INFLAMMATORY TH17 RESPONSES TO INFECTION WITH MYCOBACTERIUM AVIUM SUBSPECIES PARATUBERCULOSIS (MAP) IN CATTLE AND THEIR POTENTIAL ROLE IN THE DEVELOPMENT OF JOHNE’S DISEASE By Justin Lee DeKuiper A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of Animal Science – Doctor of Philosophy 2019 ABSTRACT INFLAMMATORY TH17 RESPONSES TO INFECTION WITH MYCOBACTERIUM AVIUM SUBSPECIES PARATUBERCULOSIS (MAP) IN CATTLE AND THEIR POTENTIAL ROLE IN THE DEVELOPMENT OF JOHNE’S DISEASE By Justin Lee DeKuiper Mycobacterium avium subspecies paratuberculosis (MAP) causes a chronic inflammatory gastrointestinal disease of ruminants known as Johne’s disease (JD). MAP primarily colonizes the ileum of ruminants, leading to reduced nutrient absorption, chronic diarrhea, and eventually death. The percentage of dairy operations infected MAP in the US may have risen to as much as 91% from earlier reports of 68%. In correspondence, an estimated dairy industry loss has increased from $200 million to $1.5 billion was due to JD. Control of JD is difficult largely due to insensitive diagnostic tools, a long subclinical stage of infection, and lack of effective vaccines. Correlates of protection are lacking in model systems of JD and the sources of inflammation due to JD are not well characterized. Inside macrophages, MAP survives and replicates. Monocyte-derived macrophages (MDMs), peripheral blood mononuclear cells (PBMCs), and various T-cells interact with MAP. Commonly studied immune responses, such as the Th1/Th2 paradigm, do not adequately explain host responses to MAP. The major remaining knowledge gaps in MAP immunopathogenesis include key inflammatory responders in the ileum, and how naïve T-cell responsive choice (Th1, Th2, Th17) is influenced. A potential role for non-classical immune responses to MAP, such as that mediated by Th17 cells, has been suggested. Indeed, MAP antigens induce mRNAs encoding the cytokines IL-23 and IL-17A in bovine PBMCs. IL-23 and IL-17A production are both associated with Th17-like immune responses. Th17 cells are also defined by surface expression of the IL-23 receptor (IL-23R). The mean relative percent (MRP) of T cell subtypes expressing IL-23R was determined by flow cytometry and indicated an increase in mean relative percent (MRP) of T cells (CD4+, CD8+, and TCR1+) in peripheral blood mononuclear cells (PBMCs) of JD+ cows with high and low expression of IL-23R (IL-23RHigh and IL-23RLow) when compared to JD- cows. Although MAP stimulates PBMCs to secrete IL-17A in-vitro, there were no differences in IL-17A levels between subclinical JD+ and JD- cows when analyzed by ELISA. Plasma with low JD+ score values had significantly more IL-17A when compared to plasma with high JD+ score values, establishing a moderate correlation between JD+ score and IL-17A. However, overall plasma from JD+ cows had significantly less IL-17A than plasma from JD- cows. Unlike IL-17A, IL-23 was greater in plasma from JD+ cows than in JD- cows. Evaluating the relationship that MAP is having on APCs and CD3+ in relation to Th17 cytokines (IL-23, IL-17A, IL-17F, IL-22, IL-27) as well as Th1 cytokine IFNγ, CD3+ were stimulated with MAP in the presence of APCs (MDMs or B cells) or alone and evaluated by RT-qPCR. MAP stimulation significantly increased production of mRNA encoding Th17 cytokines and IFNγ in CD3+ T cells regardless of APCs. However, the presence of MDMs significantly increase the quantity of mRNA. Lastly, we observed that αβ T cells (mostly CD4+) are responsible for production of Th17 cytokines as an early response to MAP in the absence of APCs. Our data suggests that Th17-like cells may indeed play a role in early immune responses to MAP infection and development or control of JD. Understanding the influences and potential novel mechanisms of an inflammatory pathway during MAP infection could be exploited for treatment, prevention, or diagnosis of JD. TABLE OF CONTENTS LIST OF TABLES ............................................................................................................ vii LIST OF FIGURES ........................................................................................................ viii KEY TO SYMBOLS AND ABBREVIATIONS .................................................................. xi INTRODUCTION ............................................................................................................. 1 INFLAMMATORY TH17 RESPONSES TO INFECTION WITH MYCOBACTERIUM AVIUM SUBSPECIES PARATUBERCULOSIS (MAP) IN CATTLE AND THEIR POTENTIAL ROLE IN DEVELOPMENT OF JOHNE’S DISEASE .............................................................................................................. 1 Introduction ........................................................................................................... 1 The Th17 Response ............................................................................................. 3 Th17 Cytokines ..................................................................................................... 6 MAP Driving Th17 ................................................................................................ 7 Final Thoughts ...................................................................................................... 9 Acknowledgements .............................................................................................. 9 APPENDIX .................................................................................................................... 11 REFERENCES .............................................................................................................. 13 CHAPTER 1 .................................................................................................................. 21 MYCOBACTERIUM AVIUM SP. PARATUBERCULOSIS (MAP) INDUCES IL- 17A PRODUCTION IN BOVINE PERIPHERAL BLOOD MONONUCLEAR CELLS (PBMCS) AND ENHANCES IL-23R EXPRESSION IN-VIVO AND IN- VITRO ................................................................................................................ 21 Abstract .............................................................................................................. 21 Introduction ......................................................................................................... 22 Materials and Methods ....................................................................................... 25 Study Animals .......................................................................................... 25 Preparation of PBMCs, MDMs, and Treatments ...................................... 26 Cell Surface Staining and Flow Cytometry ............................................... 27 MAP Culture ............................................................................................ 28 ELISA and Statistics ................................................................................ 29 Johne’s Disease Diagnosis ...................................................................... 30 Results ............................................................................................................... 31 Effect of MAP-infected MDMs on IL-17A secretion in PBMC cultures from JD- cows .................................................................................................. 31 Secretion of IL-17A by PBMCs from JD+ and JD- cows stimulated with M. paratuberculosis (MAP) ........................................................................... 32 IL-23 receptor (IL23R) expression on T cells from Johne’s-test positive (JD+) and test negative cows (JD-) .......................................................... 33 iv Johne’s-test positive cows (JD+) have less circulating IL-17A than Johne’s-test negative cows (JD-) ............................................................. 34 Discussion .......................................................................................................... 36 Acknowledgments .............................................................................................. 41 APPENDICES ............................................................................................................... 42 APPENDIX A: Table 2 ........................................................................................ 43 APPENDIX B: Table 3 ........................................................................................ 44 APPENDIX C: Table 4 ........................................................................................ 45 APPENDIX D: Figure 1 ....................................................................................... 46 APPENDIX E: Figure 2 ....................................................................................... 47 APPENDIX F: Figure 3A ..................................................................................... 48 APPENDIX G: Figure 3B .................................................................................... 49 APPENDIX H: Figure 4A .................................................................................... 50 APPENDIX I: Figure 4B ...................................................................................... 51 APPENDIX J: Figure 4C ..................................................................................... 52 APPENDIX K: Figure 4D .................................................................................... 53 APPENDIX L: Figure 5 ....................................................................................... 54 APPENDIX M: Figure 6 ...................................................................................... 55 APPENDIX N: Figure 7 ....................................................................................... 56 REFERENCES .............................................................................................................. 57 CHAPTER 2 .................................................................................................................. 63 MYCOBACTERIUM AVIUM SUBSPECIES PARATUBERCULOSIS (MAP) DRIVES AN INNATE TH17-LIKE T CELL RESPONSE REGARDLESS OF THE PRESENCE OF ANTIGEN-PRESENTING CELLS ............................................ 63 Abstract .............................................................................................................. 63 Introduction ......................................................................................................... 64 Materials and methods ....................................................................................... 67 Study animals .......................................................................................... 67 Plasma samples ...................................................................................... 67 Preparations of T cells, B cells, and MDMs ............................................. 68 MAP culture and treatment ...................................................................... 69 Cell surface staining and flow cytometry for culture validation ................. 70 RNA Extraction and RT-qPCR ................................................................. 71 ELISA and statistics ................................................................................. 71 Results ............................................................................................................... 72 CD3+ T cells co-cultured with MDMs ....................................................... 72 CD3+ T cells co-cultured with sIgM+ B cells ............................................ 73 CD3+ T cell culture .................................................................................. 74 αβ T cells are the main producers of IL-17A and IL-22 in the absence of APCs........................................................................................................ 75 CD3+ T cells also express IFNγ mRNA, but not at levels observed for IL- 17A or IL-17F mRNAs.............................................................................. 76 Increased IL-23 levels in JD ELISA test positive cows (JD+) ................... 76 Discussion .......................................................................................................... 77 Acknowledgments .............................................................................................. 82 v APPENDICES ............................................................................................................... 83 APPENDIX A: Table 5 ........................................................................................ 84 APPENDIX B: Table 6 ........................................................................................ 85 APPENDIX C: IL-23 ............................................................................................ 86 APPENDIX D: IL-17a .......................................................................................... 87 APPENDIX E: IL-22 ............................................................................................ 88 APPENDIX F: IL-23 ............................................................................................ 89 APPENDIX G: IL-17a ......................................................................................... 90 APPENDIX H: IL-22 ............................................................................................ 91 APPENDIX I: IL-17a ........................................................................................... 92 APPENDIX J: IL-22 ............................................................................................ 93 APPENDIX K: IL-23 ............................................................................................ 94 APPENDIX L: IL-17a .......................................................................................... 95 APPENDIX M: IL-22 ........................................................................................... 96 APPENDIX N: IL-23 ............................................................................................ 97 APPENDIX O: CD4+ .......................................................................................... 98 APPENDIX P: CD8+ ........................................................................................... 99 APPENDIX Q: TCR1+ (GD) ............................................................................. 100 APPENDIX R: CD3+ T cells ............................................................................. 101 APPENDIX S: Simple linear regression of Correlation ..................................... 102 APPENDIX T: Plasma IL-23 by MAP Infection Status ...................................... 103 APPENDIX U: MDM / Bound CD3+ IL-17A ...................................................... 104 APPENDIX V: Separated CD3+ IL-17A ............................................................ 105 APPENDIX W: MDM / Bound CD3+ IL-23 ........................................................ 106 APPENDIX X: Separated CD3+ IL-23 .............................................................. 107 APPENDIX Y: MDM / Bound CD3+ IL-22 ......................................................... 108 APPENDIX Z: Separated CD3+ IL-22 .............................................................. 109 REFERENCES ............................................................................................................ 110 CHAPTER 3 ................................................................................................................ 118 FUTURE DIRECTIONS OF JOHNE’S DISEASE RESEARCH AND THE CROHN’S DISEASE CONNECTION ................................................................ 118 Introduction ....................................................................................................... 118 Johne’s disease vs. Crohn’s disease ................................................................ 120 Future directions ............................................................................................... 123 Expected results ............................................................................................... 126 Potential problems and diagnostic opportunities .............................................. 127 APPENDICES ............................................................................................................. 129 APPENDIX A: Table 7 ...................................................................................... 130 APPENDIX B: Table 8 ...................................................................................... 131 APPENDIX C: Table 9 ...................................................................................... 132 REFERENCES ............................................................................................................ 134 vi LIST OF TABLES Table 1: MAP-induced cytokine expression during subclinical and clinical stages of disease. ......................................................................................................................... 12 Table 2: Primary antibodies. Immuno-staining of T-cell specific surface markers and Th17 signature receptors and cytokines. ....................................................................... 43 Table 3: Secondary antibodies. Immuno-staining of T-cell specific surface markers and Th17 signature receptors and cytokines. ....................................................................... 44 Table 4: 2-Way ANOVA using Tukey's Multiple Comparison Test to compare all the means to each other. Reporting p-values. ..................................................................... 45 Table 5: Primary and Secondary antibody list. .............................................................. 84 Table 6: Taqman primer list. .......................................................................................... 85 Table 7: Average Sensitivities and Specificities between MAP diagnostic tests. Adopted from Collins, 2006. ...................................................................................................... 130 Table 8: Homing signals to the gut indicating Th17 homing to the Ileum. ................... 131 Table 9: The argument, MAP and Crohn's Disease. Adopted from Rosenfeld, 2010. ........................................................................................................................... 132 vii LIST OF FIGURES Figure 1: Experimental set-up for analysis effects of MDMs on IL-17A production by PBMCs during MAP stimulation. ................................................................................... 46 Figure 2: Effect of MAP-infected MDMs on IL-17A secretion in PBMC cultures from JD- cows. ............................................................................................................................. 47 Figure 3A: Experimental set-up for analysis of PBMCs from JD+ and JD- cows stimulated with M. paratuberculosis (MAP). .................................................................. 48 Figure 3B: Secretion of IL-17A by PBMCs from JD+ and JD- cows stimulated with M. paratuberculosis (MAP). ................................................................................................ 49 Figure 4A: Flow cytometry gating strategy example. ..................................................... 50 Figure 4B: Untreated JD+ vs. JD- IL-23RLow. ................................................................ 51 Figure 4C: Untreated JD+ vs. JD- IL-23RHigh. ...............................................................................................52 Figure 4D: Surface expression of IL-23RLow on JD- T cells after stimulation with MAP antigen. ......................................................................................................................... 53 Figure 5: Plasma IL-17A levels of cows based on environmental MAP status of farm and JD status. ...................................................................................................................... 54 Figure 6: Plasma IL-17A levels of cows based on IDEXX Johne’s ELISA score. .......... 55 Figure 7: Correlation of JD+ IDEXX ELISA score and IL-17A plasma ELISA concentration. ................................................................................................................ 56 Figure 8A: Relative abundance of IL-23 mRNA of cultures containing MDMs co-cultured with autologous CD3+ T cells and stimulated with MAP. .............................................. 86 Figure 8B: Relative abundance of IL-17A mRNA of cultures containing MDMs co- cultured with autologous CD3+ T cells and stimulated with MAP. ................................. 87 Figure 8C: Relative abundance of IL-22 mRNA of cultures containing MDMs co-cultured with autologous CD3+ T cells and stimulated with MAP. .............................................. 88 Figure 9A: Relative abundance of IL-23 mRNA of cultures containing sIgM+ B cells co- cultured with autologous CD3+ T cells and stimulated with MAP. ................................. 89 viii Figure 9B: Relative abundance of IL-17A mRNA of cultures containing sIgM+ B cells co- cultured with autologous CD3+ T cells and stimulated with MAP. ................................. 90 Figure 9C: Relative abundance of IL-22 mRNA of cultures containing sIgM+ B cells co- cultured with autologous CD3+ T cells and stimulated with MAP. ................................. 91 Figure 10A: Relative abundance of IL-17A mRNA of cultures containing CD3+ T cells and stimulated with MAP. .............................................................................................. 92 Figure 10B: Relative abundance of IL-22 mRNA of cultures containing CD3+ T cells and stimulated with MAP. ..................................................................................................... 93 Figure 10C: Relative abundance of IL-23 mRNA of cultures containing CD3+ T cells and stimulated with MAP. ..................................................................................................... 94 Figure 11A: Relative abundance of IL-17A mRNA of CD3+ cells, MDM/CD3+, and sIgM+/CD3+ cultures stimulated with MAP. .................................................................. 95 Figure 11B: Relative abundance of IL-22 mRNA of CD3+ cells, MDM/CD3+, and sIgM+/CD3+ cultures stimulated with MAP. .................................................................. 96 Figure 11C: Relative abundance of IL-23 mRNA of CD3+ cells, MDM/CD3+, and sIgM+/CD3+ cultures stimulated with MAP. .................................................................. 97 Figure 12A: Relative abundance of Th17 mRNA encoding IL-17A, IL-22 and IL-23 from CD4+ cell cultures stimulated with MAP or left unstimulated. ....................................... 98 Figure 12B: Relative abundance of Th17 mRNA encoding IL-17A, IL-22 and IL-23 from CD8+ cell cultures stimulated with MAP or left unstimulated. ....................................... 99 Figure 12C: Relative abundance of Th17 mRNA encoding IL-17A, IL-22 and IL-23 from TCR1+ (γδ) cell cultures stimulated with MAP or left unstimulated. ............................ 100 Figure 13A: Relative abundance of mRNA encoding IL-17F (Th17), IL-27 (anti-Th17) and IFNγ from CD3+ cell cultures stimulated with MAP or left unstimulated. .............. 101 Figure 13B: Correlation between quantities of mRNA encoding IL-17F and IL-27 from CD3+ cell cultures stimulated with MAP or left unstimulated. ..................................... 102 Figure 14: Plasma IL-23 levels of cows based on IDEXX Johne’s ELISA status. ....... 103 Figure S1A: Relative abundance of IL-17A mRNA of MDMs and bound CD3+ T cells after co-culture, stimulation with MAP, and unbound cells were rinsed away. ............ 104 Figure S1B: Relative abundance of IL-17A mRNA of unbound CD3+ T cells after co- culture, stimulation with MAP. ..................................................................................... 105 ix Figure S1C: Relative abundance of IL-23 mRNA of MDMs and bound CD3+ T cells after co-culture, stimulation with MAP, and unbound cells were rinsed away. ..................... 106 Figure S1D: Relative abundance of IL-23 mRNA of unbound CD3+ T cells after co- culture, stimulation with MAP. ..................................................................................... 107 Figure S1E: Relative abundance of IL-22 mRNA of MDMs and bound CD3+ T cells after co-culture, stimulation with MAP, and unbound cells were rinsed away. ..................... 108 Figure S1F: Relative abundance of IL-22 mRNA of unbound CD3+ T cells after co- culture, stimulation with MAP. ..................................................................................... 109 x KEY TO SYMBOLS AND ABBREVIATIONS < ......................................................................................................................... Less than ≤ ....................................................................................................... Less than or equal to ° ......................................................................................................................... Degree(s) ANOVA .............................................................................................. Analysis of variance bTb .................................................................................. Mycobacterium bovis (M. bovis) C.............................................................................................................................Celsius CD ............................................................................................................ Crohn’s disease CD4 ............................................................................................................. Helper T cells CD8 ................................................................................................................ Killer T cells ELISA ..................................................................... Enzyme-linked immunosorbent assay GD ............................................................................................. γδ (Gamma Delta) T cells Hr .............................................................................................................................. Hour IBD .......................................................................... Irritable/Inflammatory bowel disorder IFN ..................................................................................................................... Interferon IgG1 ...................................................................................................... Immunoglobin G1 IgG2a .................................................................................................. Immunoglobin G2a IgG2b .................................................................................................. Immunoglobin G2b IgM .......................................................................................................... Immnuoglobin M IL ....................................................................................................................... Interleukin JD ............................................................................................................ Johne’s Disease LPMC ........................................................................... Lamina propria mononuclear cells xi MACS ................................................................................ Magnetic-activated cell sorting MAP ............... Mycobacterium avium subspecies paratuberculosis (M. paratuberculosis) MDBK ............................................................................. Madin-Darby bovine kidney cells MDCK ............................................................................. Madin-Darby canine kidney cells MDMs ............................................................................. Monocyte derived macrophages Min ..........................................................................................................................Minute MOI ................................................................................................ Multiplicity of infection ms ...........................................................................................................................Mouse NAHMS ............................................................ National animal health monitoring system Nil ................................................................................................................. Unstimulated OADC .................................................................................... Oleic acid dextrose catalase OD ............................................................................................................. Optical density PBMCs ...................................................................... Peripheral blood mononuclear cells PBS .......................................................................................... Phosphate buffered saline PPDj ......................................................................... Purified protein derivative of Johne’s PWM ................................................................................................... Pokeweed mitogen T1 ............................................................................................................... Treatment one T2 ............................................................................................................... Treatment two T3 .............................................................. Treatment three (A combination of T1 and T2) Tb ............................................................... Mycobacterium tuberculosis (M. tuberculosis) TCR1 ......................................................................................... γδ (Gamma Delta) T cells Th .............................................................................................. Helper T cell effector type TNF ................................................................................................. Tumor necrosis factor xii TNFR2 ............................................................................ Tumor necrosis factor receptor 2 UC ........................................................................................................... Ulcerative colitis α ........................................................................................................................ Alpha/anti αβ ..................................................................................................................... Alpha beta γδ ................................................................................................................. Gamma delta xiii INTRODUCTION INFLAMMATORY TH17 RESPONSES TO INFECTION WITH MYCOBACTERIUM AVIUM SUBSPECIES PARATUBERCULOSIS (MAP) IN CATTLE AND THEIR POTENTIAL ROLE IN DEVELOPMENT OF JOHNE’S DISEASE Justin L. DeKuiper and Paul M. Coussens Introduction Johne’s disease (JD) in cattle is of considerable concern to the dairy industry. JD is caused by Mycobacterium avium subspecies paratuberculosis (MAP), an intracellular pathogen. Dairy farmers lose an average of $200 million to $1.5 billion each year from increased culling, decreased production and increased testing associated with Johne’s disease (Lombard, 2013). In 2007, the estimated US national herd infection rate was 68% as determined by the National Animal Health Monitoring System (NAHMS; APHIS, 2007). This rate increased dramatically to about 91% from 2007 to 2013, though some differences in testing methods exist between these time points (Lombard, 2013). MAP primarily affects the ileum of cattle, causing inflammation and disruption of the intestinal lining, leading to incurable diarrhea and subsequently reducing the ability of infected animals to absorb nutrients. Healthy ileal tissue contains continuous villus structures lined with epithelial cells. As MAP infection progresses, inflammation increases, large numbers of infiltrating macrophages are typically found associated with sites of MAP infection, and the apparent number of villus structures decreases (Roussey, 2016). Previous studies have outlined some aspects of MAP infection and 1 potential immune responses against MAP using both peripheral blood mononuclear cells (PBMCs) and infected tissues (reviewed in Coussens, 2001; Stabel, 2006). Most of these studies focused on classical Th1-like and Th2-like immune responses, with a few studies addressing  T cells and regulatory T cells (Kabara, 2010; Roussey, 2014; Koets, 2015; Ganusov, 2015). Many studies have focused on the link between macrophages, the preferred host cell for intracellular MAP infection, and T cells. It has been suggested that immune cells from subclinical MAP infected cows initially respond to MAP antigens with a classical Th1-like immune response, including production of IFN. Clinical stages of Johne’s disease (JD) appear to coincide with a shift to a classical, humoral Th2-like immune response (Ganusov, 2015; Koets, 2015), including production of anti-MAP IgG antibodies. Currently, mechanisms resulting in early inflammatory responses to MAP and those detectable during the sub-clinical stage of infection in the ileum are not well understood. The initial Th1-like response does indeed appear to decline over time as primarily measured by IFN production from resident T cells. Mechanisms responsible for this loss in IFN production are not clear, but it has been suggested that both regulatory T cells and T cell exhaustion might be contributing factors (Roussey, 2014; Ganusov, 2015; Roussey, 2016). Lack of IFN production in MAP infected tissues and in peripheral T cells responding to MAP antigens is not accompanied in a loss of tissue inflammation. Even during this phase of infection (typically referred to as late subclinical or early clinical), inflammation of MAP infected tissues is quite profound. Although one might suggest that Tumor necrosis factor alpha (TNF) could be an important mediator of inflammation within MAP infected tissues, as in some human Crohn’s disease 2 tissues, TNF does not appear to be as important with MAP in cattle (Khalifeh and Stabel 2004). Thus, at sites of MAP infection, the abundance of both TNF and IFN is not necessarily consistent with the degree of inflammation observed (Roussey, 2016; Khalifeh and Stabel 2004; Coussens, 2004). Understanding what factors might be driving inflammation in Johne’s disease associated lesions therefore requires us to look beyond the classic Th1-like response mediators often associated with such processes. One possibility for unchecked inflammation may be the presence of large amounts of IL- 1 and IL-1 described previously (Aho, 2003; Sommer, 2009; Chiang, 2007). Another possibility is the potential development of a non-classical and highly proinflammatory Th17-like response. In this review we summarize current knowledge of the inflammatory response to Johne’s disease caused by MAP infection in cattle. We review evidence that Th17 cells and the cytokines which promote Th17 cell development may be associated with MAP-induced inflammation independent of the role of classical Th1-like cytokines, such as IFN and TNF. The Th17 Response T cells are influenced to differentiate into specific subtypes by secreted proteins (mainly cytokines) from antigen presenting cells (APCs) and epithelial cells. T cells respond to local cytokine combinations and co-stimulatory factors by developing into cytotoxic cells, helper T cells leading to enhanced humoral immunity, regulatory T cells, or into more complex pro-inflammatory mediators, including Th17 cells. Th17 responses are so called due to the production of the cytokine interleukin-17a (IL-17A) by T cells. The Th17 pathway is typically outside the normal pathogen response paradigm, but has been shown to be responsible for clearing extracellular pathogens and inducing 3 inflammation (Zielinski, 2012; Santarlasci, 2009). Th17 cells originate from a distinct subset of helper T cells and their activity can dampen other T-cell response pathways (Santarlasci, 2009; Harrington, 2005; Cosmi, 2008). In some cases, other cell types may be coerced to adopt a Th17-like phenotype. These include a subset of γδ T cells (Peckham, 2014) and even cells that are terminally differentiated, such as activated regulatory T cells (T-reg) (Bhaskaran, 2015). T cells may also adopt a Th17 phenotype following direct stimulation of Toll-Like Receptors 2 and 4 (TLR-2, 4) on the T cell surface (Nyirenda, 2011; Bhaskaran, 2015), which are thought to have involvement with Mycobacterial infection (Mucha, 2009; Sánchez, 2010). Unlike αβ T cells, γδ T cells do not require antigen presentation via the major histocompatibility complex (MHC) for stimulation to produce IL-17A or other cytokines (Baldwin, 2014). The unique T cell receptor (TCR) on γδ T cells allows direct stimulation by many antigens (Zeng, 2012; Baldwin, 2014). Also, γδ T cells do not require IL-23 for initial production of IL-17A (Lee, 2015). However, continued IL-17A secretion by γδ T cells such as those seen by Baldwin (Baldwin, 2014) may require IL-23 acting on IL-23R positive cells. Tissue- resident IL-23R+ γδ T cells could thus be a source of IL-17A during early infection with MAP, either with or without IL-23 production. During tissue injury, IL-23R+ γδ T cells have been shown to induce inflammation and limit permeability of intestinal epithelium (Lee, 2015). Our own work and that of others has shown that γδ T cells respond to MAP antigen stimulation and are present in MAP infected ileal tissues. Furthermore, the intestinal homing molecule CCR6 is expressed on the surface on the surface of both γδ T cells (Zeng, 2012; Lugering, 2005; Lyadova, 2015) and CD4+ Th17 cells (Duhen and Campbell 2014; Lee, 2015). Lastly, mRNA encoding lymphocyte chemotactic factor 4 CCL20 is also upregulated in MAP-infected tissues (Khare, 2012). Together CCR6 and CCL20 expression would support migration of these Th17-like cells to intestinal tissues, mucosal sites, and Peyer’s patches (Duhen and Campbell 2014; Lim, 2008; Lugering, 2005; Rout 2016), the primary sites of MAP infection. During the initial 18 hours of infection with MAP, expression of mRNAs encoding cytokines that could direct T cells to differentiate into a Th17-like phenotype, are upregulated. This has been documented for PBMCs exposed to MAP, as well as in MAP-infected macrophages (IL-6 and IL-23) (Roussey, 2014; Dudemaine, 2014). In lesions with low MAP burdens (Grade 1), a similar pattern of mRNA expression is observed (primarily IL-17A and IL-6) (Roussey, 2016). In tissues that were heavily infected with MAP, a Th17 associated IL-17A response was not detected. This suggests that a non-classical Th17-like response by T cells, could play a role in early stage lesions of MAP-infected tissues from JD+ cows, but production of IL-17A declines with advancing disease, much like the decline in classical Th1 responses. The phenotype of both γδ T cells and αβ T cells depends, in part, on the local mix of cytokines. APCs (Siakavellas and Bamias 2012; Dudemaine, 2014) and epithelial cells (Lee, 2017; Seydel, 1997; Ghadimi, 2012) can all produce Th17 directive cytokines, such as IL-23, IL-6, IL-1β. The presence of IL-23, IL-6, and IL-1β leads to the development of a Th17-like phenotype in naïve T cells and secretion of Th17 cytokines (Neurath 2007; Siakavellas and Bamias 2012; Passos, 2010). Our recent work has shown that both the Th17 directing cytokine IL-23 and potential Th17 cells expressing the IL-23 receptor (IL-23R) are more abundant in peripheral blood of Johne’s disease positive cows than in blood from healthy cows. Further confirmation of MAP’s influence 5 on IL-23R expression in T cells was observed after treating PBMCs from healthy cows with MAP in-vitro. MAP was capable of stimulating expression of IL-23R as observed by increases in the mean relative percent of T cells expressing IL-23R following exposure to MAP (DeKuiper and Coussens, 2019). Th17 Cytokines Cytokines responsible for driving Th17 immune responses are capable of affecting other pathways in the immune system and can have implications for the surrounding tissues. The primary cytokines involved in Th17 cell differentiation are IL-6, TGFβ, IL-23, and IL-1β (Bettelli, 2008; McGeachy, 2009), while IL-21 appears to be responsible for amplifying Th17 cells following differentiation (Nurieva, 2007). Most Th17 driving cytokines can exert deleterious effects if their expression is not tightly controlled. For example, continuous stimulation by IL-23 can cause inflammation (Lee, 2017; Paustian, 2017) and depletion of type 3 innate leukocyte cells (ILC3) in the proximal small intestines of mice (Paustian, 2017). Inhibiting IL-23 promotes an anti- inflammatory gut environment, diminishing symptoms from irritable bowel disorder (IBD) in mice (Maxwell et al, 2015). Thus, in intestinal tissues, IL-23 appears to be predominately a destructive inflammation inducing cytokine. Similarly, IL-1β is important in mounting a proper Th17 response against M. tuberculosis but can lead to inflammation and tissue injury. IL-6 exhibits a dual nature, demonstrating anti- inflammatory or regulatory properties in muscle and other tissues, but in intestinal tissues, macrophage associated IL-6 causes disruption in tissue integrity (Gabay, 2010; Dinarello 2011; Sosenko, 2006). Clearly, failure to properly limit and control expression of one or more of these Th17-driving cytokines could lead to the type of tissue injury 6 and rampant inflammation observed in clinical Johne’s disease. In recent studies, we have observed enhanced serum IL-23 levels and decreased IL-17A levels in blood from cows with Johne’s disease relative to serum healthy controls (DeKuiper and Coussens, 2019). Immune cells responding to Th17 instructive cytokines often secrete both IL-17 and IL-22. Colonic epithelial cells respond to IL-22 by increasing Claudin-2, thus increasing cell permeability of the tissue (Wang, 2017). However, IL-22 is most often found to have a protective role. For example, application of IL-22 to mice with induced colitis caused tissues to revert to a normal phenotype and protects them from irritable bowel disease (Aden, 2016; Zenewicz, 2008). M. tuberculosis studies have shown IL-22 is important for macrophagic phagosomal fusion and M. tuberculosis growth inhibition (Dhiman, 2009). One possibility for the co-production of IL-17A and Il-22 would be to limit the tissue damage and inflammation produced by IL-17A promoting cytokines such as IL-1 and IL-23. MAP Driving Th17 It is clear that, in the initial 18 hours, expression of mRNAs encoding cytokines that direct T-cell responsiveness and differentiation to a Th17-like response (IL-6, IL-1, IL-23) are upregulated in PBMCs exposed to MAP and MAP-infected macrophages (Roussey, 2014). Enhanced expression of mRNAs encoding IL-17A and IL-6 was also observed in early-stage (Grade 1) lesions from MAP infected cows (Roussey, 2016). The initial increase in mRNAs encoding IL-6 and IL-17A in early stage MAP ileal tissue lesions decreases as MAP infection and lesion scores (Grade 2-4) increase (Roussey, 2016). This scenario is similar to what is observed with challenge infection of calves 7 with Mycobacterium bovis (M. bovis). In this model, mRNA encoding IL-17A is upregulated in lung granulomas with low bacterial burden (early stage; Grade 1) compared to granulomas with high bacterial burdens (Grade 3 and 4) (Palmer, 2016). In both models, considering the relationship between Th17 promoting cytokines and inflammation, it is possible that continual IL-23 expression promotes tissue injury and inflammation. Eventual loss of IL-17A production as lesion burdens increase could be attributed to T-cell exhaustion from prolonged IL-23 exposure (Paustian, 2017). To our knowledge, this possibility has not yet been tested in either MAP lesions or in lesions from M. bovis infections. However, as noted above, we have observed enhanced IL-23 and reduced IL-17A levels in serum from cows with Johne’s disease relative to serum from controls (DeKuiper and Coussens, 2019). In vitro, monocyte-derived macrophages (MDMs) upregulate IL-23, IL-6, and IL- 1β mRNA when exposed to MAP (Dudemaine, 2014). MDM expression of both IL-1 and IL-1β mRNA and protein can be quite significant following infection with MAP (Chiang, 2007). Monocytes, precursors to macrophages and dendritic cells, can produce IL-23 when exposed to Mtb (Stephen-Victor, 2016). Dendritic cells produce IL- 23 in the presence of M. bovis antigen and Mtb (Szpakowski, 2015; Thacker, 2009; Stephen-Victor, 2016). In addition to antigen presenting cells (APCs), such as macrophages and dendritic cells, IL-23 may originate from epithelial cells (Lee, 2017; Seydel, 1997) at sites of MAP infection in intestinal tissues. While it has been established that Madin–Darby bovine kidney (MDBK) epithelial cells upregulate IL-6 when infected with MAP (Everman, 2015), expression of IL-23 from these model cells has not been explored in the context of MAP exposure. 8 Final Thoughts There is now ample evidence that Th17 cells and IL-17A are important pieces of the immune response to MAP and other mycobacteria. What is not yet clear is the precise role these cells might play in both control of infection and development of disease. On one hand, IL-17A secretion might provide an initial inflammatory response that helps to clear or limit infection. There is evidence that both  T cells and  T cells might participate in this early response. However, continued expression of IL-17A and the Th17 driving cytokines IL-1 and IL-23 may lead to progressive inflammation and disease symptoms. Importantly, inflammation derived from epithelial or macrophage IL- 23 and IL-1 would not be controlled by T cell exhaustion or other T cell regulatory mechanisms that would reduce IL-17A and IFN in late stage infection with MAP. Thus, loss of the classic Th1 cytokines IFN and TNF, as observed in advanced Johne’s disease, nor loss of IL-17A expression as observed in our recent studies (Table 1), would not reduce IL-23 or IL-1 driven inflammation at sites of infection. Clearly, the role of Th17 associated cytokines in development of clinical Johne’s disease warrants further study. In addition, the rapid and regulated expression of IL-17A in early infection as one mechanism that aides in clearance or control of MAP and other mycobacteria in some animals is worthy of additional studies, particularly as a potential marker of vaccine induced protective immune responses. Acknowledgements The authors gratefully acknowledge editorial assistance from Dr. Melinda R. Wilson. Financial support from the United States Department of Agriculture National 9 Institute of Food and Agriculture (grants 2012-67011-19936, and 2013-68004-20371), the Michigan Animal Agriculture Alliance, and Michigan AgBioResearch is also gratefully acknowledged. 10 APPENDIX 11 Table 1: MAP-induced cytokine expression during subclinical and clinical stages of disease. Cytokines Activated by Expressed in Tissue or PBMC? Found in sub- clinical? Found in clinical? Reference TNFa MAP Both Yes No IFNg MAP Both Yes No Roussey, 2014; Roussey, 2016 Roussey, 2014; Roussey, 2016 IL-1β MAP Macrophages/Tissue Yes Unknown Kabara, 2010; IL-6 MAP PBMC/Macrophage/E pithelial/Tissue Yes No IL-10 MAP Macrophages/Tissue Yes No Coussens, 2004 Roussey, 2014; Murphy, 2007; Everman, 2015; Roussey, 2016 Janagama, 2006; Roussey, 2016 IL-23 MAP PBMC/Macrophage Yes Unknown Roussey, 2014; Dudemaine, 2014 IL-22 IL-23 Unknown Unknown Unknown IL-17A MAP/IL-23 Both Yes No Roussey, 2014; Roussey, 2016; DeKuiper, 2019 12 REFERENCES 13 REFERENCES Aden, K., Rehman, A., Falk-Paulsen, M., Secher, T., Kuiper, J., Tran, F., … Rosenstiel, P. (2016). Epithelial IL-23R Signaling Licenses Protective IL-22 Responses in Intestinal Inflammation. Cell Reports, 16(8), 2208–2218. https://doi.org/10.1016/j.celrep.2016.07.054 Aho, A. D., McNulty, A. M., & Coussens, P. M. (2003). 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Coussens Abstract Johne’s disease (JD) is a chronic inflammatory gastrointestinal disease of ruminants caused by Mycobacterium avium subspecies paratuberculosis (MAP). Control of JD is difficult largely due to insensitive diagnostic tools, a long subclinical stage of infection, and lack of effective vaccines. Correlates of protection are lacking in model systems of JD and the sources of inflammation due to JD are not well characterized. Commonly studied immune responses, such as the Th1/Th2 paradigm, do not adequately explain host responses to MAP. A potential role for non-classical immune responses to MAP, such as that mediated by Th17 cells, has been suggested. Indeed, MAP antigens induce mRNAs encoding the cytokines IL-23 and IL-17A in bovine peripheral blood mononuclear cells (PBMCs). IL-23 and IL-17A production have both been associated with Th17-like immune responses. Th17 cells are also defined by surface expression of the IL-23 receptor (IL-23R). To determine the relative prevalence of potential Th17 cells in PBMCs from MAP test positive and MAP test negative cows, PBMCs were isolated and analyzed by immunostaining and flow cytometry. Fresh PBMCs from MAP test positive cows (n=12) contained a significantly higher proportion 21 of IL-23R positive cells in populations of CD4+, CD8+, and TCR1+ (γδ) T cells than in cells from MAP test negative cows (n=12; p<0.05). Treatment with MAP antigens increased the percentage of all T cell subsets with surface expression of IL-23R when compared to untreated (n=12; p<0.05) cells. ELISA results for IL-17A secretion revealed a higher concentration of IL-17A secreted from PBMCs treated with MAP antigen (n=20) than from PBMCs not treated with MAP antigens (n=20) (p<0.001), regardless of the JD test status of source cows. Also, we observed a moderate negative correlation between JD diagnostic scores for JD+ cows and plasma IL-17A concentration (n=42; r= -0.437; p-value<0.004). Plasma with low and mid JD- scores (n=31; n=9; 0.1≤X<0.3) had significantly more IL-17A when compared to plasma with high JD- scores (n=10; 0.3≤X<0.46; p-values < 0.05). Similarly, plasma with low JD+ score values (0.55≤X<1.0; n=9) had significantly more IL-17A when compared to plasma with high JD+ score values (X≥2.0; n=21; p<0.05). Overall, plasma from JD+ cows (0.55 2.00; n=18) for a total of 6 groups (3 in each JD status group). Indeed JD- cows generally had significantly higher IL-17A in circulation than JD+ cows (Mann-Whitney; p-value < 0.05) (Fig.6). However, Welch’s ANOVA analysis revealed significant differences amongst the groups (p-value < 0.01) (Fig. 6). Using Games-Howell’s multiple comparisons test, plasma from low JD- and mid JD- cows had significantly higher levels of IL-17A than plasma from high JD- cows (p- values = 0.07) (Fig.6). Within the JD+ group, cows with low JD+ OD scores had significantly more IL-17A in plasma than samples from the high JD+ cows (p-value < 0.01). High JD- scored samples include those from “JD suspect” cows, as defined above. The high JD- revealed significantly less IL-17A than low JD+ (p-value < 0.05). The low JD- also had significantly more IL-17A compared to high JD+ (p-value < 0.05). To further analyze the trend seen within the disease groups, we conducted a correlation study between JD plasma ELISA OD scores and the amount of IL-17A within the sample. Circulating IL-17A levels were found to be moderately negatively correlated 35 with JD+ cow OD scores as they increase (Pearson score ® = -0.437; p-value < 0.004) (Fig. 7). No correlation was seen within the JD- group (not shown). Discussion Th17-like cells may be of potential importance in development of, or host response to Johne’s disease (Roussey, 2014, Roussey, 2016). The precise role of Th17 responses in Johne’s disease are not clear at present and the existing literature can be conflicting. For example, IL-17A plays a protective role in some inflammatory diseases (Maxwell, 2015, Lee, 2015) and in mycobacterial infections when expressed early following infection (Palmer, 2016, Khader, 2011). However, chronic inflammation, a distinct feature of MAP infection, can also be induced by IL-17A (Leppkes, 2009) and by the Th17 promoting cytokine IL-23 (Neurath, 2007, 2019). Promotion of a Th17-like phenotype over other T-cell phenotypes can be influenced by the immediate cytokine environment with IL-1β, IL-6, IL-23, and TGF-β all promoting a Th17-like phenotype in responsive T cells (Passos, 2010, Neurath, 2007, Santarlasci, 2009, Nyirenda, 2011). MAP is known to upregulate expression of mRNAs encoding IL-6, IL-23, IL-1β, and TGF-β as early as 1-hour post infection in MAP-infected MDMs (Dudemaine, 2014), suggesting that the local cytokine environment near sites of MAP infection would tend to promote Th17 T cell development. One untested possibility is that IL-17A is helpful during early MAP infection, but failure to tightly regulated expression of IL-17A or IL-23 could lead to inflammation characteristic of clinical JD. In this study, we sought to determine if the major cytokine product of Th17-like T cells, IL-17A was associated with MAP infection and Johne’s disease. Previous work 36 from our laboratory suggested that PBMCs from both MAP-positive (JD+) and MAP- negative (JD-) cows upregulated Mrna encoding IL-17A when exposed to MAP-infected MDMs (Roussey, 2014). These studies support the notion that MAP might be a general stimulator of IL-17A Mrna expression from PBMCs. To extend these observations to IL- 17A protein secretion, we focused on co-cultures of MDMs (from JD- cows) and autologous PBMCs in the presence of different MAP treatments (Fig. 1). All treatments with MAP were able to significantly increase IL-17A production from PBMCs when compared to samples not exposed to MAP. A lack of differences between treatments indicated that prior exposure of PBMCs to MAP-infected MDMs was not required for live MAP bacteria to stimulate IL-17A secretion (Fig. 2). We therefore simplified our model to treatment of PBMCs with MAP only, without co-culture with autologous MDMs (Fig. 3A). Data presented in this report demonstrate that MAP is indeed capable of increasing IL-17A levels produced by PBMCs. This effect is not dependent on the disease status of PBMC source cows, nor on the specific strain of MAP used. Our results thus suggest that MAP acts as a general and stimulator of IL-17A secretion from PBMCs. As to the possible mechanisms of MAP-induced IL-17A secretion, there is the possibility that monocytes or B cells in PBMCs are taking up MAP and presenting it to T cells. MAP may also be inducing IL-17A secretion more directly via a mechanism similar to Toll-like receptor (TLR) signaling. Experiments with purified populations of T cells and TLR blocking reagents will help to define these mechanisms. In either case, our results suggest that IL-17A may be part of an innate immune response to MAP. 37 T cells known to produce IL-17A, including during mycobacterial infections, are CD4+, CD8+, and γδ T cells (Roussey, 2014, Srenathan, 2016, Steinbach, 2016). Of note, γδ T cells (including IL-23R+ γδ T cells) can produce IL-17A independent of IL-23 and are an innate source of IL-17A (Lee et al 2015, Zeng, 2012). However, long term expression of IL-17A in all cells studied to date requires IL-23 acting through IL-23R (Khader, 2011). Increased production of IL-23 has been observed in MDM cells infected with not only MAP (Dudemaine, 2014) but Mycobacterium avium isolates as well (Agdestein, 2014). Thus, macrophages could act as one source of IL-23 that would help promote IL-17A production in local regions of MAP infection associated with Johne’s disease. Although not in vivo, previous work in our laboratory noted that Mrna encoding IL-23 was upregulated in PBMCs stimulated with MAP relative to appropriate control cells (Roussey, 2014). Since Mrna increases do not always translate into enhanced protein production, additional experiments examining IL-23 and IL-17A secretion seem warranted. In the current study, we demonstrated that the mean relative percent (MRP) of CD4+, CD8+ and TCR1+ (γδ) T cells expressing IL-23RLow and IL-23RHigh was greater in untreated PBMCs from JD+ cows than in untreated PBMCs from JD- cows (Fig. 4B & 4C). This novel data suggests that natural MAP infection causes circulating T cells to upregulate IL-23R expression. The significant increase of IL-23R on T cells from JD+ cows may suggest that JD+ T cells would be more responsive to IL-23. We also found that MAP antigen stimulation enhanced the MRP of IL-23RLow T cells in PBMCs from JD- cows up to levels observed in circulating cells from JD+ cows. Thus providing evidence that MAP can indeed upregulate IL-23R expression on T cells. Although it did 38 not appear that differences in IL-23R expression translated into more IL-17A production by JD+ PBMCs in our limited studies, examination of MAP-induced lesions may be more informative. It is unclear at this point what the difference is between cells with IL- 23RHigh and IL-23RLow, but the level of IL-23R expression may be related to their specific function, as noted in other studies (Liang, 2013; Sivanesan, 2016). In addition, longer time studies and IL-23R expression as well as studies using recombinant IL-23 to stimulate cells from JD- and JD+ cows with subsequent measure of IL-17A secretion should be informative. We lastly sought to understand the significance of IL-17A in plasma as it pertains to Johne’s disease status. Overall, the mean IL-17A level in plasma from JD+ cows is significantly less than in plasma from JD- cows. Also, IL-17A levels have a moderate negative correlation with increasing JD+ MAP ELISA scores (Fig. 7). This correlation is also observed in the different JD+ disease score groups (Fig. 6). Roussey et al. (2014) demonstrated that CD4+ T cells from subclinical cows responded to MAP-infected macrophages in part by upregulating IL-17A Mrna expression, however, CD4+ T cells from clinical cows did not. This was further demonstrated in ileal tissues in which increasing MAP-burdens in lesions lead to decreasing levels of Mrna encoding IL-17A (Roussey et al 2016). Thus, our results would appear to be entirely consistent with other reports. Dudemaine et al. (2014), demonstrated that cows with double MAP positive scores (fecal and serum) had significantly more IL-17A in plasma, while cows with only single MAP-positive scores (fecal) showed significantly less IL-17A than double negatives and double positives. Typically, the very late stages of Johne’s disease are accompanied by a loss in both Th1 and Th2 responses. With this in mind, it is possible 39 that the double positive JD cows studied by Dudemaine et al. (2014) represent an earlier infection stage than the single positives. Future studies will focus on which cells are responsible for either an innate or acquired Th17-like response to MAP, including potential MAP responding T cells and the potential role of epithelial cells, particularly with regard to IL-23 production. Studies within lesions from infected cows should also prove valuable in discerning the role of Th17 cells in MAP infections and Johne’s disease. To our knowledge, this study is one of the first to look at IL-23R as an expression marker in natural infection and in response to MAP antigens in culture. Previous studies have concluded that Th17 related mRNAs encoding IL-23 and IL-17A are increased in PBMCs and MDMs treated with MAP (Roussey, 2014; Dudemaine, 2014). Our data now confirm that IL-17A protein is indeed upregulated in cells from both JD- and JD+ cows in response to MAP antigen stimulation. Although IL-17A does not seem to be a good potential indicator of JD disease status for serum based diagnostic tools, it does add a potential layer of difference between subclinical cows (MAP infected) and cows who are clinically diagnosed. A major limitation of our work thus far is not identifying the mechanism of MAP-induced IL-17A secretion in cells from JD- and JD+ cows. Another unanswered question is the effect that IL-23 will have on cells from JD+ versus JD- cows, where our data suggests significant differences in IL-23R expression. Finally, it will be of interest to translate findings presented in this report to tissues with defined lesion scores from MAP infected cows. 40 Acknowledgements The authors gratefully acknowledge Dr. Juan Pedro Steibel for statistical consulting and contributions from other members of the Molecular Pathogenesis Laboratory, particularly Ashley Greenlick, Monika Dziuba, and Caitlin Ancel. The authors would also like to thank the owners and managers of the commercial dairy operations that participated in this study. This work was supported by the United States Department of Agriculture and the National Institute of Food and Agriculture (grants 2012-67011-19936, and 2013-68004-20371). 41 APPENDICES 42 APPENDIX A Table 2: Primary antibodies. Immuno-staining of T-cell specific surface markers and Th17 signature receptors and cytokines. Antibody CD4 CD8 TCR1 IL-23R Dilution 1:100 1:100 1:100 1:100 Company WSU WSU WSU Sigma Specificity Clone Bovine Bovine Bovine Human CACT138A BAQ111A GB21A 3D7 Isotype IgG1 IgM IgG2b IgG2ak 43 APPENDIX B Table 3: Secondary antibodies. Immuno-staining of T-cell specific surface markers and Th17 signature receptors and cytokines. Antibody FITC R-PE PE-Cy7 AF647 Dilution 1:1000 1:200 1:200 1:1000 Company eBioscience eBioscience Invitrogen Invitrogen Specificity Clone α ms-IgG2b m2b-25g4 α ms-IgG2a γ2a α ms-IgM α ms-IgG1 m1-14d12 γ1 44 APPENDIX C Table 4: 2-Way ANOVA using Tukey's Multiple Comparison Test to compare all the means to each other. Reporting p-values. Tukey’s Multiple Comparison Test JD- Nil V JD- MAP JD- Nil V JD- MAP-K10 JD- Nil V JD+ MAP JD- Nil V JD+ MAP-K10 JD+ Nil V JD+ MAP JD+ Nil V JD+ MAP-K10 JD+ Nil V JD- MAP JD+ Nil V JD- MAP-K10 p-value 0.025 0.014 0.056 0.018 0.070 0.022 0.032 0.018 45 APPENDIX D Figure 1: Experimental set-up for analysis effects of MDMs on IL-17A production by PBMCs during MAP stimulation. MDMs cocultured with autologous PBMCs and their respective treatments. Treatment 1 (T1) is defined as an initial infection of MDMs with MAP (MOI: 20), 20-hours prior to the addition of PBMCs. Treatment 2 (T2) is defined as an uninfected MDM culture with a stimulation of MAP (MOI: 2) for 18 hours after the initial 20 hours and addition of PBMCs. Treatment 3 (T3) is defined as the combination of T1 and T2. Supernatants were collected after a total of 38 hours in culture for IL-17A ELISA. 46 APPENDIX E Figure 2: Effect of MAP-infected MDMs on IL-17A secretion in PBMC cultures from JD- cows. IL-17A ELISA of supernatants from cocultures of MDMs with autologous PBMCs after 38-hour total culture with their respective treatments (Fig. 1). n=5/treatment. Analyzed by Kruskal-Wallace and Dunn’s multiple comparison test. * = p-value < 0.10. ** = p-value < 0.05. Error bars = SEM. Grubb’s Test = no outliers found in any group. 47 APPENDIX F Figure 3A: Experimental set-up for analysis of PBMCs from JD+ and JD- cows stimulated with M. paratuberculosis (MAP). PBMCs from JD+ and JD- cows were cultured and left untreated or treated with PWM (25 μg/mL), purified protein derivative of Johne’s (PPDj; 10 μg/mL), or MAP (MOI: 2) for 18 hours. After incubation supernatants were collected and froze at -80°C for later IL-17A ELISA analysis. PBMCs were stained for surface markers (CD4, CD8, TCR1, and IL-23R) analyzed using flow cytometry (Tables 1 and 2). 48 APPENDIX G Figure 3B: Secretion of IL-17A by PBMCs from JD+ and JD- cows stimulated with M. paratuberculosis (MAP). IL-17A ELISA of PBMC supernatants after 18-hour stimulation with their respective treatments. Different letters indicate significant differences (Table 3); p-values by two-way ANOVA and Tukey’s multiple comparison test. Error bars = SEM. 49 APPENDIX H Figure 4A: Flow cytometry gating strategy example. Strategy includes no stain control (not shown), negative primary control (secondary only), and a negative IL-23R control (all other 1° Ab and all 2° Ab). This same strategy is used for gating of the other T cell subtypes and IL-23R. The IL-23R gating strategy within the T cell sub type populations uses the left gate as IL-23RLow and the right gate as IL-23RHigh. All events outside of the PBMC gate are considered debris. 50 APPENDIX I Figure 4B: Untreated JD+ vs. JD- IL-23RLow. The mean relative percent (MRP) of T cell subtypes (CD4, GD, CD8) expressing a low level of IL-23R (IL-23RLow) in unstimulated PBMCs from JD+ (n = 12, 12, 11 respectively) and JD- (n = 11, 12, 11 respectively) cows.* = p-value < 0.1. ** = p-value < 0.05. *** = p-value < 0.01. Error bars = SEM. Student T-test was used when all assumptions were met; Welch’s T-test was used when samples passed normality testing but had unequal variance; Mann-Whitney test was used when normality of the sample was not met in one sample or both. 51 APPENDIX J Figure 4C: Untreated JD+ vs. JD- IL-23RHigh. The mean relative percent (MRP) of T cell subtypes (CD4, GD, CD8) expressing a high level of IL-23R (IL-23RHigh) in unstimulated PBMCs from JD+ (n = 12, 12, 11 respectively) and JD- (n = 11, 12, 12 respectively) cows. * = p-value < 0.1. ** = p-value < 0.05. *** = p-value < 0.01. Error bars = SEM. Student T-test was used when all assumptions were met; Welch’s T-test was used when samples passed normality testing but had unequal variance; Mann- Whitney test was used when normality of the sample was not met in one sample or both. 52 APPENDIX K Figure 4D: Surface expression of IL-23RLow on JD- T cells after stimulation with MAP antigen. The mean relative percent (MRP) of T cell subtypes (CD4, GD, CD8) expressing a low level of IL-23R (IL-23RLow) in stimulated PBMCs from JD- (n = 12/treatment/group respectively) compared to their respective unstimulated controls (n = 11, 12, 11 respectively). Untreated JD+ T cells (grey) is included as a visual reference for comparison only. * = p-value < 0.1. ** = p-value < 0.05. *** = p-value < 0.01. Error bars = SEM. Student T-test was used when all assumptions were met; Welch’s T-test was used when samples passed normality testing but had unequal variance; Mann- Whitney test was used when normality of the sample was not met in one sample or both. 53 APPENDIX L Figure 5: Plasma IL-17A levels of cows based on environmental MAP status of farm and JD status. IL-17A concentration (pg/mL) in plasma by ELISA. JD+ cows (grey; n=19) are from the same environmentally MAP+ commercial farm as the JD- cows (solid black; n=18). JD- cows from an environmentally MAP- farm (black and white horizontal stripe; n=18) are from the Michigan State University Dairy Cattle Research and Teaching Center. ** = p-value < 0.05. Error bars = SEM. Samples were Log10 transformed then Kruskal-Wallis and Dunn’s multiple comparison were used for statistical analysis. 54 APPENDIX M Figure 6: Plasma IL-17A levels of cows based on IDEXX Johne’s ELISA score. IL- 17A concentrations (pg/mL) circulating in the plasma from the periphery of by ELISA.. Low JD- (x<0.2; n=50). Mid JD- (0.2 2.0; n=18). Overall JD- cows (n=68). Overall JD+ cows (n=40). * = p-value < 0.1. ** = p-value < 0.05. *** = p- value < 0.01. Error bars = SEM. Cow n is based on available stocked plasma samples. Welch’s ANOVA and Games-Howell's multiple comparisons test were used in the observation of score groups. Mann-Whitney test was used between overall JD- and JD+ groups. 55 APPENDIX N Figure 7: Correlation of JD+ IDEXX ELISA score and IL-17A plasma ELISA concentration. Linear regression and correlation analysis. Pearson score (r) = -0.44. p- value < 0.004. n=42. 56 REFERENCES 57 REFERENCES Agdestein, A., Jones, A., Flatberg, A., Johansen, T. B., Heffernan, I. A., Djønne, B., … Olsen, I. (2014). Intracellular growth of Mycobacterium avium subspecies and global transcriptional responses in human macrophages after infection. 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M., & Jones, G. J. (2016). CD4+ and πσT Cells are the main Producers of IL-22 and IL-17A in Lymphocytes from Mycobacterium bovis- infected Cattle. Scientific Reports, 6(May), 1–10. https://doi.org/10.1038/srep29990 Waters, W. R., Maggioli, M. F., Palmer, M. V., Thacker, T. C., McGill, J. L., Vordermeier, H. M., … Larsen, M. H. (2016). Interleukin-17A as a Biomarker for Bovine Tuberculosis. Clinical and Vaccine Immunology, 23(2), 168–180. https://doi.org/10.1128/CVI.00637-15 Zeng, X., Wei, Y. L., Huang, J., Newell, E. W., Yu, H., Kidd, B. A., … Chien, Y. H. (2012). γδ T Cells Recognize a Microbial Encoded B Cell Antigen to Initiate a Rapid Antigen-Specific Interleukin-17 Response. Immunity, 37(3), 524–534. https://doi.org/10.1016/j.immuni.2012.06.011 62 CHAPTER 2 MYCOBACTERIUM AVIUM SUBSPECIES PARATUBERCULOSIS (MAP) DRIVES AN INNATE TH17-LIKE T CELL RESPONSE REGARDLESS OF THE PRESENCE OF ANTIGEN-PRESENTING CELLS Justin L. DeKuiper, Hannah E. Cooperider, Noah Lubben, Caitlin M. Ancel, and Paul M. Coussens Abstract Johne’s disease (JD) is a chronic inflammatory gastrointestinal disease of ruminants caused by Mycobacterium avium subspecies paratuberculosis (MAP). Control of JD is difficult largely due to insensitive diagnostic tools, a long subclinical stage of infection, and lack of effective vaccines. Correlates of protection are lacking in model systems of JD and the sources of inflammation due to JD are not well characterized. As an alternative to commonly studied immune responses, such as the Th1/Th2 paradigm, a non-classical Th17 immune response to MAP, has been suggested. Indeed, MAP antigens induce mRNAs encoding the Th17 associated cytokines IL-17A, IL-17F, IL-22, IL-23, IL-27, and IFNγ in CD3+ T cell cultures as determined by RT-qPCR. Although not as robust as when cultured with MDMs, MAP is able to stimulate the upregulation of these cytokines from sorted CD3+ T cells in the absence of APCs. CD4+ and CD8+ T cells are the main contributors of IL-17A and IL- 22 in the absence of APCs. However, MAP stimulated monocyte derived macrophages (MDMs) are the main contributor of IL-23. In-vivo, JD+ cows have more circulating IL-23 63 JD- cows, suggesting this proinflammatory cytokine may be important in the etiology of JD. Our data suggests that Th17-like cells and associated cytokines may indeed play an important role in immune responses to MAP infection and development or control of JD. Introduction Mycobacterium avium subspecies paratuberculosis (MAP) is the causative agent for the clinical onset of Johne’s Disease (JD) in ruminants. MAP infection of the ileum reduces the ability of an animal to absorb nutrients through inflammation and disruption of the intestinal lining, as well as chronic diarrhea. Clinical JD leads to reduced milk production, culling, and/or premature death. The cumulative effects of JD are a growing concern to both animal welfare and the dairy industry. According to the 2007 National Animal Health Monitoring System (NAHMS; APHIS, 2007), the percentage of dairy operations infected with MAP in the US was approximately 68% and may now be as high as 91% (Lombard, 2013). JD economically represents a $200 million to $1.3 billion annual loss to the US dairy industry (Garcia, 2015). The cumulative effects of a long subclinical stage of infection, a lack of effective vaccines and insensitive diagnostic tools have made controlling JD difficult. Defining protective immune responses to MAP has also been difficult. Recent studies suggest that MAP may induce an early or pre-clinical Th17-like immune response (DeKuiper, 2019) in addition to the traditional Th1 and Th2 responses that have been extensively studied in both experimental and natural infections with MAP (Zurbrick, 1988; Coussens, 2004; Jun, 2009; Begg, 2011; Ganusov, 2015; Hempel, 2015; Koets, 2015; Weiss, 2002) Th17 cells produce IL-17A, IL-17F, and IL-22 in response to IL-23 acting through the IL-23 receptor (IL-23R) (Ahern, 2010) on αβ and γδ T cell surfaces (Korn, 2009; Cua 64 2010; DeKuiper, 2019). Antagonistic to Th17, typically, IL-27 and proinflammatory IFNγ are inhibitors of the Th17 pathway (Harrington, 2005; Diveu, 2009). IL-27 limits IL-17 responses through the inhibition of RORγ(c) (Diveu, 2009) and is a known inhibitor of Th17 in humans and mice (Batten, 2006; Liu, 2011), while IFNγ inhibits IL-23R expression (Harrington, 2005). IL-22 and IL-17F have a synergistic effect with IL-17A (Dixon, 2016; Leppkes, 2009). Although IL-17A has a much more profound effect on epithelial cells than IL-17F (Bougorn, 2011), IL-17F is increased in IL-17A deficient mice and can cause inflammation in the absence of IL-17A (Leppkes, 2009). Th17 cells are not limited and produce other Th class cytokines, such as IFNγ and IL-10 (Zielinski, 2012). Th17 cells co-expressing IFNγ and IL-17A also express IL-17F (Schmidt, 2013). Support for the notion that MAP stimulates a Th17 response can be found in studies with other Mycobacteria, such as M. tuberculosis and M. bovis, which are known to stimulate Th17 cytokines (Khader, 2011; Steinbach, 2016). The expression of IL-17A, IL-17F, IL-22 and IL-27 is considered essential in a protective vaccine against M. bovis (Waters, 2016). IFNγ was also positively correlated with the expression of IL-17A in M. bovis vaccination studies (Waters, 2016). The expression of IL-27, along with IL-17, is also associated with M. tuberculosis in humans (Basile, 2011; Jurando, 2012; Torrado, 2015). In current literature discussing MAP infection, the mean relative percent of IL- 23R expressing CD4+, CD8+, and TCR1+ cells are increased in MAP-ELISA test positive cows (JD+) relative to ELISA negative cows (JD-) (DeKuiper, 2019). Stimulation of PBMCs from healthy JD-cows with MAP increased the mean relative percent of T cell subtypes that expressed IL-23R (DeKuiper, 2019). Furthermore, mRNAs encoding 65 cytokines that direct T cells toward a proinflammatory Th17-like phenotype are upregulated in subclinical JD+ and even JD- PBMCs exposed to MAP in culture, including IL-6, IL-1, IL-23, and IL-17A (Roussey, 2014; DeKuiper, 2019). This is also true in MAP-infected monocyte-derived macrophages (MDMs) (primarily IL-1β, IL6, and IL-23) (Dudemaine, 2014). The Th17 signature cytokine IL-17A mRNA expression is upregulated in naïve helper T cells (CD4+ CD25-) from healthy JD- cows when these cells are co-cultured with autologous MAP-infected MDMs (Roussey, 2014). These same cytokine mRNAs, particularly IL-17A and IL-6, are upregulated in early stage MAP-infected lesions (Grade 1) (Roussey, 2016), but not in late infection/clinical Johne’s disease lesions suggesting that Th17 response to MAP are subject to the same T cell exhaustion that occurs with Th1-like cells in late clinical JD (Okagawa, 2015). The same pattern of IL-17A expression is found in plasma from cows with JD ELISA scores correlating to progressing stages of infection (DeKuiper, 2019). Both IL-23 and IL-17A are specific to a Th17 response, yet neither protein has been conclusively analyzed in bovine antigen presenting cells and T-cells. The intent of this study was to examine Th17 or Th17-like responses to MAP in CD3+ T cells, determine which T cell subtypes are responsible for producing Th17 associated cytokines, and determine if antigen presenting cells (APCs) are necessary for T cell expression of Th17 cytokines during MAP stimulation. Since αβ T cells are MHC restricted we not only included MDMs as a potential MHC source, but B cells as another possible source of antigen presentation to T cells as well (Ma, 2012). Thus, we hope to provide new insight into potential markers for protective immune responses against JD and begin to understand the cause of chronic inflammation in MAP infected tissues via the IL-23/Th17 inflammatory pathway. 66 Materials and methods Study animals In this study we used mature Johne’s ELISA test negative (JD-) Holstein cows with a similar lactation number (2nd–3rd lactation). MAP environmental contamination had been assessed on each enrolled farm via fecal sampling from the barn floor for MAP (by fecal PCR) as described previously (Frie, 2017). JD- cows were sourced from the Michigan State University Dairy Cattle Research and Teaching Center based on an extremely low herd prevalence of MAP infection combined with MAP-negative environmental testing results. All protocols for animal handling, use, and sampling were reviewed and approved by the Michigan State University Animal Use and Care Committee. Plasma samples Plasma used for IL-23 ELISA were obtained from banked plasma stocks, stored at -80°C for less than 5-years. JD ELISA scores were determined at the time of collection following manufacturer’s protocol and guidelines of commercially available IDEXX ELISA for serum samples from cows. These assays were conducted by AntelBio, Northstar Cooperative Laboratories (Grand Ledge, Michigan). All Associated numbers and coordinating JD status ranges are predetermined by the manufacturer of the assay. The diagnostic IDEXX MAP Antibody ELISA used to analyze plasma samples suggests that a S/P score (referred as OD in this paper) greater than 0.55 is considered a JD positive sample, samples scoring under 0.45 OD are considered negative, and OD values in-between are labeled as suspect. 67 Preparation of T cells, B cells, and MDMs Whole blood (30mL) was collected by coccygeal venipuncture into 10ml Vacutainer tubes containing the anticoagulant acid citrate dextrose (ACD) using 21 gauge (ga) double-sided needles. PBMCs were isolated from whole blood using a standard Percoll gradient centrifugation protocol (Frie et al., 2017). PBMCs were counted using a Beckman Coulter Counter and plated at a density of 4.2x106 in a 96- well flat bottom culture plate to generate a plating density of about 42K monocyte derived macrophages (MDMs; estimating 10% of PBMCs). Monocytes were allowed 6 hours to adhere to plate surfaces. Cells were then rinsed (4x) with warm PBS to remove non-adherent cells followed by a 5-day incubation period to allow differentiation into macrophages. CD3+ T cells were isolated from PBMCs using positive selection of the CD3 surface molecule and magnetic-activated cell sorting (MACS) microbeads (α-ms IgG1; Miltenyi Biotec) as directed by the manufacturer. Briefly, PBMCs were incubated with an α-bovine CD3 mouse IgG1 antibody (Table 1) at 4°C for 30min followed by rinses (3x). The cells were then incubated with α-ms IgG microbeads (#130-048-402; Miltenyi Biotec) at 4°C for 30min followed by rinses (3x), resuspension, and placement in MS Columns (#130-042-201; Miltenyi Biotec) on an OctoMACS magnet. Once in the columns the cells were rinsed (3x) to remove any non-CD3+ cells. MS columns were then removed from the magnet to allow CD3+ cells to rinse from the column. Cells were then counted again using a Beckman Coulter Counter and added to MDMs or cultured alone. B cell and T cell co-cultures were generated as described above, with the addition of an α-bovine sIgM mouse IgG1 antibody (Table 1) to the incubation with the CD3 antibody. Isolated CD4, CD8, and TCR1 T cells (Table 1) were positively selected 68 as described above for CD3. All incubations were completed in RPMI 1640 media with 10% fetal bovine serum, 1% penicillin/streptomycin, and 1% Fungizone at 39°C and 5% CO2 for 18h unless otherwise noted. MAP culture and treatment Isolated MAP cultures (American Type Culture Collection Strain #19698 (Murphy, 2006; Kabara, 2010; Roussey, 2014,) were grown in Middlebrook 7H9 media with 10% oleic acid dextrose catalase (OADC) with 0.2% Mycobactin J supplementation. MAP cultures were maintained at 38°C and kept in log phase until use. MAP was purified from cultures essentially as described previously (Janagama, 2006). Briefly, once cultures reached an OD 600nm of 0.6, MAP was removed from culture media by centrifugation. Pellets were resuspended and rinsed (3x) with warm PBS using repeated cycles through a 25g syringe to reduce MAP cell clumps. MAP cell concentrations were estimated as described previously (Janagama, 2006) where an OD 600nm of 0.3 is set approximately equal to 109 bacteria/ml. Potential contamination of MAP cultures was monitored prior to use by inoculation of brain-heart infusion media with MAP culture aliquots and incubation for 72h at 37°C. As a positive control, separate cultures of cells were treated with 25μg/mL of the general T cell stimulant pokeweed mitogen (PWM). Test cultures received a treatment of intact MAP at a multiplicity of infection (MOI) of 2. CD3+ cultures had an additional test group which received 10μg/mL purified protein derivative of Johne’s (PPDj). After incubation, cultures were pelleted at 600 ×g for 5min at 4°C to aspirate the supernatant. The cells were then rinsed in cold PBS and spun down again. After removing the PBS, cells were frozen at -80°C until downstream processing for RNA or protein. 69 A set of isolated CD3+ T cells were cocultured with MDMs. After treatment, the nonadherent or unbound cells (CD3+) were separated from the MDMs by aspiration and collection of subsequent PBS rinses (3X). EDTA was not used in the PBS as it may have removed the MDMs from the plate surface as well. CD3 and CD14 primers (Table 2) were used to analyze the purity of separated cultures by RT-qPCR. The resulting cultures included one containing well separated CD3+ T (~90%) cells and another with both MDMs and CD3+ T cells which were most likely still bound to adhered MDM cells. Cell surface staining and flow cytometry for culture validation For cell surface staining, isolated cells were washed in PBS and pelleted at 600 ×g for 5min at 4°C and then resuspended in primary antibody cocktail diluted in sterile First Wash Buffer (1X PBS with 10% acid citrate dextrose, 2% heat-inactivated horse serum (Gibco), 0.09% sodium azide) (Table 1). Plates were incubated at 4°C for 30min, washed, and pelleted. Supernatants were aspirated from the wells and cells were resuspended in secondary antibody cocktail diluted in sterile First Wash Buffer (Table 1). Incubation of cells with secondary antibody for 30min at 4°C was followed by another wash. Cells were then pelleted, resuspended and fixed in 1X PBS containing 4% paraformaldehyde for 10min at 4°C. Fixed cells were washed, pelleted, resuspended in Second Wash Buffer (90% 1X PBS, 10% acid citrate dextrose, 0.09% sodium azide), and analyzed by flow cytometry (Accuri C6, Becton Dickinson USA, New Jersey). If immediate analysis was unable to be performed, plates were briefly stored at 4°C. PBMCs were identified on the basis of forward and side scatter properties with log scaling on both the x and y-axis. Primary gating regarded all events outside of the PBMC gate as cellular debris. Secondary positive gating strategies calculated the purity 70 of the populations by the mean relative percent (MRP) of cells with positive surface marker staining using FCS Express 4 analytical software. Briefly, MRP of T cell subtype surface markers within gated cells were analyzed by log side scatter on the y-axis and fluorescence intensity on the x-axis. Unstained (without antibodies) and negative primary control (secondary antibodies only) were also included on each plate to determine final gating. RNA Extraction and RT-qPCR Total RNA was extracted from cultures containing CD3+ using RNeasy Mini Kit (Qiagen) as per the manufacturer’s instruction. Due to the lower culture sizes of the T cell isotypes, Arcturus PicoPure RNA isolation kit (Applied Biosystems) was used to isolate total RNA from these samples as per the manufacturer’s instruction with DNA digestion using RNase free DNase set (Qiagen). RNA was quantified using a NanoDrop. Synthesis of cDNA followed isolation using a High Capacity cDNA Reverse Transcription kit (Applied Biosystems) following manufacturer’s instruction. RT-qPCR was performed in triplicate for all genes. PPIA was used as housekeeping gene for IL- 17A, IL-22, and IL-23 (Table 1). TBP was later used as a housekeeping gene for IL-17F, IL-27, and IFNγ. Taqman reagents were used for all RT-qPCR experiments (Table 2) using an Applied Biosystems 7500 qPCR instrument and 7500 software v2.0.6. ELISA and statistics Plasma samples were diluted 1:2 and analyzed for circulating IL-23 using pre- made plates and solutions (MyBioSource) following the manufacturer’s recommended protocol. All ELISA analytics were performed at 450nm using a Molecular Devices 71 Spectra Max M5 and SoftMax Pro 6.5.1 analytical software. IL-23 plasma cytokine levels and cultures with CD3+ T cells were analyzed using Kruskal-Wallis and Dunn’s multiple comparison tests. Isolated T cell subtype culture medians were compared Wilcoxon Mann Whitney tests. Statistical significance was set at p ≤ 0.05 (α=0.05; chance of making a Type I error). All statistics were computed using Prism GraphPad statistical analysis software. Results CD3+ T cells co-cultured with MDMs To further understand the relationship between MDMs plus CD3+ T cells and IL- 23, IL-17A, and IL-22 in regard to a MAP stimulated Th17 response, 5-day old MDMs were co-cultured with autologous MAC sorted CD3+ T cells and stimulated with MAP antigen for 18-hours. RNA was extracted from each culture, pooled, and analyzed. MAP stimulation (n=8) significantly increased mRNA encoding the Th17 promoting cytokine IL-23 when compared to unstimulated control cultures (n=8; p < 0.001; Fig. 8A). PPDj (n=8). PPDj was able to significantly upregulate IL-23 mRNA when compared to unstimulated control cultures (p < 0.05), but to a lesser extent than live MAP (Fig. 8A). Th17 specific mRNA encoding cytokine IL-17A and the Th17 associated cytokine IL-22 were also significantly upregulated when compared to control cultures (n=7 and n=6 respectively). IL-17A mRNA increased 118-fold in MAP treated cultures relative to control untreated cultures (Fig. 8B) and IL-22 increased 369-fold in MAP treated cultures relative to control untreated cultures (Fig. 8C) (n=7 and n=6; p < 0.001 and 0.01 respectively). PPDj also tended to increase IL-17A and IL-22 mRNA levels (3.1 and 3.6- fold, respectively) in cocultures relative to untreated cells, however these differences 72 were not statistically significant in this test. All analyses were completed using Kruskal- Wallis and Dunn’s multiple comparison test. One extreme outlier was identified for IL-23 within the PPDj group by Grubb’s outlier test and is not included in this analysis. A smaller set of MDM plus CD3+ T cell cocultures were rinsed to remove any unbound CD3+ T cells before RNA isolation. When compared to their respective unstimulated controls, MAP stimulated MDM cultures with remaining bound CD3+ T cells demonstrated a mean 440-fold increase of IL-17A mRNA relative to untreated cells (p < 0.05; n=3; Fig. S1A). Separated MAP stimulated CD3+ T cells contained 29-fold more IL-17A mRNA than untreated CD3+ T cells (p < 0.05; n=3; Fig. S1B). These increased levels of IL-17A were similar to MAP treated CD3+ only cultures as shown in Figure 10A). IL-23 mRNA was significantly upregulated in MAP-stimulated cultures containing MDMs by a mean of 22-fold relative to untreated cultures (p < 0.05; n=3; Fig. S1C). However, the separated MAP stimulated CD3+ T cells exhibited only a median 2.2-fold (p = 0.06; n=4; Fig. S1D) increase in IL-23 mRNA relative to untreated CD3+ T cells. IL- 22 mRNA was increased 104-fold (p < 0.05; n=4; Fig. S1E) in separated MAP- stimulated CD3+ cultures but not in the MDMs with bound CD3+ coculture (p = 0.07; n=4; Fig. S1F). PPDj did not have as profound of an effect on IL-23 mRNA production as it did with IL-17A and IL-22 (p ≤ 0.14; Fig. S1A-F). CD3+ T cells co-cultured with sIgM+ B cells B cells have the capacity to be antigen presenting cells (Ma, 2012). In lieu of this characteristic, autologous sIgM B cells and CD3+ T cells were MAC sorted, co-cultured, and stimulated with MAP antigen for 18-hours as before, control cultures were not treated with MAP. Unlike the cultures with MDMs, mRNA encoding IL-23 was not 73 upregulated in MAP (n=7) or PPDj (n=6) stimulated cultures when compared to their respective untreated control cultures (n=7; Fig. 9A). However, like the MDM co-culture, IL-17A mRNA was increased 127-fold in MAP-stimulated cultures when compared to untreated control cultures (n=7; p < 0.001; Fig. 9B). Similarly, mRNA encoding IL-22 was upregulated 11-fold in MAP-stimulated cultures compared to untreated controls (n=7; p < 0.01; Fig. 9C). The increase in IL-22 mRNA was significant, but numerically much less than observed with MAP stimulated MDM co-cultures. Again, PPDj slightly increased IL-17A (n=6) and IL-22 (n=6) mRNA, but these differences were not significant (5.4 and 1.4-fold respectively). One extreme outlier was identified in all PPDj-stimulated groups by Grubb’s outlier test and these values are not included in this analysis. CD3+ T cell culture Because γδ T cells can be induced to express IL-17A without the aid of traditional APCs (Zeng, 2012) and the possibility that MAP stimulation of αβ T cells via TCR independent mechanisms could also trigger IL-17A production, we next examined CD3+ MAC sorted T cells stimulated with MAP for 18-hours in the absence of any appreciable B cells or monocytes. When cultured without the presence of traditional APCs, CD3+ T cells respond to MAP in a Th17-like manner. Th17 specific mRNA encoding cytokines IL-17A (n=8; Fig. 10A) and IL-22 (n=8; Fig. 10B) are upregulated in MAP-stimulated CD3+ cultures when compared to unstimulated cultures (p < 0.01). IL- 23 mRNA is significantly increased by 2.2-fold in CD3+ T cell cultures treated with MAP (n=8) when compared to control cultures (n=8; p < 0.01; Fig. 10C). When compared to MAP stimulated MDM plus CD3+ T cell and sIgM+ B cell plus CD3+ T cell co-cultures, 74 the average fold-change of IL-17A mRNA in MAP stimulated CD3+ only cultures relative to controls was lower (p = 0.13 and 0.15 respectively; Fig. 11A). Likewise, the mean fold-change for IL-22 mRNA was significantly lower in MAP stimulated CD3+ only cultures compared to that observed in MDM plus CD3+ T cell co-cultures (p < 0.05) but was similar to IL-22 mRNA levels seen in sIgM+ B cell plus CD3+ cell co-cultures (Fig. 11B). The mean IL-23 mRNA fold-change was significantly lower when compared to the 14-fold increase seen with MAP-stimulated MDM plus CD3+ T cell co-cultures (p = 0.05; Fig. 11C). However, this was not significantly different when compared to the 1.3-fold change in sIgM+ B cell and CD3+ T cell co-cultures (Fig. 9A). One extreme outlier was identified in all MAP-stimulated groups and the PPDj group of IL-23 by Grubb’s outlier test and is therefore not included in this analysis αβ T cells are the main producers of IL-17A and IL-22 in the absence of APCs Although γδ T cells are able to produce IL-17A in the absence of APCs (Zeng, 2012), CD4+ and CD8+ T cells appear to be the main producers of IL-17A in response to MAP without the aid of traditional APCs (Fig. 12A and 12B). MAP stimulated CD4+ T cells (n=5) increased expression of mRNA encoding IL-17A with a median 23-fold increase compared to unstimulated CD4+ T cells (n=5; p <0.01; Fig. 12A). MAP stimulated CD8+ T cells (n=4) upregulated IL-17A mRNA with a median 130-fold increase compared to non-stimulated cells (n=4; p < 0.05; Fig. 12B). IL-22 mRNA was also significantly increased (182-fold) in MAP stimulated CD4+ T cells (n=4) compared to non-stimulated cells (n=4; p < 0.05; Fig. 12A), while MAP stimulated CD8+ cells (n=3) tended to express more (median 79-fold) IL-22 mRNA relative to unstimulated CD8+ cells (p = 0.10; Fig. 12B and 12C). Rather surprisingly, γδ T cells (TCR1+) did not 75 significantly increase IL-17A mRNA (n=3; median 1.6-fold) or IL-22 mRNA (n=3; median 6.7-fold; Fig. 12C) in response to MAP stimulation relative to untreated control γδ T cells. IL-23 mRNA was significantly increased in CD4+ T cells (n=6; median 1.6-fold), but not in CD8+ T cells (n=4; median 1.6-fold) or γδ T cells (n=4; median 1.5-fold) compared to respective controls. Grubb’s test identified one extreme outlier in the γδ T cell IL-23 assays that was not included in the results. CD3+ T cells also express enhanced levels of IFNγ mRNA, but not at levels observed for IL-17A or IL-17F mRNAs MAP stimulated CD3+ T cells also upregulate Th17 specific mRNA encoding cytokine IL-17F by a mean 26-fold when compared to unstimulated samples (p = 0.02; n=10; Fig. 13A). MAP stimulated CD3+ T cells also upregulate mRNA encoding IL-27 slightly by a mean 2.24-fold (p = 0.0006; n=10; Fig. 13A) and IFNγ at a median 7.7-fold (p=0.0135; n=10; Fig. 13A). IL-27 and IL-17F share a moderate negative correlation (Spearman r = -0.59) but this is only near significant with the sample size we currently have (n=10; p = 0.081; Fig.13B). Increased IL-23 levels in JD ELISA test positive cows (JD+) To help better understand IL-23’s importance on the prevalence of IL-17A as previously reported in DeKuiper, 2019, we looked into plasma IL-23 levels as it corresponds to infection status. Indeed JD+ cows had significantly more IL-23 in circulation than JD- cows (Unpaired Student T Test; p < 0.05) (Fig. 14). There were no significant differences in circulating IL-23 levels in JD+ cows separated in groups according to increasing JD ELISA test score OD values (data not shown). 76 Discussion Th17 cells and cytokines are likely playing a major role in MAP infection and Johne’s disease (DeKuiper, 2019). While it is not yet clear what precise roles Th17 responses and their associated cytokines might have in the progression of Johne’s disease, data presented herein and elsewhere (Dudemaine, 2014; Roussey, 2014, Roussey, 2016; DeKuiper, 2019, 2019) suggest that Th17 responses are an important feature of immune cell reactions to MAP. IL-17A is a protective cytokine in some inflammatory diseases (Maxwell, 2015; Lee, 2015) and in mycobacterial infections following early infection (Khader, 2011; Palmer, 2016). Chronic inflammation, a distinct feature of MAP infection and Johne’s disease, could be induced by either IL-17A (Leppkes, 2009) or IL-23 (Neurath, 2007, 2019). However, IL-17A may play more of a protective role, as it seems to be upregulated in early infection, but is down regulated in late clinical disease, similar to what is observed with IFN (Roussey, 2014, 2016; DeKuiper, 2019). Evidence suggests these patterns of expression may be due to T cell exhaustion in clinical Johne’s disease (Koets, 2015; Okagawa, 2015). Continuous production of IL-23 from, for example, MAP infected macrophages, could cause epithelial dysregulation and inflammation which are hallmarks of clinical Johne’s disease (Neurath, 2007, 2019). MAP is known to upregulate expression of mRNAs encoding IL-6, IL-23, IL-1β, and TGF-β as early as 1-hour post infection in MAP-infected MDMs (Dudemaine et al. 2014), suggesting that the local cytokine environment near sites of MAP infection would tend to promote development of Th17 cells. MAP infection appears to increase the mean relative percent of T cells expressing IL-23R (DeKuiper, 2019). Indeed, CD3+ T 77 cells dramatically increase mRNA production for Th17 cytokines such as IL-17A, IL-22, and IL-23 in the presence of MAP and MDMs. It is likely that a majority of the IL-23 mRNA expression we observed is from MAP stimulated or infected MDMs and not T cells, given the IL-23 mRNA levels from CD3+ T cell only cultures and cocultures containing CD3+ T cells and B cells are not significant or only slightly increased relative to control untreated cells. We did observe a slight increase in IL-23 mRNA in CD3+ only cultures, likely due to the known properties of Th1 and Th2, cells that can express IL-23 mRNA (Oppmann, 2000). In experimentally and naturally infected JD+ cows, B cell counts are increased, including activated and memory B cells (Stabel, 2011; Frie, 2017). B-cells are also able to produce IL-23 through BCR signaling (Gagro, 2006). However, the increase of IL-17A mRNA in sIgM+ B cell plus CD3+ T cell cocultures did not seem to be coincident with elevated IL-23 mRNA levels, as it was in MDM plus CD3+ T cell cultures. There was little or no increase in IL-23 or IL-22 mRNAs in cocultures containing CD3+ T cells and B cells. Our results do not rule out a potential role for IL-17A produced by either B cells or by γδ T cells during early responses to MAP infection. B cells can produce IL-17A independently of IL-23 (Bermejo, 2013) and this has importance in response to other pathogens (Bermejo, 2013). Similarly, prior research has shown that γδ T cells also respond during early infection with IL-17A production after direct stimulation of their TCR and without restriction of MHC activation (Zeng 2012). This “innate” IL-17A production may be of limited quantity and/or duration (Baldwin, 2014). While γδ T-cells do not require IL-23 for initial production of IL-17A (Lee, 2015), continued IL-17A secretion may require IL-23 (Baldwin, 2014). 78 In the current study, we cultured CD3+ T cells without appreciable traditional APCs present. Indeed, CD3+ T cells were able to increase IL-17A (19-fold), IL-22 (9.4- fold) and IL-23 (2.2-fold) mRNA levels in the absence of traditional APCs. Macrophages are a major supplier of IL-23 (Dudemaine, 2014). As seen in Baldwin et al. (Baldwin, 2014) the production quantity of IL-17A and IL-22 is reliant on the presence of APCs or the quantity of IL-23. The loss of macrophages in our CD3+ cultures resulted in a reduction of IL-23 mRNA and consequently, a reduction IL-17A mRNA and IL-22 mRNA. This is largely noticeable in comparison to the cultures containing MDMs to cultures containing only CD3+ T cells. Additionally, cultures containing B cells were unable to upregulate IL-23 mRNA in comparison to CD3+ T cell cultures and thus unable to increase IL-22 mRNA either. IL-17A was unaffected, however, this may likely be due to B cell’s ability to secrete IL-17A through pathogen stimulation (Bermejo, 2013) rather than IL-23 stimulating IL-23R on the T cell surface. We sought to determine which CD3+T cell subtypes were responsible for producing IL-17A, IL-22, and perhaps IL-23. To this end we sorted T cell subgroups representing CD4+, CD8+, and TCR1+ (γδ) T cells. These sorted T cell subtypes were cultured with MAP, but without added MDM cells and analyzed to determine which subtypes were contributing to production of Th17 cytokines during MAP stimulation. We found that αβ T cells were the main producers IL-17A and IL-22 mRNAs, with CD4+ cells producing only small amounts of IL-23 mRNA. CD8+ cells exhibited a larger increase in IL-17A mRNA (129-fold) than either CD4+ cells (23-fold) or γδ T cells (1.55- fold) relative to unstimulated cells. However, CD4+ cells produced more IL-22 mRNA in response to MAP (182-fold) than either CD8+ (79-fold) or γδ T cells (6.67-fold), relative 79 to unstimulated cells. These results clearly indicate that T cells can directly respond to MAP with production of Th17 associated cytokine mRNA production without significant APCs. Our study is not the first to show CD4+ cells expressing a Th17 phenotype without the direction or activation of APCs (St. Leger, 2018). Other studies have indicated that naïve αβ T cells can produce IL-17A and IL-22 without IL-23 as part of an innate response (St. Leger, 2018). In another study αβ T cells produced IL-23 mRNA with direct stimulation (Oppmann, 2000). While we expected to observe significant IL- 17A mRNA production by isolated γδ T cells, this was not the case. By isolating γδ T cells from other T cell subtypes, we may have hindered their innate potential to respond to MAP in a Th17 manner as seen in other studies (Do, 2012). An alternative explanation is that γδ T cells tend to produce significant amounts of both IFNγ and IL- 27, which are inhibitors of IL-17A production. Cells and tissues infected with M. tuberculosis or M. bovis upregulate IL-17A, IL- 22, and IL-23 expression. These infected tissues also exhibit increased levels of IL-17F as well as the Th17 inhibitors IL-27 and IFNγ (Waters, 2016; Basile, 2011; Jurado, 2012; Torrado, 2015). MAP-stimulated CD3+ T cells seem to follow this same pattern of expression. CD3+ T cells respond to MAP stimulation with increased production of both IL-17A and IFN mRNA (mean 11-fold relative to unstimulated cells). Our data does not allow a distinction between non-classical Th17 cells that also express IFNγ and traditional Th1-like T cells. Contrary to decreased IL-17A levels in the periphery of cows with high JD+ ELISA scores (DeKuiper, 2019), IL-23 is still elevated in plasma of JD+ cows. IL-23 is also a proinflammatory cytokine that can cause inflammation and dysregulation 80 (Maxwell, 2015; Lee, 2017; Paustian, 2017). Overall, JD+ cows have more IL-23 in the periphery than JD- cows. Our results also suggest that macrophages exposed to MAP are likely a significant source of IL-23. It is well known that as JD progresses, more macrophages migrate to the tissue and are infected with MAP (Roussey, 2016). If these macrophages are stimulated to produce IL-23 mRNA and protein, as seen in this study and others (Dudemaine, 2014), the continued expression of IL-23 seen here could be a cause for the advanced inflammation (Lee, 2017; Paustian, 2017) and depletion of T cells observed in tissues of advanced JD (Roussey, 2016; Paustian et al., 2017). MAP induces CD3+ T cells to adopt a Th17-like phenotype in even in the absence of APCs as evidenced by enhanced production of IL-17A, IL-17F, IL-22, and IL-23. However, continued or robust expression of Th17 cytokines IL-17A and IL-22 during early infection are reliant of the presence of IL-23, which could be supplied by MAP infected macrophages. MAP stimulated expression of cytokine mRNA follows similar patterns seen by other mycobacterial infections, including significant increases in IL-22 mRNA and increased expression of mRNA encoding IFNγ and IL-27 (Waters, 2016; Basile, 2011; Jurado, 2012; Torrado, 2015). sIgM+ B cells are also a likely source of innate MAP-stimulated IL-17A, although further exploration is warranted. Additionally, MAP studies focusing on IL-27 should also be considered. M. tuberculosis studies demonstrated an increased protective role by IL-17A, as well as an increased percentage of CCR6+/CD4+ T cells and IL-17+/CD4+ T cells in mice that were IL-27 receptor (R) deficient (Erdmann, 2018). Neutralizing IL-27 also induced lysosomal acidification in macrophages infected with M. tuberculosis and increased mycobacterial killing (Jung, 2014; Sharma, 2014). IL-27 was a source of immune suppression in M. 81 leprae infection (Teles, 2015). It is possible that the inflammation observed in JD may, at least in part, be attributed to Th17 related cytokines, specifically IL-23. Future studies concerning JD should be directed into further exploration of these cytokines. Acknowledgements The authors would like to thank the Michigan State University Dairy Cattle Teaching and Research Center for their continued support. This work was supported by the United States Department of Agriculture and the National Institute of Food and Agriculture (grants 2012-67011-19936, and 2013-68004-20371). Support from the Michigan Alliance for Animal Agriculture and MSU AgBioResearch is also acknowledged. 82 APPENDICES 83 APPENDIX A Table 5: Primary and Secondary antibody list. Antibody CD4 Dilution 1:100 Company WSU CD8 1:100 WSU TCR1 1:100 WSU 1:100 1:100 CD3 sIgM Fluorophore Dilution FITC R-PE PE-Cy7 1:1000 1:200 1:200 WSU WSU Company eBioscience Mouse Invitrogen Mouse eBioscience Mouse m2b-25g4 γ2a m1-14d12 Bovine Specificity Clone IL11A Bovine CACT138A BAQ111A CACT80C GB21A CACTB814 MM1A BIG73A Bovine Bovine Specificity Clone Bovine Isotype IgG2a IgG1 IgM IgG1 IgG2b IgG1 IgG1 IgG1 Target IgG2b IgG2a IgM 84 APPENDIX B Table 6: Taqman primer list. Cytokine Assay ID Entrez Gene ID IL-22 IL-17A IL-23A IL-17F IL-27 IFNg CD3d CD14 PPIA TBP Bt03261459_m1 Bt03210252_m1 Bt04284624_m1 Bt04309062_m1 Bt04298832_m1 Bt03212723_m1 Bt03225300_m1 Bt03212325_g1 Bt03224615_g1 Bt03241948_m1 507778 282863 511022 506030 614927 281237 281053 281048 281418 516578 85 APPENDIX C Figure 8A: Relative abundance of IL-23 mRNA of cultures containing MDMs co- cultured with autologous CD3+ T cells and stimulated with MAP. CD3+ T cells were cultured with 5-day old MDMs and stimulated with PPDj, MAP (MOI of 2), or left unstimulated for 18 hours. Subsequent RNA extraction and qPCR results are shown. MAP stimulated cultures showed a significant upregulation of IL-23 as well as PPDj by Kruskal-Wallis and Dunn’s multiple comparison tests. n=8/group. *p<0.05. **p<0.01. ***p<0.001. 86 APPENDIX D Figure 8B: Relative abundance of IL-17A mRNA of cultures containing MDMs co- cultured with autologous CD3+ T cells and stimulated with MAP. CD3+ T cells were cultured with 5-day old MDMs and stimulated with PPDj, MAP (MOI of 2), or left unstimulated for 18 hours. Subsequent RNA extraction and qPCR results are shown. MAP stimulated cultures showed a significant upregulation of IL-17A while PPDj did not by Kruskal-Wallis and Dunn’s multiple comparison tests. n=7/group. *p<0.05. **p<0.01. ***p<0.001. 87 APPENDIX E Figure 8C: Relative abundance of IL-22 mRNA of cultures containing MDMs co- cultured with autologous CD3+ T cells and stimulated with MAP. CD3+ T cells were cultured with 5-day old MDMs and stimulated with PPDj, MAP (MOI of 2), or left unstimulated for 18 hours. Subsequent RNA extraction and qPCR results are shown. MAP stimulated cultures showed a significant upregulation of IL-22 while PPDj did not by Kruskal-Wallis and Dunn’s multiple comparison tests. n=6/group. *p<0.05. **p<0.01. ***p<0.001. 88 APPENDIX F Figure 9A: Relative abundance of IL-23 mRNA of cultures containing sIgM+ B cells co-cultured with autologous CD3+ T cells and stimulated with MAP. CD3+ T cells were cultured with sIgM+ B cells and stimulated with PPDj, MAP (MOI of 2), or left unstimulated for 18 hours. Subsequent RNA extraction and qPCR results are shown. MAP and PPDj stimulated cultures did not show upregulation of IL-23 by Kruskal-Wallis and Dunn’s multiple comparison tests. n=6/group. *p<0.05. **p<0.01. ***p<0.001. 89 APPENDIX G Figure 9B: Relative abundance of IL-17A mRNA of cultures containing sIgM+ B cells co-cultured with autologous CD3+ T cells and stimulated with MAP. CD3+ T cells were cultured with sIgM+ B cells and stimulated with PPDj, MAP (MOI of 2), or left unstimulated for 18 hours. Subsequent RNA extraction and qPCR results are shown. MAP stimulated cultures showed significant upregulation of IL-17A by Kruskal-Wallis and Dunn’s multiple comparison tests. n=7/group. *p<0.05. **p<0.01. ***p<0.001. 90 APPENDIX H Figure 9C: Relative abundance of IL-22 mRNA of cultures containing sIgM+ B cells co-cultured with autologous CD3+ T cells and stimulated with MAP. CD3+ T cells were cultured with sIgM+ B cells and stimulated with PPDj, MAP (MOI of 2), or left unstimulated for 18 hours. Subsequent RNA extraction and qPCR results are shown. MAP stimulated cultures showed significant upregulation of IL-22 by Kruskal-Wallis and Dunn’s multiple comparison tests. n=7/group. *p<0.05. **p<0.01. ***p<0.001. 91 APPENDIX I Figure 10A: Relative abundance of IL-17A mRNA of cultures containing CD3+ T cells and stimulated with MAP. CD3+ T cells were cultured independently and stimulated with PPDj, MAP (MOI of 2), or left unstimulated for 18 hours. Subsequent RNA extraction and qPCR results are shown. MAP stimulated cultures showed significant upregulation of IL-17A by Kruskal-Wallis and Dunn’s multiple comparison tests. n=8/group. *p<0.05. **p<0.01. ***p<0.001. 92 APPENDIX J Figure 10B: Relative abundance of IL-22 mRNA of cultures containing CD3+ T cells and stimulated with MAP. CD3+ T cells were cultured independently and stimulated with PPDj, MAP (MOI of 2), or left unstimulated for 18 hours. Subsequent RNA extraction and qPCR results are shown. MAP stimulated cultures showed significant upregulation of IL-22 by Kruskal-Wallis and Dunn’s multiple comparison tests. n=8/group. *p<0.05. **p<0.01. ***p<0.001. 93 APPENDIX K Figure 10C: Relative abundance of IL-23 mRNA of cultures containing CD3+ T cells and stimulated with MAP. CD3+ T cells were cultured independently and stimulated with PPDj, MAP (MOI of 2), or left unstimulated for 18 hours. Subsequent RNA extraction and qPCR results are shown. MAP stimulated cultures showed significant upregulation of IL-23 by Kruskal-Wallis and Dunn’s multiple comparison tests. n=8/group. *p<0.05. **p<0.01. ***p<0.001. 94 APPENDIX L Figure 11A: Relative abundance of IL-17A mRNA of CD3+ cells, MDM/CD3+, and sIgM+/CD3+ cultures stimulated with MAP. CD3+ T cell cultures with and without APCs were stimulated with MAP for 18 hours. Subsequent RNA extraction and qPCR results are shown. APC containing cultures demonstrated the most upregulation of IL- 17A by Kruskal-Wallis and Dunn’s multiple comparison tests. n=7-8/group. 95 APPENDIX M Figure 11B: Relative abundance of IL-22 mRNA of CD3+ cells, MDM/CD3+, and sIgM+/CD3+ cultures stimulated with MAP. CD3+ T cell cultures with and without APCs were stimulated with MAP for 18 hours. Subsequent RNA extraction and qPCR results are shown. MDM containing cultures demonstrated the most upregulation of IL- 22 by Kruskal-Wallis and Dunn’s multiple comparison tests. n=7-8/group. *p<0.05. **p<0.01. ***p<0.001. 96 APPENDIX N Figure 11C: Relative abundance of IL-23 mRNA of CD3+ cells, MDM/CD3+, and sIgM+/CD3+ cultures stimulated with MAP. CD3+ T cell cultures with and without APCs were stimulated with MAP for 18 hours. Subsequent RNA extraction and qPCR results are shown. MDM containing cultures demonstrated the most upregulation of IL- 23 by Kruskal-Wallis and Dunn’s multiple comparison tests. n=7-8/group. 97 APPENDIX O Figure 12A: Relative abundance of Th17 mRNA encoding IL-17A, IL-22 and IL-23 from CD4+ cell cultures stimulated with MAP or left unstimulated. Isolated CD4+ T cell cultures were stimulated with MAP or left unstimulated for 18hrs. Subsequent RNA extraction and qPCR results are shown. CD4+ T cells demonstrated significant upregulation of all three cytokines (IL-17A, IL-23, and IL-22) by Wilcoxon Mann Whitney tests. n=4-6/group. *p<0.05. **p<0.01. ***p<0.001. 98 APPENDIX P Figure 12B: Relative abundance of Th17 mRNA encoding IL-17A, IL-22 and IL-23 from CD8+ cell cultures stimulated with MAP or left unstimulated. Isolated CD8+ T cell cultures were stimulated with MAP or left unstimulated for 18hrs. Subsequent RNA extraction and qPCR results are shown. CD8+ T cells demonstrated significant upregulation of IL-17A and near significant for IL-22 by Wilcoxon Mann Whitney tests. However, IL-23 was not. n=3-4/group. *p<0.05. **p<0.01. ***p<0.001. 99 APPENDIX Q Figure 12C: Relative abundance of Th17 mRNA encoding IL-17A, IL-22 and IL-23 from TCR1+ (γδ) cell cultures stimulated with MAP or left unstimulated. Isolated CD8+ T cell cultures were stimulated with MAP or left unstimulated for 18hrs. Subsequent RNA extraction and qPCR results are shown. γδ T cells did not demonstrat significant upregulation of any of the cytokines by Wilcoxon Mann Whitney tests. One extreme outlier was found by Grubb’s test in the “IL-23” group and is not included in this test. n=3-4/group. *p<0.05. **p<0.01. ***p<0.001. 100 APPENDIX R Figure 13A: Relative abundance of mRNA encoding IL-17F (Th17), IL-27 (anti- Th17) and IFNγ from CD3+ cell cultures stimulated with MAP or left unstimulated. Isolated CD3+ T cell cultures were stimulated with MAP or left unstimulated for 18hrs. Subsequent RNA extraction and qPCR results are shown. CD3+ T cells demonstrated significant upregulation of mRNA for Th17 cytokine IL-17F, Th1 cytokine IFNγ, and Th17 inhibitory cytokine IL-27 by Wilcoxon Mann Whitney tests. n=10/group. *p<0.05. **p<0.01. ***p<0.001. 101 APPENDIX S Figure 13B: Correlation between quantities of mRNA encoding IL-17F and IL-27 from CD3+ cell cultures stimulated with MAP or left unstimulated. Isolated CD3+ T cell cultures were stimulated with MAP or left unstimulated for 18hrs. Subsequent RNA extraction and qPCR results are shown. IL-17F may share a moderate negative correlation with IL-27 by Spearman correlation test. n=10. r = -0.59. p = 0.081. 102 APPENDIX T Figure 14: Plasma IL-23 levels of cows based on IDEXX Johne’s ELISA status. IL- 23 concentrations (pg/mL) circulating in the plasma from the periphery of by ELISA. Overall JD- cows (n=46). Overall JD+ cows (n=30). * p<0.05. Error bars = SEM. Cow n is based on available stocked plasma samples. Unpaired T test was used between overall JD- and JD+ groups. 103 APPENDIX U Figure S1A: Relative abundance of IL-17A mRNA of MDMs and bound CD3+ T cells after co-culture, stimulation with MAP, and unbound cells were rinsed away. CD3+ T cells were cultured with 5-day old MDMs and stimulated with PPDj, MAP (MOI of 2), or left unstimulated for 18 hours. Unbound CD3+ T cells were rinsed and analyzed separately. Subsequent RNA extraction and qPCR results are shown. MAP stimulated cultures showed a significant upregulation of IL-17A by Kruskal-Wallis and Dunn’s multiple comparison tests. n=3/group. *p<0.05. **p<0.01. ***p<0.001. 104 APPENDIX V Figure S1B: Relative abundance of IL-17A mRNA of unbound CD3+ T cells after co-culture, stimulation with MAP. CD3+ T cells were cultured with 5-day old MDMs and stimulated with PPDj, MAP (MOI of 2), or left unstimulated for 18 hours. Unbound CD3+ T cells were separated by rinse. Subsequent RNA extraction and qPCR results are shown. MAP stimulated cultures showed a significant upregulation of IL-17A by Kruskal-Wallis and Dunn’s multiple comparison tests. n=4/group. *p<0.05. **p<0.01. ***p<0.001. 105 APPENDIX W Figure S1C: Relative abundance of IL-23 mRNA of MDMs and bound CD3+ T cells after co-culture, stimulation with MAP, and unbound cells were rinsed away. CD3+ T cells were cultured with 5-day old MDMs and stimulated with PPDj, MAP (MOI of 2), or left unstimulated for 18 hours. Unbound CD3+ T cells were rinsed and analyzed separately. Subsequent RNA extraction and qPCR results are shown. MAP stimulated cultures showed a significant upregulation of IL-23 by Kruskal-Wallis and Dunn’s multiple comparison tests. n=3/group. *p<0.05. **p<0.01. ***p<0.001. 106 APPENDIX X Figure S1D: Relative abundance of IL-23 mRNA of unbound CD3+ T cells after co- culture, stimulation with MAP. CD3+ T cells were cultured with 5-day old MDMs and stimulated with PPDj, MAP (MOI of 2), or left unstimulated for 18 hours. Unbound CD3+ T cells were separated by rinse. Subsequent RNA extraction and qPCR results are shown. MAP stimulated cultures showed an upregulation of IL-23 by Kruskal-Wallis and Dunn’s multiple comparison tests. n=4/group. *p<0.05. **p<0.01. ***p<0.001. 107 APPENDIX Y Figure S1E: Relative abundance of IL-22 mRNA of MDMs and bound CD3+ T cells after co-culture, stimulation with MAP, and unbound cells were rinsed away. CD3+ T cells were cultured with 5-day old MDMs and stimulated with PPDj, MAP (MOI of 2), or left unstimulated for 18 hours. Unbound CD3+ T cells were rinsed and analyzed separately. Subsequent RNA extraction and qPCR results are shown. MAP stimulated cultures showed an upregulation of IL-22 by Kruskal-Wallis and Dunn’s multiple comparison tests. n=3/group. *p<0.05. **p<0.01. ***p<0.001. 108 APPENDIX Z Figure S1F: Relative abundance of IL-22 mRNA of unbound CD3+ T cells after co- culture, stimulation with MAP. CD3+ T cells were cultured with 5-day old MDMs and stimulated with PPDj, MAP (MOI of 2), or left unstimulated for 18 hours. Unbound CD3+ T cells were separated by rinse. Subsequent RNA extraction and qPCR results are shown. MAP stimulated cultures showed a significant upregulation of IL-22 by Kruskal- Wallis and Dunn’s multiple comparison tests. n=4/group. *p<0.05. **p<0.01. ***p<0.001. 109 REFERENCES 110 REFERENCES Ahern, P. P., Schiering, C., Buonocore, S., McGeachy, M. J., Cua, D. J., Maloy, K. J., & Powrie, F. (2010). Interleukin-23 Drives Intestinal Inflammation through Direct Activity on T Cells. Immunity, 33(2), 279–288. https://doi.org/10.1016/j.immuni.2010.08.010 APHIS. 2007. 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Infection and Immunity, 56(7), 1692–1697. 117 CHAPTER 3 FUTURE DIRECTIONS OF JOHNE’S DISEASE RESEARCH AND THE CROHN’S DISEASE CONNECTION Introduction A national loss of $200 million to $1.5 billion per year to the dairy industry (Garcia, 2015) is attributed to a national herd infection rate of up to 91% by Mycobacterium avium sp. paratuberculosis (MAP) (Lombard, 2013), the causative agent of Johne’s disease (JD). JD affects the ileum of cattle by inducing inflammation and disruption of the intestinal lining, reducing nutrient absorption, thus negatively affecting animal health and productivity (review, Coussens, 2004). Furthermore, the significant variation in time between infection and diagnosis results in lost productivity and a progressive decline in animal health. Continued study of the innate immune response to MAP may improve current, severely lacking, early diagnostic testing for JD. Increased levels of Th17 and Th1 secreted cytokines may be of use for new early diagnostic markers as a tool for control (DeKuiper, 2019, submitted). MAP is a zoonotic bacterium and is thought to contribute to decreased human health by causing or exasperating some cases of Crohn’s disease (CD) (Kuenstner, 2017). Although MAP’s involvement in CD is still somewhat controversial, MAP may play a pathogenic role in human health by contributing to CD (Hermon, 2009; Naser, 2014). Approximately 1 in 500 people are affected by inflammation and discomfort caused by CD (Kappelman, 2007). Research in support of MAP as a causative agent for CD is largely based on MAP-bacterial culture or MAP-PCR positivity from CD patient tissue (Bull, 2003; Naser, 2014). Though 118 both supportive (mostly molecular) and contradictory evidence (mostly correlative) data for a role of MAP in CD exist (Table 5) (Rosenfeld, 2010), the consensus of the MAP community is that the bulk of research supports MAP as one potential causative agent in CD (Kuenstner, 2017). It is undeniable that some CD patients treated with anti-MAP antibodies had improved outcomes (as high as 84%) compared to treatment with anti- TNF antibodies (ranges below 39%) (Prantera, 1989; Borody, 2007; Chamberlin, 2011). MAP has been shown to survive some methods of pasteurization and has been found in calf milk replacer (Grant, 2017), as well as meat and other products, such as milk, infant formula, yoghurt, and cheese (Millar, 1996; Hastings, 2001; Grant, 2002; Donaghy, 2010; Van Brandt, 2011; Paolicchi, 2012; Galiero,2015; Savi, 2015; Botsaris, 2016; Acharya, 2017 Lorencova, 2019) – suggesting that human exposure to MAP via consumption of bovine products is possible. Consequently, reducing and controlling MAP in cows will reduce human exposure, MAP-associated Crohn’s disease, and potentially other irritable bowel diseases. The previous chapters have outlined alternative mechanisms MAP uses to induce inflammation and may provide insight into new targets for diagnosis of MAP infection in cattle, correlates of protection for new MAP vaccines, and possibly treatment of MAP-induced Crohn’s disease. Furthermore, inflammation of the ileum, diarrhea, and malabsorption of nutrients, as well as the cytokine milieu during active or progressed stages of either disease are all phenotypes that both JD and CD share, thus making MAP infection and cows (JD) an excellent model for human disease (CD). 119 Johne’s disease vs. Crohn’s disease Regardless of MAP’s direct involvement in CD, CD patients express a very similar profile of cytokines to that which we have seen in our studies with MAP infection (Siakavellas, 2012; DeKuiper, submitted). Our work assists in making a stronger case for using JD in cattle as a model system in which to study the genetics, immunopathology, and new treatments for CD. Traditionally, CD patients use anti-TNF therapies, and these provide relief from symptoms in some cases, but do not often clear the disease. Patients who respond well to anti-TNF therapies have significantly more Tumor necrosis factor receptor 2 (TNFR2) than non-responders (Schmitt, 2017). Patients who do not respond to anti-TNF therapies have an increase of α4β7 (a gut homing integrin) positive T cells which are both TNFR2 and IL-23R positive, positive for both Th1 markers (T-Bet/IFNγ) and Th17 markers (RORγt/IL-17A), and expand in the presence of IL-23 and TNF blockers (Schmitt, 2017). The α4β7 integrin is expressed by Th17 and Th17-like cells (Duhen, 2014; Rout, 2016; Table #). Similar to advanced stages of JD, CD patients have an increase in CD14+ macrophages within affected intestinal tissues (Kamada, 2008; Roussey, 2016; Schmitt, 2017). As we have demonstrated here, these cells may produce substantial IL-23 and therefore activate IL- 23R+ T cells, thus perpetuating inflammation even in the face of T cell exhaustion (Schmitt, 2017). IL-23 rather than IL-17A is likely the main candidate for inflammation observed in the later stages of JD (Roussey, 2016), as it has shown to be a contributor for induced colitis in mice (Uhlig, 2006). IL-17A appears to have a protective role in some mycobacterial infections in human and cattle (Khader, 2011; Steinbach, 2016), as well as in ulcerative colitis (UC), inflammatory bowel disease (IBD) and CD in humans 120 (Wang, 2018; Hohenberger, 2018; Smith, 2019). Some control patients receiving IL-17A blockers were soon after diagnosed with Crohn’s-like disease and put on IL-23 blockers (Smith, 2019), while another study showed swift inception of fulminant colitis upon receiving an IL-17A blocker (Wang. 2018). Sera IL-17A is also observed to be much lower in advanced JD (DeKuiper, 2019), as well as in CD patients when compared to healthy controls or patients in CD remission (Sahin, 2014). Similarly, mouse models of UC and CD demonstrated that knocking of IL-17A or IL-17RA perpetuated inflammation (Maxwell, 2015; Hohenberger, 2018). When inhibiting IL-23, inflammation was reduced in mouse UC/CD models (Maxwell, 2015). In non-TNF-responding human CD patients, IL-23 blockade promoted maintenance or remission (Sandborn, 2012). However, treatment of IL-23 may need to be directed at mitigating IL-23 and not abolishing it in MAP infection, as IL-23 is essential for a protective Th17 recall response in M. tuberculosis and long-term containment of bacteria (Monin, 2015; Khader, 2011). Protective amounts of IL-17A may also be reliant on IL-23 levels (Khader, 2011; Eken, 2014; DeKuiper, submitted). Early or subclinical infections with MAP can be difficult to diagnose and often do not display overt disease or bacterial shedding (Magombedze, 2017). Similarly, CD and UC patients may have periods of inactivity or apparent remission (Fujino, 2002). During times of inactivity, IL-17A+ cells are less abundant than they are during periods of active disease, including both CD3+ IL-17A+ T cells and CD68+ IL17A+ monocytes/macrophages (Fujino, 2002). It may be possible that macrophages are contributing to IL-17A mRNA levels observed in the CD3+ plus MDM cultures described in Chapter 2. However, contribution of IL-17A mRNA by MAP-infected macrophages is 121 unlikely due to the lack of IL-17A mRNA in clinical JD tissues and sera (Roussey, 2016; DeKuiper, 2019). These tissues contain numerous macrophages both with and without internalized acid-fast bacilli (Roussey, 2016). Similar to IL-17A and IL-23, another relatively unexplored Th17 cytokine in JD is IL-22. In the absence of IL-23R, mice are protected from innate immune cell directed colitis, however, IL-22 can stimulate an innate-induced colitis (Eken, 2014). MAP stimulates a greater relative increase of early IL-22 mRNA from CD3+ T cells than IL- 17A mRNA (DeKuiper, submitted). IL-22 is also increased in sites of intestinal inflammation of the ileum from CD an UC patients (Brand, 2006; Bassolas, 2018). Sera from these patients contain more IL-22 than sera from healthy controls (Schmechel, 2008), despite observed decreases in IL-17A and IL-23R in CD and UC biopsies (Bassolas, 2018). IL-22 has demonstrated protective properties in IBD (Zenewicz, 2008; Pickert, 2009, Aden, 2016; Mizoguchi, 2017). However, like IL-17A and IL-23, IL-22 can exacerbate inflammation (Nagalakshmi, 2004; Beatty, 2007; Durant, 2010; Eken, 2014), potentially indirectly through the stimulation of Claudin-2 (Wang, 2017) as described later. Additionally, IL-22 level differences in sera of CD patients are correlated with IL- 23R genotypes and susceptibility to CD. IL-22’s relationship with IL-23R is further demonstrated in IL-23R-/- mice, which are observed to have a significant decrease in sera IL-22 (Maxwell, 2015). IL-22’s role in MAP infection is still unclear, but expression of this cytokine may depend both upon expression of IL-23 from macrophages and the activity of IL-23R positive T cells. Our work presented in Chapter 2 suggests that CD3+ T cell only cultures express far less IL-22 mRNA than CD3+ T cells cocultured with MDM cells. 122 Future directions To address remaining gaps in knowledge concerning sources of JD specific inflammation, targets for early diagnostics, and furthering the inflammatory connection between JD and human IBDs, future MAP studies may also include epithelial response signals (i.e. Claudin-2). Claudin-2 is a pore-forming tight junction protein (Furuse, 1999; Colegio, 2002) and is upregulated in intestinal epithelium with increased presence of IL- 22 (Wang, 2017), which is known to have increased mRNA levels in CD/UC tissues and MAP stimulated T cells (Brand, 2006; Bassolas, 2018; DeKuiper, submitted). In active CD, claudin-2 expression increases while non-pore-forming claudin-5 and -8 expression is decreased (Zeissig, 2007). Similarly, claudin-2 expression is also correlated to disease severity in UC and is a biomarker for disease pathogenesis (Randall, 2016). Claudin-2 is not normally expressed in the gastrointestinal mucosa (Nichols, 2004). However, claudin-2 is expressed in follicle associated epithelium of Peyer’s patches (Tamagawa, 2003). One may hypothesize that increasing IL-22, and potentially claudin- 2, may be one mechanism that MAP utilizes to provide escape after replication, as both are shown to an important part in innate clearance of enteric pathogens (Tsai, 2017). Several epithelial cell types also do not normally express claudin-2 in-vitro, including commercial lines such as MDCKs (Singh, 2004; Ohkubo, 2004; Turksen, 2004). MDCKs as well as MDBK could be used as part of potential in-vitro models defining JD related inflammation, as well as using ileal loop models as an in-vivo model (Momotani, 1988; Adams, 2010; Khare, 2016). Prior in-vivo studies with MAP burdened ileal tissue lesions (Grades 0 to 1) demonstrate that proinflammatory IL-6 and IL-17A mRNA expression initially increase 123 then decrease as MAP infection and lesion score increase (Grade 2-4) (Roussey, 2016). Similarly, when challenging calves with M. bovis, mRNA encoding IL-17A is upregulated in lung granulomas with low MAP burden (early stage; Grade 1) when compared to late-stage granulomas with high bacterial burden, (Grade 3 and 4) (Palmer, 2016; Roussey, 2016). Loss of IL-17A production could be attributed to T cell exhaustion, which has also been proposed to explain the apparent reduction in classical Th1 responses in late stage Johne’s disease (Paustian, 2017; Okagawa, 2015; Roussey, 2016). Inflammation, such as that seen in higher MAP infected tissues, could occur with continual IL-23 expression (Paustian, 2017) and this concept involving IL-23 is yet to be explored in MAP infected tissues. Further support in IL-23’s pathogenetic involvement in JD can be demonstrated by similarities found in the intestinal CD14+ macrophages and lamina propria mononuclear cells (LPMCs) from CD patients compared to MAP-infected MDMs and MAP-stimulated PBMCs/CD3+ T cells. In the presence the IFNγ secreted by LPMCs, IL- 23 producing macrophages can become IL-23 hyperproducing (Kamada, 2008). Similarly, IL-23 mRNA increases in the presence of MAP (Dudemaine, 2014), while PBMCs and CD3+ T cells stimulated with MAP also upregulate IFNγ mRNA (Roussey, 2014; DeKuiper, submitted), thus creating a similar environment for IL-23 hyperproduction. Both CD and JD have an increase of macrophages in the inflamed tissue (Kamada, 2008; Roussey, 2016; Schmitt, 2017), and perhaps IL-23. This could lead to IL-23 induced T cell exhaustion and inflammation as mentioned earlier. Current literature has much to offer JD therapeutic potentials through human inflammatory bowel diseases, such as IL-23, although may be too expensive for use on commercial 124 farms. JD and CD have very similar cytokine profiles as well as disease attributes, suggesting JD as a potential animal disease model for human inflammatory bowel diseases, such as CD. The next aim should be designed to test the in-vitro results from chapters 1 and 2 of this Thesis against in-vivo natural representation of inflamed MAP-infected ileal tissues and of the surrounding lymph nodes. Observations should confirm or refute in- vitro representation for Johne’s disease and MAP infection. This can be approached with analysis of frozen tissue stocks already available from naturally infected JD+ and JD- cows. These samples of lesions of the ileum have been previously graded as to increasing levels of MAP burden and inflammation (Grades 0 to 4). In this scheme, grade 0 lesions show no sign of being infected and no inflammation while grade 4 lesions have a high MAP burden and are highly inflamed (Roussey, 2016). Lesions analyzed by immunohistochemistry and immunofluorescence staining for Th17-related cytokines as well as their receptors on epithelial and tissue-present T-cells would give insight to which cytokines are contributing the most and where. However, since cytokines are secreted, Western blotting could be used instead of microscopy-based detection. In support of these studies, prior work on CD, UC, and IBD has shown the expression and importance of IL-21R, IL-23R, IL-17R as well as Th17 inhibitor IL-27 and claudin-2 as related to disease and inflammation (Zeissig, 2007; Schmechel, 2008; Wang, 2014; Randall, 2016; Nemeth, 2017; Schmitt, 2017; Hohenberger, 2018; Holm, 2018; Wang, 2018). Where antibody reagents are not available, mRNA analysis of frozen tissues by RT-qPCR could provide additional insight. Lesions with low MAP- burden are indicative of early disease and tend to show the greatest in mRNA from T- 125 cells in the ileum with declining amounts as lesions become more severe. In addition to banked tissues, ileal loop models could also be used. While late JD show a decrease of γδ T cells in the ileum compared to early JD, as well as cell exhaustion in the ileum of later stages of JD (Koets, 2015; Okagawa, 2015; Roussey, 2016), γδ T-cells remain abundant in the mesenteric lymph nodes (Roussey, 2016). Although γδ T cells may not have shown much direct-stimulated activity in earlier studies with MAP (DeKuiper, submitted), this may have been a factor of the experimental design to isolate γδ T cells from other T cell types (Do, 2012) as mentioned in Chapter 2. The increase in mean relative percent (MRP) of JD- γδ T cells expressing IL-23R in PBMCs when treated with MAP is greater than that of their untreated controls, as well as a greater in MRP JD+ cows than JD- (DeKuiper, 2019), suggesting γδ T cells may still play a role in JD and IL-23. However, much is still unknown of their involvement during MAP infection and should still be included in future studies. Expected results Research suggests that potential Th17-like cells may target the gut/ileum at a higher rate than other T-cell effector subtypes based on their expressed homing markers (Table 3). I would expect IL-17A will be upregulated in lesion scores with low MAP burden (Grades 1), while downregulated in high MAP burdened lesion scores (Grades 3 and 4). I would expect to find an increase in IL-23 in high MAP burdened lesions with a concurrent decrease of IL-17A. I expect an increase of IL-23R and IL-22R mRNA expression in tissues, as well as an increase in claudin-2, confirming a response to Th17 cytokines. Using ileal loop models and current anti-IL-23, CD therapeutics, or 126 similar anti-IL-23 approaches, will reduce inflammation in MAP-infected tissues and potentially help clear bacteria with the proper time of application and dosage. Potential problems and diagnostic opportunities Some bovine antibodies may lack in commercial availability, so if immuno- staining protocols are insufficient, RNAseq may be a better way of analysis, although more costly. Stocked samples have extracted, and quantified RNA and cDNA stocks stored at -80°C to use if any potential problems arise with protein isolation. Samples are readily availability to obtain freshly extracted RNA samples from frozen graded lesions similar as noted in prior studies (Roussey, 2016), as well as obtaining fresh samples through collaborations, which may be used to perform or expand data sets. The earlier chapters outlined in this dissertation have generated enough interest for good collaboration and our work is in line with interests in other groups (Stabel, 2019). Current therapies for IL-23 would be far too expensive and thus not practical for use in any type of therapy for JD. However, understanding the pathogenesis associated with Johne’s disease may have implications for improving early diagnostic markers and sensitivity (Table 1) due to intermittent MAP shedding activity (Nielsen, 2006, 2006; Magombedze, 2017). Current assessment tools (tests) for diagnosis of MAP infection are inadequate. Although highly specific to MAP antigen, the sensitivity of MAP diagnostic tests comes with a cost in a high rate for false negatives (Table 1). MAP cultures from fecal samples can be falsely negative due to lack of bacterial shedding at the time of collection and are further handicapped by the months of time needed to generate results. Serum testing is quick but less sensitive than fecal culture due to the potential lack of MAP activity at the 127 time of sampling. My research may not only provide alternative treatment options but a potential for a more sensitive tool in early diagnosis as well. In summary, in-vivo and in- vitro models described in this chapter will help better understand the disease, subsequently increasing animal and human welfare and decreasing the direct cost of disease worldwide. 128 APPENDICES 129 APPENDIX A Table 7: Average Sensitivities and Specificities between MAP diagnostic tests. Adopted from Collins, 2006. Test (by cow) * Bacterial Culture of Fecal samples PCR assay of Fecal samples ELISA on Serum or Milk Evaluation of Biopsy Specimens Necropsy Sensitivity of Specificity for test (%) 60±5 MAP (%) 99.9 ± 0.1 30±5 30±5 90.5±5 100 99.5 ± 0.5 99.0 ± 1.0 100 100 *Test sensitivity and specificity are an average from literature Adopted from Collins, 2006 130 APPENDIX B Table 8: Homing signals to the gut indicating Th17 homing to the Ileum. Homing Th1 CCR6 Th17 (Duhen, 2014) Dual Tregs (Duhen, 2014) (Lim, 2008) CXCR3 (Duhen, (Duhen, 2014) 2014) (Lim, 2008) (Rout, 2016) (Lim, 2008) (Rout, 2016) (Rout, 2016) (Duhen, 2014) (Rout, 2016) (Zeng, 2012) CCR2/4/5/7 CD69 CD161 Integrin α4β7 (Duhen, 2014) Vγ7 expressed by γδ T-cells Migration Peyers Patch(Lugering, 2005), Skin and Mucosal sites(Lydova, 2015) Mucosal Tissue(Rout, 2016) Intestine(Kleinschek, 2009) Gut homing(Rout, 2016) lymphocytes of small intestine(Zeng, 2012) 131 APPENDIX C Table 9: The argument, MAP and Crohn's Disease. Adopted from Rosenfeld, 2010. The evidence supporting MAP as a cause of Crohn’s Disease 1. The similarity between Johne’s Disease and CD (McKenna, 2006) 2. MAP has been found in milk and water supplies and is capable of surviving commercial pasteurization methods (McKenna, 2006; Grant, 2002; Millar, 1996) 3. MAP has been detected in the tissues and blood of CD patients with a greater frequency than those without CD (Naser, 2004; Sanderson, 1992; Hulten, 2001; Bently, 2008) 4. Positive antibodies to MAP antigens in the blood of CD patients compared with controls (Naser, 2000; Olsen, 2001) 5. Detection of MAP in human breast milk from patients with CD (Naser, 2000) 6. The gene NOD2/CARD15 has previously been shown to be a susceptibility gene for the development of CD (Ogura, 2001; Goyette, 2007). NOD2/CARD15 mutations result in a defective innate response to MAP The evidence not supporting MAP as a cause of Crohn’s Disease 1. Humans exposed to animals infected with MAP do not show a higher prevalence of CD (Jones, 2006) 2. MAP has been isolated from individuals without CD, albeit in smaller numbers (Bull, 2003; Sechi, 2005). This would suggest that MAP is at least, not a sufficient cause for CD and that other factors are necessary to induce disease organisms causes CD (Abubakar, 2007) 3. There is a lack of evidence that consumption of food containing MAP organisms causes CD (Abubakar, 2007) 4. There is no evidence to support the increased transmission of MAP and CD to offspring despite the report of MAP cultured from breast milk of MAP-infected mothers with CD (Sartor, 2005) 5. CD responds to immunosuppressive therapy, such as corticosteroids, which has been associated with decreased levels of MAP DNA (Autschbach, 2005). 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