Low dose T. muris infection alters the lung immune microenvironment  To characterize the cross-mucosal effect of a localized intestinal immune response, we employed the Trichuris muris model of intestinal infection 177. T. muris infection is a powerful model to study the development of protective Th2 cell-dependent and non-protective Th1 cell-mediated immune responses in vivo. After 21 days of a low dose infection, C57BL/6 mice develop an adaptive anti-T. muris response, characterized by high levels of Th1-cell derived IFN-, resulting in a persistent worm burden (Figure 2.1A). We analyzed the distal mucosal lung tissue for changes in cytokine gene expression by qRT-PCR and found no differences in Th2 43  cell-associated cytokines, such as Il5 (Il4 and Il13 were undetectable), between control and T. muris-infected mice (Figure 2.1B); however, we found significantly increased expression of Ifng and Il10 in the lungs of T. muris-infected mice compared to uninfected mice. These changes in cytokine gene expression in the lungs of T. muris-infected mice correlated with similar increases in IFN-γ and IL-10 protein levels when measured by ELISA of ex vivo stimulated lung cell cultures (Figure 2.1C). From the lungs of T. muris-infected mice only IFN-γ, but not IL-5 nor IL-10, was detectable at low levels in response to T. muris-specific antigen (Figure 2.1D). Despite these changes in lung cytokine expression in T. muris-infected mice, there was no evidence of inflammation or pathology in the lungs of T. muris-infected mice compared to naive controls by histology and based on total cells and cell composition in bronchoalveolar lavage (BAL) fluid (Figure 2.1E). Thus, a low dose intestinal immune response is sufficient to alter the lung immune microenvironment by upregulating IFN-γ and IL-10 expression without causing overt lung tissue pathology. 44   Figure 2.1 Low dose intestinal infection with T. muris alters the lung immune microenvironment. (A) Enumerated cecal worm burdens. (B) Lung mRNA expression of Il5, Ifng, and Il10 normalized relative to Actb. (C) ELISA of IL-5, IFN-γ, and IL-10 from ex vivo unstimulated and polyclonally stimulated lung cell cultures. (D) ELISA of IFN-γ from T. muris-antigen- (Tm Ag) stimulated lung cell cultures. (E) Lung histology visualized using 20X magnification and total and differential bronchoalveolar lavage (BAL) counts. Data are means ± SEM, pooled from 3 independent experiments, n=6 mice per group (A-E). Sections are representative of 3 independent experiments (E). (Ctrl: control; Tm: T. muris-infected; nd: not detected). *p < 0.05, **p < 0.01, ***p < 0.001.  2.3.2 Lung CD4+ T cells increase IFN-γ production during low dose T. muris infection  To determine the cellular source of the infection-induced IFN-γ and IL-10 in the lung, we performed ex vivo culture stimulations of lung cells to characterize cytokine expression by key cell populations. As several recent studies have identified populations of lung CD4+ T cells that produce both IFN- and IL-10 178–180, we examined whether CD4+ T cells were the dominant  45  source of these cytokines. Following stimulation of lung cells with anti-CD3/CD28 antibodies, we found that a significant frequency of lung CD4+ T cells produced IFN-γ, with the highest levels in mice infected with T. muris (Figure 2.2A). In contrast, CD4+ T cells produced low levels of IL-10 in the lungs of either uninfected or T. muris-infected mice (Figure 2.2B) and we observed a negligible population of IFN-/IL-10 double-positive CD4+ T cells (data not shown). To better assess IL-10 expression without cell stimulation, we performed fluorescence-activated cell sorting of IFN-γ+ and IFN-γ− CD4+ T cells from the lungs of T. muris-infected IFN-γ reporter (GREAT) mice 181 followed by gene expression analysis ex vivo. We confirmed that IFN--producing CD4+ T cells appeared to be Th1 cells that expressed Tbx21 (encoding T-box transcription factor Tbx21/Tbet) and Ifng but did not express increased levels of Il10 (Figure 2.2C). Although CD8+ T cells have also been shown to produce IFN- and IL-10 following viral infection of the lung, we failed to detect differences in CD8+ T cell-derived IFN- or IL-10 (data not shown). Together, these results show that the intestinally-restricted T. muris infection results in a potent Th1 cell response in the lungs but that CD4+ T cells are likely not the main source of IL-10.    46   Figure 2.2 Lung CD4+ T cells upregulate IFN-γ expression but not IL-10 during low dose T. muris infection. Frequencies of CD4+ lymphocytes expressing (A) IFN-γ and (B) IL-10 in the lung ex vivo after polyclonal stimulation. (C) mRNA expression of Tbx21, Ifng, and Il10 (relative to Actb) in IFN-γ+ and IFN-γ− FACS-sorted lung CD4+ T cells from GREAT mice. Data are means ± SEM, representative of 3 independent experiments, n=3-5 mice per experiment (A, B) or representative of 2 independent experiments, n=3 mice per experiment (C). (Ctrl: control; Tm: T. muris-infected; ns: not significant). *p < 0.05, **p < 0.01.  2.3.3 Lung myeloid cells increase IL-10 expression during low dose T. muris infection  Since we saw no differences in T cell-derived IL-10, we hypothesized that innate cells were producing IL-10 in the lungs of T. muris-infected mice. We therefore characterized possible innate cellular sources of IL-10 induced in the lungs of T. muris-infected mice ex vivo using a fluorescent IL-10 reporter (Vert-X) 173 mouse strain crossed onto a Rag1−/− background. Compared to naive control mice, T. muris-infected Rag1−/− Vert-X mice had a significant increase in the frequency and total number of IL-10+ cells in the lung (Figure 2.3A). There was a significant expansion of CD11b+CD11c+ myeloid cells producing IL-10 in T. muris-infected mice compared to uninfected controls while we saw no differences in IL-10 production between 47  CD11b−CD11c+ populations in terms of absolute numbers (Figure 2.3B). Further phenotypic analysis showed that lung IL-10+ myeloid cells from T. muris infected mice were predominantly Ly6G−Ly6C− (Figure 2.3C) and were largely negative for surface markers associated with alveolar and interstitial macrophages such as Siglec F and F4/80 (Figure 2.3D). Lung IL-10+ myeloid cells from T. muris infected mice also had a reduction in MHC-II (Figure 2.3E). Upon examining individual cell types for IL-10 production, we found that lung CD11c+B220+ cells, resembling plasmacytoid DCs (pDCs), had the most significant increase in IL-10 production during T. muris infection (Figure 2.3F). These data show that lung IL-10+ myeloid cells that are predominantly CD11b+CD11c+, including a subset of pDC-like cells, emerge during low dose T. muris intestinal infection, independent of adaptive immunity.  48   Figure 2.3 Low dose T. muris infection induces expression of IL-10 in lung myeloid cells. (A) Total lung IL-10+ cell frequencies and absolute numbers in Rag1−/− VertX (IL-10 reporter) mice either uninfected or infected with low dose T. muris. (B) Frequencies and absolute numbers of live CD45+-gated CD11b−CD11c+ and CD11b+CD11c+ cell populations comprising the total IL-10+ cells in the lung. Surface marker analysis of Ly6G vs. Ly6C (C), Siglec F vs. F4/80 (D), and MHC-II vs. CD11c (E) of CD11b+CD11c+IL-10+ gated 49  cells. (F) Gating of total CD11c+B220+ cells and IL-10+ frequencies. Data are means ± SEM, representative of 3 independent experiments, n=5-6 mice per experiment (A-F). (Ctrl: control; Tm: T. muris-infected). **p < 0.01, ***p < 0.001.   2.3.4 Low dose T. muris infection protects against innate-mediated allergic airway inflammation  We hypothesized that the immunological changes in the lung elicited by our low dose intestinal helminth infection could affect the development of type 2 inflammatory responses, such as allergic airway inflammation (AAI). To directly address this, C57BL/6 mice were infected with a low dose of T. muris and challenged with papain intranasally (Figure 2.4A). Acute airway challenge with papain causes an early activation of ILC2s that promotes eosinophil recruitment and mucus hyperproduction 70. Papain treatment did not affect infection dynamics as low dose infections persisted following papain exposure (Figure 2.4B). BAL fluid analysis showed that papain induced an influx of inflammatory cells in the airways of uninfected mice (Figure 2.4C). In contrast, mice infected with T. muris had a significant reduction in airway cell influx after papain challenge. While uninfected mice displayed airway eosinophilia after papain challenge, infected mice were significantly protected from eosinophil infiltration. Further, T. muris-infected mice had a reduction in lung tissue eosinophils after papain challenge. Papain-induced goblet cell hyperplasia and mucus production (inferred from PAS staining) were also reduced in T. muris-infected mice (Figure 2.4D). Expression of the Th2 cell-associated cytokine Il5 in the lung was increased in control mice but was not induced in T. muris-infected mice after papain challenge, correlating with the reduction in eosinophil recruitment to the airways and lungs (Figure 2.4E). Consistent with our previous results, T. muris-infected mice had 50  significantly increased expression of lung Ifng and Il10 after papain airway challenge. Despite this T. muris-mediated protection from papain-induced AAI, we found no differences in the induction of lung ILC2 cells in response to papain in terms of absolute numbers (Figure 2.4F). Additionally, the capacity for lung ILC2 cells to produce IL-13 in response to papain was unaffected by T. muris infection (Figure 2.4G). Thus, changes in the lung immune microenvironment during low dose intestinal infection with T. muris are associated with protection against papain-induced AAI.   Figure 2.4 Low dose T. muris infection suppresses papain-induced allergic airway inflammation. (A) Experimental timecourse of intestinal T. muris infection and airway challenge with papain. (B) Enumerated cecal worm burdens. (C) Total bronchoalveolar lavage (BAL) cell, BAL eosinophil, and lung tissue eosinophil counts. (D) PAS-stained sections of lungs, visualized using 20X magnification. (E) Lung mRNA expression of Il5, Ifng, and Il10 normalized relative to Actb. (F) Gating of live CD45+lineage−CD90+ cells, further gated on CD25+ and ST2+ to identify ILC2 cells in uninfected or T. muris-infected mice challenged with papain with absolute numbers  51  shown. (G) IL-13 production by ILC2 cells after ex vivo lung cell stimulation after papain exposure; naive (no papain) is shown as a gray dashed line, Ctrl is shown as a solid black line, and T. muris-infected is shown as a dotted black line. Data are means ± SEM, pooled from 3 independent experiments, n=3-15 mice per group (B-G). Sections are representative of 3 independent experiments (D). (N: naive; Ctrl: control; Tm: T. muris-infected, ns: not significant). *p < 0.05, **p < 0.01, ***p < 0.001.  2.3.5 Low dose T. muris infection protects against acute airway inflammation in the absence of CD4+ T cells  Given that we saw an increase in lung CD4+ Th1 cells following infection with T. muris, we wanted to test whether these cells were necessary for protection against papain-induced AAI. T. muris-infected mice were treated with a depleting monoclonal antibody against CD4 prior to papain challenge (Figure 2.5A). Strikingly, T. muris-infected mice depleted of CD4+ cells were protected from papain-induced AAI, based on a persistent reduction of airway eosinophilia compared to uninfected control mice (Figure 2.5B). Gene expression analysis showed that infected, CD4-depleted mice had a significant reduction in Ifng expression in the lungs but still maintained increased expression of Il10 when compared to non-depleted mice (Figure 2.5C). These results confirm a role for lung CD4+ T cells in IFN- production following T. muris infection and suggested that Th1 cell-expressed IFN- is not absolutely required for infection-induced protection of AAI. Further, these results confirm that CD4+ T cells are not the source of IL-10 in the lung.  52   Figure 2.5 Depletion of CD4+ T cells does not reverse T. muris-mediated protection from papain-induced allergic airway inflammation. (A) Experimental timecourse of CD4 depletion during the infection/airway challenge model. (B) Bronchoalveolar lavage (BAL) eosinophil counts. (C) Lung mRNA expression of Cd4, Ifng, and Il10 (relative to Actb). Data are means ± SEM, representative of 2 independent experiments, n=3-6 mice per experiment (B, C). (Ctrl: control; Tm: T. muris-infected; pap: papain). *p < 0.05, **p < 0.01, ***p < 0.001.     53  2.3.6 Rag1−/− mice are partially protected from acute innate-mediated airway inflammation during low dose T. muris infection  T. muris-infected mice that were CD4-depleted were protected from AAI despite a significant reduction in Ifng expression in the lungs of T. muris-infected CD4-depleted mice compared to non-depleted mice; however, lung Ifng levels were still elevated compared to uninfected controls which may have represented lingering effects of IFN-γ induction in other cell types prior to CD4 depletion as IFN-γ can exhibit positive feedback signaling 182. Therefore, we next ascertained whether the entire adaptive immune system was necessary in our experimental model. To do so, we performed our T. muris-infection/papain challenge model using Rag1−/− mice, which lack T cells and B cells but are still susceptible to innate-mediated type 2 inflammation. Surprisingly, T. muris-infected Rag1−/− mice had a significant reduction in total BAL cells as well as BAL eosinophils when compared to uninfected Rag1−/− mice after papain challenge (Figure 2.6A). However, despite a reduction in airway eosinophilia, histopathological analysis showed that T. muris-infected Rag1−/− mice were equally susceptible to mucus overproduction in the large airways (Figure 2.6B). Ifng expression was not induced in the lungs of T. muris-infected Rag1−/− mice after papain challenge, further confirming CD4+ T cells as the likely source (Figure 2.6C). However, we observed a significant increase in lung Il10 expression following T. muris infection, confirming an innate cell-derived source of IL-10 in the lungs during T. muris infection. These results suggest that T. muris-induced adaptive immune responses are important for regulating goblet cell hyperplasia and mucus production in the lung, but are dispensable for inhibiting eosinophil infiltration into the airways after papain exposure.  54   Figure 2.6 Low dose infection with T. muris in Rag1−/− mice partially protects from papain-induced lung inflammation. (A) Total bronchoalveolar lavage (BAL) cell and BAL eosinophil counts. (B) PAS-stained sections of lungs, visualized using 20X magnification. (C) Lung mRNA expression of Ifng and Il10 (relative to Actb). Data are means ± SEM, representative of 3 independent experiments, n=4-5 mice per experiment (A-C). Sections are representative of 3 independent experiments (B). (Ctrl: control; Tm: T. muris-infected). **p < 0.01.  2.3.7 Neutralization of IL-10 activity in T. muris-infected Rag1−/− mice during acute airway inflammation  Since we observed a consistent upregulation of IL-10 in the lungs of T. muris-infected C57BL/6 and Rag1–/– mice, we hypothesized that blocking IL-10 activity would restore susceptibility to eosinophilia in infected Rag1−/− mice after papain challenge. To do so, we injected mice with α-IL-10R antibodies every 3 days starting on day 6 post-infection, before airway challenge (Figure 2.7A). Following induction of acute lung inflammation with papain, we found that compared to control and IgG1-treated T. muris-infected (Tm IgG1) mice, α-IL-10R-treated T. muris-infected (Tm α-IL-10R) mice trended toward an increase in total airway  55  cellular infiltration as measured by total BAL cell counts comprised of mainly neutrophils (Figure 2.7B). However, blockade of IL-10R did not reverse suppression of eosinophilia. Consistent with our previous results, histological examination revealed no significant differences in terms of mucus hyperproduction in papain-challenged mice, with neither worm infection nor IL-10R blockade having an effect (Figure 2.7C). Together, these data show that while IL-10 did not play a role in limiting eosinophil infiltration in our model, there was a trend in IL-10 playing a role in limiting neutrophil accumulation in the airways during acute AAI.   Figure 2.7 Blockade of IL-10R in T. muris-infected Rag1−/− mice following papain challenge. (A) Timecourse of low dose infection with T. muris and blockade of IL-10R. (B) Total bronchoalveolar lavage (BAL) cell, BAL eosinophil, and BAL neutrophil counts. (C) PAS-stained sections of lungs, visualized using 20X magnification. Data are means ± SEM, pooled from 2 independent experiments, n= 6-8 mice per group (B, C). Sections are representative of 2 independent experiments (C). (Ctrl: control; Tm: T. muris-infected). *p < 0.05.  56  2.3.8 Low dose T. muris infection protects against the house dust mite model of asthma  We next wanted to test whether intestinal helminth infection could also abrogate the development of house dust mite (HDM)-induced AAI. HDM antigen, which displays similar protease activity to papain, has been used extensively as an animal model of asthma. Unlike papain, HDM-induced AAI involves priming and challenge phases of exposure to HDM antigen with a more prominent role for adaptive immune responses. Thus, we infected C57BL/6 mice with T. muris either ten days before the sensitization phase (Tm d-10) or on the first day of sensitization (Tm d0) (Figure 2.8A). We found that after challenge with HDM antigen, control and T. muris d0 mice had an increase in total BAL cells and eosinophils while T. muris d-10 mice had significantly fewer total cells and eosinophils in the BAL fluid (Figure 2.8B). Histologically, control and T. muris d0 mice after HDM antigen challenge had an increase in cellular infiltration around airways as well as evidence of mucus hyperproduction in bronchioles; in contrast, the lungs of T. muris d-10 mice largely resembled naive lungs with a reduction in both airway cellular infiltration and mucus production (Figure 2.8C). Gene expression analysis of the lungs showed a significant increase in Ifng expression in both groups of mice T. muris-infected mice following HDM antigen challenge (Figure 2.8D); there were no significant differences found between control mice and infected mice in terms of Il10 gene expression. Based on these results, we conclude that T. muris-mediated protection from HDM antigen-induced airway disease likely involves an effect during the early sensitization phase of the model. 57   Figure 2.8 Time dependence of low dose T. muris-mediated suppression of house dust mite antigen-induced allergic airway inflammation. (A) Timecourse of low dose infection either before (Tm d-10) or concurrent (Tm d0) with intranasal (i.n.) sensitization with house dust mite (HDM) antigen. (B) Total bronchoalveolar lavage (BAL) cell and BAL eosinophil counts. (C) PAS-stained sections of lungs, visualized using 20X magnification. (D) Lung mRNA expression of Ifng and Il10 (relative to Actb). Data are means ± SEM, representative of 3 independent experiments, n= 3-5 mice per experiment (B-D). Sections are representative of 3 independent experiments (C). (Ctrl: control; Tm: T. muris-infected; nd: not detected). *p < 0.05, **p < 0.01.    58  2.4 Discussion Low dose infections with T. muris cause a localized intestinal Th1 cell response, resulting in parasite persistence in most strains of wild type mice 177. Since low dose T. muris infections have been shown to induce increased intestinal levels of IFN-γ and IL-10 183,184, we assessed whether levels of these cytokines were also affected distally in the lungs. We found that T. muris infection was sufficient to significantly increase IFN-γ and IL-10 expression in the lung, in the absence of any signs of airway pathology. Our follow-up analyses determined that T. muris infection-induced lung IFN-γ was predominately produced by CD4+ Th1 cells while IL-10 was mainly myeloid cell-derived. Furthermore, these infection-induced changes in the lung immune microenvironment correlated with complete resistance to innate-mediated type 2 responses to the protease allergen papain. Our results suggest that both adaptive and innate responses were required for mediating infection-induced distal suppression of AAI. IFN-γ from Th1 cells may inhibit the development of Th2 cell responses during AAI but its role has been controversial and context-dependent 185–187. It has been reported that IFN-γ can negatively regulate goblet cell function during AAI 188,189, a role that is consistent with our results showing a consistent infection-induced upregulation of lung IFN-γ correlating with reductions in goblet cell-derived mucus production during AAI. In parallel, IL-10 has pleiotropic effects on all immune cells and during AAI, can be considered to be protective 60,190; for example, IL-10 can regulate Th2 cell cytokine responses and have a negative effect on eosinophil function and survival 191–193. To tease out the individual role of these cytokines, we showed that T. muris-infected Rag1–/– mice, which had elevated lung IL-10 in the absence of lung IFN-γ responses, were not fully protected from AAI. In addition, we blocked IL-12, a cytokine that potently drives IFN-γ expression, and found this was not 59  sufficient to reverse protection from AAI during T. muris infection (Supplemental Figure A.1). Thus, infection-induced IFN- and IL-10 have non-redundant roles in the inhibition of AAI. Previously, Mohrs et al. showed that infection with a different intestinal helminth, Heligmosomoides polygyrus bakeri, which induces a mixed Th2/regulatory T (Treg) cell response, results in systemic dissemination of Th2 cells to extraintestinal sites, including the airways 194; together with our results showing accumulation of Th1 cells in the lung following T. muris infection, this suggests that specific Th cell-polarized responses can propagate from the intestine to the lung. In parallel with this Th1 response, the innate IL-10+ cells may represent a heterogeneous population of immature myeloid cells that expand during low dose T. muris infection. Further, our results demonstrate that a prominent subset of these myeloid cells were B220+, resembling plasmacytoid DCs (pDCs); pDCs have been associated with the priming of IL-10 producing Treg cells 195 but have not been described before to intrinsically produce IL-10. In the context of cross-mucosal trafficking, lung DCs can upregulate gut-homing receptors on T cells to induce migration from the lung to the gut 34. Given that we saw T. muris-antigen responding Th1 cells in the lung, T cell migration between the gut and the lung may actually be bidirectional. However, it remains to be determined whether these adaptive and innate cells are intestinally-derived that traffic to the lung or are instead locally induced de novo in the lung mucosa following systemic dissemination of helminth antigens. Additionally, future studies should determine whether T. muris infection affects hematopoiesis in the bone marrow since IFN-γ is known to profoundly impact hematopoietic progenitor cells and eosinophil differentiation 196.  It has previously been shown that intestinal H. polygyrus infection can also suppress murine models of asthma, primarily through the induction of Treg cells via a helminth-secreted 60  TGF-β analog 197. Further, soluble excretory/secretory products from H. polygyrus, when co-administered with allergens intranasally, are sufficient in suppressing type 2 responses 198. In contrast, we found that our model of intestinal T. muris infection modulating lung immune responses does not depend on Treg cell responses when considering our CD4 depletion results and after quantifying lung Treg cells following papain exposure (Supplemental Figure A.2). H. polygyrus-mediated suppression of  AAI is dependent on IL-10 116; although we found a consistent upregulation of IL-10 in the lung during T. muris infection, innate IL-10 in infected Rag1–/– mice was not sufficient to fully protect against AAI.  Instead, our results suggest non-redundant roles for both Th1 cells and innate myeloid cells in T. muris infection-mediated protection from type 2 airway inflammation. Thus, we hypothesize that the Th1 cell-associated response to T. muris gets misdirected to the lungs and in the absence of appropriate antigen, becomes self-limiting by inducing immunoregulatory responses involving IL-10, resulting in bystander immunosuppression in the local airway mucosa. In parallel to these responses, it has been shown that chronic T. muris infection causes alterations in the intestinal microbiota composition and metabolome 199 which may potentially play a role in mediating gut-to-lung immune crosstalk since this has been described to occur during H. polygyrus infection 200.  Lastly, we assessed the impact of our Th1 cell-driven infection model on an AAI model more relevant to human asthma, with a significant adaptive immune component. Complete protection from HDM antigen-induced AAI only occurred in mice infected with T. muris prior to sensitization, suggesting that timing of the infection is critical in establishing protection against asthma. Consistent with our data on early type 2 responses to papain, this likely involves a critical effect of the infection on the early phases of type 2 inflammation. This may partially explain why treatment of allergic rhinitis patients with Trichuris suis ova has been found to be 61  ineffective during the peak of allergic symptoms 201. Broadly, this suggests that Th1 cell-driven intestinal inflammation and associated immunoregulatory responses are only effective as a prophylactic means to suppress type 2 responses, as opposed to being therapeutic.  In summary, we have shown that an intestinally-restricted, Th1 cell-driven infection can potently suppress the development of innate-mediated type 2 responses in the lung, in a temporally-dependent manner. The cellular mechanisms involved display a complex, multifaceted means of host immunomodulation that involves both innate and adaptive immunity and spans distinct mucosal tissues. These converging immune pathways may highlight novel avenues of better understanding cross-mucosal immune regulation.  62  Chapter 3: Chronic Trichuris muris infection affects hematopoiesis in the bone marrow  3.1 Overview  In the preceding studies performed in chapter 2, we found that there is a dramatic effect of chronic intestinal infection with T. muris on the development of AAI in the lung. While the mechanisms of this immune regulation could involve direct cross-talk from the intestine to the lung, it is also possible that the chronic infection has broader systemic effects on the host. Therefore, it was necessary to characterize these putative infection-induced systemic effects by looking at other immune compartments. One likely site affected by chronic infections is the BM, the major site of hematopoiesis in the body. Infections are known to affect hematopoiesis as an ongoing immunological insult elicits a greater turnover of short-lived immune cells which warrants a continued replenishment from the BM 202. Therefore, studies were carried out to determine the effect of chronic T. muris infections on hematopoiesis in the BM. We found that chronic, but not acute, low dose infection with T. muris leads to profound changes in the BM in terms of total Sca-1 expression and the number and composition of hematopoietic stem cell (HSC) populations. We also show that this process is dependent on IFN-γ and is associated with the accumulation of IFN-γ+ T cells within the BM itself. Together, these results suggest that chronic intestinal infection with T. muris has systemic effects on hematopoiesis and may provide mechanistic insight into the relationship between intestinal and lung mucosal immune responses.  63  3.2 Materials and methods 3.2.1 Mice and intestinal worm infections  C57BL/6 or GREAT (IFN-γ YFP reporter) mice were infected with either no worms, a low dose (30 eggs), or a high dose (200 eggs) of T. muris by oral gavage. Infections were carried out over 21 days before sacrifice of mice by administration of CO2 gas. All mice were bred and housed in specific pathogen-free animal facilities at the UBC Biomedical Research Centre.   3.2.2 Isolation of bone marrow cells  Following sacrifice of mice, hip joints were dislocated and the right hind limbs of each mouse was dissected to isolate single femur bones, clean of any muscle and other tissues. The ends of each femur were sliced by a razor blade and a 27-gauge needle was used to flush out the bone marrow with CTCM. The bone marrow was then passed through a 70 μm cell strainer to obtain single-cell suspensions. Bone marrow samples were then red cell-lysed using ammonium chloride buffer, and suspended to a known volume before being enumerated by a hemocytometer.  Cells were then reconstituted in standard FACS buffer containing 2% FBS, 1mM EDTA, and 0.02% NaN3 in PBS prior to surface staining for flow cytometry.  3.2.3 In vivo neutralization of IFN-γ  Mice were infected with T. muris as described above. On day 4 post-infected, mice were injected i.p. with 500 μg (per injection) of either control IgG1 or anti-IFN-γ (XMG1.2) and were repeatedly injected thereafter on days 8, 12, and 16.  64  3.2.4 Colony forming unit (CFU) assay  BM cells were isolated as above using proper aseptic technique. Following manufacturer instructions, 1.5x104 cells from each sample were plated in duplicate in Methocult Media (STEMCELL M3234, Vancouver, BC) supplemented with 50 ng/ml SCF, 10 ng/ml IL-3, and 10 ng/ml IL-6. Plates were incubated for approximately 10 days prior to morphological analysis and differential counts of CFUs.  3.2.5 Antibodies and flow cytometry  All antibody dilutions and cell staining were done with PBS containing 2% FCS, 1 mM EDTA, and 0.05% sodium azide. Fixable Viability Dye eFluor 506 was purchased from eBioscience to exclude dead cells from analyses. Prior to staining, samples were Fc-blocked with buffer containing anti-CD16/32 (93, eBioscience) and 1% rat serum to prevent non-specific antibody binding. APC-conjugated anti-CD150 (TC15-12F12.2) was purchased from BioLegend. PE-conjugated anti-CD3e (2C11); PE-Cy7-conjugated anti-Sca1 (D7); eFluor 450-conjugated anti-CD8a (53-6.7), anti-CD48 (HM48-1); APC-conjugated anti-CD4 (GK1.5); and APC-eFluor 780-conjugated anti-c-kit (2B8) were purchased from eBioscience. FITC-conjugated anti-CD19 (1D3), anti-CD11b (M1/70), anti-CD11c (N418), anti-Gr1 (RB6-8C5), anti-NK1.1 (PK136), anti-Ter119 (Ter119); Pacific Blue-conjugated anti-CD45 (I3/2), anti-CD11b (M1/70); and Alexa Fluor 647-conjugated anti-CD11c (N418) were produced in-house (AbLab Biologics, UBC). Data were acquired on an LSR II flow cytometer (BD Biosciences) and analyzed with FlowJo software (TreeStar).  65  3.2.6 Statistics  Data are presented as means ± S.E.M. Statistical significance between two groups was determined by the Student’s t-test while comparisons between 3 or more groups were made by ANOVA with a Bonferroni post-hoc test using GraphPad Prism software.  Results were considered statistically significant with a *p < 0.05, **p < 0.01, or ***p < 0.001.  3.3 Results 3.3.1 Chronic T. muris infection induces increased Sca-1 expression on hematopoietic cells and increases LSK cells in the BM Hematopoiesis in the BM is the essential process where HSCs give rise to all blood cell lineages which includes erythrocytes, myeloid cells, and lymphoid cells. While hematopoiesis is continuously needed during homeostasis, the replenishment of immune cells during an inflammatory response is particularly crucial. For example, early during many infections there is a rapid generation and mobilization of neutrophils from the BM 203. Further, Pro-inflammatory cytokines, such as interferons and TNF-α, can directly induce HSC proliferation and differentiation 204. However, under conditions of chronic inflammation, IFN-γ can have a negative impact on HSC proliferation and self-renewal and thus may play a critical role in hematopoietic stress 205. Previous studies have implicated IFN-γ affecting the BM such as during viral lung infections and during colitis 206,207. However, the role of IFN-γ in modulating hematopoiesis in the context of intestinal infections has not been thoroughly investigated.  To assess the systemic impact of localized intestinal immune responses on hematopoiesis, we employed the T. muris model of helminth infection. The T. muris model is a well-characterized system that allows the induction of either a Th1 cell-driven persistent infection 66  (low dose) or a Th2 cell-driven acute infection (high dose) in the large intestine of most strains of wild-type mice 136. Using this infection model, we could compare the effects of intestinal Th1 and Th2 cell-dependent immune responses on hematopoiesis. As expected, low dose (LD) T. muris infection of C57BL/6 mice resulted in a persistent worm burden 21 days post-infection (Figure 3.1A). In contrast, high dose (HD) T. muris infection resulted in few or no detectable worms in the cecum, consistent with a protective Th2 cell response. Isolation and enumeration of total BM cells from these mice showed that LD, but not HD T. muris infection resulted in a significant reduction in total cellularity in the BM (Figure 3.1B). Next, we assessed whether specific changes to the composition of major cell populations in the BM occur in response to LD or HD T. muris infections. Since LD T. muris infection is associated with an IFN-γ-driven Th1 cell responses 136 and that IFN-γ has been shown to induce the expression of stem cells antigen-1 (Sca-1) on immune cells 208, we assessed Sca-1 expression on hematopoietic cells in the BM of infected mice. The frequency and absolute number of total CD45+ BM cells expressing Sca-1 was increased 2-fold in LD T. muris-infected mice compared to naive controls (Figure 3.1C). Conversely, mice that received a HD T. muris infection did not have elevated Sca-1 expression on hematopoietic cells in the BM. We further analyzed lineage-negative cell populations in the BM and found that only LD T. muris infection caused a significant increase in the frequency and absolute number of Sca-1+c-Kit+ (LSK) cells compared to naive and HD T. muris-infected mice (Figure 3.1D). Together, these data show that a persistent LD T. muris infection induces significant and broad changes in the hematopoietic compartment in the BM that are absent after a transient HD T. muris infection.  67     Figure 3.1 Induction of Sca-1 expression and LSK cells in the BM during low dose T. muris infection. (A) Cecal worm counts of either uninfected control (Ctrl), low dose T. muris-infected (LD), and high dose T. muris-infected (HD) mice. (B) Quantification of total BM cells. (C) Frequency and absolute number of live-gated CD45+Sca-1+ cells in the BM. (D) Gating strategy to identify lineage−Sca-1+c-Kit+ (LSK) in the BM with frequencies and absolute numbers shown. Data are means ± SEM and are representative of 3 independent experiments (n=4-5 mice per experiment) (A-D). *p < 0.05, ***p < 0.001.   68  3.3.2 Chronic T. muris infection alters bone marrow HSC and MPP populations and is dependent on adaptive immunity  To further characterize the infection-induced changes in the BM during LD T. muris infection, we analyzed progenitor and stem cell populations within the LSK population using the signaling lymphocytic activation molecule (SLAM) family markers CD48 and CD150 - multipotent progenitor (MPP) cells are enriched within the CD48−CD150− population while HSCs can be identified as being CD48−CD150+ 209. We found that LD T. muris-infected mice had significant expansion of CD48−CD150− MPPs in the BM compared to naive mice (Figure 3.2A). In parallel, the frequency of BM CD48−CD150+ HSCs was significantly reduced during LD T. muris infection. To test the functional effects of these changes in the BM, we performed in vitro colony forming assays. BM from LD T. muris-infected mice had significantly higher total colony forming unit (CFU) counts (Figure 3.2B). There were no significant differences found in terms of monoblast (CFU-M) or granulocyte (CFU-G) colony counts; however, BM from LD T. muris-infected mice yielded more precursor CFU-GM (granulocyte, monocyte) colonies. Therefore, LD T. muris infection alters primitive progenitor populations in the BM that correlate with an increase in myeloid colony forming potential in vitro. As LD T. muris infection is associated with a Th1 cell response 177, we next tested whether the effects we saw were indeed dependent on adaptive immunity by analyzing the BM of infected Rag1−/− mice which lack T and B cells. In contrast to infected WT mice, LD T. muris-infected Rag1−/− mice did not show an expansion of MPPs in the BM when compared to uninfected controls (Figure 3.2C). However, there was still a significant reduction in the frequency of HSCs among the LSK cell population in the BM of infected Rag1−/− mice. Thus, 69  LD T. muris infection alters MPP and HSC populations in the hematopoietic compartment of the BM and is dependent on the presence of adaptive immune cells.   Figure 3.2 Low dose T. muris infection alters hematopoietic progenitor populations in the BM. (A) Gating strategy to identify LSK-gated CD48−CD150− MPPs and CD48−CD150+ HSCs and respective frequencies of LSK cells in the BM of either uninfected control (Ctrl) or low dose T. muris-infected (LD) mice. (B) Colony forming unit (CFU) assay of BM cells showing total and differential counts of CFU-M, CFU-G, and CFU-GM colonies. (C) Frequencies of LSK-gated CD48−CD150− MPPs and CD48−CD150+ HSCs in either uninfected Rag1−/− or LD Rag1−/− mice. Data are means ± SEM and are representative of 3 independent experiments (n=4 mice per experiment) (A) or 2 independent experiments (n=3-4 mice per experiment) (B-C). *p < 0.05, **p < 0.01.    70  3.3.3 Changes in hematopoiesis during chronic T. muris infection is IFN-γ-dependent  Since we found that changes in the BM in response to LD T. muris infection depended on adaptive immunity, we sought to determine whether the infection-mediated effects on hematopoiesis were indeed driven by the Th1 cell-associated cytokine IFN-γ. To directly test this, we neutralized IFN-γ systemically during the course of the infection using monoclonal antibody treatment. Blockade of IFN-γ did not cause mice to clear a LD T. muris infection, as infected mice treated with control IgG antibodies or anti-IFN- had comparable worm burdens (Figure 3.3A). Analysis of the BM revealed that IFN-γ blockade abrogated the increase in total Sca-1 expression on hematopoietic cells during LD T. muris infection (Figure 3.3B). Further, the infection-induced increase in LSK cells, in terms of frequencies and absolute numbers, was completely reversed upon blockade of IFN-γ (Figure 3.3C). As shown in Figure 3.3D, the expansion of CD48−CD150− MPPs in the BM during LD T. muris infection was reversed in infected mice treated with α-IFN-γ which resembled naive MPP frequencies. Further, the frequency of CD48−CD150+ HSCs was also restored upon IFN-γ blockade in infected mice. Thus, alterations in hematopoietic progenitor populations in the BM, elicited by LD T. muris intestinal infection, is completely dependent on IFN-γ signalling.  71   Figure 3.3 Alterations in BM hematopoietic progenitor populations during low dose T. muris infection is IFN-γ-dependent. (A) Cecal worm counts of either uninfected control (Ctrl), low dose T. muris-infected/control IgG treated (LD IgG), or low dose T. muris-infected/α-IFN-γ treated (LD α-IFN-γ) mice. (B) Frequencies of total Sca-1+CD45+ in the BM. (C) Gating strategy to quantify frequencies and absolute numbers of LSK cells in the BM. (D) Frequencies of LSK-gated CD48−CD150− MPPs and CD48−CD150+ HSCs. Data are means ± SEM and are representative of 3 independent experiments (n=3-4 mice per experiment) (A-D). *p < 0.05, **p < 0.01, ***p < 0.001.  72  3.3.4 A mixed population of IFN-γ-expressing T cells accumulate in the BM during chronic T. muris infection  Given that we found that IFN-γ was playing a major role in influencing hematopoiesis during LD T. muris infection, we next assessed whether IFN-γ-expressing cells are found in the BM. To do this, we infected IFN-γ-YFP reporter (GREAT) mice with LD T. muris and characterized IFN-γ expression in the BM. Compared to naive controls, LD T. muris-infected mice had a small yet significant increase in IFN-γ+ cells in the BM (Figure 3.4A). Additionally, the IFN-γ-producing BM cells from LD T. muris-infected mice also had a significantly higher mean fluorescent intensity (MFI) compared to controls. Upon examination of specific cell types in the BM, we found that the majority of cells with increased IFN-γ expression during infection were CD3+ lymphocytes (Figure 3.4B). IFN-γ expression was significantly increased in CD4+, CD8+, and CD4−CD8− double negative (DN) T cell populations in the BM of LD T. muris-infected mice compared to BM from naive mice. These IFN-γ+ T cells in the BM were also significantly increased in terms of absolute numbers during LD T. muris infection (data not shown). In contrast, NK cells showed no significant differences in IFN-γ expression. Therefore, a mixed population of IFN-γ+ T cells accumulate in the BM during persistent LD T. muris intestinal infection.  73   Figure 3.4 Low dose T. muris infection induces an accumulation of IFN-γ+ T cells in the BM. (A) Frequency and MFI of total IFN-γ+ cells of either uninfected control (Ctrl) or low dose T. muris-infected (LD) IFN-γ-YFP reporter mice. (B) Frequencies of IFN-γ+ cells gated on either CD3+CD4+, CD3+CD8+, CD3+CD4−CD8−, or CD3−NK1.1+ cells. Data are means ± SEM and are representative of 3 independent experiments (n=3-4 mice per experiment) (A-B). **p < 0.01, ***p < 0.001.  3.3.5 IFN-γ-expressing T cells in the BM display activated and central memory phenotypes during chronic T. muris infection  We next phenotyped surface marker expression on IFN-γ+ T cells in the BM of LD T. muris-infected mice. The majority (> 90%) of IFN-γ-expressing CD4+ and CD8+ T cells in the BM were predominately TCR-β+ but were of comparable frequency between LD T. muris-infected and uninfected mice (Figure 3.5A). Conversely, IFN-γ+ DN T cells could be split into both TCR-β+ and TCR-γδ+ populations (2:1 ratio respectively) and were of similar frequencies 74  between LD T. muris-infected and naive controls. Analysis of CD44 and CD62L expression showed that IFN-γ+ CD4+ T cells in the BM from both LD T. muris-infected and naive mice were mainly CD44+CD62L−, consistent with an activated phenotype (Figure 3.5B). However, for both IFN-γ+ CD8+ and DN T cells, there was a significant increase in the frequency of CD44+CD62L+ cells in the BM of LD T. muris-infected mice, indicative of an increase in central memory type cells. Thus, LD T. muris infection induces a mixed population of T cells expressing IFN-γ that display activated and central memory phenotypes in the BM.     75  Figure 3.5 Activated and central memory-like phenotype of IFN-γ+ T cells in the BM of low dose T. muris-infected mice. (A) Frequencies of TCR-β+ and TCR-γδ+ cells of CD4+IFN-γ+ gated, CD8+IFN-γ+ gated, or CD4−CD8−IFN-γ+ gated cells in the BM of either uninfected control (Ctrl) or low dose T. muris-infected (LD) mice. (B) Frequencies and quantification of CD44 and CD62L expression on CD4+IFN-γ+ gated, CD8+IFN-γ+ gated, or CD4−CD8−IFN-γ+ gated cells in the BM. Data are means ± SEM and are representative of 2 independent experiments (n=3 mice per experiment) (A-B). **p < 0.01, ***p < 0.001.  3.4 Discussion  In the present study, we have shown that a persistent, low burden infection with the intestinal helminth T. muris broadly influences hematopoietic progenitor populations in the BM. Although there have not been any previous studies on the direct effects of intestinal helminths on the BM, it has been shown that chronic intestinal inflammation can profoundly influence hematopoiesis; during colitis, pro-inflammatory cytokines can induce the dysregulated proliferation of HSCs and can drive myelopoiesis that further exacerbate chronic intestinal inflammation 207. This is consistent with our results showing an expansion of MPPs in the BM and associated increase in myeloid colony formation in vitro that occurs during a persistent intestinal T. muris infection. However, in contrast to colitis, chronic intestinal infection with T. muris is not associated with overt morbidity in mice, due to the induction of regulatory responses such as IL-10 140,184. Since we concurrently observed a dependence on IFN-γ signalling in our model, it is likely that pro-inflammatory cytokines induced during both colitis and intestinal helminth infection have a similar overall impact on BM responses.  With respect to other mucosal sites, Maltby et al. showed that during acute pneumonia virus (PVM) infection in the lung, an increase in myelopoiesis occurs in the BM that is IFN-γ-76  driven 206; however, upon blockade of IFN-γ in their model, viral load increased which may have confounded their findings. In contrast, when we neutralized IFN-γ during LD T. muris infection, we saw no effect on intestinal worm burden, suggesting that the infection-induced effects on the BM were primarily IFN-γ-mediated and not dependent on the presence of worms alone or other factors associated with the chronic infection. Other infection models that induce Th1 cell responses suggest a consistent role for IFN-γ in affecting BM output: IFN-γ negatively regulates granulopoiesis and promotes monopoiesis during mycobacterial infections 210 which also occurs during LCMV infection 211. However, we did not observe a specific skewing toward monopoiesis in the BM of mice chronically infected with T. muris, based on colony forming potential and analysis of mature lineages in the BM showing increases in granulocytes (data not shown). This enhanced myelopoiesis is consistent with our previous in vivo studies showing an increase in myeloid cells at peripheral sites such as the lungs during chronic T. muris infection 212. Together, these findings are consistent with the notion that the role of IFN-γ during hematopoiesis is context dependent 213 which will vary between different infection models. Additionally, IFN-γ can differentially affect BM responses by synergizing with other cytokines 214 which are likely considerably different in our model of intestinal helminth infection compared to these other experimental models. Thus, chronic T. muris infection induces myelopoiesis in the BM similar to other Th1 cell-inducing models but with differences in myeloid lineage-specific skewing.   IFN-γ has been previously shown to be elevated both in the mesenteric lymph nodes and systemically in the blood during chronic T. muris infection 215. Our results suggest that in addition to this systemic induction of IFN-γ, IFN-γ+ T cells also accumulate in the BM during the chronic intestinal helminth infection. The accumulation of activated T cells expressing IFN-γ 77  in the BM has been previously described 205, and has been shown to occur in the contexts of T cell-driven colitis 207 and anti-tumor responses 216. Indeed, the BM is considered a preferential site in which memory T cells home 217, consistent with our results showing that both CD8+ and DN IFN-γ+ T cells in the BM display a  central memory phenotype during chronic T. muris infection. DN T cells have roles in various Th1 cell-mediated responses including intracellular bacterial and viral infections 218,219. Further, during intracellular infection with Leishmania major, DN T cells that can rapidly produce IFN-γ are critical for primary and secondary responses and display memory-like cell properties 220. This is in line with our results showing that central memory-like DN T cells had the largest increase in IFN-γ production during a Th1 cell-associated chronic T. muris infection. Future studies will focus on determining whether these BM-residing T cells expressing IFN-γ are key in modulating hematopoietic progenitor populations during chronic T. muris infection.   Overall, our data show for the first time that a chronic intestinal infection with T. muris influences hematopoiesis in the BM via the induction IFN-γ and is associated with activated and memory-like T cells expressing IFN-γ accumulating in the BM. Additionally, our results highlight that intestinal worm infection-induced effects on the BM are mechanistically consistent with a variety of distinct Th1 cell-associated responses.  78  Chapter 4: The RA-metabolizing enzyme Cyp26b1 controls Th cell differentiation and function  4.1 Overview  Chapters 2 and 3 focused on the modulation of Th1 and Th2 cell responses by intestinal helminth infections. To investigate other arms of Th cell responses, we explored the role of a specific dietary immunomodulatory, RA, in controlling mucosal T cell responses. RA is an important mediator of intestinal immunity 221, particularly with respect to T cell differentiation and function 222. However, the processes that control RA signaling and responsiveness in T cells, such as RA metabolism are poorly defined. Cyp26b1 is the primary negative regulator of RA signaling that is highly induced in T cells upon stimulation with RA 169, but the role of Cyp26b1 has not be well characterized in vivo. Therefore, to better assess the function of Cyp26b1 in T cells, mice were generated harboring a conditional KO of Cyp26b1 specifically in T cells. Developmentally we found that mice with T cells deficient in Cyp26b1 still had normal lymphoid development and had no overt health defects in the steady state. However, upon isolation and in vitro polarization of CD4+ T cells, we found that Cyp26b1-deficient T cells had a higher propensity to differentiate into both iTreg and Th17 cells. We also show that Cyp26b1 is important in the effector function of T cells during the induction of experimental colitis in vivo. Together these results highlight an important role of Cyp26b1 in modulating T cell differentiation, function, and ability to cause disease during chronic intestinal inflammation.   79  4.2 Materials and methods 4.2.1 Mice  Cyp26b1fl/fl mice were originally derived and obtained from Dr. Martin Petkovich (Queens University) 223. Cyp26b1fl/fl mice were crossed with Cd4-Cre mice to generate T cell-specific Cyp26b1−/− knockout mice. Rag1−/− mice were originally obtained from Jackson Labs. All mice were bred and housed in specific pathogen-free animal facilities at the UBC Biomedical Research Centre.   4.2.2 In vitro T cell polarization assays  Spleens and lymph nodes were passed through 70 µm strainers and pooled before isolation of CD4+ T cells using EasySep mouse CD4+ enrichment kits on a RoboSep (STEMCELL, Vancouver, BC). Cells were enumerated by hemocytometer and 5x105 cells per well were cultured onto plates coated with 1 µg/ml α-CD3/α-CD28 under either iTreg cell- (10 ng/ml IL-2 and TGF-β), or Th17 cell- (1 ng/ml TGF-β, 10 ng/ml IL-1β, IL-6, IL-23, TNF-α, 10 µg/ml α-IL-4 and α-IFN-γ) polarizing conditions for 6 days at 37° C and 4% CO2; CTCM was the primary cell culture medium whereas X-VIVO 20 (Lonza, Basel, CH) was used for serum-free cultures.   4.2.3 Treg suppression assay  CD4+ T cells were isolated from spleens and lymph nodes as above and a PE-selection kit (STEMCELL) was used to further isolate CD25+ Treg cells. CD25− conventional T cells (Tc) were labeled with CFSE and plated in CTCM at 7x104 cells per well in the presence of 104 mouse T-activator beads (Gibco) and titrated with 2-fold increments of Treg cells (Tc:Treg from 80  32:1 to 1:1). Percent suppression of Tc cells was measured by flow cytometry and was normalized to unsuppressed control samples.  4.2.4 In vitro stimulation of CCR9 expression  CD4+ T cells were isolated from spleens and lymph nodes as above and stimulated with plate-bound anti-CD3/CD28 (1 μg/ml) with 10 ng/ml IL-2, in the presence or absence of 10 nM ATRA for 2 days at 37° C and 4% CO2. Cells were re-plated in the absence of antibodies and maintained on the same concentration of IL-2 and ATRA for another 2 days of cell culture prior to surface staining of CCR9 expression for flow cytometry.  4.2.5 T cell proliferation assay CD4+ T cells were isolated from spleens and lymph nodes as above and suspended in PBS containing 5% FBS. An aliquot of cells was saved to serve as unlabeled controls. 10x106 cells/ml were incubated with 1.25 µM CFSE at room temperature for 5 min. Immediately following CFSE-labeling, the reaction was quenched with pure FBS and the cells were subsequently washed with 5% FBS-PBS. Cells were plated at a density of 5x105 cells per well in a 96-well tissue culture plate under T cell polarizing conditions described above.  4.2.6 Antibodies and flow cytometry Cells were stimulated with PMA (50 ng/ml), ionomycin (750 ng/ml), and Brefeldin A (10 µg/ml) and stained for flow cytometry using the Foxp3/intracellular staining kit and fixable viability dyes (eBioscience); antibodies used: CD4 (GK1.5), CD25 (PC61.5), CCR9 (eBio-CW1.2), Foxp3 (FJK-16s), IFN-γ (XMG1.2), IL-17a (eBio1787), and LPAM-1 (integrin α4β7) 81  (DATK-32) (eBioscience). Samples were analyzed using a LSR-II (BD Biosciences) and FlowJo software (Tree Star).  4.2.7 RNA isolation and quantitative real-time PCR  RNA was isolated from colonic tissues by mechanical disruption and the TRIzol method according to the manufacturer's instructions (Ambion, Austin, TX). RNA was purified from isolated CD4+ T cells using RNeasy mini kits (Qiagen).  Reverse transcription was used to generate cDNA and qPCR was performed using SYBR green primer sets.  Reactions were run on an ABI 7900 real-time PCR machine (Applied Biosystems).  Samples were normalized relative to beta actin (Actb). Primers and sequences used are shown in Table 4.1.  Primer Forward (5’-3’) Reverse (5’-3’) Cyp26b1 GCAAGATCCTACTGGGCGAAC TTGGGCAGGTAGCTCTCAAGT Cd4 CTTCGCAGTTTGATCGTTTTGAT CCGGACTGAAGGTCACTTTGA Il17a AGCAGCGATCATCCCTCAAAG TCACAGAGGGATATCTATCAGGGTC Ifng GGATGCATTCATGAGTATTGCC CCTTTTCCGCTTCCTGAGG Tnfa CATCTTCTCAAAATTCGAGTGACAA TGGGAGTAGACAAGGTACAACCC Foxp3 CCCAGGAAAGACAGCAACCTT TTCTCACAACCAGGCCACTTG Batf CACAGAAAGCCGACACCCTT GCTGTTTGATCTCTTTGCGGA  Itga4 GATGCTGTTGTTGTACTTCGGG ACCACTGAGGCATTAGAGAGC Itgb7 ACCTGAGCTACTCAATGAAGGA CACCGTTTTGTCCACGAAGG Ccr9 CTGGTATTGCACAAGAGTGAAGA CCACACTGATGCACATGATGA  Table 4.1 Primer sets used for qRT-PCR analysis.  82  4.2.8 T cell transfer model of colitis  Spleens and peripheral lymph nodes from donor mice were processed for CD4+ T cell isolation. 4.5x105 FACS-sorted CD4+CD45RBhiCD25− T cells were transferred into recipient Rag1−/− mice i.p. to induce colitis. Weight loss and disease progression were monitored each week. Weight loss of 20% was considered the humane endpoint. Colons were scored for gross pathology by the presence of intestinal wall thickening (rigid appearance and texture), shortening of length, and presence of bloody stool. Proximal colons were processed for histological hematoxylin and eosin staining and blindly scored for disease (inflammatory cell infiltration, loss of epithelial architecture, thickening of colonic wall) as previously described 224. Spleens and mesLN cells were stimulated with 1 µg/ml α-CD3/α-CD28 overnight, then stimulated with PMA, ionomycin, and Brefeldin-A for 5 hours, and were intracellularly stained for flow cytometry.  4.2.9 Statistics  Results are presented as mean ± SEM.  Statistical significance between two groups was determined by unpaired Student’s t-test while comparisons between 3 or more groups were made by ANOVA with a Bonferroni post-hoc test using Prism software (GraphPad).  Results were considered significant with a P value of <0.05.   83  4.3 Results 4.3.1 Expression of Cyp26b1 is dispensable for T cell development  Mice with a germline deletion of Cyp26b1 display severe bone and limb abnormalities and die in utero 225. In order to assess the role of Cyp26b1 in adult T cells, we generated mice with a T cell-specific deletion of Cyp26b1 by breeding Cyp26b1fl/fl mice with mice expressing the Cre recombinase under the control of the Cd4 promoter/enhancer (here termed Cyp26b1−/− mice). Cyp26b1−/− mice developed normally into adult-hood, displayed no gross defects and were born with expected Mendelian ratios compared to Cyp26b1fl/fl littermates. We failed to observe any differences in the frequency of CD4+ and CD8+ single-positive, or CD4+CD8+ double-positive thymocytes in Cyp26b1−/− mice (Figure 4.1). Further, Cyp26b1−/− mice had equivalent frequencies of CD4+ and CD8+ cells in the spleen and mesenteric lymph nodes (mesLN) compared to Cyp26b1fl/fl mice. Thus, Cyp26b1 is not required for naïve T cell development in the thymus or periphery.   Figure 4.1 Cyp26b1 is dispensable for normal lymphoid development. 84  Cyp26b1 was specifically deleted in T cells. Thymus, spleen and mesenteric lymph nodes (mesLNs) from Cyp26b1fl/fl and Cyp26b1−/− mice were analyzed for CD4+ and CD8+ cell frequencies by flow cytometry. Data are from one representative experiment of 2 independent experiments (n = 3–4 per experiment).  4.3.2 Naturally-occurring Treg cells are normal in Cyp26b1−/− mice  The role of RA signaling in the development of thymic-derived nTreg cell development has not been examined in detail, although RAR-activating retinoids have been shown to be produced within the thymus 226. We examined the frequency and function of nTreg cells in Cyp26b1−/− mice. We observed equivalent frequencies of nTreg cells in the spleens of Cyp26b1fl/fl and Cyp26b1−/− mice (Figure 4.2A), suggesting that RA signaling is not a major determinant of nTreg cell development. Further, Cyp26b1 was dispensable for the suppressive ability of nTreg cells (Figure 4.2B), as nTreg cells from either Cyp26b1fl/fl or Cyp26b1−/− mice were able to suppress effector T cell proliferation equivalently. Thus, Cyp26b1-dependent RA metabolism is not required for nTreg cell development and suppressive function.  85   Figure 4.2 Cyp26b1 is not required for nTreg cell development and suppressive function. T cells were isolated from spleens of Cyp26b1fl/fl and Cyp26b1−/− mice. (A) CD4+Foxp3+CD25+ nTreg cell frequencies were determined by flow cytometry. (B) Purified CD4+CD25+ nTreg were cocultured with CFSE-labeled CD4+CD25− conventional T (Tc) cells at increasing ratios and suppression of Tc cells was measured by flow cytometry. Data in (A) are from one representative experiment of 3 independent experiments (n = 3-4 per experiment); Data in (B) are from a single experiment.  4.3.3 Cyp26b1 modulates iTreg and Th17 cell polarization in vitro  RA plays an important role in the differentiation of naive CD4+ T cells into iTreg and Th17 cells 162,164,227. To directly test whether Cyp26b1-dependent regulation of RA signaling controls iTreg or Th17 cell differentiation, we stimulated CD4+ T cells from Cyp26b1fl/fl and Cyp26b1−/− mice under iTreg cell- and Th17 cell-promoting conditions. Increased expression of Cyp26b1 was observed in Th17 cells, with a lower expression in iTreg cells (Figure 4.3A). Following stimulation under Th17 cell-promoting conditions, we observed a marked increased frequency of IL-17a-producing CD4+ T cells in the absence of Cyp26b1 (Figure 4.3B). These 86  results are consistent with the expression pattern of Cyp26b1 and suggest that induction of Cyp26b1 is required for limiting Th17 cell differentiation. Surprisingly, we also found that the absence of Cyp26b1 resulted in heightened frequencies of CD4+CD25+Foxp3+ iTreg cells (Figure 4.3C), despite the low levels of Cyp26b1 expression observed in iTreg cells. Foxp3+ cells tended to be increased overall under serum-free conditions in the absence of retinoids which is likely due to differences in media composition that may favor T cell proliferation. These results suggest that metabolism of RA is important for limiting iTreg and Th17 cell responses. However, we had not added any exogenous RA to these cultures, suggesting that low levels of serum retinoids affect iTreg and Th17 cell differentiation in the absence of Cyp26b1. To directly test this, we repeated the experiment in serum-free media. Under these conditions, we found equivalent frequencies of iTreg cells and Th17 cells following stimulation of CD4+ T cells from both Cyp26b1fl/fl and Cyp26b1−/− mice. Thus, Cyp26b1 regulates RA signaling in T cells and is critical for limiting iTreg and Th17 cell differentiation. Further, these results suggest that physiological levels of RA in naïve cells impacts effector T cell differentiation.      87     Figure 4.3 Cyp26b1 limits iTreg and Th17 cell differentiation in vitro. CD4+ T cells were isolated from Cyp26b1fl/fl and Cyp26b1−/− mice and cultured in iTreg cell- or Th17 cell-promoting conditions, in either serum-containing media or in serum-free media. (A) Gene expression of Cyp26b1 (normalized relative to Actb) was measured by qRT-PCR. (B) Frequencies of IL-17a+ Th17 cells and (C) Foxp3+ CD25+ iTreg cells were determined by flow cytometry. Data in (A) represent mean±SEM of 4 independent experiments; Data in (B) and (C) are from one representative experiment of 4 independent experiments.  88  4.3.4 Cyp26b1-deficient T cells fail to induce intestinal inflammation   Based on our in vitro results demonstrating a role for Cyp26b1 in limiting iTreg and Th17 cell differentiation, we next examined the role of Cyp26b1 in T cell differentiation in vivo. We employed a well-characterized model of T cell-dependent intestinal inflammation 99. Transfer of CD4+CD45RBhighCD25− naïve T cells isolated from Cyp26b1fl/fl mice into immunodeficient Rag1−/− mice resulted in significant weight loss and morbidity associated with intestinal inflammation (Figure 4.4A, B). In contrast, transfer of T cells isolated from Cyp26b1−/− mice resulted in significantly attenuated disease progression, including decreased weight loss and less severe intestinal inflammation. In contrast to our in vitro results, polyclonal stimulation of cells isolated from mesLNs or spleens resulted in no striking differences in the production of IL-17a by Cyp26b1fl/fl and Cyp26b1−/− T cells (Figure 4.4C). In addition, the frequency of Foxp3+ Treg cells in the mesLN and spleen of Rag1−/−mice that received either Cyp26b1fl/fl and Cyp26b1−/− T cells were also similar (Figure 4.4D). Despite equivalent numbers of Cyp26b1fl/fl and Cyp26b1−/− CD4+ T cells in the intestine as measured by Cd4 gene expression (Figure 4.4E), we observed reduced levels of Il17a gene expression in Rag1–/– mice that received Cyp26b1−/− T cells (Figure 4.4F). Further, consistent with reduced disease, we observed a trend in decreased expression of the pro-inflammatory cytokines Ifng and Tnfa in the intestine of Rag1−/− mice that received Cyp26b1−/− T cells (Figure 4.4G). The reduced capacity of Cyp26b1−/− T cells to cause disease was not due to an increase in Treg cells in the intestine, based on Foxp3 gene expression (Figure 4.4H). Thus, Cyp26b1 is critical for the development of pathological T cell responses in the intestine.  89    Figure 4.4 Cyp26b1-deficient T cells fail to promote intestinal inflammation following adoptive transfer into Rag1−/− mice. CD4+CD45RBhighCD25− naive effector T cells from Cyp26b1fl/fl and Cyp26b1−/− mice were transferred i.p. into Rag1−/−  mice. (A) Weight loss was monitored over the entire disease course. (B) Histological sections of proximal colons stained with hematoxylin and eosin were scored for pathology and colons were scored for gross pathology. Cells from spleens and mesLNs were polyclonally stimulated overnight and stained for (C) IL-17a and (D) 90  Foxp3 and CD25, then measured by flow cytometry. Gene expression of (E) Cd4, (F) Il17a, (G) Ifng and Tnfa, and (H) Foxp3 (normalized relative to Actb) in proximal colons was measured by qRT-PCR. Data in (A–H) are representative of one of 2 independent experiments (n = 7-9 per experiment). *p < 0.05.  4.3.5 Cyp26b1-deficient T cells do not have altered expression of intestinal homing molecules  We next assessed whether Cyp26b1−/− T cells have an impaired ability to express intestinal homing molecules as a possible reason for why these T cells failed to cause disease in our colitis transfer model. Isolated Cyp26b1fl/fl and Cyp26b1−/− T cells showed no difference in integrin α4β7 surface expression after stimulation with or without AtRA (Figure 4.5A). Isolated Cyp26b1fl/fl and Cyp26b1−/− T cells were also stimulated to express the RA-inducible chemokine receptor CCR9 but showed no differences in surface expression upon stimulation with AtRA (Figure 4.5B). Further, Cyp26b1fl/fl and Cyp26b1−/− T cells polarized to Th17 and Treg cell lineages showed no differences in gene expression of the homing molecules α4β7 and CCR9 or the transcription factor BATF recently been shown to regulate RA-induced expression of these homing molecules 228 (Figure 4.5C). Thus, Cyp26b1 in T cells does not regulate RA-induced expression of intestinal homing molecules.  91    Figure 4.5 Deficiency in Cyp26b1 does not alter expression of intestinal homing molecules on T cells. (A) CD4+ T cells were isolated from Cyp26b1fl/fl and Cyp26b1−/− mice and stimulated with α-CD3/CD28 and IL-2 with or without 10 nM AtRA and α4β7 integrin expression was measured by flow cytometry. (B) CD4+ T cells were isolated from Cyp26b1fl/fl and Cyp26b1−/− mice were transiently stimulated with α-CD3/CD28 and IL-2 in the presence or absence of 10 nM AtRA to induce CCR9 expression measured by flow cytometry. (C) Gene expression of Batf, Itga4, Itgb7, and Ccr9 was measured in isolated Cyp26b1fl/fl and Cyp26b1−/− T cells polarized under Th17- and Treg-promoting conditions (normalized relative to Actb). Data in (A) are from a single experiment n=4; Data in (B–C) are representative of 3 independent experiments, n=3 mice per experiment.  92  4.4 Discussion  The Cyp26 family of enzymes is critical for limiting RA responses in vivo. For example, lack of Cyp26 during embryogenesis results in severe developmental defects 167,229. RA signaling in T cells has been shown to be regulated by Cyp26b1 169. Our study aimed to better characterize how control of RA metabolism and responsiveness by Cyp26b1 affects effector T cell differentiation and function in vivo. To do so, we generated a T cell-specific conditional knockout of Cyp26b1 in mice. It is known that RA signaling is not required for normal hematopoiesis but can regulate precursors of the myeloid compartment 230. On the other hand, deficiency in RAR signaling in T cells leads to significant activation defects 166. However, an involvement of Cyp26b1-dependent RA metabolism during T cell development has not been investigated previously. It is known that infants exposed to retinoids in utero have been shown to develop malformations of various organs including the thymus 231. We did not observe any gross developmental defects in Cyp26b1−/− mice, which were completely viable throughout adulthood. In terms of lymphoid development, Cyp26b1−/− mice displayed normal levels of CD4+ and CD8+ populations in the thymus, spleen and mesLN. Thus, our results suggest that RA metabolism in T cells has little effect on normal lymphoid development.  RA has paradoxical roles in controlling the balance between iTreg and Th17 cell differentiation: RA promotes Treg cell differentiation at the expense of Th17 cell development 227,232, yet Th17 cells require physiological concentrations of RA for development and migration to the intestine 164. Another study recently found that RA and TGF-β together induce histone modifications at the Foxp3 locus that promote the stability of iTreg cells 233. Furthermore, the RA-inducible microRNA miR-10a was found to be expressed in both nTreg and iTreg cells, playing an important role in blocking the plasticity of Treg cells 163. Deficiency in Cyp26b1 led 93  to increased frequencies of both iTreg and TH17 cells suggesting that Cyp26b1 plays a role in limiting the differentiation of these T cell lineages. This effect was dependent on the presence of retinoids since serum-free culture conditions eliminated these differences, identifying a role for serum retinoids in T cell differentiation in the absence of Cyp26b1 in vitro. Based on our findings, we propose that Cyp26b1 can alter T cell sensitivity to endogenous retinoids during iTreg and TH17 cell differentiation and plays a role in determining T cell responsiveness to RA.  We also identified a role for T cell-intrinsic expression of Cyp26b1 in vivo. We found that transfer of Cyp26b1−/− T cells into Rag1−/− mice led to a profound reduction in intestinal inflammation as compared to Rag1−/− mice transferred with Cyp26b1fl/fl T cells. Interestingly, we found no differences in the frequencies of iTreg or Th17 cells in the mesLN and spleen. However, despite equivalent levels of CD4+ cells in the intestine, based on Cd4 gene expression and comparable CD4+ cell frequencies (data not shown) in the mesLN, we saw a reduction in the expression of Il17a, Ifng and Tnfa in the colon of Rag1−/− mice transferred with Cyp26b1−/− T cells but no difference in levels of Foxp3 gene expression. Thus, together with our observed comparable suppressive function between Cyp26b1fl/fl and Cyp26b1−/−  Treg cells in vitro, Treg function in our T cell transfer colitis model is likely negligible. The transcription factor BATF was recently shown to regulate the RA-inducible Th cell expression of intestinal homing receptors 228. To assess whether Cyp26b1−/− T cells have an impaired migratory ability, we characterized the ability of Cyp26b1−/− T cells to express RA-inducible intestinal homing molecules integrin α4β7 and the chemokine receptor CCR9 but found no altered capacity of surface expression. Similarly, expression of BATF, Itga4, Itgb7, and Ccr9 were comparable in Cyp26b1fl/fl and Cyp26b1−/−  T cells polarized to become TH17 and Treg cells in vitro. Taken together, our results suggest that the role of RA metabolism in T cell function occurs 94  predominantly in the intestinal tissues, consistent with the high levels of RA synthesizing enzymes expressed in the intestine 153,154,234. Thus, the role of Cyp26b1 in promoting the development of T cell-mediated pathology in the intestine does not involve shifting the Treg/Th17 balance but potentially involves modulating effector T cell function at the site of inflammation.  In closing, we have shown that Cyp26b1 can limit the differentiation of iTreg and Th17 cells and is differentially expressed by these lineages to fine tune RA responsiveness. Cyp26b1 in T cells was demonstrated to be required for the development of T cell-mediated chronic inflammation in the colon, potentially by regulating T cell effector function in the intestinal tissue. Thus, Cyp26b1 may serve as a novel therapeutic target to treat inflammatory bowel disease.   95  Chapter 5: Conclusions  5.1 Research summary and significance Summarized below are the major conclusions reached for this dissertation and overarching research significance with respect to the central hypothesis: Th1-driven intestinal helminth infections can impinge on distal Th2 cell responses and that RA metabolism can modulate mucosal T cell responses during homeostasis and disease.  Chapter 2: Alteration of the lung immune microenvironment by the intestinal helminth Trichuris muris 1. Chronic intestinal infection with T. muris induces distal immunological changes in the lungs. 2. Chronic T. muris infection causes an increase in CD4+ T cell-derived IFN-γ and myeloid cell-derived IL-10 in the lungs. 3. Chronic T. muris infection abrogates the development of protease allergen-mediated AAI in the lungs. 4. T. muris-mediated protection from AAI is dependent on both adaptive and innate immune responses. Studies in chapter 2 investigated the effects of intestinal helminth infections on the development of allergic immune responses in the lung. Broadly, our results reiterate the ability of parasitic organisms to potently immunomodulate host responses and particularly responses that are distal to the site of infection. Indeed, this work contributes to our overall understanding of inter-mucosal immune responses, an area that, as of 2016, is poorly characterized in the scientific 96  literature. We showed for the first time that a regulated Th1 cell-associated response during an intestinal helminth infection can impinge on type 2 immune responses in the airways, independent of Treg cell responses. This is in contrast to other previously described intestinal helminth infection models where Treg cells are commonly implicated in protecting against airway disease 235. Furthermore, this is a particularly novel aspect of chronic T. muris infections that has not been described before in the literature. Additionally, no other study has explored the effects of adaptive immune responses to helminth worms on the early, innate-dependent phases of AAI. Overall, our data support the central hypothesis of the dissertation that intestinal helminths can modulate distal mucosal T cell responses. However, we revealed that this cross-mucosal immune regulation also involves regulatory myeloid cells of the innate immune system that can express the anti-inflammatory cytokine IL-10. Our characterization of IL-10-producing myeloid cells in the lung during chronic T. muris infection, and particularly the identification of IL-10+ plasmacytoid DCs, is also a novel aspect of this work. In addition, our model suggests that the timing of these immune responses originating in the intestine is another key factor in distally regulating responses in the lung. Thus, our model highlights a complex interplay between distinct mucosal tissues that involves non-redundant roles for adaptive and innate immune cells during helminth-mediated regulation of allergic responses.  In terms of the broader implications of this work, these results further support the concept of manipulating mucosal immune cross-talk as a therapeutic means to treat T cell-mediated allergic disease. Studies have already been conducted to test the efficacy of live intestinal helminths in humans to treat allergic disease 236. However, therapies need not necessarily involve the use of live helminth infections which have been largely met with inconsistent results in these clinical trials in addition to safety concerns 237. Indeed, our results using the HDM model of 97  asthma suggest that chronic helminth infections may only be a preventative measure for allergic disease rather than being curative of pre-existing disease. This is in line with more recent iterations of the Hygiene Hypothesis which proposes that infectious stimuli encountered during a critical period of immune priming early in life, prior to the exposure of allergens, are what drive protective responses against allergic disease 110. Therefore, it is likely that our results would better serve as a model to identify drug-targetable immune pathways that bypass the difficulties with live infections to treat allergic disease.  Chapter 3: Chronic Trichuris muris infection affects hematopoiesis in the bone marrow 1. Chronic but not acute intestinal T. muris infection affects the function of hematopoietic progenitor cells in the BM. 2. Chronic T. muris infection functionally stimulates myelopoiesis in the BM. 3. Alterations in hematopoiesis elicited by chronic T. muris infection are largely mediated by IFN-γ. 4. T cells expressing IFN-γ accumulate in the BM during chronic T. muris infection. Overall, the studies in chapter 3 demonstrate that there are broad, systemic effects of chronic intestinal helminth infections with respect to host hematopoietic processes occurring in the BM. With respect to the field at large, this is the first study to investigate the impact of intestinal helminths on the BM. Further, the novelty of these findings are also bolstered by our use of the T. muris model of infection where we were able to compare the effects of intestinal Th1 and Th2 cell responses on hematopoiesis.  The pro-inflammatory cytokine IFN-γ is widely recognized to potently affect hematopoietic responses during chronic inflammation and infections 196. IFN-γ produced during 98  Th1 responses has been previously established to favour granulopoiesis in the BM in experimental models ranging from chronic colitis to mycobacterial lung infections 207,238. Thus, this work provides further evidence that mucosal Th1 cell responses, mounted during a variety of inflammatory contexts, have convergent mechanistic effects on hematopoiesis in the BM.  Furthermore, these results offer a potential underlying cellular mechanism for the findings presented in chapter 2, whereby the BM may serve as an intermediate response site between the intestine and the lungs during chronic helminth infection. Therefore, these results provide further support to the central hypothesis of this dissertation, adding another layer of complexity where intestinal helminths may indirectly modulate cross-mucosal immune responses via the BM.   Chapter 4: The RA-metabolizing enzyme Cyp26b1 controls Th cell differentiation and function 1. Cyp26b1 is dispensable for normal lymphoid cell development. 2. Cyp26b1 is differentially induced during iTreg and Th17 cell differentiation. 3. Cyp26b1 is essential for the development of T cell-driven colitis. The critical role of RA in regulating intestinal immunity, mucosal T cell responses, and the induction of Treg cells is well-established 239. However, prior to our studies, characterization of the T cell intrinsic regulation of RA was limited to only a single study by Takeuchi et al. in 2011 which tested the role of RA metabolizing enzymes in vitro using inhibitor compounds 169. While this study established Cyp26b1 as the primary cytochrome p450 enzyme in T cells induced by RA signalling, more studies were needed to investigate the functional significance of this enzyme with regards to T cell differentiation and function. Our studies were the first to test the role Cyp26b1 specifically in CD4+ T cells, through the use of T cell conditional KO mice, 99  enabling us to determine an in vivo role for T cell-intrinsic Cyp26b1 while circumventing the off-target effects of pharmacological inhibitors used by prior studies. The role of RA during Th cell differentiation has been controversial – one study previously suggested that RA favours iTreg cell differentiation at the expense of the Th17 cell lineage 232 – while another study later proposed that RA is indispensable for Th17 cell function in vivo 164. In our work, we found that in vitro iTreg and Th17 cells differentially upregulate Cyp26b1, possibly as a means to fine tune the cellular levels of RA for each lineage. While we saw enhanced iTreg and Th17 cell differentiation in the absence of Cyp26b1 in vitro, our in vivo results from the T cell transfer colitis model did not suggest increased iTreg or Th17 cell differentiation after transfer of Cyp26b1−/− T cells. The precise role of RA is further complicated by the fact that RA signalling is indispensable for general effector T cell differentiation and function 165. However, perturbing the regulation of RA by deficiency in Cyp26b1 resulted in effector T cells being unable to cause disease in our colitis model. Together, this suggests that the role of RA in modulating T cell differentiation and function is context dependent and likely changes in response to different inflammatory and cytokine milieus. In sum, this work supports the central hypothesis of this dissertation that RA metabolism can modulate mucosal T cell responses. However, a study that was published concurrently with our work reported that human polymorphism in the CYP26B1 gene is associated with an increased risk for CD 240. Given these paradoxical findings in addition to the diverse role of RA in immunity, future work must be conducted before Cyp26b1 can be considered a potential therapeutic target for treating IBD.   100  5.2 Study limitations Although many of the goals and expectations for this dissertation project were met, a number of study limitations and caveats must be acknowledged. Most of these issues concern the nature of the animal models, mechanistic scope of our models, and the clinical translatability of experimental results derived from these animal studies. The mouse strains and disease models used in our studies have certain limitations that certainly impact the interpretability of the data. The use of multiple mouse strains would strengthen our studies by ensuring our results are not dependent on a particular genetic background. However, all studies conducted in chapters 2-4 only used mice that were on the C57BL/6 genetic background. The sole use of C57BL/6-derived mice in these studies was primarily justified by the fact that most of available reporter strains are typically bred on this genetic background and the genetic tools, such as Cre mice, were only derived on the C57BL/6 background. Consequently, studies using the lung inflammation models in chapter 2 could not include any measures of airway resistance, a key disease indicator for airway remodeling during asthma 241. This is due to previous reports that C57BL/6 mice are very poor responders to methacholine challenge, a conventional assay used to measure airway hyperreactivity 242,243. Thus, these limitations for chapter 2 prevent us from describing our experimental results in the context of a true asthma model but rather as a model of AAI. Therefore, our models need to be repeated using mouse strains that are amenable to standard methods of measuring airway resistance, such as Balb/c mice. Other non-invasive methods to study airway remodeling have been developed, such as in vivo micro computed tomography 244, which would also help circumvent these limitations. 101  Other deficiencies in this dissertation derive from the complexity of the experimental models used that limited our ability to define heavily detailed mechanisms in these studies. This is exemplified in chapters 2-3 where we were unable to provide an exact linear mechanism of immune regulation during chronic T. muris infection. One reason for this is the involvement of the cytokines IFN-γ and IL-10 which have pleiotropic effects on the immune system and are also known to display positive and negative feedback regulation 245,246, hence making it difficult to define the absolute causality of the responses induced in our model. As opposed to systemically neutralizing these cytokines, using mice with cell-specific KO of the IFN-γ or IL-10 receptors could define a more detailed role for these cytokines in our model. As well, the mechanisms upstream of the induction of these pleiotropic cytokines should be investigated. A detailed mechanism for the studies in chapter 4 was not described since we did not characterize intracellular RA levels or changes in RA-regulated gene expression in Cyp26b1-deficienct T cells. RA-responsiveness in these T cells could be correlated with specific effector responses, cytokine production, or activation state by using RARE reporter mice 247. High throughput sequencing methods could also be used to characterize RA-regulated gene expression in Cyp26b1-deficienct T cells. However, such analyses would nevertheless be difficult to interpret since a broad number of genes are regulated by RA which can involve both direct (RARE regulated) or indirect target gene regulation 248,249. Finally, the overall clinical translatability of the results obtained through this work is difficult to determine without future studies and validation in non-murine systems. This is a general limitation for all animal models of disease since human and mouse immune systems have evolved in drastically distinct contexts and have been exposed to different evolutionary 102  pressures. Since many of the studies in this dissertation used T. muris, a mouse-specific parasite, our results must be carefully validated in other systems for clinical relevance.   5.3 Future directions As the results in this dissertation generated further insight into the regulation of mucosal immune responses, many key questions remain that should be addressed in future studies. Outlined below are only some of the more general questions along with suggested experimental approaches to address them.   What are the specific innate immune mechanisms involved in preventing airway allergen sensitization during chronic T. muris infection?   The findings in Chapter 2 suggested that a chronic Th1 cell-associated intestinal helminth infection impinges on the early sensitization phase of AAI, as evidenced by the fact that timing of the infection was critical in preventing HDM asthma. This suggests that innate immune mechanisms must be involved early on that ultimately determine susceptibility to type 2 inflammation in the lung. ILC2s are important mediators of early type 2 immune responses to protease allergens in the lung 70. While our data suggested that lung ILC2 number and function were not impaired during chronic T. muris infection, it is possible that our use of ex vivo stimulation assays to measure ILC2 cytokine production may have overridden potential defects that were present in vivo. Further it has been shown that IFN-γ can directly antagonize ILC2 function in vivo 250 and thus it would be highly informative to perform our infection/airway challenge model using mice that have IFN-γ receptor-deficient ILC2s. Upstream of lung ILC2 activation is the release of alarmin cytokines such as IL-33 by damaged epithelial cells 50. 103  Interestingly, it has been shown that excretory/secretory products from H. polygyrus are sufficient to prevent AAI in the lung by blocking IL-33 release 198. Therefore, assessing the regulation of IL-33 in the lung during chronic T. muris infection would further elucidate potential innate immune mechanisms involved in protection from AAI.  What is the role of cell migration during gut-to-lung immune responses during T. muris infection?  As we saw an increase in Th1 cells and IL-10+ myeloid cells in lung during chronic T. muris infection (Chapter 2), it is of great interest to determine the precise origin of these cells. These immune cells could possibly be gut-derived that migrate to other mucosal sites during infection. Alternatively, these immune cells could be induced de novo in the lung in response to systemically disseminated inflammatory signals from the gut such as cytokines, chemokines, and worm antigens. One approach to address this would be to block lymphocyte egress from lymphoid tissues using the sphingosine-1-phosphate receptor modulator, FTY720 (Fingolimod) 251 during chronic T. muris infection. This would allow one to test whether the sequestration of T cells in the lymph nodes during intestinal helminth infection affects gut-to-lung immune responses. Another approach would be to use KO mice deficient in specific homing receptors and integrins to further test the role for leukocyte migration in our model.      104  How persistent and transmissible are the changes in the BM during chronic T. muris infection?   The effects of chronic T. muris infection on the BM occurred at the most primitive level of hematopoiesis, affecting HSCs and MPP cells (Chapter 3). It would be interesting to determine the kinetics and persistence of these infection-induced changes in the BM by investigating longer time courses and by manipulating worm burdens by drug-mediated expulsion of the worms. Additionally, one could further assess the transmissibility of these effects on the BM by performing transplant of BM cells from infected donor mice to lethally-irradiated recipients. As well, long-term limiting dilution analysis would allow us to better characterize HSC frequencies and functional potential 252 that may be altered during chronic T. muris infection. This would also be a means to test whether such a BM transplant is sufficient to mediate protection against AAI in the lung seen in Chapter 2. However, it is conceivable that the absence of pro-inflammatory signals generated by the physical presence of worms would abrogate the effects on hematopoiesis.  What are the specific BM-localized effects of chronic T. muris infection?  In Chapter 3, we found that chronic T. muris infection elicited an accumulation of a mixed population of IFN-γ-expressing T cells in the BM. A particularly important question to address is what is the specific function of these BM-localizing T cells? While some of these cells resembled activated effector Th1 cells, a significant proportion of these T cells had a central memory-like phenotype which fits with the notion that the BM is a preferential niche for migratory memory T cells 217. As IFN-γ played a critical role in mediating alterations in hematopoiesis in our model, it would be informative, albeit technically challenging to determine if IFN-γ is acting on the BM systemically through the circulation or whether these BM IFN-γ+ T 105  cells directly regulate hematopoietic progenitors. One experimental approach to test this would be to isolate these IFN-γ+ T cells from the BM of mice chronically infected with T. muris and transplant them into uninfected recipient mice to determine whether they indeed preferentially home to the BM and if they are sufficient to alter hematopoiesis. Additionally, it is necessary to tease out in our model whether IFN-γ is acting directly on HSCs or on other cell types in the BM, such as stromal cells, to influence hematopoiesis. This could be achieved by the targeted deletion of the IFN-γ receptor in various niche stromal cell populations in the BM, such as reticular cells, endothelial cells, and mesenchymal progenitors 253.    Does the microbiota play a key role in driving these cross-mucosal immune responses during T. muris infection?  The intestinal microbiota has an undisputed role in modulating host immunity in a variety of contexts, including susceptibility to asthma 254. Since intestinal helminths share a niche with commensals, there are conceivable interactions between these organisms that could influence host immune responses. For T. muris specifically, it has been shown that parasite egg hatching is induced by interaction with specific bacteria in the intestine 255 and that chronic infection correlates with shifts in the intestinal microbiota composition 199,256. Microbial-derived metabolites, such as short-chain fatty acids, have also been shown to potently modulate host immune responses and affect allergic immune responses in the airways 257. Thus, experiments could be performed using a variety of antibiotic treatments during chronic T. muris infection to test whether the perturbation of the microbiota plays a key role in mediating cross-mucosal immune responses.  106  Are RA-dependent processes influenced by chronic helminth infection?  As of 2015, only a few studies have investigated the role of RA during chronic helminth infection – chronic T. muris infection negatively impacts the synthesis of RA by intestinal DCs 258 and treatment with the RAR agonist Am80 exacerbates intestinal inflammation during T. muris infection 259. Thus, there is some evidence for a relationship between RA and chronic helminth infection. Given that RA is implicated in T cell homing to the intestine from the lungs 260, can modulate BM responses 221 and regulate eosinophil survival 261,262, it would be interesting to test whether RA-regulated processes are implicated in our model of cross-mucosal immune regulation. Some approaches would be to use available RAR agonist and antagonist drugs or conditional KO of RARs and Cyp26 in specific immune lineages during our chronic T. muris infection model.  What is the role of Cyp26b1 in other mucosal T cell-driven diseases?  We found a role for Cyp26b1 in modulating Th cell differentiation and effector T cell responses during colitis in chapter 4. Given the diverse functions of RA, it would be informative to investigate other mucosal inflammatory contexts in which Cyp26b1 could play a role in T cell responses. This could include employing infections models using the Cyp26b1ΔT mice that generate Th cell lineage-biasing responses such as towards helminths (Th2 cell-driven) and viruses (Th1 cell-driven). Additionally, it would be worthwhile to test the role of Cyp26b1 in T cells during autoimmune and allergic models of disease as a means to better characterize the broader functions of this enzyme in a variety of inflammatory milieus.   107  5.4 Concluding remarks  The findings presented in this dissertation highlight the complex mechanisms that can regulate mucosal T cell responses during homeostasis and disease. Thus, the major argument of this work is that intestinal immune responses are immensely responsive to environmental signals, be it infections or dietary factors, that can have profound effects on mucosal T cell-mediated responses both locally and systemically.108  Bibliography 1.  Turner, J.R. Intestinal mucosal barrier function in health and disease. 2009. Nat. Rev. Immunol. 9:799–809. 2.  Neurath, M.F. Cytokines in inflammatory bowel disease. 2014. Nat. Rev. Immunol. 14:329–42. 3.  Mason, K.L., Huffnagle, G.B., Noverr, M.C., & Kao, J.Y. Overview of gut immunology. 2008. Adv. Exp. Med. Biol. 635:1–14. 4.  McGhee, J.R., & Fujihashi, K. Inside the mucosal immune system. 2012. PLoS Biol. 10:e1001397. 5.  Kim, Y.S., & Ho, S.B. Intestinal goblet cells and mucins in health and disease: recent insights and progress. 2010. Curr. Gastroenterol. Rep. 12:319–30. 6.  Guarner, F., & Malagelada, J.-R. Gut flora in health and disease. 2003. Lancet. 361:512–9. 7.  Ganz, T. Defensins: antimicrobial peptides of innate immunity. 2003. Nat. Rev. Immunol. 3:710–20. 8.  Hayday, A.C. Gammadelta T cells and the lymphoid stress-surveillance response. 2009. Immunity. 31:184–96. 9.  Coombes, J.L., & Powrie, F. Dendritic cells in intestinal immune regulation. 2008. Nat. Rev. Immunol. 8:435–46. 10.  Mowat, A.M. Anatomical basis of tolerance and immunity to intestinal antigens. 2003. Nat. Rev. Immunol. 3:331–41. 11.  Brandtzaeg, P. Mucosal immunity: induction, dissemination, and effector functions. 2009. Scand. J. Immunol. 70:505–15. 12.  Macpherson, A.J., McCoy, K.D., Johansen, F.-E., & Brandtzaeg, P. The immune geography of IgA induction and function. 2008. Mucosal Immunol. 1:11–22. 13.  Stone, K.D., Prussin, C., & Metcalfe, D.D. IgE, mast cells, basophils, and eosinophils. 2010. J. Allergy Clin. Immunol. 125:S73–80. 14.  Kolaczkowska, E., & Kubes, P. Neutrophil recruitment and function in health and inflammation. 2013. Nat. Rev. Immunol. 13:159–75. 15.  Mowat, A.M., & Bain, C.C. Mucosal macrophages in intestinal homeostasis and inflammation. 2011. J. Innate Immun. 3:550–64. 16.  Eberl, G., Colonna, M., Di Santo, J.P., & McKenzie, A.N.J. Innate lymphoid cells: A new paradigm in immunology. 2015. Science. 348:1–8. 17.  Zhu, J., Yamane, H., & Paul, W.E. Differentiation of effector CD4 T cell populations. 2010. Annu. Rev. Immunol. 28:445–89. 109  18.  Korn, T., Bettelli, E., Oukka, M., & Kuchroo, V.K. IL-17 and Th17 Cells. 2009. Annu. Rev. Immunol. 27:485–517. 19.  Sakaguchi, S., Miyara, M., Costantino, C.M., & Hafler, D.A. FOXP3+ regulatory T cells in the human immune system. 2010. Nat. Rev. Immunol. 10:490–500. 20.  Lee, G.R., Kim, S.T., Spilianakis, C.G., Fields, P.E., & Flavell, R.A. T helper cell differentiation: regulation by cis elements and epigenetics. 2006. Immunity. 24:369–79. 21.  Wilson, C.B., Rowell, E., & Sekimata, M. Epigenetic control of T-helper-cell differentiation. 2009. Nat. Rev. Immunol. 9:91–105. 22.  Jones, B., & Chen, J. Inhibition of IFN-gamma transcription by site-specific methylation during T helper cell development. 2006. EMBO J. 25:2443–52. 23.  Chang, S., & Aune, T.M. Dynamic changes in histone-methylation “marks” across the locus encoding interferon-gamma during the differentiation of T helper type 2 cells. 2007. Nat. Immunol. 8:723–31. 24.  Wei, G., Wei, L., Zhu, J., Zang, C., Hu-Li, J., Yao, Z., Cui, K., Kanno, Y., Roh, T.-Y., Watford, W.T., Schones, D.E., Peng, W., Sun, H.-W., Paul, W.E., O’Shea, J.J., & Zhao, K. Global mapping of H3K4me3 and H3K27me3 reveals specificity and plasticity in lineage fate determination of differentiating CD4+ T cells. 2009. Immunity. 30:155–67. 25.  Mestecky, J. The common mucosal immune system and current strategies for induction of immune responses in external secretions. 1987. J. Clin. Immunol. 7:265–76. 26.  Holmgren, J., & Czerkinsky, C. Mucosal immunity and vaccines. 2005.  27.  Pasetti, M.F., Simon, J.K., Sztein, M.B., & Levine, M.M. Immunology of gut mucosal vaccines. 2011. Immunol. Rev. 239:125–48. 28.  Czerkinsky, C., & Holmgren, J. Enteric vaccines for the developing world: a challenge for mucosal immunology. 2009. Mucosal Immunol. 2:284–7. 29.  Keely, S., Talley, N.J., & Hansbro, P.M. Pulmonary-intestinal cross-talk in mucosal inflammatory disease. 2012. Mucosal Immunol. 5:7–18. 30.  Penders, J., Stobberingh, E.E., van den Brandt, P.A., & Thijs, C. The role of the intestinal microbiota in the development of atopic disorders. 2007. Allergy. 62:1223–36. 31.  Russell, S.L., Gold, M.J., Hartmann, M., Willing, B.P., Thorson, L., Wlodarska, M., Gill, N., Blanchet, M.-R., Mohn, W.W., McNagny, K.M., & Finlay, B.B. Early life antibiotic-driven changes in microbiota enhance susceptibility to allergic asthma. 2012. EMBO Rep. 13:440–7. 32.  Russell, S.L., Gold, M.J., Reynolds, L.A., Willing, B.P., Dimitriu, P., Thorson, L., Redpath, S.A., Perona-Wright, G., Blanchet, M.-R., Mohn, W.W., Finlay, B.B., & McNagny, K.M. Perinatal antibiotic-induced shifts in gut microbiota have differential effects on inflammatory lung diseases. 2015. J. Allergy Clin. Immunol. 135:100–9. 33.  Round, J.L., & Mazmanian, S.K. Inducible Foxp3+ regulatory T-cell development by a commensal bacterium of the intestinal microbiota. 2010. Proc. Natl. Acad. Sci. U. S. A. 110  107:12204–9. 34.  Ruane, D., Brane, L., Reis, B.S., Cheong, C., Poles, J., Do, Y., Zhu, H., Velinzon, K., Choi, J.-H., Studt, N., Mayer, L., Lavelle, E.C., Steinman, R.M., Mucida, D., & Mehandru, S. Lung dendritic cells induce migration of protective T cells to the gastrointestinal tract. 2013. J. Exp. Med. 210:1871–88. 35.  Bonnegarde-Bernard, A., Jee, J., Fial, M.J., Aeffner, F., Cormet-Boyaka, E., Davis, I.C., Lin, M., Tomé, D., Karin, M., Sun, Y., & Boyaka, P.N. IKKβ in intestinal epithelial cells regulates allergen-specific IgA and allergic inflammation at distant mucosal sites. 2014. Mucosal Immunol. 7:257–67. 36.  Nemoto, Y., Kanai, T., Takahara, M., Oshima, S., Okamoto, R., Tsuchiya, K., Matsumoto, S., & Watanabe, M. Th1/Th17-mediated interstitial pneumonia in chronic colitis mice independent of intestinal microbiota. 2013. J. Immunol. 190:6616–25. 37.  Wang, H., Liu, J.-S., Peng, S.-H., Deng, X.-Y., Zhu, D.-M., Javidiparsijani, S., Wang, G.-R., Li, D.-Q., Li, L.-X., Wang, Y.-C., & Luo, J.-M. Gut-lung crosstalk in pulmonary involvement with inflammatory bowel diseases. 2013. World J. Gastroenterol. 19:6794–804. 38.  Ober, C., & Yao, T.-C. The genetics of asthma and allergic disease: a 21st century perspective. 2011. Immunol. Rev. 242:10–30. 39.  Cho, J.H., & Brant, S.R. Recent insights into the genetics of inflammatory bowel disease. 2011. Gastroenterology. 140:1704–12. 40.  Jenerowicz, D., Silny, W., Dańczak-Pazdrowska, A., Polańska, A., Osmola-Mańkowska, A., & Olek-Hrab, K. Environmental factors and allergic diseases. 2012. Ann. Agric. Environ. Med. 19:475–81. 41.  Molodecky, N.A., & Kaplan, G.G. Environmental risk factors for inflammatory bowel disease. 2010. Gastroenterol. Hepatol. (N. Y). 6:339–46. 42.  Holgate, S.T. Epithelium dysfunction in asthma. 2007. J. Allergy Clin. Immunol. 120:1233–44; quiz 1245–6. 43.  Lambrecht, B.N., & Hammad, H. The immunology of asthma. 2014. Nat. Immunol. 16:45–56. 44.  Bisgaard, H., & Szefler, S. Prevalence of asthma-like symptoms in young children. 2007. Pediatr. Pulmonol. 42:723–8. 45.  Eder, W., Ege, M.J., & von Mutius, E. The asthma epidemic. 2006. N. Engl. J. Med. 355:2226–35. 46.  Barnes, P.J. Glucocorticosteroids: current and future directions. 2011. Br. J. Pharmacol. 163:29–43. 47.  Holgate, S.T., & Polosa, R. Treatment strategies for allergy and asthma. 2008. Nat. Rev. Immunol. 8:218–30. 48.  Hammad, H., & Lambrecht, B.N. Dendritic cells and epithelial cells: linking innate and 111  adaptive immunity in asthma. 2008. Nat. Rev. Immunol. 8:193–204. 49.  Saenz, S.A., Taylor, B.C., & Artis, D. Welcome to the neighborhood: epithelial cell-derived cytokines license innate and adaptive immune responses at mucosal sites. 2008. Immunol. Rev. 226:172–90. 50.  Salimi, M., Barlow, J.L., Saunders, S.P., Xue, L., Gutowska-Owsiak, D., Wang, X., Huang, L.-C., Johnson, D., Scanlon, S.T., McKenzie, A.N.J., Fallon, P.G., & Ogg, G.S. A role for IL-25 and IL-33-driven type-2 innate lymphoid cells in atopic dermatitis. 2013. J. Exp. Med. 210:2939–50. 51.  Kim, B.S., Siracusa, M.C., Saenz, S.A., Noti, M., Monticelli, L.A., Sonnenberg, G.F., Hepworth, M.R., Van Voorhees, A.S., Comeau, M.R., & Artis, D. TSLP elicits IL-33-independent innate lymphoid cell responses to promote skin inflammation. 2013. Sci. Transl. Med. 5:170ra16. 52.  Ito, T., Wang, Y.-H., Duramad, O., Hori, T., Delespesse, G.J., Watanabe, N., Qin, F.X.-F., Yao, Z., Cao, W., & Liu, Y.-J. TSLP-activated dendritic cells induce an inflammatory T helper type 2 cell response through OX40 ligand. 2005. J. Exp. Med. 202:1213–23. 53.  Monticelli, L.A., Sonnenberg, G.F., & Artis, D. Innate lymphoid cells: critical regulators of allergic inflammation and tissue repair in the lung. 2012. Curr. Opin. Immunol. 24:284–9. 54.  Kouro, T., & Takatsu, K. IL-5- and eosinophil-mediated inflammation: from discovery to therapy. 2009. Int. Immunol. 21:1303–9. 55.  Cohn, L., Elias, J.A., & Chupp, G.L. Asthma: mechanisms of disease persistence and progression. 2004. Annu. Rev. Immunol. 22:789–815. 56.  Lund, S., Walford, H.H., & Doherty, T.A. Type 2 Innate Lymphoid Cells in Allergic Disease. 2013. Curr. Immunol. Rev. 9:214–21. 57.  Durrant, D.M., & Metzger, D.W. Emerging roles of T helper subsets in the pathogenesis of asthma. 2010. Immunol. Invest. 39:526–49. 58.  Kips, J., Brusselle, G., Joos, G., Peleman, R., Tavernier, J., Devos, R., & Pauwels, R. Interleukin-12 inhibits antigen-induced airway hyperresponsiveness in mice. 1996. Am. J. Respir. Crit. Care Med. 153:535–9. 59.  van Oosterhout, A.J.M., & Bloksma, N. Regulatory T-lymphocytes in asthma. 2005. Eur. Respir. J. 26:918–32. 60.  Akdis, C.A., & Akdis, M. Mechanisms of immune tolerance to allergens: role of IL-10 and Tregs. 2014. J. Clin. Invest. 124:4678–80. 61.  He, S., Tong, Q., Bishop, D.K., & Zhang, Y. Histone methyltransferase and histone methylation in inflammatory T-cell responses. 2013. Immunotherapy. 5:989–1004. 62.  Corren, J., Lemanske, R.F., Hanania, N.A., Korenblat, P.E., Parsey, M. V, Arron, J.R., Harris, J.M., Scheerens, H., Wu, L.C., Su, Z., Mosesova, S., Eisner, M.D., 112  Bohen, S.P., & Matthews, J.G. Lebrikizumab treatment in adults with asthma. 2011. N. Engl. J. Med. 365:1088–98. 63.  Torgerson, T.R., & Ochs, H.D. Regulatory T cells in primary immunodeficiency diseases. 2007. Curr. Opin. Allergy Clin. Immunol. 7:515–21. 64.  Duan, W., & Croft, M. Control of regulatory T cells and airway tolerance by lung macrophages and dendritic cells. 2014. Ann. Am. Thorac. Soc. 11 Suppl 5:S306–13. 65.  Curotto de Lafaille, M.A., Kutchukhidze, N., Shen, S., Ding, Y., Yee, H., & Lafaille, J.J. Adaptive Foxp3+ regulatory T cell-dependent and -independent control of allergic inflammation. 2008. Immunity. 29:114–26. 66.  Ray, A., Khare, A., Krishnamoorthy, N., Qi, Z., & Ray, P. Regulatory T cells in many flavors control asthma. 2010. Mucosal Immunol. 3:216–29. 67.  Thorburn, A.N., & Hansbro, P.M. Harnessing regulatory T cells to suppress asthma: from potential to therapy. 2010. Am. J. Respir. Cell Mol. Biol. 43:511–9. 68.  Kumar, R.K., Herbert, C., & Foster, P.S. The “classical” ovalbumin challenge model of asthma in mice. 2008. Curr. Drug Targets. 9:485–94. 69.  Novey, H.S., Marchioli, L.E., Sokol, W.N., & Wells, I.D. Papain-induced asthma--physiological and immunological features. 1979. J. Allergy Clin. Immunol. 63:98–103. 70.  Halim, T.Y.F., Krauss, R.H., Sun, A.C., & Takei, F. Lung natural helper cells are a critical source of Th2 cell-type cytokines in protease allergen-induced airway inflammation. 2012. Immunity. 36:451–63. 71.  Gøtzsche, P.C., & Johansen, H.K. House dust mite control measures for asthma: systematic review. 2008. Allergy. 63:646–59. 72.  Gold, M.J., Antignano, F., Halim, T.Y.F., Hirota, J.A., Blanchet, M.-R., Zaph, C., Takei, F., & McNagny, K.M. Group 2 innate lymphoid cells facilitate sensitization to local, but not systemic, TH2-inducing allergen exposures. 2014. J. Allergy Clin. Immunol. 133:1142–8. 73.  Sartor, R.B. Mechanisms of disease: pathogenesis of Crohn’s disease and ulcerative colitis. 2006. Nat. Clin. Pract. Gastroenterol. Hepatol. 3:390–407. 74.  Bernstein, C.N., Wajda, A., Svenson, L.W., MacKenzie, A., Koehoorn, M., Jackson, M., Fedorak, R., Israel, D., & Blanchard, J.F. The epidemiology of inflammatory bowel disease in Canada: a population-based study. 2006. Am. J. Gastroenterol. 101:1559–68. 75.  Molodecky, N.A., Soon, I.S., Rabi, D.M., Ghali, W.A., Ferris, M., Chernoff, G., Benchimol, E.I., Panaccione, R., Ghosh, S., Barkema, H.W., & Kaplan, G.G. Increasing incidence and prevalence of the inflammatory bowel diseases with time, based on systematic review. 2012. Gastroenterology. 142:46–54.e42; quiz e30. 76.  Hendrickson, B.A., Gokhale, R., & Cho, J.H. Clinical aspects and pathophysiology of inflammatory bowel disease. 2002. Clin. Microbiol. Rev. 15:79–94. 113  77.  Soucy, G., Wang, H.H., Farraye, F.A., Schmidt, J.F., Farris, A.B., Lauwers, G.Y., Cerda, S.R., Dendrinos, K.G., & Odze, R.D. Clinical and pathological analysis of colonic Crohn’s disease, including a subgroup with ulcerative colitis-like features. 2012. Mod. Pathol. 25:295–307. 78.  Langan, R.C., Gotsch, P.B., Krafczyk, M.A., & Skillinge, D.D. Ulcerative colitis: diagnosis and treatment. 2007. Am. Fam. Physician. 76:1323–30. 79.  DeRoche, T.C., Xiao, S.-Y., & Liu, X. Histological evaluation in ulcerative colitis. 2014. Gastroenterol. Rep. 2:178–92. 80.  Maul, J., Loddenkemper, C., Mundt, P., Berg, E., Giese, T., Stallmach, A., Zeitz, M., & Duchmann, R. Peripheral and intestinal regulatory CD4+CD25high T cells in inflammatory bowel disease. 2005. Gastroenterology. 128:1868–78. 81.  Shimomura, Y., Mizoguchi, E., Sugimoto, K., Kibe, R., Benno, Y., Mizoguchi, A., & Bhan, A.K. Regulatory role of B-1 B cells in chronic colitis. 2008. Int. Immunol. 20:729–37. 82.  Darrasse-Jèze, G., Deroubaix, S., Mouquet, H., Victora, G.D., Eisenreich, T., Yao, K., Masilamani, R.F., Dustin, M.L., Rudensky, A., Liu, K., & Nussenzweig, M.C. Feedback control of regulatory T cell homeostasis by dendritic cells in vivo. 2009. J. Exp. Med. 206:1853–62. 83.  Pache, I., Rogler, G., & Felley, C. TNF-alpha blockers in inflammatory bowel diseases: practical consensus recommendations and a user’s guide. 2009. Swiss Med. Wkly. 139:278–87. 84.  Geremia, A., Arancibia-Cárcamo, C. V, Fleming, M.P.P., Rust, N., Singh, B., Mortensen, N.J., Travis, S.P.L., & Powrie, F. IL-23-responsive innate lymphoid cells are increased in inflammatory bowel disease. 2011. J. Exp. Med. 208:1127–33. 85.  Peluso, I., Pallone, F., & Monteleone, G. Interleukin-12 and Th1 immune response in Crohn’s disease: pathogenetic relevance and therapeutic implication. 2006. World J. Gastroenterol. 12:5606–10. 86.  Heller, F., Florian, P., Bojarski, C., Richter, J., Christ, M., Hillenbrand, B., Mankertz, J., Gitter, A.H., Bürgel, N., Fromm, M., Zeitz, M., Fuss, I., Strober, W., & Schulzke, J.D. Interleukin-13 is the key effector Th2 cytokine in ulcerative colitis that affects epithelial tight junctions, apoptosis, and cell restitution. 2005. Gastroenterology. 129:550–64. 87.  Harbour, S.N., Maynard, C.L., Zindl, C.L., Schoeb, T.R., & Weaver, C.T. Th17 cells give rise to Th1 cells that are required for the pathogenesis of colitis. 2015. Proc. Natl. Acad. Sci.201415675. 88.  Hueber, W., Sands, B.E., Lewitzky, S., Vandemeulebroecke, M., Reinisch, W., Higgins, P.D.R., Wehkamp, J., Feagan, B.G., Yao, M.D., Karczewski, M., Karczewski, J., Pezous, N., Bek, S., Bruin, G., Mellgard, B., Berger, C., Londei, M., Bertolino, A.P., Tougas, G., & Travis, S.P.L. Secukinumab, a human anti-IL-17A monoclonal antibody, for moderate to severe Crohn’s disease: unexpected results of a 114  randomised, double-blind placebo-controlled trial. 2012. Gut. 61:1693–700. 89.  O’Connor, W., Kamanaka, M., Booth, C.J., Town, T., Nakae, S., Iwakura, Y., Kolls, J.K., & Flavell, R.A. A protective function for interleukin 17A in T cell-mediated intestinal inflammation. 2009. Nat. Immunol. 10:603–9. 90.  Mayne, C.G., & Williams, C.B. Induced and natural regulatory T cells in the development of inflammatory bowel disease. 2013. Inflamm. Bowel Dis. 19:1772–88. 91.  Boehm, F., Martin, M., Kesselring, R., Schiechl, G., Geissler, E.K., Schlitt, H.-J., & Fichtner-Feigl, S. Deletion of Foxp3+ regulatory T cells in genetically targeted mice supports development of intestinal inflammation. 2012. BMC Gastroenterol. 12:97. 92.  Buckner, J.H. Mechanisms of impaired regulation by CD4(+)CD25(+)FOXP3(+) regulatory T cells in human autoimmune diseases. 2010. Nat. Rev. Immunol. 10:849–59. 93.  Hardenberg, G., Steiner, T.S., & Levings, M.K. Environmental influences on T regulatory cells in inflammatory bowel disease. 2011. Semin. Immunol. 23:130–8. 94.  Wang, J., Ioan-Facsinay, A., van der Voort, E.I.H., Huizinga, T.W.J., & Toes, R.E.M. Transient expression of FOXP3 in human activated nonregulatory CD4+ T cells. 2007. Eur. J. Immunol. 37:129–38. 95.  Yan, Y., Kolachala, V., Dalmasso, G., Nguyen, H., Laroui, H., Sitaraman, S. V, & Merlin, D. Temporal and spatial analysis of clinical and molecular parameters in dextran sodium sulfate induced colitis. 2009. PLoS One. 4:e6073. 96.  Siegmund, B., Fantuzzi, G., Rieder, F., Gamboni-Robertson, F., Lehr, H., Hartmann, G., Dinarello, C., Endres, S., & Eigler, A. Neutralization of interleukin-18 reduces severity in murine colitis and intestinal IFN-gamma and TNF-alpha production. 2001. Am. J. Physiol. Integr. Comp. Physiol. 281:R1264–73. 97.  Podolsky, D.K. Lessons from genetic models of inflammatory bowel disease. Acta Gastroenterol. Belg. 60:163–5. 98.  Büchler, G., Wos-Oxley, M.L., Smoczek, A., Zschemisch, N.-H., Neumann, D., Pieper, D.H., Hedrich, H.J., & Bleich, A. Strain-specific colitis susceptibility in IL10-deficient mice depends on complex gut microbiota-host interactions. 2012. Inflamm. Bowel Dis. 18:943–54. 99.  Ostanin, D. V, Bao, J., Koboziev, I., Gray, L., Robinson-Jackson, S. a, Kosloski-Davidson, M., Price, V.H., & Grisham, M.B. T cell transfer model of chronic colitis: concepts, considerations, and tricks of the trade. 2009. Am. J. Physiol. Gastrointest. Liver Physiol. 296:G135–46. 100.  Izcue, A., Coombes, J.L., & Powrie, F. Regulatory T cells suppress systemic and mucosal immune activation to control intestinal inflammation. 2006. Immunol. Rev. 212:256–71. 101.  Powrie, F., Leach, M.W., Mauze, S., Menon, S., Barcomb Caddle, L., & Coffman, R.L. Inhibition of Thl responses prevents inflammatory bowel disease in scid mice reconstituted with CD45RBhi CD4+ T cells. 1994. Immunity. 1:553–62. 115  102.  Cox, F.E.G. History of human parasitology. 2002. Clin. Microbiol. Rev. 15:595–612. 103.  Hotez, P.J., Bundy, D.A.P., Beegle, K., Brooker, S., Drake, L., de Silva, N., Montresor, A., Engels, D., Jukes, M., Chitsulo, L., Chow, J., Laxminarayan, R., Michaud, C., Bethony, J., Correa-Oliveira, R., Shuhua, X., Fenwick, A., & Savioli, L. Helminth infections: soil-transmitted helminth infections and schistosomiasis. 2006 [cited 2015 May 28]. Disease Control Priorities in Developing Countries. 2nd ed. 104.  Strunz, E.C., Addiss, D.G., Stocks, M.E., Ogden, S., Utzinger, J., & Freeman, M.C. Water, sanitation, hygiene, and soil-transmitted helminth infection: a systematic review and meta-analysis. 2014. PLoS Med. 11:e1001620. 105.  McSorley, H.J., & Maizels, R.M. Helminth infections and host immune regulation. 2012. Clin. Microbiol. Rev. 25:585–608. 106.  Maizels, R.M., Hewitson, J.P., & Smith, K.A. Susceptibility and immunity to helminth parasites. 2012. Curr. Opin. Immunol. 24:459–66. 107.  Allen, J.E., & Wynn, T.A. Evolution of Th2 immunity: a rapid repair response to tissue destructive pathogens. 2011. PLoS Pathog. 7:e1002003. 108.  McSorley, H.J., & Maizels, R.M. Helminth infections and host immune regulation. 2012. Clin. Microbiol. Rev. 25:585–608. 109.  McSorley, H.J., Hewitson, J.P., & Maizels, R.M. Immunomodulation by helminth parasites: defining mechanisms and mediators. 2013. Int. J. Parasitol. 43:301–10. 110.  Okada, H., Kuhn, C., Feillet, H., & Bach, J.-F. The “hygiene hypothesis” for autoimmune and allergic diseases: an update. 2010. Clin. Exp. Immunol. 160:1–9. 111.  Weinstock, J. V., & Elliott, D.E. Helminth infections decrease host susceptibility to immune-mediated diseases. 2014. J. Immunol. 193:3239–47. 112.  Wammes, L.J., Mpairwe, H., Elliott, A.M., & Yazdanbakhsh, M. Helminth therapy or elimination: epidemiological, immunological, and clinical considerations. 2014. Lancet. Infect. Dis. 14:1150–62. 113.  Summers, R.W., Elliott, D.E., Urban, J.F., Thompson, R., & Weinstock, J. V. Trichuris suis therapy in Crohn’s disease. 2005. Gut. 54:87–90. 114.  Fleming, J.O. Helminth therapy and multiple sclerosis. 2013. Int. J. Parasitol. 43:259–74. 115.  Johnston, M.J.G., MacDonald, J.A., & McKay, D.M. Parasitic helminths: a pharmacopeia of anti-inflammatory molecules. 2009. Parasitology. 136:125–47. 116.  Kitagaki, K., Businga, T.R., Racila, D., Elliott, D.E., Weinstock, J. V, & Kline, J.N. Intestinal helminths protect in a murine model of asthma. 2006. J. Immunol. 177:1628–35. 117.  Liu, Q., Sundar, K., Mishra, P.K., Mousavi, G., Liu, Z., Gaydo, A., Alem, F., Lagunoff, D., Bleich, D., & Gause, W.C. Helminth infection can reduce insulitis and type 1 diabetes through CD25- and IL-10-independent mechanisms. 2009. Infect. Immun. 77:5347–58. 116  118.  Hang, L., Setiawan, T., Blum, A.M., Urban, J., Stoyanoff, K., Arihiro, S., Reinecker, H.-C., & Weinstock, J. V. Heligmosomoides polygyrus infection can inhibit colitis through direct interaction with innate immunity. 2010. J. Immunol. 185:3184–9. 119.  Donskow-Łysoniewska, K., Krawczak, K., & Doligalska, M. Heligmosomoides polygyrus: EAE remission is correlated with different systemic cytokine profiles provoked by L4 and adult nematodes. 2012. Exp. Parasitol. 132:243–8. 120.  Mangan, N.E., van Rooijen, N., McKenzie, A.N.J., & Fallon, P.G. Helminth-modified pulmonary immune response protects mice from allergen-induced airway hyperresponsiveness. 2006. J. Immunol. 176:138–47. 121.  Elliott, D.E., Li, J., Blum, A., Metwali, A., Qadir, K., Urban, J.F., & Weinstock, J. V. Exposure to schistosome eggs protects mice from TNBS-induced colitis. 2003. Am. J. Physiol. Gastrointest. Liver Physiol. 284:G385–91. 122.  Zaccone, P., Fehérvári, Z., Jones, F.M., Sidobre, S., Kronenberg, M., Dunne, D.W., & Cooke, A. Schistosoma mansoni antigens modulate the activity of the innate immune response and prevent onset of type 1 diabetes. 2003. Eur. J. Immunol. 33:1439–49. 123.  La Flamme, A.C., Ruddenklau, K., & Backstrom, B.T. Schistosomiasis decreases central nervous system inflammation and alters the progression of experimental autoimmune encephalomyelitis. 2003. Infect. Immun. 71:4996–5004. 124.  Espinoza-Jiménez, A., Rivera-Montoya, I., Cárdenas-Arreola, R., Morán, L., & Terrazas, L.I. Taenia crassiceps infection attenuates multiple low-dose streptozotocin-induced diabetes. 2010. J. Biomed. Biotechnol. 2010:850541. 125.  Park, H.-K., Cho, M.K., Choi, S.H., Kim, Y.S., & Yu, H.S. Trichinella spiralis: infection reduces airway allergic inflammation in mice. 2011. Exp. Parasitol. 127:539–44. 126.  Saunders, K.A., Raine, T., Cooke, A., & Lawrence, C.E. Inhibition of autoimmune type 1 diabetes by gastrointestinal helminth infection. 2007. Infect. Immun. 75:397–407. 127.  Khan, W.I., Blennerhasset, P.A., Varghese, A.K., Chowdhury, S.K., Omsted, P., Deng, Y., & Collins, S.M. Intestinal nematode infection ameliorates experimental colitis in mice. 2002. Infect. Immun. 70:5931–7. 128.  Wu, Z., Nagano, I., Asano, K., & Takahashi, Y. Infection of non-encapsulated species of Trichinella ameliorates experimental autoimmune encephalomyelitis involving suppression of Th17 and Th1 response. 2010. Parasitol. Res. 107:1173–88. 129.  Wohlleben, G., Trujillo, C., Müller, J., Ritze, Y., Grunewald, S., Tatsch, U., & Erb, K.J. Helminth infection modulates the development of allergen-induced airway inflammation. 2004. Int. Immunol. 16:585–96. 130.  Bouchery, T., Kyle, R., Ronchese, F., & Le Gros, G. The differentiation of CD4(+) T-helper cell subsets in the context of helminth parasite infection. 2014. Front. Immunol. 5:487. 131.  Mishra, P.K., Patel, N., Wu, W., Bleich, D., & Gause, W.C. Prevention of type 1 diabetes through infection with an intestinal nematode parasite requires IL-10 in the 117  absence of a Th2-type response. 2013. Mucosal Immunol. 6:297–308. 132.  Hewitson, J.P., Grainger, J.R., & Maizels, R.M. Helminth immunoregulation: the role of parasite secreted proteins in modulating host immunity. 2009. Mol. Biochem. Parasitol. 167:1–11. 133.  Bethony, J., Brooker, S., Albonico, M., Geiger, S.M., Loukas, A., Diemert, D., & Hotez, P.J. Soil-transmitted helminth infections: ascariasis, trichuriasis, and hookworm. 2006. Lancet. 367:1521–32. 134.  Behnke, J., & Wakelin, D. Survival of Trichuris-muris in wild populations of its natural hosts. 1973. Parasitology. 67:157–64. 135.  Grencis, R.K., Humphreys, N.E., & Bancroft, A.J. Immunity to gastrointestinal nematodes: mechanisms and myths. 2014. Immunol. Rev. 260:183–205. 136.  Klementowicz, J.E., Travis, M.A., & Grencis, R.K. Trichuris muris: a model of gastrointestinal parasite infection. 2012. Semin. Immunopathol. 34:815–28. 137.  Else, K., Hultner, L., & Grencis, R. Cellular immune-responses to the murine nematode parasite Trichuris-muris: differential induction of Th-cell subsets in resistant versus susceptible mice. 1992. Immunology. 75:232–7. 138.  Bhardwaj, E.K., Else, K.J., Rogan, M.T., & Warhurst, G. Increased susceptibility to Trichuris muris infection and exacerbation of colitis in Mdr1a-/- mice. 2014. World J. Gastroenterol. 20:1797–806. 139.  Hurst, R.J.M., & Else, K.J. Trichuris muris research revisited: a journey through time. 2013. Parasitology. 140:1325–39. 140.  Schopf, L.R., Hoffmann, K.F., Cheever, A.W., Urban, J.F., & Wynn, T.A. IL-10 is critical for host resistance and survival during gastrointestinal helminth infection. 2002. J. Immunol. 168:2383–92. 141.  Fraser, P.D., & Bramley, P.M. The biosynthesis and nutritional uses of carotenoids. 2004. Prog. Lipid Res. 43:228–65. 142.  Li, Y., Wongsiriroj, N., & Blaner, W.S. The multifaceted nature of retinoid transport and metabolism. 2014. Hepatobiliary Surg. Nutr. 3:126–39. 143.  Rhinn, M., & Dollé, P. Retinoic acid signalling during development. 2012. Development. 139:843–58. 144.  Hall, J.A., Grainger, J.R., Spencer, S.P., & Belkaid, Y. The role of retinoic acid in tolerance and immunity. 2011. Immunity. 35:13–22. 145.  Humphrey, J.H., West, K.P., & Sommer,  a. Vitamin A deficiency and attributable mortality among under-5-year-olds. 1992. Bull. World Health Organ. 70:225–32. 146.  Semba, R.D., Muhilal, Scott, A.L., Natadisastra, G., Wirasasmita, S., Mele, L., Ridwan, E., West, K.P., & Sommer, A. Depressed immune response to tetanus in children with vitamin A deficiency. 1992. J. Nutr. 122:101–7. 118  147.  Sommer, A., Tarwotjo, I., Hussaini, G., & Susanto, D. Increased mortality in children with mild vitamin A deficiency. 1983. Lancet (London, England). 2:585–8. 148.  Villamor, E., & Fawzi, W.W. Vitamin A supplementation: implications for morbidity and mortality in children. 2000. J. Infect. Dis. 182 Suppl :S122–33. 149.  von Boehmer, H. Oral tolerance: is it all retinoic acid? 2007. J. Exp. Med. 204:1737–9. 150.  McCullough, F.S., Northrop-Clewes, C.A., & Thurnham, D.I. The effect of vitamin A on epithelial integrity. 1999. Proc. Nutr. Soc. 58:289–93. 151.  Cassani, B., Villablanca, E.J., De Calisto, J., Wang, S., & Mora, J.R. Vitamin A and immune regulation: role of retinoic acid in gut-associated dendritic cell education, immune protection and tolerance. 2012. Mol. Aspects Med. 33:63–76. 152.  Belatik, A., Hotchandani, S., Bariyanga, J., & Tajmir-Riahi, H.A. Binding sites of retinol and retinoic acid with serum albumins. 2012. Eur. J. Med. Chem. 48:114–23. 153.  Iwata, M. Retinoic acid production by intestinal dendritic cells and its role in T-cell trafficking. 2009. Semin. Immunol. 21:8–13. 154.  Molenaar, R., Knippenberg, M., Goverse, G., Olivier, B.J., de Vos, A.F., O’Toole, T., & Mebius, R.E. Expression of retinaldehyde dehydrogenase enzymes in mucosal dendritic cells and gut-draining lymph node stromal cells is controlled by dietary vitamin A. 2011. J. Immunol. 186:1934–42. 155.  Coleman, M.M., Ruane, D., Moran, B., Dunne, P.J., Keane, J., & Mills, K.H.G. Alveolar macrophages contribute to respiratory tolerance by inducing FoxP3 expression in naive T cells. 2013. Am. J. Respir. Cell Mol. Biol. 48:773–80. 156.  Lee, H.-P., Casadesus, G., Zhu, X., Lee, H., Perry, G., Smith, M.A., Gustaw-Rothenberg, K., & Lerner, A. All-trans retinoic acid as a novel therapeutic strategy for Alzheimer’s disease. 2009. Expert Rev. Neurother. 9:1615–21. 157.  Majumdar, A., Petrescu, A.D., Xiong, Y., & Noy, N. Nuclear translocation of cellular retinoic acid-binding protein II is regulated by retinoic acid-controlled SUMOylation. 2011. J. Biol. Chem. 286:42749–57. 158.  Leid, M., Kastner, P., & Chambon, P. Multiplicity generates diversity in the retinoic acid signalling pathways. 1992. Trends Biochem. Sci. 17:427–33. 159.  Guo, Y., Lee, Y.-C., Brown, C., Zhang, W., Usherwood, E., & Noelle, R.J. Dissecting the role of retinoic acid receptor isoforms in the CD8 response to infection. 2014. J. Immunol. 192:3336–44. 160.  Iwata, M., Hirakiyama, A., Eshima, Y., Kagechika, H., Kato, C., & Song, S.-Y. Retinoic acid imprints gut-homing specificity on T cells. 2004. Immunity. 21:527–38. 161.  Kim, M.H., Taparowsky, E.J., & Kim, C.H. Retinoic Acid Differentially Regulates the Migration of Innate Lymphoid Cell Subsets to the Gut. 2015. Immunity. 162.  Benson, M.J., Pino-Lagos, K., Rosemblatt, M., & Noelle, R.J. All-trans retinoic acid mediates enhanced T reg cell growth, differentiation, and gut homing in the face of high 119  levels of co-stimulation. 2007. J. Exp. Med. 204:1765–74. 163.  Takahashi, H., Kanno, T., Nakayamada, S., Hirahara, K., Sciumè, G., Muljo, S. a, Kuchen, S., Casellas, R., Wei, L., Kanno, Y., & O’Shea, J.J. TGF-β and retinoic acid induce the microRNA miR-10a, which targets Bcl-6 and constrains the plasticity of helper T cells. 2012. Nat. Immunol. 13:587–95. 164.  Wang, C., Kang, S.G., HogenEsch, H., Love, P.E., & Kim, C.H. Retinoic acid determines the precise tissue tropism of inflammatory Th17 cells in the intestine. 2010. J. Immunol. 184:5519–26. 165.  Pino-Lagos, K., Guo, Y., Brown, C., Alexander, M.P., Elgueta, R., Bennett, K. a, De Vries, V., Nowak, E., Blomhoff, R., Sockanathan, S., Chandraratna, R. a, Dmitrovsky, E., & Noelle, R.J. A retinoic acid-dependent checkpoint in the development of CD4+ T cell-mediated immunity. 2011. J. Exp. Med. 208:1767–75. 166.  Hall, J. a, Cannons, J.L., Grainger, J.R., Dos Santos, L.M., Hand, T.W., Naik, S., Wohlfert, E. a, Chou, D.B., Oldenhove, G., Robinson, M., Grigg, M.E., Kastenmayer, R., Schwartzberg, P.L., & Belkaid, Y. Essential role for retinoic acid in the promotion of CD4(+) T cell effector responses via retinoic acid receptor alpha. 2011. Immunity. 34:435–47. 167.  Pennimpede, T., Cameron, D. a, MacLean, G. a, Li, H., Abu-Abed, S., & Petkovich, M. The role of CYP26 enzymes in defining appropriate retinoic acid exposure during embryogenesis. 2010. Birth Defects Res. A. Clin. Mol. Teratol. 88:883–94. 168.  Thatcher, J.E., & Isoherranen, N. The role of CYP26 enzymes in retinoic acid clearance. 2009. Expert Opin. Drug Metab. Toxicol. 5:875–86. 169.  Takeuchi, H., Yokota, A., Ohoka, Y., & Iwata, M. Cyp26b1 regulates retinoic acid-dependent signals in T cells and its expression is inhibited by transforming growth factor-β. 2011. PLoS One. 6:e16089. 170.  Figueiredo, C.A., Barreto, M.L., Rodrigues, L.C., Cooper, P.J., Silva, N.B., Amorim, L.D., & Alcantara-Neves, N.M. Chronic intestinal helminth infections are associated with immune hyporesponsiveness and induction of a regulatory network. 2010. Infect. Immun. 78:3160–7. 171.  Crompton, D.W.T., & Nesheim, M.C. Nutritional impact of intestinal helminthiasis during the human life cycle. 2002. Annu. Rev. Nutr. 22:35–59. 172.  McSorley, H.J., O’Gorman, M.T., Blair, N., Sutherland, T.E., Filbey, K.J., & Maizels, R.M. Suppression of type 2 immunity and allergic airway inflammation by secreted products of the helminth Heligmosomoides polygyrus. 2012. Eur. J. Immunol. 42:2667–82. 173.  Madan, R., Demircik, F., Surianarayanan, S., Allen, J.L., Divanovic, S., Trompette, A., Yogev, N., Gu, Y., Khodoun, M., Hildeman, D., Boespflug, N., Fogolin, M.B., Gröbe, L., Greweling, M., Finkelman, F.D., Cardin, R., Mohrs, M., Müller, W., Waisman, A., Roers, A., & Karp, C.L. Nonredundant roles for B cell-derived IL-10 in immune counter-regulation. 2009. J. Immunol. 183:2312–20. 120  174.  Taylor, B.C., Zaph, C., Troy, A.E., Du, Y., Guild, K.J., Comeau, M.R., & Artis, D. TSLP regulates intestinal immunity and inflammation in mouse models of helminth infection and colitis. 2009. J. Exp. Med. 206:655–67. 175.  Arora, S., McDonald, R.A., Toews, G.B., & Huffnagle, G.B. Effect of a CD4-depleting antibody on the development of Cryptococcus neoformans-induced allergic bronchopulmonary mycosis in mice. 2006. Infect. Immun. 74:4339–48. 176.  Hadidi, S., Antignano, F., Hughes, M.R., Wang, S.K.H., Snyder, K., Sammis, G.M., Kerr, W.G., McNagny, K.M., & Zaph, C. Myeloid cell-specific expression of Ship1 regulates IL-12 production and immunity to helminth infection. 2012. Mucosal Immunol. 5:535–43. 177.  Cliffe, L.J., & Grencis, R.K. The Trichuris muris system: a paradigm of resistance and susceptibility to intestinal nematode infection. 2004. Adv. Parasitol. 57:255–307. 178.  Cope, A., Le Friec, G., Cardone, J., & Kemper, C. The Th1 life cycle: molecular control of IFN-γ to IL-10 switching. 2011. Trends Immunol. 32:278–86. 179.  Freitas do Rosário, A.P., Lamb, T., Spence, P., Stephens, R., Lang, A., Roers, A., Muller, W., O’Garra, A., & Langhorne, J. IL-27 promotes IL-10 production by effector Th1 CD4+ T cells: a critical mechanism for protection from severe immunopathology during malaria infection. 2012. J. Immunol. 188:1178–90. 180.  Owens, B.M.J., Beattie, L., Moore, J.W.J., Brown, N., Mann, J.L., Dalton, J.E., Maroof, A., & Kaye, P.M. IL-10-producing Th1 cells and disease progression are regulated by distinct CD11c+ cell populations during visceral leishmaniasis. 2012. PLoS Pathog. 8:e1002827. 181.  Reinhardt, R.L., Liang, H.-E., & Locksley, R.M. Cytokine-secreting follicular T cells shape the antibody repertoire. 2009. Nat. Immunol. 10:385–93. 182.  Boehm, U., Klamp, T., Groot, M., & Howard, J.C. Cellular responses to interferon-gamma. 1997. Annu. Rev. Immunol. 15:749–95. 183.  Bancroft, A.J., Else, K.J., & Grencis, R.K. Low-level infection with Trichuris muris significantly affects the polarization of the CD4 response. 1994. Eur. J. Immunol. 24:3113–8. 184.  Foth, B.J., Tsai, I.J., Reid, A.J., Bancroft, A.J., Nichol, S., Tracey, A., Holroyd, N., Cotton, J.A., Stanley, E.J., Zarowiecki, M., Liu, J.Z., Huckvale, T., Cooper, P.J., Grencis, R.K., & Berriman, M. Whipworm genome and dual-species transcriptome analyses provide molecular insights into an intimate host-parasite interaction. 2014. Nat. Genet. 46:693–700. 185.  Wild, J.S., Sigounas,  a, Sur, N., Siddiqui, M.S., Alam, R., Kurimoto, M., & Sur, S. IFN-gamma-inducing factor (IL-18) increases allergic sensitization, serum IgE, Th2 cytokines, and airway eosinophilia in a mouse model of allergic asthma. 2000. J. Immunol. 164:2701–10. 186.  Wong, C.K., Ho, C.Y., Ko, F.W.S., Chan, C.H.S., Ho,  a. S.S., Hui, D.S.C., & Lam, 121  C.W.K. Proinflammatory cytokines (IL-17, IL-6, IL-18 and IL-12) and Th cytokines (IFN-γ, IL-4, IL-10 and IL-13) in patients with allergic asthma. 2001. Clin. Exp. Immunol. 187.  Mitchell, C., Provost, K., Niu, N., Homer, R., & Cohn, L. IFN-γ acts on the airway epithelium to inhibit local and systemic pathology in allergic airway disease. 2011. J. Immunol. 187:3815–20. 188.  Ford, J.G., Rennick, D., Donaldson, D.D., Venkayya, R., McArthur, C., Hansell, E., Kurup, V.P., Warnock, M., & Grünig, G. Il-13 and IFN-gamma: interactions in lung inflammation. 2001. J. Immunol. 167:1769–77. 189.  Contreras-Ruiz, L., Ghosh-Mitra, A., Shatos, M.A., Dartt, D.A., & Masli, S. Modulation of conjunctival goblet cell function by inflammatory cytokines. 2013. Mediators Inflamm. 2013:636812. 190.  Böhm, L., Maxeiner, J., Meyer-Martin, H., Reuter, S., Finotto, S., Klein, M., Schild, H., Schmitt, E., Bopp, T., & Taube, C. IL-10 and regulatory T cells cooperate in allergen-specific immunotherapy to ameliorate allergic asthma. 2014. J. Immunol. 194:887–97. 191.  Takanaski, S. Interleukin 10 inhibits lipopolysaccharide-induced survival and cytokine production by human peripheral blood eosinophils. 1994. J. Exp. Med. 180:711–5. 192.  Chung, F. Anti-inflammatory cytokines in asthma and allergy: interleukin-10, interleukin-12, interferon-gamma. 2001. Mediators Inflamm. 10:51–9. 193.  Ogawa, Y., Duru, E., & Ameredes, B. Role of IL-10 in the resolution of airway inflammation. 2008. Curr. Mol. Med. 8:437–45. 194.  Mohrs, K., Harris, D.P., Lund, F.E., & Mohrs, M. Systemic dissemination and persistence of Th2 and type 2 cells in response to infection with a strictly enteric nematode parasite. 2005. J. Immunol. 175:5306–13. 195.  Ito, T., Yang, M., Wang, Y.-H., Lande, R., Gregorio, J., Perng, O.A., Qin, X.-F., Liu, Y.-J., & Gilliet, M. Plasmacytoid dendritic cells prime IL-10-producing T regulatory cells by inducible costimulator ligand. 2007. J. Exp. Med. 204:105–15. 196.  de Bruin, A.M., Voermans, C., & Nolte, M.A. Impact of interferon-γ on hematopoiesis. 2014. Blood. 124:2479–86. 197.  Grainger, J.R., Smith, K.A., Hewitson, J.P., Mcsorley, H.J., Harcus, Y., Filbey, K.J., Finney, C.A.M., Greenwood, E.J.D., Knox, D.P., Wilson, M.S., Belkaid, Y., Rudensky, A.Y., & Maizels, R.M. Helminth secretions induce de novo T cell Foxp3 expression and regulatory function through the TGF-beta pathway. 2010. J. Exp. Med. 207:2331–41. 198.  McSorley, H.J., Blair, N.F., Smith, K.A., McKenzie, A.N., & Maizels, R.M. Blockade of IL-33 release and suppression of type 2 innate lymphoid cell responses by helminth secreted products in airway allergy. 2014. Mucosal Immunol. 7:1068–78. 199.  Houlden, A., Hayes, K.S., Bancroft, A.J., Worthington, J.J., Wang, P., Grencis, R.K., & Roberts, I.S. Chronic Trichuris muris infection in C57BL/6 mice causes significant 122  changes in host microbiota and metabolome: effects reversed by pathogen clearance. 2015. PLoS One. 10:e0125945. 200.  Zaiss, M.M., Rapin, A., Lebon, L., Dubey, L.K., Mosconi, I., Sarter, K., Piersigilli, A., Menin, L., Walker, A.W., Rougemont, J., Paerewijck, O., Geldhof, P., McCoy, K.D., Macpherson, A.J., Croese, J., Giacomin, P.R., Loukas, A., Junt, T., Marsland, B.J., & Harris, N.L. The intestinal microbiota contributes to the ability of helminths to modulate allergic inflammation. 2015. Immunity. In press: 201.  Bourke, C.D., Mutapi, F., Nausch, N., Photiou, D.M.F., Poulsen, L.K., Kristensen, B., Arnved, J., Rønborg, S., Roepstorff, A., Thamsborg, S., Kapel, C., Melbye, M., & Bager, P. Trichuris suis ova therapy for allergic rhinitis does not affect allergen-specific cytokine responses despite a parasite-specific cytokine response. 2012. Clin. Exp. Allergy. 42:1582–95. 202.  Zaretsky, A.G., Engiles, J.B., & Hunter, C.A. Infection-induced changes in hematopoiesis. 2014. J. Immunol. 192:27–33. 203.  Furze, R.C., & Rankin, S.M. Neutrophil mobilization and clearance in the bone marrow. 2008. Immunology. 125:281–8. 204.  Schuettpelz, L.G., & Link, D.C. Regulation of hematopoietic stem cell activity by inflammation. 2013. Front. Immunol. 4:204. 205.  de Bruin, A.M., Demirel, Ö., Hooibrink, B., Brandts, C.H., & Nolte, M.A. Interferon-γ impairs proliferation of hematopoietic stem cells in mice. 2013. Blood. 121:3578–85. 206.  Maltby, S., Hansbro, N.G., Tay, H.L., Stewart, J., Plank, M., Donges, B., Rosenberg, H.F., & Foster, P.S. Production and differentiation of myeloid cells driven by proinflammatory cytokines in response to acute pneumovirus infection in mice. 2014. J. Immunol. 193:4072–82. 207.  Griseri, T., McKenzie, B.S., Schiering, C., & Powrie, F. Dysregulated hematopoietic stem and progenitor cell activity promotes interleukin-23-driven chronic intestinal inflammation. 2012. Immunity. 37:1116–29. 208.  Zhao, X., Ren, G., Liang, L., Ai, P.Z., Zheng, B., Tischfield, J.A., Shi, Y., & Shao, C. Brief report: interferon-gamma induces expansion of Lin(-)Sca-1(+)C-Kit(+) Cells. 2010. Stem Cells. 28:122–6. 209.  Oguro, H., Ding, L., & Morrison, S.J. SLAM family markers resolve functionally distinct subpopulations of hematopoietic stem cells and multipotent progenitors. 2013. Cell Stem Cell. 13:102–16. 210.  Murray, P.J., Young, R.A., & Daley, G.Q. Hematopoietic remodeling in interferon-gamma-deficient mice infected with mycobacteria. 1998. Blood. 91:2914–24. 211.  de Bruin, A.M., Libregts, S.F., Valkhof, M., Boon, L., Touw, I.P., & Nolte, M.A. IFNγ induces monopoiesis and inhibits neutrophil development during inflammation. 2012. Blood. 119:1543–54. 212.  Chenery, A.L., Antignano, F., Burrows, K., Scheer, S., Perona-Wright, G., & Zaph, 123  C. Low dose intestinal Trichuris muris infection alters the lung immune microenvironment and can suppress allergic airway inflammation. 2015. Infect. Immun. 213.  Askenasy, N. Interferon and tumor necrosis factor as humoral mechanisms coupling hematopoietic activity to inflammation and injury. 2015. Blood Rev. 29:11–5. 214.  Caux, C., Moreau, I., Saeland, S., & Banchereau, J. Interferon-gamma enhances factor-dependent myeloid proliferation of human CD34+ hematopoietic progenitor cells. 1992. Blood. 79:2628–35. 215.  Dénes, A., Humphreys, N., Lane, T.E., Grencis, R., & Rothwell, N. Chronic systemic infection exacerbates ischemic brain damage via a CCL5 (regulated on activation, normal T-cell expressed and secreted)-mediated proinflammatory response in mice. 2010. J. Neurosci. 30:10086–95. 216.  Schmitz-Winnenthal, F.H., Volk, C., Z’graggen, K., Galindo, L., Nummer, D., Ziouta, Y., Bucur, M., Weitz, J., Schirrmacher, V., Büchler, M.W., & Beckhove, P. High frequencies of functional tumor-reactive T cells in bone marrow and blood of pancreatic cancer patients. 2005. Cancer Res. 65:10079–87. 217.  Di Rosa, F., & Pabst, R. The bone marrow: a nest for migratory memory T cells. 2005. Trends Immunol. 26:360–6. 218.  Cowley, S.C., Hamilton, E., Frelinger, J.A., Su, J., Forman, J., & Elkins, K.L. CD4-CD8- T cells control intracellular bacterial infections both in vitro and in vivo. 2005. J. Exp. Med. 202:309–19. 219.  Neyt, K., GeurtsvanKessel, C.H., & Lambrecht, B.N. Double-negative T resident memory cells of the lung react to influenza virus infection via CD11chi dendritic cells. 2015. Mucosal Immunol. 220.  Mou, Z., Liu, D., Okwor, I., Jia, P., Orihara, K., & Uzonna, J.E. MHC class II restricted innate-like double negative T cells contribute to optimal primary and secondary immunity to Leishmania major. 2014. PLoS Pathog. 10:e1004396. 221.  Kim, C.H. Roles of retinoic acid in induction of immunity and immune tolerance. 2008. Endocr. Metab. Immune Disord. Drug Targets. 8:289–94. 222.  Elias, K.M., Laurence, A., Davidson, T.S., Stephens, G., Kanno, Y., Shevach, E.M., & O’Shea, J.J. Retinoic acid inhibits Th17 polarization and enhances FoxP3 expression through a Stat-3/Stat-5 independent signaling pathway. 2008. Blood. 111:1013–20. 223.  MacLean, G., Li, H., Metzger, D., Chambon, P., & Petkovich, M. Apoptotic extinction of germ cells in testes of Cyp26b1 knockout mice. 2007. Endocrinology. 148:4560–7. 224.  Antignano, F., Burrows, K., Hughes, M.R., Han, J.M., Kron, K.J., Penrod, N.M., Oudhoff, M.J., Wang, S.K.H., Min, P.H., Gold, M.J., Chenery, A.L., Braam, M.J.S., Fung, T.C., Rossi, F.M. V, McNagny, K.M., Arrowsmith, C.H., Lupien, M., Levings, M.K., & Zaph, C. Methyltransferase G9A regulates T cell differentiation during murine intestinal inflammation. 2014. J. Clin. Invest. 124:1945–55. 225.  Yashiro, K., Zhao, X., Uehara, M., Yamashita, K., Nishijima, M., Nishino, J., Saijoh, 124  Y., Sakai, Y., & Hamada, H. Regulation of retinoic acid distribution is required for proximodistal patterning and outgrowth of the developing mouse limb. 2004. Dev. Cell. 6:411–22. 226.  Kiss, I., Rühl, R., Szegezdi, É., Fritzsche, B., Tóth, B., Pongrácz, J., Perlmann, T., Fésüs, L., & Szondy, Z. Retinoid receptor-activating ligands are produced within the mouse thymus during postnatal development. 2008. Eur. J. Immunol. 38:147–55. 227.  Mucida, D., Park, Y., Kim, G., Turovskaya, O., Scott, I., Kronenberg, M., & Cheroutre, H. Reciprocal TH17 and regulatory T cell differentiation mediated by retinoic acid. 2007. Science. 317:256–60. 228.  Wang, C., Thangamani, S., Kim, M., Gu, B.-H., Lee, J.H., Taparowsky, E.J., & Kim, C.H. BATF is required for normal expression of gut-homing receptors by T helper cells in response to retinoic acid. 2013. J. Exp. Med. 210:475–89. 229.  Dranse, H.J., Sampaio, A. V, Petkovich, M., & Underhill, T.M. Genetic deletion of Cyp26b1 negatively impacts limb skeletogenesis by inhibiting chondrogenesis. 2011. J. Cell Sci. 124:2723–34. 230.  Collins, S.J. The role of retinoids and retinoic acid receptors in normal hematopoiesis. 2002. Leukemia. 16:1896–905. 231.  Lammer, E.J., Chen, D.T., Hoar, R.M., Agnish, N.D., Benke, P.J., Braun, J.T., Curry, C.J., Fernhoff, P.M., Grix, A.W., Lott, I.T., Richard, J.M., & Sun, S.C. Retinoic acid embryopathy. 1985. N. Engl. J. Med. 313:837–41. 232.  Xiao, S., Jin, H., Korn, T., Liu, S.M., Oukka, M., Lim, B., & Kuchroo, V.K. Retinoic acid increases Foxp3+ regulatory T cells and inhibits development of Th17 cells by enhancing TGF-beta-driven Smad3 signaling and inhibiting IL-6 and IL-23 receptor expression. 2008. J. Immunol. 181:2277–84. 233.  Lu, L., Ma, J., Li, Z., Lan, Q., Chen, M., Liu, Y., Xia, Z., Wang, J., Han, Y., Shi, W., Quesniaux, V., Ryffel, B., Brand, D., Li, B., Liu, Z., & Zheng, S.G. All-trans retinoic acid promotes TGF-β-induced Tregs via histone modification but not DNA demethylation on Foxp3 gene locus. 2011. PLoS One. 6:e24590. 234.  Niederreither, K., Fraulob, V., Garnier, J.-M., Chambon, P., & Dollé, P. Differential expression of retinoic acid-synthesizing (RALDH) enzymes during fetal development and organ differentiation in the mouse. 2002. Mech. Dev. 110:165–71. 235.  Smits, H.H., Everts, B., Hartgers, F.C., & Yazdanbakhsh, M. Chronic helminth infections protect against allergic diseases by active regulatory processes. 2010. Curr. Allergy Asthma Rep. 10:3–12. 236.  Bourke, C.D., Mutapi, F., Nausch, N., Photiou, D.M.F., Poulsen, L.K., Kristensen, B., Arnved, J., Rønborg, S., Roepstorff, A., Thamsborg, S., Kapel, C., Melbye, M., & Bager, P. Trichuris suis ova therapy for allergic rhinitis does not affect allergen-specific cytokine responses despite a parasite-specific cytokine response. 2012. Clin. Exp. Allergy. 42:1582–95. 125  237.  Helmby, H. Human helminth therapy to treat inflammatory disorders- where do we stand? 2015. BMC Immunol. 16:12. 238.  Baldridge, M.T., King, K.Y., Boles, N.C., Weksberg, D.C., & Goodell, M.A. Quiescent haematopoietic stem cells are activated by IFN-gamma in response to chronic infection. 2010. Nature. 465:793–7. 239.  Brown, C.C., & Noelle, R.J. Seeing through the dark: New insights into the immune regulatory functions of vitamin A. 2015. Eur. J. Immunol. 45:1287–95. 240.  Fransén, K., Franzén, P., Magnuson, A., Elmabsout, A.A., Nyhlin, N., Wickbom, A., Curman, B., Törkvist, L., D’Amato, M., Bohr, J., Tysk, C., Sirsjö, A., & Halfvarson, J. Polymorphism in the Retinoic Acid Metabolizing Enzyme CYP26B1 and the Development of Crohn’s Disease. 2013. PLoS One. 8:e72739. 241.  Kuperman, D.A., Huang, X., Koth, L.L., Chang, G.H., Dolganov, G.M., Zhu, Z., Elias, J.A., Sheppard, D., & Erle, D.J. Direct effects of interleukin-13 on epithelial cells cause airway hyperreactivity and mucus overproduction in asthma. 2002. Nat. Med. 8:885–9. 242.  Sahu, N., Morales, J.L., Fowell, D., & August, A. Modeling susceptibility versus resistance in allergic airway disease reveals regulation by Tec kinase Itk. 2010. PLoS One. 5:e11348. 243.  Kelada, S.N.P., Wilson, M.S., Tavarez, U., Kubalanza, K., Borate, B., Whitehead, G.S., Maruoka, S., Roy, M.G., Olive, M., Carpenter, D.E., Brass, D.M., Wynn, T.A., Cook, D.N., Evans, C.M., Schwartz, D.A., & Collins, F.S. Strain-dependent genomic factors affect allergen-induced airway hyperresponsiveness in mice. 2011. Am. J. Respir. Cell Mol. Biol. 45:817–24. 244.  Lederlin, M., Ozier, A., Dournes, G., Ousova, O., Girodet, P.-O., Begueret, H., Marthan, R., Montaudon, M., Laurent, F., & Berger, P. In vivo micro-CT assessment of airway remodeling in a flexible OVA-sensitized murine model of asthma. 2012. PLoS One. 7:e48493. 245.  Schroder, K., Hertzog, P.J., Ravasi, T., & Hume, D.A. Interferon-gamma: an overview of signals, mechanisms and functions. 2004. J. Leukoc. Biol. 75:163–89. 246.  Saraiva, M., & O’Garra, A. The regulation of IL-10 production by immune cells. 2010. Nat. Rev. Immunol. 10:170–81. 247.  Mic, F.A., Haselbeck, R.J., Cuenca, A.E., & Duester, G. Novel retinoic acid generating activities in the neural tube and heart identified by conditional rescue of Raldh2 null mutant mice. 2002. Development. 129:2271–82. 248.  Dawson, H.D., Collins, G., Pyle, R., Key, M., Weeraratna, A., Deep-Dixit, V., Nadal, C.N., & Taub, D.D. Direct and indirect effects of retinoic acid on human Th2 cytokine and chemokine expression by human T lymphocytes. 2006. BMC Immunol. 7:27. 249.  Balmer, J.E. Gene expression regulation by retinoic acid. 2002. J. Lipid Res. 43:1773–808. 126  250.  Moro, K., Kabata, H., Tanabe, M., Koga, S., Takeno, N., Mochizuki, M., Fukunaga, K., Asano, K., Betsuyaku, T., & Koyasu, S. Interferon and IL-27 antagonize the function of group 2 innate lymphoid cells and type 2 innate immune responses. 2016. Nat. Immunol. 17:76–86. 251.  Chun, J., & Hartung, H.-P. Mechanism of action of oral fingolimod (FTY720) in multiple sclerosis. Clin. Neuropharmacol. 33:91–101. 252.  Purton, L.E., & Scadden, D.T. Limiting factors in murine hematopoietic stem cell assays. 2007. Cell Stem Cell. 1:263–70. 253.  Greenbaum, A., Hsu, Y.-M.S., Day, R.B., Schuettpelz, L.G., Christopher, M.J., Borgerding, J.N., Nagasawa, T., & Link, D.C. CXCL12 in early mesenchymal progenitors is required for haematopoietic stem-cell maintenance. 2013. Nature. 495:227–30. 254.  Frei, R., Lauener, R.P., Crameri, R., & O’Mahony, L. Microbiota and dietary interactions: an update to the hygiene hypothesis? 2012. Allergy. 67:451–61. 255.  Hayes, K.S., Bancroft,  a J., Goldrick, M., Portsmouth, C., Roberts, I.S., & Grencis, R.K. Exploitation of the intestinal microflora by the parasitic nematode Trichuris muris. 2010. Science. 328:1391–4. 256.  Holm, J.B., Sorobetea, D., Kiilerich, P., Ramayo-Caldas, Y., Estellé, J., Ma, T., Madsen, L., Kristiansen, K., & Svensson-Frej, M. Chronic Trichuris muris Infection Decreases Diversity of the Intestinal Microbiota and Concomitantly Increases the Abundance of Lactobacilli. 2015. PLoS One. 10:e0125495. 257.  Trompette, A., Gollwitzer, E.S., Yadava, K., Sichelstiel, A.K., Sprenger, N., Ngom-Bru, C., Blanchard, C., Junt, T., Nicod, L.P., Harris, N.L., & Marsland, B.J. Gut microbiota metabolism of dietary fiber influences allergic airway disease and hematopoiesis. 2014. Nat. Med. 20:159–66. 258.  Hurst, R.J.M., & Else, K.J. The retinoic acid-producing capacity of gut dendritic cells and macrophages is reduced during persistent T. muris infection. 2013. Parasite Immunol. 35:229–33. 259.  Hurst, R.J.M., De Caul, A., Little, M.C., Kagechika, H., & Else, K.J. The retinoic acid receptor agonist Am80 increases mucosal inflammation in an IL-6 dependent manner during Trichuris muris infection. 2013. J. Clin. Immunol. 33:1386–94. 260.  Ruane, D., Brane, L., Reis, B.S., Cheong, C., Poles, J., Do, Y., Zhu, H., Velinzon, K., Choi, J.-H., Studt, N., Mayer, L., Lavelle, E.C., Steinman, R.M., Mucida, D., & Mehandru, S. Lung dendritic cells induce migration of protective T cells to the gastrointestinal tract. 2013. J. Exp. Med. 210:1871–88. 261.  Ueki, S., Nishikawa, J., Yamauchi, Y., Konno, Y., Tamaki, M., Itoga, M., Kobayashi, Y., Takeda, M., Moritoki, Y., Ito, W., & Chihara, J. Retinoic acids up-regulate functional eosinophil-driving receptor CCR3. 2013. Allergy. 68:953–6. 262.  Ueki, S., Mahemuti, G., Ito, W., Kobayashi, Y., Chiba, T., Oyamada, H., Kamada, 127  Y., Yamaguchi, K., Takeda, M., Kayaba, H., & Chihara, J. Retinoic Acids Induce Eosinophil Survival and MCP-1 Production. 2007. J. Allergy Clin. Immunol. 119:S220.    128  Appendices Appendix A  chapter 2 supplemental material A.1 Supplemental figures   Figure A.1 Neutralization of IL-12 does not reverse T. muris infection-mediated protection from papain-induced allergic airway inflammation. (A) Ex vivo intracellular staining of IFN-γ from lung CD4+ cells following papain exposure. (B) Quantification of total lung eosinophils following papain exposure. Data are means ± SEM, representative of 2 independent experiments, n= 3 mice per experiment (A-B). (Ctrl: papain/IgG1 treated; Tm: T. muris-infected papain/IgG1 treated; Tm/αIL-12: T. muris-infected/anti-IL-12 treated/papain treated).     129   Figure A.2 T. muris infection does not affect lung Treg cells. CD4+ -gated frequencies and absolute numbers of lung Foxp3+CD25+ Treg cells after T. muris infection and papain exposure. Data are means ± SEM, representative of 2 independent experiments, n= 3 mice per experiment. (N: naive, no papain; Ctrl: papain treated; Tm: T. muris-infected papain treated).  

Cite

Citation Scheme:

        

Citations by CSL (citeproc-js)

Usage Statistics

Share

Embed

Customize your widget with the following options, then copy and paste the code below into the HTML of your page to embed this item in your website.
                        
                            <div id="ubcOpenCollectionsWidgetDisplay">
                            <script id="ubcOpenCollectionsWidget"
                            src="{[{embed.src}]}"
                            data-item="{[{embed.item}]}"
                            data-collection="{[{embed.collection}]}"
                            data-metadata="{[{embed.showMetadata}]}"
                            data-width="{[{embed.width}]}"
                            async >
                            </script>
                            </div>
                        
                    
IIIF logo Our image viewer uses the IIIF 2.0 standard. To load this item in other compatible viewers, use this url:
http://iiif.library.ubc.ca/presentation/dsp.24.1-0300054/manifest

Comment

Related Items