The Role of Dendritic Cells in Promoting Adaptive ... · It was nice of them to recruit me, an...

The Role of Dendritic Cells in Promoting Adaptive Antiviral Immune Responses at the Intestinal Mucosa

by

Tian Sun

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy

Department of Immunology University of Toronto

© Copyright by Tian Sun, 2017

ii

The Role of Dendritic Cells in Promoting Adaptive Antiviral Immune

Responses at the Intestinal Mucosa

Tian Sun

Doctor of Philosophy

Department of Immunology

University of Toronto

2017

Abstract

The immune system of the gastrointestinal tract must be tightly regulated to limit inflammatory

responses towards innocuous food and commensal antigens while allowing for rapid

development of effector responses against invading pathogens. Highly specialized antigen

presenting cells, such as dendritic cells (DCs), play an essential role in balancing the regulatory

and inflammatory responses in the gut. Although multiple DC subsets have been described in

both lymphoid and non-lymphoid tissues, we know little about which DC subset(s) provoke

antiviral responses within the gut, as well as what DC-intrinsic pathways are needed for optimal

CD8+ T cell responses against viral infection at the mucosa. Herein, using an infection model

with rotavirus (RV), a double-stranded RNA virus with a small intestinal tropism, I demonstrated

that BATF3-dependent DCs are required for generating optimal RV-specific CD8+ T-cell

responses in adult mice. However, a significant amount of RV-specific CD8+ T cells are still

detectable in the small intestinal lamina propria (SILP) of Batf3-/- adult mice, suggesting the

existence of compensatory cross-presentation machinery in the absence of BATF3-dependent

DCs. Interestingly, BATF3-dependent DCs are absolutely needed for RV-specific CD8+ T-cell

responses in neonatal mice. Furthermore, a decreased Th1 response and an increased Th17 in

both adult and neonatal Batf3-/- mice is observed upon RV infection, although local and systemic

iii

RV-specific immunoglobulin A (IgA) production kinetics and titers are unimpaired in these

mice. Our lab previously showed that DC-intrinsic lymphotoxin beta receptor (LTR) signaling

pathway is important for optimal CD8+ T-cell expansion in responses to foreign protein antigens

in the periphery. Moreover, Lta-/- mice display prolonged RV antigen shedding and delayed anti-

RV IgA response. To determine whether LTR signaling pathway in DC can impact on anti-RV

adaptive immune responses, I challenged Ltbr-/- WT bone marrow chimeric mice with RV. I

found that LTR signaling in the radio-sensitive is dispensable for RV clearance. In Ltbr-/-WT

chimeric mice, increased interferon gamma-secreting RV-specific CD8+ T cells and polyclonal

Th1 cells are present, accompanied by a decrease in interleukin 17-producing polyclonal CD4+ T

cells in the SILP. In spite of this altered cytokine profile, the local and systemic RV-specific IgA

response is unperturbed. Taken together, these studies contribute towards our understanding of

DCs in regulating antiviral adaptive immune responses in the intestine.

iv

Acknowledgments

I would like to start by acknowledging my co-supervisors Drs. Jennifer Gommerman and Dana

Philpott. It was nice of them to recruit me, an international student with almost zero background

on immunology, into their research teams. Jen is always supportive and patient with my

elementary-level written English. I learned a lot about how to ask questions, how to use precise

language and how to address scientific questions from her. She was also very generous to offer

me an opportunity to go to Kenya for a human study. I’m grateful that I grasped this chance, not

only to gain some experience working with human gut biopsies, but also to explore the

mysterious African continent and visit Masai Mara. Dana is a lovely lady who always

encourages me to take my time and do what I like to do. Although I spent less time in her

laboratory, she never complains about that. I still remember the first conference I attended in

Quebec City, we sat next to each other on the plane (what a coincidence!) and the conversation

we had calmed me down. Both Jen and Dana set a good model of how to be female scientists

while balancing work and family. I learned a lot from them and I’m still learning.

I gratefully acknowledge the contribution of my committee members Drs. Thierry Mallevaey and

Ken Croitoru. Thank you for always bringing helpful ideas to the table, but also being friendly

and supportive. Your feedback and guidance helped me shape my mind and drove me to look

further. I would like to thank my previous supervisor Dr. Erguang Li in Nanjing University, who

provided me with helpful suggestions and recommended me to land here in Toronto.

I gratefully acknowledge the DCM animal facility staff, in particular Stacy Nichols for her

dedication to animal care and the little chats. I thank the MSB flow cytometry staff Dionne

White for her advice and expertise and being patient with my countless clogging issues:

acquiring “glue-like” gut cells isn’t always smooth and easy, but I’m grateful to have her help

me unclog the machine. I thank the Immunology Office personnel, Sherry Kuhn, William Hsia,

Lynne Omoto, graduate assistant Kate Sedor and undergraduate assistant Anna Frey for

providing a friendly working atmosphere.

I found support and friendship in past and present Gommerman lab members: Dr. Dennis Ng, Dr.

Georgina Galicia-Rosas, Dr. Bryant Boulianne, “Dr. Natalia Pikor, Dr. Elisa Porfilio, Dr.

Blandine Maitre, Jennifer Yam” (The Feb Girls!), Leslie Leung, Lesley Ward, Dr. Gary Chao,

Dr. Olga Rojas, Dr. Conglei Li, Dennis Lee, Albert Nguyen, Eric Cao, Evelyn Lam and Dr.

v

Valeria Ramaglia- I thank you all for your support and friendship. In particular, I would like to

thank Olga for her guidance of how to do gut preps, how to operate the flow cytometer and how

to perform ELISA, which is extremely helpful. I couldn’t have done my PhD without her.

I found support and friendship in past and present Philpott lab members, in particular Dr. Kaoru

Geddes, Dr. Matthew Sorbara, Dr. Susan Robertson, Dr. Dave Prescott, Dr. Juliana Rocha,

Charles Maisonneuve, Elisabeth Foerster, Nichole Escalante, Ashleigh Goethel and some

smiling faces from Girardin lab. Thank you for the support, kind words, science brainstorming

and friendship.

I thank all the other colleagues from Department of Immunology on 7th floor of MSB, for sharing

the equipment, reagents and protocols. In particular, I would like to mention Angela Zhou from

Watts lab. We are in the same year and both of us can be found in the mouse house, or in the

flow lab, or on the way to the mouse house/flow lab. I enjoyed talking with her about global

political news, the culture difference and negative results.

I would like to thank my friends who made my life outside of the lab colorful. It was Joshua

Moreau and Eric Gracey who put up a running club when I first joined the department. Since

then, I started running in the city to see the neighborhoods and various trekking trails. Then came

Liu Zhang, Hang Zhou, and other friends, who trained with me and together we ran some 10k

and half-marathons for the last few years. Finally came Dr. Fei Luo, my old roommate, who is an

incredible running buddy and we trained and ran the full marathon together! It’s been a long

journey so far, but I enjoyed the experience and scenery along the way. So many other friends

have helped me along this journey, I want to thank you all!

Lastly, I’m grateful to my parents who are living on the other side of the earth, for their

unconditional love and support throughout this adventure. I thank them for letting their only

child go to a foreign country and pursue her PhD in immunology. I’m thankful to the freedom

and independence they endowed, which are invaluable to me. Finally, I would like to thank

Chikin Kuok, my husband and science buddy, for delivering coffee and snacks when I have late-

night experiments and for cheering me up when I’m in a bad mood. His support has been

steadfast and fun, and it’s my fortune to have him with me.

vi

Table of Contents

Acknowledgments.......................................................................................................................... iv

Table of Contents ........................................................................................................................... vi

List of Abbreviations ..................................................................................................................... ix

List of Tables ............................................................................................................................... xiv

List of Figures ................................................................................................................................xv

Manuscripts Arising from this Thesis ......................................................................................... xvii

Chapter 1 ..........................................................................................................................................1

Introduction .................................................................................................................................1

1.1 Dendritic cells, a heterogeneous population ........................................................................2

1.1.1 Dendritic cells bridge the innate and adaptive immune response ............................2

1.1.2 A brief historical perspective on DC .......................................................................2

1.1.3 Conventional DCs ....................................................................................................3

1.1.4 Plasmacytoid DCs ....................................................................................................5

1.1.5 Monocyte-derived DCs ............................................................................................5

1.1.6 Langerhans Cells ......................................................................................................6

1.2 DC ontogeny and development ............................................................................................6

1.2.1 Cytokine control of the DC lineage .........................................................................8

1.2.1.1 FLT3 ligand ...............................................................................................8

1.2.1.2 CSF-1 (M-CSF) and CSF-2 (GM-CSF) ....................................................8

1.2.2 Transcriptional control of the DC lineage ...............................................................9

1.2.2.1 Transcription factors affecting multiple DC lineages ...............................9

1.2.2.2 Transcription factors affecting the CD8+CD11b- DCs and

CD103+CD11b- DCs ...............................................................................10

1.2.2.3 Transcription factors affecting the CD8-CD11b+ DCs and

CD103+CD11b+ DCs ...............................................................................12

1.2.3 Tools for studying DC biology ..............................................................................14

1.2.3.1 In vitro culture systems ...........................................................................14

1.2.3.2 Depletion of DCs by DTR/DTA systems ................................................14

1.2.3.3 Cre strains for conditional deletion of genes in DCs ...............................16

1.2.3.4 Transcription factor-based depletion of DCs ..........................................17

1.3 Effect of age on DC phenotype ..........................................................................................18

vii

1.4 The role of DC in intestinal health and disease .................................................................19

1.4.1 The intestinal environment and barrier function ....................................................19

1.4.2 Anatomy of the intestinal wall ...............................................................................21

1.4.3 Lymphoid structures in the intestine ......................................................................22

1.4.3.1 Peyer’s patches ........................................................................................23

1.4.3.2 The small intestinal lamina propria .........................................................25

1.4.3.3 Mesenteric lymph nodes ..........................................................................25

1.4.4 Regulation of intestinal homeostasis by DCs ........................................................26

1.4.4.1 Sampling and antigen uptake from the intestinal lumen .........................26

1.4.4.2 DC maturation .........................................................................................27

1.4.5 T-cell priming and induction of adaptive immune responses by DC ....................28

1.4.5.1 Signal 1 and Signal 2 ...............................................................................30

1.4.6 Signal 3: Determining T-cell differentiation into an effector cell .........................31

1.4.6.1 Expression of Lymphotoxin and LTR signaling ...................................32

1.4.6.2 LTβR signaling is required for the organogenesis of secondary

lymphoid tissues and the maintenance of lymphoid tissue

microarchitecture .....................................................................................33

1.4.6.3 LTβR signaling in regulating immune responses ....................................35

1.4.6.4 LTR signaling in intestinal disease .......................................................38

1.4.7 Fates of mucosal immune responses - Fate 1: Oral Tolerance ..............................40

1.4.8 Fate 2: Immune response against harmful pathogens ............................................41

1.4.9 Fate 3: Immune response against self-antigens .....................................................42

1.4.10 Rotavirus infection model ......................................................................................43

1.4.10.1 Epidemiology and RV vaccines ..............................................................43

1.4.10.2 Rotavirus ..................................................................................................44

1.4.10.3 Pathogenesis ............................................................................................45

1.4.10.4 Host immunity to RV infection ...............................................................46

1.4.10.5 Contribution of maternal effects on RV immune responses ...................51

1.5 Human cDC .......................................................................................................................52

1.6 Summary ............................................................................................................................53

Chapter 2 ........................................................................................................................................54

Methods and Materials for Chapter 3 and Chapter 4 ................................................................54

Chapter 3 ........................................................................................................................................61

viii

Intestinal BATF3-dependent dendritic cells are required for optimal antiviral T-cell

responses in adult and neonatal mice ........................................................................................61

3.1 Abstract ..............................................................................................................................62

3.2 Introduction ........................................................................................................................63

3.3 Results ................................................................................................................................66

3.3.1 Depletion of ZBTB46-dependent cDCs is not affected by RV infection ..............66

3.3.2 ZBTB46-dependent cDCs are required for anti-RV CD8+ T-cell responses .........70

3.3.3 Adult and neonatal Batf3-/- mice exhibit similar deficiencies in

CD103+CD11b- cDCs both at steady state and during RV infection .....................73

3.3.4 Adult and neonatal mice have distinct DC requirements for mounting RV-

specific CD8+ T-cell responses ..............................................................................76

3.3.5 Polyclonal antiviral Th1 responses in neonatal mice are BATF3-dependent ........80

3.3.6 Intact local and systemic anti-RV IgA responses in cDC-deficient mice ..............80

3.3.7 CD103+CD11b+ DCs are not required for mounting anti-RV adaptive immune

responses ................................................................................................................82

3.4 Discussion ..........................................................................................................................86

Chapter 4 ........................................................................................................................................94

LTR deficiency in radio-sensitive compartments results in skewed T-cell responses

during a mucosal viral infection ................................................................................................94

4.1 Abstract ..............................................................................................................................95

4.2 Introduction ........................................................................................................................96

4.3 Results ................................................................................................................................99

4.3.1 Ltbr-/- chimeric mice clear RV with normal kinetics .............................................99

4.3.2 Ltbr-/- chimeric mice generate more IFN-secreting CD8+ T cells during

primary RV infection .............................................................................................99

4.3.3 Polyclonal CD4+ T cell cytokine profiles are skewed in the Ltbr-/- chimeric

mice at steady state and after RV infection .........................................................103

4.3.4 Ltbr-/- chimeric mice generate a normal intestinal IgA response to RV ..............106

4.4 Discussion ........................................................................................................................109

Chapter 5 ......................................................................................................................................113

Discussion and Future Directions ...........................................................................................113

References ....................................................................................................................................129

ix

List of Abbreviations

ADP Adenosine diphosphate

AID Activation-induced deaminase

ALDH Aldehyde dehydrogenase

AMP Antimicrobial peptide

AP Activator protein

APC Antigen presenting cell

APRIL A proliferation-inducing ligand

ATP Adenosine triphosphate

2m Beta 2-microglobulin

BAC Bacterial artificial chromosome

BAFF B-cell activating factor

BATF3 Basic leucine zipper ATF-like transcription factor 3

BCL-6 B-cell CLL/Lymphoma 6

Blimp-1 B lymphocyte–induced maturation protein-1

BM Bone marrow

BMDC Bone marrow-derived dendritic cell

BST2 Bone marrow stromal antigen 2

BTLA B- and T-lymphocyte attenuator

CCL C-C motif chemokine ligand

CCR C-C motif chemokine receptor

CD Crohn’s disease

CD40L CD40 ligand

cDC Classical dendritic cell

CDP Common DC progenitor

CLEC C-type lectin

CLP Common lymphoid progenitor

CLR C-type lectin receptor

CMP Common myeloid progenitor

CP Cryptopatch

CSL CBF1-suppressor of hairless-Lag1

CSR Class switch recombination

CTL Cytotoxic T lymphocyte

CTLA-4 Cytotoxic T-lymphocyte-associated protein 4

CXCL C-X-C motif chemokine ligand

CXCR C-X-C motif chemokine receptor

CX3CR1 C-X3-C motif chemokine receptor 1 or fractalkine receptor

DAMP Damage-associated molecular pattern

DC Dendritic cell

DCM Division of comparative medicine

x

DcR3 Decoy receptor 3

DD50 50% diarrhea dose

DN DC Double negative dendritic cell

dsRNA Double-stranded RNA

DTA Diphtheria toxin A-chain

DTR Diphtheria toxin receptor

DTx Diphtheria toxin

ELISA Enzyme-Linked Immunosorbent Assay

ELISPOT Enzyme-Linked ImmunoSpot

ENS Enteric nervous system

ER Endoplasmic reticulum

ESAM Endothelial cell-selective adhesion molecule

FAE Follicle-associated epithelium

FcRI The high-affinity IgE receptor

FcRn Neonatal Fc receptor

FcγRI Fc-gamma receptor 1

FDC Follicular dendritic cell

FLT3 Fms-like tyrosine kinase 3

FRC Fibroblast reticular cell

GALT Gut-associated lymphoid tissue

GC Germinal center

GF Germ-free

GFP Green fluorescent protein

GM-CSF/CSF-2 Granulocyte/macrophage colony-stimulating factor

GM-CSFR Granulocyte–macrophage colony-stimulating factor receptor

HVEM Herpesvirus entry mediator

HIV Human immunodeficiency virus

HLA Human leukocyte antigen

HLH Helix-loop-helix

HMGB1 High mobility group box 1 protein

HSC Hematopoietic stem cell

HSP Heat shock protein

HSV Herpes simplex virus

IBD Inflammatory bowel disease

ICAM-1 Intercellular adhesion molecule 1

ICN Intracellular domain of Notch

ICOS-L Inducible costimulator-ligand

Id2 Inhibitor of DNA binding 2

iDC Inflammatory dendritic cell

IDO Indoleamine 2,3-dioxygenase

IEC Intestinal epithelial cell

IEL Intraepithelial lymphocyte

xi

IFN Interferon

IFR Interfollicular region

Ig Immunoglobulin

IKK IkB kinase

IL Interleukin

ILC Innate lymphoid cell

ILF Isolated lymphoid follicle

iNOS Inducible nitric-oxide synthase

IRES Internal ribosome entry site

IRF Interferon-regulatory factor

ISG Interferon stimulated gene

ISRE IFN sequence response element

KLF4 Kruppel-like factor 4

KO Knockout

LAMP2 Lysosome-associated membrane protein 2

LC Langerhans cell

LCMV Lymphocytic choriomeningitis virus

LIGHT Homologous to lymphotoxin, inducible expression, competes

with herpes simplex virus (HSV) glycoprotein D for HSV entry

mediator, a receptor expressed on T lymphocytes

LN Lymph node

LP Lamina propria

LPS Lipopolysaccharide

LT Lymphotoxin alpha

LT Lymphotoxin beta

LTR Lymphotoxin beta receptor

LTi Lymphoid tissue inducer

Lto Lymphoid tissue organizing

M cell Microfold cell

mAb Monoclonal antibody

MAdCAM-1 Mucosal vascular addressin cell adhesion molecule 1

MALT Mucosal associated lymphoid tissue

MAPK Mitogen-activated protein kinase

MAVS Mitochondrial antiviral signaling protein

M-CSF/CSF-1 Macrophage colony stimulating factor

MDA5 Melanoma differentiation-associated protein 5

MDP Macrophage and dendritic cell precursor

MHC Major histocompatibility complex

MHCII Class II major histocompatibility complex

MHV-68 Murine gammaherpesvirus 68

MIP-1 Macrophage inflammatory protein-1 beta or CCL4

MMTV Mouse mammary tumor virus

xii

MNV Murine norovirus

moDC Monocyte-derived dendritic cell

MyD88 Myeloid differentiation primary response protein 88

MZ Marginal zone

NALT Nasal associated lymphoid tissue

NF-κB Nuclear factor-κB

NIK NF-κB-inducing kinase

NK cell Natural killer cell

NLR NOD-like receptor

NOD Nucleotide-binding oligomerization domain

NP 4-hydroxy-3-nitrophenyl acetyl

NSP Non-structural protein

NZB New Zealand Black

OVA Ovalbumin

PAMP Pathogen-associated molecular pattern

pDC Plasmacytoid dendritic cell

PNAd Peripheral node addressin

PP Peyer’s patch

PRR Pattern recognition receptor

RA Retinoic acid

Rag Recombinase-activating gene

RANK Receptor activator of nuclear factor kappa-B or TRANCE

Receptor

RANKL Receptor activator of nuclear factor kappa-B ligand or TRANCE

RBP-J Recombination-signal-binding protein-J

RegIII Regenerating islet-derived protein III

RIG-I Retinoid acid-inducible gene I

RLR Retinoic acid-inducible gene I-like receptor

ROR Retinoic acid- related orphan receptor

RRV Rhesus rotavirus

RV Rotavirus

SCID Severe combined immunodeficiency

SED Subepithelial dome

SFB Segmented filamentous bacteria

SHM Somatic hypermutation

Siglec-H Sialic acid-binding immunoglobulin-like lectin H

SILP Small intestinal lamina propria

SILT Solitary isolated lymphoid tissue

SIRP Signal-regulatory protein alpha or CD172

ssRNA Single-stranded RNA

STAT Signal transducer and activator of transcription

TAP Transporter associated with antigen processing

xiii

TCR T-cell receptor

Tfh Follicular helper T cell

TG2 Transglutaminase 2

TGF- Transforming growth factor beta

Th T helper

Tip DC Tumor-necrosis factor alpha- and inducible nitric-oxide

synthase- producing dendritic cell

TLR Toll-like receptor

TNF Tumor-necrosis factor

TNFSF Tumor-necrosis factor superfamily

TRAF6 Tumor necrosis factor receptor-associated factor 6

Treg Regulatory T cell

TRIF TIR-domain-containing adaptor-inducing interferon beta

TSLP Thymic stromal lymphopoietin

UC Ulcerative colitis

UTR Untranslated region

UV Ultraviolet

VCAM-1 Vascular cell adhesion molecule 1

VP Viral protein

WT Wild type

XCR1 XC-chemokine receptor 1

ZBTB Broad Complex, Tramtrack, Bric-a-Brac, and Zinc Finger family

member

xiv

List of Tables

Table 1-1 Comparison of different cDC subsets............................................................................. 3

Table 1-2 Comparison of different transcription factor knockout mice ....................................... 17

Table 2-1 Mouse strains ................................................................................................................ 55

Table 2-2 List of buffers and solutions used for cell isolation and culture ................................... 59

Table 2-3 Primer sets .................................................................................................................... 60

Table 5-1 Alterations of different DC types, plasma cells and IgA in different chimeras ......... 121

xv

List of Figures

Figure 1-1 DC hematopoiesis ......................................................................................................... 7

Figure 1-2 Layers of the intestinal wall ........................................................................................ 22

Figure 1-3 Gut-associated lymphoid tissues in the small intestine ............................................... 23

Figure 1-4 The Lymphotoxin system and the LTR signaling pathway. ..................................... 33

Figure 1-5 The role of LTR signaling in the periphery and the intestine ................................... 39

Figure 1-6 Rotavirus structure ...................................................................................................... 45

Figure 3-1 Conventional DC populations in DTx-treated Zbtb46-DTR WT chimeric mice.... 67

Figure 3-2 Gating strategy for SILP single cell populations. ....................................................... 69

Figure 3-3 ZBTB46-dependent cDCs are required to prime RV-specific CD8+ T cells. ............. 71

Figure 3-4 Tetramer staining and ICS of SILP CD8+ T cells in the SILP of Zbtb46-DTRWT

chimeric mice. ....................................................................................................................... 72

Figure 3-5 Antigen presenting cells in the SILP of adult and neonatal Batf3-/- mice at steady state

and upon RV infection. ......................................................................................................... 74

Figure 3-6 Kinetics of absolute numbers of DCs and DC subsets in the SILP and MLNs of

Batf3+/- and Batf3-/- mice. ...................................................................................................... 75

Figure 3-7 CD103+CD11b- DCs are required for optimal anti-RV CD8+ T-cell responses in

Batf3-/- adult and neonatal mice. ........................................................................................... 77

Figure 3-8 SILP CD8+ T-cell responses (absolute numbers) in adult Batf3-/- mice. .................... 79

Figure 3-9 Alteration of Th1 and Th17 responses in adult and neonatal Batf3-/- mice. ............... 81

Figure 3-10 cDCs are dispensable for the induction and maintenance of local and systemic

antiviral IgA. ......................................................................................................................... 83

Figure 3-11 CD103+CD11b+ cDCs are not required for anti-RV CD8+ T-cell responses in the

SILP of adult and neonatal huLangerin-DTA mice. ............................................................. 84

Figure 3-12 Similar anti-RV CD8+ T cell responses are observed in CD11c-DTRWT and

Zbtb46-DTRWT chimeric mice. ....................................................................................... 87

Figure 3-13 Innate immune responses in neonatal Batf3+/- and Batf3-/- mice with and without RV

infection. ............................................................................................................................... 91

Figure 4-1 LTR deficiency in the radio-sensitive compartment does not affect local viral

clearance. ............................................................................................................................ 100

Figure 4-2 Gating strategy for the SILP CD4+ and CD8+ T cells. .............................................. 101

xvi

Figure 4-3 Frequency and proliferation status of CD8+ T cells in RV-infected Ltbr-/- chimeric

mice. .................................................................................................................................... 102

Figure 4-4 Assessment of IFN-producing CD8+ T cells in Ltbr-/- chimeric mice induced by

mitogen or viral peptide stimulation. .................................................................................. 105

Figure 4-5 Expansion of polyclonal CD4+ T cells in response to RV infection is increased, and

CD4+ T cell cytokine profiles are skewed in Ltbr-/- chimeric mice. ................................... 107

Figure 4-6 LTR signaling pathway in radio-sensitive compartments is dispensable for local

antiviral IgA production. ..................................................................................................... 108

Figure 5-1 Th1 and Th17 responses in Zbtb46-DTRWT chimeric mice. ............................... 115

Figure 5-2 Frequency of pDC in the SILP at 7 d.p.i. .................................................................. 119

Figure 5-3 Fecal and serum RV-specific IgA responses in CD11c-DTR chimeric mice. .......... 120

Figure 5-4 Contrasting views on the role of LT12 in IgA class switch. ................................ 123

xvii

Manuscripts Arising from this Thesis

Tian Sun, Olga L. Rojas, Conglei Li, Lesley A. Ward, Dana J. Philpott, Jennifer L.

Gommerman. Intestinal Batf3-dependent dendritic cells are required for optimal antiviral T-cell

responses in adult and neonatal mice. Mucosal Immunol. 2017 May; 10(3):775-788.

Tian Sun, Olga L. Rojas, Conglei Li, Dana J. Philpott, Jennifer L. Gommerman. Hematopoietic

LTβR deficiency results in skewed T cell cytokine profiles during a mucosal viral infection. J

Leukoc Biol. 2016 Jul; 100(1):103-110

1

Chapter 1

Introduction

2

1.1 Dendritic cells, a heterogeneous population

1.1.1 Dendritic cells bridge the innate and adaptive immune response

Productive immune responses are accompanied by signals that alert the immune system to

danger, which include damage-associated molecular patterns (DAMPs) and pathogen-associated

molecular patterns (PAMPs) that activate pattern recognition receptors (PRRs) (Janeway and

Medzhitov, 2002; Matzinger, 2002). Following their activation, PRRs provide signals to the host

indicating the presence of a microbial infection or cell damage. PRR signals trigger innate

immune responses in specialized cells called dendritic cells (DCs). DCs are sentinel cells

existing throughout the body: from lymphoid organs like spleen and lymph nodes (LNs), to non-

lymphoid tissues such as skin, lung, reproductive tract and intestine. These widespread stellate-

shaped cells serve as the bridge between the innate and adaptive immune response via a process

called antigen presentation. Indeed, they are considered the "professional" antigen presenting

cells or APCs of the immune system.

Following activation by PAMPs/DAMPs, DCs downregulate antigen-uptake capacity and

migrate from the inflamed tissues to the closest draining LNs via chemotaxis. Within LNs, DCs

process and present immunogenic peptides to cells of the adaptive immune system (T cells) in

the context of major histocompatibility complex (MHC) molecules. The adaptive immune

system then responds to the infection or damage by initiating clonal selection of appropriate

lymphocytes. These lymphocytes eliminate any pathogen or damage that has not been taken care

of by early innate immune responses and provides immunologic memory for long-term

protection. Therefore, the central orchestrating cell to these complex steps is the DC. In this

thesis, my aim was to obtain a better understanding of this specialized cell type in the context of

the small intestinal viral infection in mice. Such new knowledge will inform the rational

development of mucosal vaccines.

1.1.2 A brief historical perspective on DC

DCs were first described in the suprabasal region of the epidermis by a medical student in

Germany, Paul Langerhans, who thought they were part of the nervous system (Langerhans,

1868). It was not until 1973 that Ralph Steinman and Zanvil Cohn at the Rockefeller University

3

identified DCs as accessory cells with unique morphology in the mouse spleen (Steinman and

Cohn, 1973). Most importantly, Steinman and colleagues described DCs as potent T cell-

stimulating cells and they found DCs to be at least 100 times more effective at priming T cells

than macrophages (Nussenzweig et al., 1980; Steinman et al., 1983). Their discovery began the

modern era of the DC biology and Ralph Steinman was awarded the Nobel prize in Physiology

or Medicine in 2011.

We now know that there are four types of DCs: conventional or classical DCs (cDCs),

plasmacytoid DCs (pDCs), Langerhans cells (LCs), and monocyte-derived DCs (moDCs). Each

type of DCs is briefly described below.

1.1.3 Conventional DCs

cDCs exhibit superior capacity for taking up, processing and presenting antigens to naïve T cells.

They express high levels of CD11c and class II MHC (MHCII) and can be further divided based

on the surface markers of CD8 and CD11b in lymphoid tissues (Vremec et al., 2000; Vremec et

al., 1992), or CD103 and CD11b in nonlymphoid tissues (Ginhoux et al., 2009). Different cDC

subsets require distinct genetic factors for their development and display unique gene-expression

profiles and functions (Miller et al., 2012). Some features of different cDC subsets are descripted

below (Table 1-1).

XC-chemokine receptor 1 (XCR1) and signal-regulatory protein alpha (SIRPor CD172; a

receptor for the signal-regulatory protein CD47 which are expressed in a mutually exclusive

manner, have been recently shown to be superior markers for distinguishing cDC subsets,

replacing CD8 and CD11b, respectively (Bachem et al., 2010; Gurka et al., 2015). However, to

remain consistent in the following chapters, I will still use CD103/CD8 and CD11b to describe

cDC subsets.

Table 1-1 Comparison of different cDC subsets

cDC CD11c+MHCII+lin(F4/80, CD3B220)-

Subsets in

lymphoid

tissues

CD8+CD11b- CD8-CD11b+ CD8-CD11b-

4

Subsets in

nonlymphoid

tissues

CD103+CD11b- CD103+CD11b+ CD103-CD11b+/- (?)

Additional

markers

XCR1,

CLEC9A/DNGR-1,

CD24, DEC-205,

Langerin/CD207

SIRPCD172 CD4,

CLEC4A4

SIRP

Transcription

control

ZBTB46, IRF8,

BATF3, Id2, BCL-6,

Nfil3

(Murphy et al., 2016)

ZBTB46 and IRF4;

partially by KLF4, RBP-J

(Murphy et al., 2016)

Partially by IRF4 and

IRF8

(Tamura et al., 2005)

Funct

ions

Anti

gen

pre

senta

tion

MHC-I machinery

(viral, tumor and

intracellular bacterial

antigens)

(den Haan et al., 2000;

Edelson et al., 2011;

Hildner et al., 2008)

MHC-II machinery

(extracellular bacterial

and fungal antigens);

neonatal Fc receptor

(FcRn)- mediated cross

presentation

(Baker et al., 2011)

Antigen presentation

to CD8+ T cells in the

lung and the gut

(Ballesteros-Tato et

al., 2014;

Ballesteros-Tato et

al., 2010; Belz et al.,

2004; Fleeton et al.,

2004)

Cyto

kin

e pro

duct

ion a

nd r

elat

ed f

un

ctio

ns

IL-12

Th1 differentiation;

anti-parasite defense;

suppress helminth-

driven Th2 response

(Everts et al., 2016;

Martínez-López et al.,

2015; Mashayekhi et

al., 2011)

IL-13; IL-6 and IL-23

Th2 polarization; Th17

differentiation; induction

of IL-22 secretion

(Heink et al., 2017;

Kinnebrew et al., 2012;

Persson et al., 2013b;

Plantinga et al., 2013;

Schlitzer et al., 2013;

Tumanov et al., 2011;

Williams et al., 2013)

TNF and IL-23; IL-

12

Th17 differentiation

in vitro

(Coombes et al.,

2007; Iwasaki and

Kelsall, 2001; Scott

et al., 2015)

TGF and RA; IDO

Individually redundant but are together required to

maintain intestinal Treg homeostasis

(Coombes et al., 2007; Matteoli et al., 2010; Welty

et al., 2013)

Skin CD103- DCs

produce RA and

induce Treg

(Guilliams et al.,

2010)

Induct

ion o

f T

-

cell

hom

ing

Induce gut tropism to

OT-I T cells; but

expression of gut-

homing receptors on

homeostatic T cells is

Confer gut tropism to

differentiating T cells in

vitro; but expression of

gut-homing receptors on

homeostatic CD4+ T cells

Intestinal lymphatic

CD103-DC is ability

to confer gut tropism

to differentiating T

cells in vitro

(Cerovic et al., 2013)

5

normal in mice lacking

this DC subset

(Cerovic et al., 2015;


Ohta et al., 2016)

is normal in mice lacking

this DC subset

(Cerovic et al., 2013;

Persson et al., 2013b)

Individually redundant but are together required to

imprint gut-homing receptors on Treg cells

(Welty et al., 2013)

\ O

ther

funct

ions

Maintain intestinal LP

T cell and IEL

homeostasis

(Luda et al., 2016;

Muzaki et al., 2016)

Promote IgA+ B-cell

responses in the PPs

(Reboldi et al., 2016; Sato

et al., 2003)

Produce osteopontin

during colitis

(pathogenic)

(Kourepini et al.,

2014)

1.1.4 Plasmacytoid DCs

pDCs are a unique DC subset that specializes in producing type I interferons (IFNs) in response

to viruses (Cella et al., 1999; Siegal et al., 1999). They accumulate mainly in the blood and

lymphoid tissues and enter the LNs through the blood circulation. The markers most commonly

used to identify pDCs in mice are CD11c, B220, Ly6C, bone marrow stromal antigen 2 (BST2,

or CD317) and sialic acid-binding immunoglobulin-like lectin H (Siglec-H) (Swiecki and

Colonna, 2015). The recognition of viruses or self-nucleic acids by pDCs is mainly mediated by

toll-like receptor (TLR) 7 and TLR9, resulting in their secretion of type I IFNs via the myeloid

differentiation primary response protein 88 (MyD88)- interferon-regulatory factor (IRF) 7

pathway, as well as their production of pro-inflammatory cytokines and chemokines via the

MyD88-nuclear factor-κB (NF-κB) pathway (Gilliet et al., 2008). In addition, pDCs can act as

APCs as they express MHCII molecules as well as the co-stimulatory molecules CD40, CD80

and CD86, and can present antigens to CD4+ T cells, albeit not as efficiently as cDCs

(Villadangos and Young, 2008).

1.1.5 Monocyte-derived DCs

MoDCs or inflammatory DCs (iDCs) are a DC subset derived from monocytes that infiltrate into

inflamed tissues. Differentiation of monocytes into DCs in vitro and in vivo was first described

by Randolph and colleagues (Randolph et al., 1998; Randolph et al., 1999). moDCs are primarily

recognized as MHCII+CD11b+CD11c+F4/80+Ly6C+ DCs (León et al., 2007). Subsequently,

6

markers such as mannose receptor, granulocyte–macrophage colony-stimulating factor receptor

(GM-CSFR), lysosome-associated membrane protein 2 (LAMP2), high-affinity immunoglobulin

(Ig) E receptor (FcRI) and Fc-gamma receptor 1 (FcγRI) /CD64 are also found to be expressed

by moDCs (Segura and Amigorena, 2013). One example of moDC is the tumor-necrosis factor

(TNF) and inducible nitric-oxide synthase (iNOS) -producing DCs (or Tip DCs) that are found

in the spleen of Listeria monocytogenes-infected mice, and are absent from CCR2-deficient mice

(Serbina et al., 2003). In addition to their production of inflammatory mediators, moDCs can

induce Th1/Th17-polarized CD4+ T-cell responses (Ko et al., 2014; León et al., 2007), cross-

prime antigen-specific CD8+ T cells (Aldridge et al., 2009; Le Borgne et al., 2006) and regulate

optimal IgA production in the gut (Tezuka et al., 2007). Therefore, moDCs play important roles

in both innate and adaptive immune responses.

1.1.6 Langerhans Cells

LCs are a population of mononuclear phagocytes restricted to the epidermal skin layer. LCs are

characterized phenotypically as MHCII+CD11c+langerin (CD207)hiCD11b+F4/80+CX3CR1-

(Merad et al., 2013). Interestingly and unique among DCs, under steady state conditions, LCs

proliferate in situ to form a radio-resistant, self-renewing population, whereas upon ultraviolet

(UV) light-induced skin inflammation, blood-borne LC precursors are recruited to the skin in a

CCR2-dependent manner (Merad et al., 2002). LCs can extend dendritic processes in the vertical

axis towards the stratum corneum (the outermost layer of the epidermis) to acquire antigens at or

near the external surface of the skin (Kubo et al., 2009), or in the horizontal plane of the

epidermis to sample the area of the epidermis and contact keratinocytes (Kaplan, 2010). After

antigen uptake, LCs migrate to regional LNs where they can present antigen to naïve and

memory T cells and induce Th17- or regulatory T (Treg)- cell responses but not viral specific

CD8+ T-cell responses (Allan et al., 2003; Igyarto et al., 2011; Shklovskaya et al., 2011).

1.2 DC ontogeny and development

Most DCs are short-lived hematopoietic cells that are continually replaced by blood-derived

precursors (Merad et al., 2013). Starting from hematopoietic stem cell (HSCs) in the bone

marrow (BM), the earliest committed precursors include clonal common myeloid progenitors

7

(CMPs) and common lymphoid progenitors (CLPs). CMPs subsequently develop into

granulocyte macrophage progenitors (GMPs) and both of these can give rise to macrophage and

DC precursors (MDPs) (Liu et al., 2009; Merad et al., 2013). The commitment of myeloid

precursors to the mononuclear phagocyte lineage is thought to occur at the MDP stage, as MDPs

are able to produce DCs and macrophages but lose the ability to generate granulocytes upon

adoptive transfer (Fogg et al., 2006). Common monocyte progenitors (cMoP) and common DC

progenitors (CDPs) are found to be immediately downstream of MDPs (Hettinger et al., 2013;

Liu et al., 2009). While cMoP develop into monocytes (Hettinger et al., 2013), CDPs is thought

to generate pre-cDCs and pre-pDCs, of which the latter gives rise to pDCs (Naik et al., 2007;

Onai et al., 2007).

Figure 1-1 DC hematopoiesis

cDCs develop from BM HSCs in a stepwise manner. HSCs generate CLPs and CMPs. CMPs

then develop into GMPs, both of these cells can develop into MDPs. MDPs then give rise to

cMoP and CDPs: the former develop into monocytes/macrophage lineage and the later branch

into ZBTB46-dependent pre-cDCs and pre-pDCs. Most BM pre-cDCs are still uncommitted, but

these cells gradually split up into IRF8/BATF3-dependent pre-CD8+ cDC and IRF4-dependent

pre-CD11b+ cDC. These committed pre-cDCs then leave the BM into the bloodstream and seed

the different tissues where they fully differentiate into CD8+ cDC and CD11b+ cDC. pre-pDCs

give rise to pDCs in the BM and then join the circulation.

8

Recently, a few studies have investigated heterogeneity within the pre-cDC population in more

detail and have further defined pre-CD8+ cDCs committed to CD8+ cDCs development, and

pre-CD11b+ cDCs committed to CD11b+ cDC development in the BM (Grajales-Reyes et al.,

2015; Schlitzer et al., 2015). Monocytes, pDCs and pre-cDCs then leave the BM and seed

lymphoid and non-lymphoid tissues (Diao et al., 2006; Naik et al., 2006). In these tissues, pre-

cDCs further differentiate into immature cDC subsets (Schlitzer et al., 2015) (Figure 1-1).

1.2.1 Cytokine control of the DC lineage

Many cytokines and transcription factors are required for the overall process of DC development.

Most of these are also involved in the development of other hematopoietic lineages. Some,

however, have a strong selective effect on the generation of DCs or particular DC subtypes.

Below is a summary of selected cytokines that extrinsically regulate DC lineages.

1.2.1.1 FLT3 ligand

The ligand of tyrosine kinase receptor fms-like tyrosine kinase 3 (FLT3, also termed fetal liver

kinase 2 (FLK2) or CD135), or FLT3L, is a key regulator of DC commitment in hematopoiesis

(Merad et al., 2013). FLT3 is expressed throughout DC development, including HSCs

(Adolfsson et al., 2001), a subset of CMPs (Karsunky et al., 2003) and maintained on MDPs

(Waskow et al., 2008), CDPs (Onai et al., 2007), pre-cDCs (Liu et al., 2009) and tissue cDCs,

with the exception of LCs (Bogunovic et al., 2009). Loss of Flt3 expression in hematopoietic

progenitors correlates with the loss of DC differentiation potential (Karsunky et al., 2003),

whereas enforced Flt3 expression in progenitors that lack DC potential partially restores DC

development (Onai et al., 2006). Moreover, inhibition of FLT3L leads to decreased numbers of

MDPs, CDPs and tissue cDCs and pDCs (Kingston et al., 2009; Tussiwand et al., 2005).

Conversely, injection or overexpression of FLT3L in mice leads to a dramatic expansion of

cDCs and pDCs (Manfra et al., 2003; Maraskovsky et al., 1996).

1.2.1.2 CSF-1 (M-CSF) and CSF-2 (GM-CSF)

CSF-1 or macrophage colony stimulating factor (M-CSF) is a hematopoietic factor that regulates

the survival, proliferation and differentiation of macrophages. CSF-1 receptor (CSF-1R) is

expressed on MDPs, CDPs, reduced in pre-cDCs, and lost on CD8+ (and CD103+) cDCs while

9

maintained on a subset of CD11b+ cDCs (Merad et al., 2013). Therefore, the balance between

FLT3 versus CSF-1R signals likely determines MDP progression to CDPs instead of to a

monocyte phenotype (Schmid et al., 2010). CSF-1R partly regulates the differentiation or

survival of cDCs in nonlymphoid tissue, potentially reflecting a monocytic origin of this subset

or the heterogeneity of this population (Bogunovic et al., 2009).

CSF-2 (or granulocyte/macrophage colony-stimulating factor, GM-CSF) is a hematopoietic

growth factor that controls the differentiation of the myeloid lineage. CSF-2 receptor (CSF-2R)

is expressed on MDPs, CDPs and differentiated cDCs (Kingston et al., 2009). Although CSF-2 is

a key cytokine for promoting the differentiation of mouse and human hematopoietic progenitors,

mice lacking CSF-2 or its receptor display only minor deficiencies in cDC development within

lymphoid tissues (Vremec et al., 1997). However, a reduction in the number of cDCs is found in

nonlymphoid tissues of Csf-2-/- mice, suggesting that CSF-2 is a critical regulator of cDC

maintenance in nonlymphoid tissues, but not in lymphoid organs (Greter et al., 2012).

1.2.2 Transcriptional control of the DC lineage

Pluripotent HSCs undergo progressive restriction in their lineage potential to give rise to mature,

terminally differentiated cells. The process of HSCs differentiation is thought to follow a

developmentally ordered pattern of gene expression (Shivdasani and Orkin, 1996). The

transcription factors regulating the differentiation and expansion of specific DC lineages are

described below.

1.2.2.1 Transcription factors affecting multiple DC lineages

• STAT3

Hematopoietic deletion of signal transducer and activator of transcription (STAT) 3, a

transcription factor in downstream FLT3 signaling, leads to reduced DC development in

lymphoid organs (Laouar et al., 2003), whereas overexpression and activation of STAT3 in Flt3-

deficient hematopoietic progenitors rescues both pDC and cDC differentiation potential (Onai et

al., 2006). Furthermore, the requirement of STAT3 in the FLT3 pathway is stage-specific and is

restricted to the CMP to CDP transition phase (Laouar et al., 2003). Since STAT3 participates in

10

a wide variety of physiological processes (Levy and Lee, 2002), it is not an ideal candidate to

manipulate for studying DC biology.

• ZBTB46

The transcription factor ZBTB46 (BTBD4), a Broad Complex, Tramtrack, Bric-a-Brac, and Zinc

Finger (BTB-ZF) family member, is selectively expressed by cDC lineages, vascular

endothelium and megakaryocyte-erythroid progenitors (Meredith et al., 2012a; Satpathy et al.,

2012). During DC development, ZBTB46 is first expressed in pre-cDCs and retains at a high

level of expression in downstream cDC lineages in both lymphoid and nonlymphoid tissues, but

not in the myeloid or lymphoid cell types (Figure 1-1)(Meredith et al., 2012a; Satpathy et al.,

2012). These observations suggest that ZBTB46 is a marker for cDC commitment. However,

ZBTB46 is not required for early cDC development in the BM, rather its deficiency alters the

cDC subset composition in the spleen in favor of CD8+ DCs (Meredith et al., 2012b; Satpathy

et al., 2012).

1.2.2.2 Transcription factors affecting the CD8+CD11b- DCs and CD103+CD11b- DCs

• IRF8

IRF8 (also known as IFN consensus sequence binding protein, ICSBP) plays a critical role in

myeloid cell differentiation, while inhibiting the development of granulocytes. IRF8 binds to

other members of the IRF family and to the hematopoietic-specific member of the Ets family,

PU.1, to form transcriptional complexes and activate transcription via binding to PU.1/IRF

composite sequences (Marecki et al., 1999). The expression of IRF8 is restricted to myeloid and

lymphoid cell lineages, including cells of monocyte/macrophage lineage, B lymphocytes, and

activated T cells (Tamura and Ozato, 2002).

Irf8-/- mice develop a myeloproliferative disease distinguished by excessive granulocyte

production, failure to generate adequate monocyte numbers, and lack of pDCs, CD8+ cDCs in

lymphoid tissues and CD103+ cDCs in nonlymphoid tissues (Edelson et al., 2010; Holtschke et

al., 1996; Schiavoni et al., 2002). A spontaneous point mutation (R294C) of IRF8 in BHX2 mice

also causes myeloproliferative disease and impairs the development of CD8+ and CD103+

11

cDCs without impairing pDC generation (Tailor et al., 2008; Turcotte et al., 2005). Additionally,

IRF8 controls CD8+ cDC maturation and IL-12 production (Schiavoni et al., 2002) and plays a

role in the tolerogenic functions of cDCs by regulating the expression of Indo, the gene coding

for the enzyme indoleamine 2,3-dioxygenase (IDO) (Orabona et al., 2006), as well as modulating

pDC function (Sichien et al., 2016).

• BATF3

BATF3 (also known as Jun dimerization protein p21SNFT) is highly expressed in cDCs, with

low to absent expression in other immune cells and nonimmune tissues (Hildner et al., 2008).

BATF3 is expressed in both CD8+CD11b- cDCs and CD8-CD11b+ cDCs, but BATF3 is

required only for the development of CD8+CD11b- cDCs in lymphoid tissues and CD103+

CD11b- cDCs in nonlymphoid tissues (Figure 1-1)(Edelson et al., 2010; Hildner et al., 2008).

Furthermore, the Murphy group found that after specification of pre-CD8 DCs, BATF3 is

required to maintain a high level of Irf8 auto-activation via binding to IRF8 within an Irf8

superenhancer region (Grajales-Reyes et al., 2015). Thus, auto-activation of the Irf8 gene,

promoted by BATF3, maintains the CD8+CD11b- cDC lineage. Lack of Batf3 leads to a decay

of IRF8 levels, and the CD8+CD11b- cDC lineage diverts to the CD8- CD11b+ cDC lineage

(Shortman, 2015).

• Others (Id2 and BCL-6)

Mice deficient in the helix-loop-helix (HLH) transcription factor inhibitor of DNA binding 2

(Id2) exhibit markedly reduced splenic CD8+ DCs as well as epidermal LCs (Hacker et al.,

2003). In nonlymphoid tissues such as the intestinal lamina propria (LP), CD103+CD11b- DCs

express high level of Id2, and its absence blocks the development of CD103+ CD11b- DCs

(Ginhoux et al., 2009). These data suggest Id2 plays an important role in the development of

CD8+ and CD103+ CD11b- DC subset.

Recently, comparative analysis of transcriptomes identified transcriptional repressors B-cell

CLL/Lymphoma 6 (BCL-6) in the specification of CD8+/CD103+CD11b- DCs (Watchmaker et

al., 2014). Butcher’s group found that CD8+/CD103+CD11b- DCs express higher levels of

12

BCL-6 compared to CD103+CD11b+ DCs, and Bcl6-deficient mice lack splenic CD8α+ DCs and

intestinal CD103+CD11b- DCs (Watchmaker et al., 2014).

1.2.2.3 Transcription factors affecting the CD8-CD11b+ DCs and CD103+CD11b+ DCs

• IRF4

Similar to IRF8, IRF4 interacts with PU.1 and binds to a composite PU.1/IRF DNA motif, a

sequence element containing adjacent PU.1 and IFN sequence response elements (ISRE) motifs

(Eisenbeis et al., 1995; Matsuyama et al., 1995). IRF4 is expressed in lymphoid and myeloid

compartments. Moreover, expression of IRF4 in B and T cells is essential for their function

(Marecki et al., 1999; Mittrucker et al., 1997).

Mice lacking the Irf4 have selective defects in splenic CD8-CD11b+ DCs (Suzuki et al., 2004;

Tamura et al., 2005). In nonlymphoid tissues such as the skin, IRF4 has recently been implicated

in regulating CCR7 expression on CD11b+ dermal DCs and their subsequent migration to skin-

draining LNs (Bajaña et al., 2012). In the intestine, IRF4 is important for the homeostasis of

CD103+CD11b+ cDCs at the post-precursor stage, and appears to affect cDC survival and

migration (Persson et al., 2013b; Schlitzer et al., 2013).

• RBP-J

The Notch signaling pathway is an evolutionarily conserved mechanism that regulates the

development of multiple cells and tissues (Bray, 2006). The interaction of Notch with its ligand

on a neighboring cell causes receptor cleavage that releases that intracellular domain of Notch

(ICN). ICN translocates into the nucleus and binds the transcription factor CSL (CBF1-

suppressor of hairless-Lag1), the mouse homologue of which is RBP-J (recombination-signal-

binding protein-J). The resulting ICN-RBP-J complex recruits coactivators and activates Notch-

dependent gene expression programs.

In the adaptive immune system, Notch-RBP-J signaling is essential for the commitment to and

early development of the T cell lineages, generation of marginal zone (MZ) B cells and

specification of effector T cell function (Maillard et al., 2005). In terms of DC development,

13

Notch-RBP-J signaling is essential for DC homeostasis. In particular, splenic CD8-

CD11b+endothelial cell-selective adhesion molecule (ESAM)hi DCs require Notch2 signals for

their survival and persistence in the MZ, and intestinal CD103+CD11b+ DCs require Notch2 for

their homeostasis (Caton et al., 2007; Lewis et al., 2011). Moreover, splenic Notch2-dependent

CD8-CD11b+ESAMhi DCs are selectively dependent on signaling from the lymphotoxin beta

receptor (LTR) (Satpathy et al., 2013), suggesting that Notch2 and LTR may act in sequence,

perhaps with Notch2 promoting DC interactions with cells in the MZ that produce LT12

(Murphy, 2013).

• KLF4

Zinc-finger transcription factor Kruppel-like factor 4 or KLF4 can act as a repressor or activator

of transcription and regulates cell proliferation/differentiation in the skin (Segre et al., 1999) and

the intestine (Katz et al., 2002; Kuruvilla et al., 2016). In terms of DC development, CD11c-Klf4-

/- conditional knockout (KO) mice have reduced CD11b+ cDCs in spleen (Park et al., 2012).

Moreover, in nonlymphoid tissues, KLF4 regulates the development of a subset of IRF4-

expressing CD103+CD11b+ cDCs that are required for normal priming of Th2 cell responses

(Tussiwand et al., 2015).

• Others (RelB and Blimp-1)

RelB, an NF-B family transcription factor, is expressed strongly in CD8- DC but only weakly

in CD8+ DC (Wu et al., 1998). Relb-/- mice display a severe reduction of DCs associated with

profound myeloid expansion, suggesting an essential role for RelB in the development of CD8-

CD11b+ DCs (Briseno et al., 2017; Wu et al., 1998).

B lymphocyte–induced maturation protein-1, Blimp-1 (encoded by gene Prdm1), which is

known for its role in regulating plasma cell differentiation and T-cell homeostasis and function,

is also required for DC homeostasis. By conditionally deleting Prdm1 in the pan-hematopoietic

lineage (by using Tie2-Cre), Chan et al found a selective expansion of CD8- DCs in the spleen

and the peripheral LNs (Chan et al., 2009). However, a more restricted depletion of Prdm1in the

CD11c+ compartment revealed a loss of CD103+CD11b+ DCs in the intestinal LP and the

14

mesenteric LNs (MLNs), with no accompanying DC defects in the spleen or peripheral LNs

(Watchmaker et al., 2014). More work is needed to explain this discrepancy.

1.2.3 Tools for studying DC biology

In addition to culture systems, numerous models of constitutive and inducible DC depletion have

been generated and used to identify specific functions of DC subsets. There are three main

categories of mouse models: DTR/DTA based models, Cre/flox based models and transcription

factor knockout mice. Below are summaries of in vitro DC culture systems as well as commonly

used mouse models to study DC biology.

1.2.3.1 In vitro culture systems

BM-derived DCs (BMDCs)

The relative number of DCs in vivo is low compared with most other lineages, and isolation of

sufficient numbers for the clinic or for comprehensive in vitro studies can be logistically

burdensome. Therefore, the majority of applications rely on the in vitro generation of DCs from

blood monocytes, CD34+ progenitors or BM cells with the appropriate hematopoietic growth

factors, usually GM-CSF plus IL-4 (Inaba et al., 1992; Sallusto and Lanzavecchia, 1994) or

FLT3L (Brasel et al., 2000; Naik et al., 2005). It has been reported that the GM-CSF/IL-4

BMDCs are larger and more granular and they produce more inflammatory mediators including

TNF, iNOS and CCL2 (a chemokine that attracts monocytes or basophils) upon TLR ligation.

FLT3L BMDCs tend to migrate more efficiently to draining LNs after subcutaneous injection

(Xu et al., 2007). These data suggest that the GM-CSF/IL-4 BMDCs are the approximate

equivalent of inflammatory moDCs whereas FLT3L BMDCs better represent cDCs.

1.2.3.2 Depletion of DCs by DTR/DTA systems

The first models of genetic ablation of cell lineages were transgenic mice in which cell-type-

specific promoters drive expression of the diphtheria toxin (DTx) A-chain, or DTA (Breitman et

al., 1987; Palmiter et al., 1987). This toxin disrupts protein translation by catalyzing adenosine

diphosphate (ADP)-ribosylation of poly-peptide chain elongation factor 2 and eventually leads to

cell death (Honjo et al., 1968; Robinson et al., 1974). While DTx efficiently ablates the cells in

15

which it is expressed, it can be problematic as even low levels of off-target expression can lead

to death of unintended cells or even to effects on embryogenesis or morphogenesis (Breitman et

al., 1990). Subsequent mouse models overcame this problem by expressing the human or simian

diphtheria toxin receptor (DTR) under the control of a cell-type-specific promoter, with

subsequent administration of DTx to deplete cells of interest in mice (Saito et al., 2001). As the

mouse ortholog of the DTR is orders of magnitude less sensitive to DTx, this allows for the

efficient depletion of only DTR-expressing cells and has the added benefit of allowing for

inducible depletion of these target cells rather than constitutive ablation (Durai and Murphy,

2016).

Historically, the first DTR-based model used for DC depletion was the Itgax-DTR (or CD11c-

DTR) strain, a transgenic mouse line expressing a DTR-eGFP fusion protein under the control of

the murine Itgax promoter (Jung et al., 2002). DTx administration completely depletes CD11c+

cells within 24 hr and DCs begin to reappear 3 days after DTx treatment. While this strain was

vital for early work confirming the functions of DCs in T-cell priming, several limitations have

emerged. First, repeated administration of DTx is lethal to these mice, restricting the

maintenance of DC depletion status. This lethality is likely due to off-target expression of the

DTR transgene in radio-resistant cells. This limitation can be overcome by generating chimeras

of CD11c-DTR BM into WT recipients, which tolerate repeated DTx treatment (Zammit et al.,

2005). The second caveat derives from CD11c expression by non-DCs and the depletion by DTx

of such cells, which include macrophages (Probst et al., 2005), activated CD8+ T cells (Jung et

al., 2002) and plasmablasts (Hebel et al., 2006). Ablation of these cells complicates analysis with

this strain, because phenotypes observed might result from their depletion rather than that of

DCs, and other methodologies should be used to validate findings. Lastly, CD11c-DTR mice

have been reported to display neutrophilia and monocytosis upon DTx injection (Tittel et al.,

2012; van Blijswijk et al., 2013), and reduced LN cellularity is found in CD11c-DTR (and other

DTR strains) without DTx treatment (van Blijswijk et al., 2015).

The recently developed Zbtb46-DTR mouse allows for more specific depletion of cDCs, since

ZBTB46 expression is restricted to cDCs but not other mononuclear phagocytes (Meredith et al.,

2012a). However, ZBTB46 has been shown to be expressed on radio-resistant endothelial cells,

resulting in the death of Zbtb46-DTR mice upon DTx treatment (Satpathy et al., 2012).

16

Therefore, transplant of Zbtb46-DTR BM into lethally irradiated WT recipients is required for

cDC depletion without affecting mouse viability.

Several DTA/DTR strains allow for depletion of specific DC subsets. Kaplan et al have

generated huLangerin-DTA mice using a bacterial artificial chromosome (BAC) in which an

internal ribosome entry site (IRES, an RNA element that allows for translation initiation)-DTA

was inserted into the 3’ untranslated region (UTR) of the human CD207 gene that codes for

langerin, a C-type lectin (CLEC) expressed on epidermal LCs and dermal CD8+ cDCs (Kaplan

et al., 2005). This strain has constitutive ablation of LCs as well as CD103+CD11b+ cDCs in the

small intestinal LP and MLNs (Welty et al., 2013). Clec4a4-DTR mice allow for ablation of

CD103+CD11b+ cDCs in the intestinal LP and the MLNs while depletion of these DCs in other

organs remains to be determined (Muzaki et al., 2016). On the other hand, the Clec9a-DTR

mouse (Muzaki et al., 2016) and the Xcr1-DTR mouse (Yamazaki et al., 2013) have been

generated to specifically deplete CD8+ cDCs.

1.2.3.3 Cre strains for conditional deletion of genes in DCs

The Cre-loxP system allows for conditional deletion of genes in specific cell lineages. In this

system, two 34-bp loxP sites are inserted on either side of a gene or exon, which is then said to

be “floxed”. Cell type-specific promoters are then used to express the bacteriophage P1 cre gene,

which encodes an integrase that mediates recombination between two adjacent loxP sites,

leading to deletion of the intervening DNA and inactivation of the floxed gene in Cre-expressing

cells (Sauer, 1998).

A number of Cre lines have been constructed to delete genes in DCs and DC subtypes. The

CD11c-cre strain was first widely used for depleting a gene of interest in DCs. However, similar

to the CD11c-DTR mice, CD11c-cre is active in non-DC populations. Recently, a Zbtb46-cre

line was generated by the Nussenzweig group allowing for more specific deletion of genes in

cDCs (Loschko et al., 2016). Whether Zbtb46-cre is as active in endothelial cells (as is the case

for Zbtb46 driven DTR expression) is not known. A Cre line useful for targeting particular DC

subsets is the Xcr1-cre strain, which allows for gene depletion in CD8+ cDCs (Ohta et al.,

17

2016). A Cre line for targeting CD11b+ cDCs has not been generated due to the heterogeneity of

this DC subset.

1.2.3.4 Transcription factor-based depletion of DCs

As mentioned in previous sections, transcription factors tightly control the development of DCs

and cDC subsets. Particularly, several transcription factors have been identified whose deletion

selectively depletes specific subsets of cDCs, and deletion of these transcription factors in knock-

out mice provides a useful means for studying DC function. Below is a summary of commonly

used mouse models in the field (Table 1-2).

Table 1-2 Comparison of different transcription factor knockout mice

Strain Cells depleted Caveats References

Irf8-/- CD8+ cDCs, monocytes,

pDCs, CD103+ cDCs in

nonlymphoid tissues

Myeloid neoplasia

eventually results

with age

(Aliberti et al., 2003;


Schiavoni et al.,

2002)

Irf8fl/fl Zbtb46-

cre

Irf8fl/fl CD11c-

cre

CD8+CD11b- DCs and

CD103+CD11b- cDCs in

nonlymphoid tissues

Deficient in some

intraepithelial

lymphocyte subsets

(Esterházy et al.,

2016; Luda et al.,

2016)

Id2-/- CD8+CD11b- cDCs, NK

cells, ILCs

Deficient in LTi cells

and lymphoid tissue

development

(Ginhoux et al.,

2009; Hacker et al.,

2003; Yokota et al.,

1999)

Batf3-/- CD8+CD11b- DCs and

CD103+CD11b- cDCs in

nonlymphoid tissues

This cDC subset may

develop in certain

infections in the

Batf3-/- mice

(Edelson et al., 2010;

Hildner et al., 2008;

Tussiwand et al.,

2012)

Relb-/- CD8-CD11b+ESAMhi

cDCs in the spleen

Fatal multiorgan

inflammation

(Briseno et al., 2017;

Burkly et al., 1995)

Notch2fl/fl

CD11c-cre

CD8-CD11b+ESAMhi

cDCs in the spleen, part

of CD103+CD11b+ cDCs

in the intestine

Total splenic DCs are

reduced; increased

CD103+CD11b- cDCs

in the intestine

(Lewis et al., 2011;

Satpathy et al., 2013)

18

Irf4l/fl CD11c-cre CD8-CD11b+ cDCs in

the LNs, part of

CD103+CD11b+ cDCs in

the lung and intestine

May affect cDC

functions

(Persson et al.,

2013b; Schlitzer et

al., 2013)

Klf4fl/fl CD11c-

cre

Part of CD8-CD11b+

cDCs in the LNs, part of

CD103+CD11b+ cDCs in

nonlymphoid tissues

(Park et al., 2012;

Tussiwand et al.,

2015)

1.3 Effect of age on DC phenotype

Newborn immune cells are qualitatively distinct from adult cells. Subsets of cells are present in

different proportions in neonates and adults and, among cells of the same subtype, phenotypic

differences have been described. Historically, the function of neonatal adaptive immune cells has

been considered to be immature. However, it is now clear that neonates are competent, under

certain circumstances, to mount adult-level T-cell responses in vivo (Forsthuber et al., 1996;

Ridge et al., 1996; Sarzotti et al., 1996).

Several studies have shown that the absolute number of DCs in neonatal mice is reduced by

several logs compared with adults (Dadaglio et al., 2002; Dakic et al., 2004; Sun et al., 2003).

Studies of neonatal DCs mainly focused on lymphoid tissues such as the spleen, thymus and BM.

In the spleen, a higher percentage of CD4-CD8+ cDCs and a lower percentage of CD4+CD8-

cDCs was found in neonatal mice (Dakic et al., 2004; Sun et al., 2003). Functionally, DCs in

neonatal mice have mature properties under certain conditions. For example, injection of

neonatal mice with CpG oligonucleotides (TLR9 ligand) results in the upregulation of MHCII,

CD40 and CD86 expression by CD11c+ DCs in situ. Moreover, adoptively transferred CD11c+

cells from neonatal mice can promote strong cytotoxic lymphocyte responses to lymphocytic

choriomeningitis virus (LCMV)-derived peptide in adult hosts (Sun et al., 2003). Additionally, it

has been shown that DCs expanded by injection of FLT3L in neonates can improve type I IFN

antiviral responses and IL-12-associated antibacterial immune defense (Vollstedt et al., 2003).

19

In nonlymphoid tissues such as the lung, distribution of DC subsets skews towards CD103+ DCs

in the neonatal period, and then CD11b+ DCs rapidly catch up and reach a balance with CD103+

DCs (Ruckwardt et al., 2014). Neonatal lung DCs (CD103+ and CD11b+ subsets) display a

reduced expression of co-stimulatory molecules CD86 and CD80. Moreover, signaling through

CD28 may differentially impact epitope-specific CD8+ T cells responses, and reduced expression

of CD80 and CD86 on neonatal DC constitutes one mechanism by which neonatal mice establish

an epitope hierarchy that is distinct from that of adults upon respiratory syncytial virus infection

(Ruckwardt et al., 2014).

In the intestinal compartment, neonatal mice exhibit a marked deficit in CD103+ DCs during the

first week of life, perhaps due to weak production of chemokines by neonatal intestinal epithelial

cells. The relative paucity of CD103+ DCs in the neonatal intestine contributes to the high

susceptibility to intestinal infection in neonates. For example, in the neonatal period, CD103+

DCs are key players in the innate control of Cryptosporidium parvum (a zoonotic protozoan

parasite) infection in the intestinal epithelium via the production of IL-12 and IFN (Lantier et

al., 2013). However, the role of neonatal DCs in controlling viral infections in the intestine

has not been well investigated. The role of DCs in the intestine of neonatal mice challenged

with virus will be examined in Chapter 3.

1.4 The role of DC in intestinal health and disease

1.4.1 The intestinal environment and barrier function

The mammalian intestine is a complex environment that is constantly exposed to antigens

derived from the microbiota and food that are present at very high density within the intestinal

lumen. With an estimated composition of 100 trillion cells, human symbionts outnumber host

cells and express at least 10-fold more unique genes than their host’s genome (Ley et al., 2006).

These complex communities of microbes that include bacteria, fungi, viruses, and eukaryotes

such as protozoa and helminths, provide tremendous metabolic capability and play an important

mutualistic role with the host physiology (Belkaid and Hand, 2014).

To maintain homeostasis in light of these constitutive challenges, the gut has evolved physical

and immunological strategies to prevent aberrant inflammation and achieve host-microbial

20

mutualism. The physical barrier is composed of the mucus and the glycocalyx of epithelial cells,

and the single layer of epithelial cells that form a continuous cell sheet interconnected by tight

junctions (Hansson, 2012).

• Mucus layer

Mucus is a highly regenerative lubricating glycoprotein sheet secreted by goblet cells that covers

the mucosal surface and protects epithelial cells against chemical, enzymatic, microbial, and

mechanical insult. In the small intestine the mucus is discontinuous, but in the stomach and large

intestine there are two layers, a relatively thin inner layer attached to the epithelium and a thicker

loose layer, which microbes can inhabit. Moreover, mucus provides a matrix for secretory IgA

and a rich array of antimicrobial molecules (e.g., regenerating islet-derived protein III (RegIII) ,

RegIII, defensin and lysozymes) secreted by enterocytes and Paneth cells, which continuously

trap and expel microorganisms to discourage intestinal colonization and invasion by pathogens.

Underneath the mucus layer, epithelial cells present a dense forest of highly diverse

glycoproteins and glycolipids, which form the glycocalyx (Linden et al., 2008). Together, the

mucus and the glycocalyx are constantly renewed and have the potential to rapidly adjust to

changes in the environment. In patients with inflammatory bowel disease (IBD) and colon

cancer, an altered mucus profile has been observed (Larsson et al., 2011; Rhodes, 1996).

Moreover, spontaneous colitis and colorectal cancer develops in mice that lack specific mucin

genes (Fu et al., 2011; Heazlewood et al., 2008; Van der Sluis et al., 2006; Velcich et al., 2002).

• Epithelial layer

Mucosal epithelial cells form a contiguous lining that acts as a barrier between the luminal

environment of the intestine and the interior of the host. Key to this barrier, the epithelial cell

plasma membrane is impermeable to most hydrophilic solutes in the absence of specific

transporters. Next, the paracellular pathway between cells is tightly sealed. Sealing is mediated

by apical junction complexes (tight junctions and adherens junctions) (Turner, 2009). Altered

junction complexes can cause a loss of a differentiated polarized phenotype of enterocytes

(Hermiston and Gordon, 1995a), increased gut permeability (or a “leaky” gut) (Laukoetter et al.,

2007) and gut inflammation (Clayburgh et al., 2005; Hermiston and Gordon, 1995b; Schmitz et

al., 1999).

21

In addition to epithelial cells, intraepithelial lymphocytes (IELs) reside interspersed among these

cells. In mice, IELs represent up to half the number of T cells in the organism (Rocha et al.,

1991). IELs are composed of conventional T-cell receptors (TCR) cells expressing the CD4

or the heterodimer CD8 co-receptors, and unconventional TCRcells and TCRcells

expressing CD8(Cheroutre et al., 2011. Intestinal IELs can exert beneficial roles in

preserving the integrity of the mucosal barrier and in preventing pathogen entry and spreading.

For example, T cells play a major role in limiting the entrance of commensal bacteria after

epithelial injury via the release of antimicrobial peptides (AMPs) that are induced by the

microbiota (Ismail et al., 2009). Conversely, intestinal IELs can also contribute to immune

pathology and initiate and/or exacerbate inflammatory diseases, such as IBD and Celiac disease

(Simpson et al., 1997; Sollid, 2004).

1.4.2 Anatomy of the intestinal wall

The gastrointestinal tract is composed of four layers: the innermost layer is the mucosa,

underneath this is the submucosa, followed by the muscularis propria and finally, the outermost

layer- the serosa (Figure 1-2). The structure and composition of these layers varies in different

regions of the digestive tract, depending on their function. For example, small intestinal

epithelial cells in the mucosa have finger-like projections or villi, which extend into the lumen to

maximize the surface area for nutrient absorption. In the large intestine, whose primary function

is water absorption, villi structures are absent.

The innermost mucosa layer of the gastrointestinal tract is composed of three layers: a single

layer of epithelium, connective tissue (LP) and a thin layer of muscularis (muscularis

mucosa)(Figure 1-2). The epithelium consists mostly of absorptive enterocytes, although

specialized secretive cells (e.g. mucus-secreting goblet cells and AMP-secreting Paneth cells),

intestinal stem cells and IELs can be found. The LP provides vascular support for the epithelium

and contains mucosal glands. A large number of lymphocytes reside in the LP and maintain

intestinal homeostasis. The third layer is the muscularis mucosa, which is a thin layer of smooth

muscle supporting the local movement of the mucosa.

The submucosa is a loose connective tissue layer, with transecting larger blood vessels, draining

lymphatics, and nerves, and can contain mucus secreting glands. The next muscularis propria is

22

composed of two layers: an inner circular and an outer longitudinal layer of smooth muscle

layers. These layers of smooth muscle maintain the contraction of rhythmic waves, which help to

move food down through the gut. The outermost layer serosa is covered by the visceral

peritoneum, functions as a protective barrier and is composed of avascular connective tissue and

simple squamous epithelium.

Figure 1-2 Layers of the intestinal wall

The figure represents the schematic layout of the intestinal wall.

1.4.3 Lymphoid structures in the intestine

The organized structures of the gut-associated lymphoid tissue (GALT) and the draining LNs are

the principal locations for priming adaptive immune cells in the intestine. Conversely, effector

immune cells are diffusely distributed throughout the LP and the overlying epithelium.

The GALT is comprised of Peyer’s patches (PPs) (located on the antimesenteric side of the small

intestine), caecal patches (around the ileocaecal valve) and colonic patches (located throughout

the colon and rectum), as well as smaller lymphoid aggregates (isolated lymphoid follicles or

ILFs, and cryptopatches or CPs) that are collectively termed solitary isolated lymphoid tissues

(SILTs) that distribute in both small and large intestines (Mowat and Agace, 2014).

As in other organs, DCs play an important role in immune defense against pathogens in the

intestine. DCs residing in the GALT and draining LNs (primarily mesenteric LNs, or MLNs)

have the unique property of establishing oral tolerance against food antigens and commensal

microbes. Disruption of this critical and delicate balance can result in devastating inflammatory

23

reactions such as hyper-reactivity to food components (e.g. Celiac disease (Meresse et al., 2012))

and IBD (Xavier and Podolsky, 2007). The features of the PPs, small intestinal LP and MLNs

are discussed below, together with the DC populations residing in each compartment.

Figure 1-3 Gut-associated lymphoid tissues in the small intestine

Small intestine contains the villi-crypt structure. Mucus layer that overlays the intestinal

epithelium layer provides a matrix for IgA and AMPs, to keep the microbes at bay. Underneath

the epithelium layer is the lamina propria, which is a reservoir for immune cells. Large organized

lymphoid tissues such as PPs and small lymphoid aggregates such as SILT can be found along

the small intestine. DCs located in the lamina propria constantly migrate to the draining LN (e.g.

MLNs) to prime tolerogenic or immunogenic immune responses. Gut-homing immune cells

traffic from the circulation to the intestine via HEV. PPs are covered by a specialized epithelium

called FAE, which contains M cells to transport luminal materials. Underneath the FAE is the

SED region, the B cell follicles and the IFR.

1.4.3.1 Peyer’s patches

PPs are large lymphoid structures built on a stromal scaffold, composed of aggregated lymphoid

follicles surrounded by the follicle-associated epithelium (FAE) that forms the interface between

the GALT and the luminal microenvironment. The FAE contains specialized cells named M (for

24

microfold) cells. These M cells transport luminal antigens and bacteria towards underlying

immune cells that inhibit or activate the immune response, leading to either tolerance or an

inflammatory immune response. Morphologically, PPs are separated into three main domains:

the follicular area, the interfollicular region (IFR) and the FAE (Figure 1-3)(Neutra et al., 2001).

The follicular area contains the PP lymphoid follicles with germinal center (GC) containing B

cells, follicular dendritic cells (FDCs -a mesenchymal cell that not related to DC) and

macrophages. The follicle is surrounded by the corona, or subepithelial dome (SED) containing

B cells, T cells, macrophages and DCs. DCs can also be found in the FAE zone (Jung et al.,

2010).

DCs in the PPs were first isolated and defined by Steinman and Cohn (Steinman and Cohn,

1973). Later on, distinct subsets of DCs based on their surface marker expression and

localization have been identified in PP (Iwasaki and Kelsall, 2000, 2001). All the subsets express

CD11c and MHCII but differ based on their expression of CD8 and CD11b. The

CD8+CD11b- DCs are localized within the T-cell rich IFR, while the CD8-CD11b+ DCs are

present under the FAE in the SED (Iwasaki and Kelsall, 2000). A third subset, which is CD8-

CD11blo/- (double negative, DN) DC, is also present in the SED, the IFR and within the FAE of

the PP (Figure 1-3) (Iwasaki and Kelsall, 2001). Interestingly, this DN DC subset can take up

viral antigen from infected apoptotic enterocytes for presentation to CD4+ T cells following

reovirus infection (Fleeton et al., 2004). The functions of different DC subsets are described in

Table 1-1.

The distribution of PP DC subsets is controlled by the chemokines expressed within the PP. It

has been shown that all PP DC subsets express CCR7, while its ligands CCL19 and CCL21 are

secreted by the fibroblast reticular cells (FRCs) located in the IFR and thus are chemotactic to all

PP DCs (Iwasaki and Kelsall, 2000; Link et al., 2007). It has been shown that only CD8-

CD11b+ DCs express CCR6 in addition to CCR7 and migrate toward its ligand CCL20, secreted

by the FAE overlying the SED (Cook et al., 2000; Iwasaki and Kelsall, 2000). Additionally,

CCL9, the ligand for CCR1, is secreted by FAE but not the villus enterocytes, and attracts

CD8-CD11b+ DCs toward the FAE (Zhao et al., 2003). Thus, under steady state, CCR6+ DCs

(CD8-CD11b+ DCs) are found only in the SED, but they can migrate to the FAE following oral

25

infection with Salmonella (Salazar-Gonzalez et al., 2006) or rotavirus (Lopez-Guerrero et al.,

2010).

1.4.3.2 The small intestinal lamina propria

The small intestinal LP (SILP) is located under the lining of the intestinal epithelial cells and is

enriched in lymphocytes and myeloid cells. Lymphocytes and mononuclear phagocytes in the LP

have been notoriously difficult to isolate and can only be retrieved after extensive enzymatic

digestions. On top of that, classifying mononuclear phagocyte subsets according to surface

marker profiles has been a challenge for a long time. Bona fide DCs in the intestinal LP are

CD11c+MHCII+, lack the expression of macrophage-associated markers CD64 and F4/80, and

express the transcription factor ZBTB46. Three main DC subsets have been identified in mouse

intestinal LP and they are classified on the basis of their expression of CD103 and CD11b.

Interestingly, there are marked differences in the ratio of CD103+CD11b+ and CD103+CD11b-

DCs along the length of the mouse intestine, with CD103+CD11b+ DCs making up the majority

of DCs in the SILP, but being rare in the colonic LP. By contrast, CD103+CD11b- DCs are the

major CD103+ DC subset in the colonic LP, and they are also enriched in small intestinal GALT

compared with their CD103+CD11b+ DC counterparts (Denning et al., 2011; Mowat and Agace,

2014; Persson et al., 2013a). The DC subsets in different regions of the intestine may contribute

to maintaining the balance between tolerogenic and proinflammatory immune responses (such as

the balance between Treg and Th17). The functions of different DC subsets can be found in

Table 1-1.

1.4.3.3 Mesenteric lymph nodes

MLNs, duodenopancreatic LNs (buried in the pancreas) and caudal LN (alongside the posterior

mesenteric artery) drain different segments of the intestine (Carter and Collins, 1974; Mowat and

Agace, 2014). Collectively, they are the largest LNs in the body. The development of MLNs is

distinct from PPs and other LNs, as a lack of TNF, TNFR, LT does not abolish the presence

of MLNs (Alimzhanov et al., 1997). It is proposed that these factors might have complementary

roles in MLN development (Spahn et al., 2002), and furthermore, LIGHT (homologous to

lymphotoxin, inducible expression, competes with herpes simplex virus (HSV) glycoprotein D

for HSV entry mediator, a receptor expressed on T lymphocytes) might also provide a substitute

26

for LT deficiency (Scheu et al., 2002). Accumulation of lymphocytes in the MLNs requires

both L-selectin and 47 integrin adhesion molecules (interacts with peripheral node addressin

(PNAd) and mucosal vascular addressin cell adhesion molecule 1 (MAdCAM-1), respectively),

which normally direct lymphocytes homing to peripheral and mucosal tissues, respectively

(Wagner et al., 1998).

Both migratory DCs and resident DCs can be found in the MLNs. Migratory DCs are

CD11c+MHCIIhi with an immature phenotype, while resident DCs are CD11c+MHCII+ with a

mature phenotype (Satpathy et al., 2012; Shortman and Naik, 2007). After activation, intestinal

DCs carrying luminal antigens migrate to MLNs and present these antigens to naïve T cells

(Figure 1-3). MLN DCs are important for the generation of Tregs (oral tolerance) (Matteoli et al.,

2010; Spahn et al., 2002), the class switch of B cells to IgA (indirect via transforming growth

factor beta (TGF secreted by Tregs), and induce the expression of gut-homing molecules

CCR9 and 47 on T cells and B cells (Stagg et al., 2002).

1.4.4 Regulation of intestinal homeostasis by DCs

1.4.4.1 Sampling and antigen uptake from the intestinal lumen

DCs can pick up antigen that has been transported across the intestinal epithelium through

various different routes as outlined in the following (Schulz and Pabst, 2013): 1) in the PPs, DCs

can uptake luminal antigens transported into PP by specialized M cells that are present in the

FAE (Mabbott et al., 2013); 2) goblet cells can function to shuttle low-molecular weight soluble

antigens to CD103+ DCs in the LP (McDole et al., 2012); 3) after capturing soluble antigen in

the gut lumen, CX3CR1+ macrophages transfer antigen to CD103+ DCs via a gap junction-

mediated mechanism (Mazzini et al., 2014); and 4) in certain inflammatory settings, CD103+

DCs can be recruited from the LP to insert their dendrites through the tight junctions between

enterocytes to directly sample the luminal contents (Farache et al., 2013; Jaensson et al., 2008).

Currently it is still unknown whether in the steady state these routes direct luminal antigens to

specific phagocytes and, as a consequence, fundamentally influence the nature of the immune

response directed to those antigens (Mabbott et al., 2013).

27

1.4.4.2 DC maturation

PAMPs are essential functional components of microorganisms that direct the targeted host cell

to distinguish ‘self’ from ‘non-self’ (‘stranger hypothesis’) and promote signals associated with

innate immunity (Janeway and Medzhitov, 2002). Major PAMPs are microbial nucleic acids,

including DNA (e.g. unmethylated CpG motifs), double-stranded RNA (dsRNA), single-

stranded RNA (ssRNA), as well as lipoproteins, surface glycoproteins, and cell wall components

(peptidoglycans, lipopolysaccharide (LPS) and glycosylphosphatidylinositol). DAMPs are cell-

derived molecules that can initiate and perpetuate immunity in response to trauma, ischemia,

cancer, and other settings of tissue damage in the absence of overt pathogenic infection (sterile

inflammation and ‘danger model’) (Matzinger, 2002; Tang et al., 2012). DAMPs include high

mobility group box 1 protein (HMGB1), heat shock proteins (HSPs), adenosine triphosphate

(ATP), self RNA and DNA. Both PAMPs and DAMPs are recognized by host PRRs (such as

TLRs, Nod-like receptors (NLRs) and retinoic acid-inducible gene I-like receptor (RLRs))

localized to the cell surface, the cytoplasm, and/or the intracellular vesicles.

Ligation of PRRs by PAMPs/DAMPs is the key signal to induce DC maturation. Indeed, DCs

are equipped with a battery of PRRs, including C-type lectin receptors (CLRs), mannose

receptors and TLRs (Akira et al., 2006; Geijtenbeek et al., 2004). Different DC subsets express

distinct sets of TLRs, which is likely to contribute to their functional specialization. For example,

intestinal CD103+CD11b+ DCs express TLR5 and TLR9 and produce proinflammatory cytokines

such as IL-23, IL-6 and IL-12 in response to flagellin and CpG stimulation, respectively

(Kinnebrew et al., 2012; Uematsu et al., 2008), whereas the CD103+CD11b- DCs express TLR3,

TLR7 and TLR9 and produce IL-6 and IL-12p40 but not TNF, IL-10 or IL-23 in response to

their respective TLR ligands (Fujimoto et al., 2011). In addition to PAMPs/DAMPs, exogenous

signals, such as inflammatory cytokines (e.g. TNF), CD40 ligand (CD40L) (Sallusto and

Lanzavecchia, 1994; Winzler et al., 1997), as well as by binding of complement-coated particles

through complement receptors, or antibody-coated particles through Fc receptors (Amigorena

and Bonnerot, 1999; Regnault et al., 1999) can also trigger DC maturation.

Immunogenic DC maturation is a complex process characterized by the acquisition of a number

of fundamental properties. Briefly, immature DCs triggered by PAMPs/DAMPs upregulate

28

MHCII and co-stimulatory molecules at the cell surface. These DCs migrate towards the T-cell

zones in the nearest draining LNs in a CCR7-dependent manner. These matured DCs present

antigen and prime antigen-specific naïve T cells in the T-cell zone, thus initiating the adaptive

immune response. In contrast to immunogenic maturation, a fraction of steady state DCs undergo

a constitutive maturation, termed “homeostatic maturation” (Lutz and Schuler, 2002) or “semi-

maturation” (Reis e Sousa, 2006). These DCs are believed to be tolerogenic. They display

processed self-peptide in order to delete self-reactive T cells that have escaped central tolerance

and maintain T-cell tolerance to innocuous environmental antigens through the generation of

inducible Treg.

After antigen acquisition from the intestinal lumen, mature DCs carry antigens to the MLNs for

T-cell priming (Liu and MacPherson, 1991, 1993). There are several lines of evidence that

support the concept that CD103+ DCs are the major DC subset that transports antigen from the

intestinal LP to MLNs. First, CD103+DCs are selectively reduced in the MLNs but not the LP of

CCR7-deficient mice (Johansson-Lindbom et al., 2005; Worbs et al., 2006). Second, confocal

imaging and flow cytometric analysis of intestinal lymph has demonstrated that the majority of

CD11c+ cells in the intestinal draining lymph are CD103+ DCs (Schulz et al., 2009). Finally, the

assessment of cannulated thoracic duct lymph from mice with mesenteric lymphadenectomy has

shown that CD103+CD11b+/- DCs (and a minor population of CD103- DCs) constitutively traffic

in intestinal lymph from the intestinal LP (Cerovic et al., 2013). Homeostatic migration of

intestinal DCs is apparently independent of TLR signaling and the commensal microbiota, and

may rely on a DC-inherent differentiation program and/or on a tonic release of low levels of

inflammatory cytokines in the intestine (Wilson et al., 2008). If antigen is acquired in the PP,

then DCs will migrate to the IFR to prime T cells.

1.4.5 T-cell priming and induction of adaptive immune responses by DC

Terminally differentiated or mature DCs that have downregulated antigen-sampling functions are

exceedingly potent at priming T cells. Once activated by DCs, these T cells can complete the

immune response by interacting with other cells, such as B cells for antibody formation,

macrophages for cytokine release, and cellular targets for lysis. To obtain antigenic peptide for

MHC presentation, APCs utilize three major systems:

29

MHC class II presentation pathway: This system is achieved by endocytosis and comprises a

large collection of proteases with variable substrate specificity and pH requirements.

Endocytosed proteins, whether self or foreign, endogenous or exogenous, are degraded by these

proteases in the endosomal compartments (Honey and Rudensky, 2003). In APCs, the resulting

peptides can be loaded into the peptide-binding groove of MHCII molecules and then presented

on the plasma membrane for recognition by CD4+ T cells.

MHC class I presentation pathway: The second major proteolytic system used by eukaryotic

cells is the proteasome, a multimeric complex found in the cytosol that is composed of several

proteolytic and regulatory subunits. The peptides generated by the proteasome can be

translocated by the transporter associated with antigen processing (TAP) into the endoplasmic

reticulum (ER), where they are loaded into the binding groove of newly synthesized MHC class I

molecules. The resulting MHC class I-peptide complex then follows the default secretory

pathway through the Golgi apparatus and is displayed on the plasma membrane for inspection by

CD8+ T cells.

MHC class I cross-presentation pathway: When APCs are not directly infected, they need to

acquire exogenous antigens from the infectious agent and present them on MHC class I

molecules by a third system, the cross-presentation pathway (Heath et al., 2004). A lot of work

has been done to understand the classic MHC class I and MHC class II antigen presentation

pathways, however, for this thesis, I will mainly focus on the cross-presentation pathway in the

following sections.

During cross-presentation, exogenous proteins are diverted from either the endosomal

compartment or directly from the extracellular fluid into the cytosol for processing in the

conventional MHC class I pathway (Heath and Carbone, 2001). This process has been found to

play out in the cross-presentation of HSPs (likely a receptor-mediated mechanism) (Srivastava et

al., 1994), antibody-mediated immune complexes (Regnault et al., 1999), exosomes (Wolfers et

al., 2001; Zitvogel et al., 1998), apoptotic cells (Albert et al., 1998), and particles absorbed by

macropinocytosis (Norbury et al., 1997). After phagocytosis, exogenous antigens can be

exported into the cytosol, where they are processed by the proteasome. The processed antigens

can then be loaded on MHC class I molecules in the ER or re-imported into the phagosome to be

30

loaded on MHC class I molecules (called the cytosolic pathway). Alternatively, exogenous

antigens can be directly degraded into peptides in the phagosome without the cytosolic

proteasome step, where they are then loaded onto MHC class I molecules (called the vacuolar

pathway) (Joffre et al., 2012).

Recently, it is believed that only some DC subsets can cross-present antigens efficiently. The

contribution of different DC subtypes to cross-presentation and cross-priming (the induction of

effector CD8+ T cells in vivo) varies depending on the experimental setting. Initially, CD8+

DCs were shown to be more efficient at cross-presentation than CD8- DCs in the steady state

(den Haan et al., 2000; Shortman and Heath, 2010), whereas both DC subtypes can present

antigens efficiently after receptor-mediated endocytosis (den Haan and Bevan, 2002). Other DC

subsets have also been shown to cross-present antigens efficiently: CD103+ cDCs are the most

efficient at cross-presentation within nonlymphoid tissues, such as the lungs (del Rio et al., 2007;

Desch et al., 2011), the skin (Bedoui et al., 2009) or the intestine (Cerovic et al., 2015).

1.4.5.1 Signal 1 and Signal 2

The fate of naïve T cells is determined by three signals that are provided by activated DCs. The

first signal results from the ligation of TCRs by peptide antigens presented by MHC class I or II

molecules on the cell surface of DCs, thus directing an antigen-specific response. Signal 1 alone

is thought to promote naïve T-cell inactivation by anergy, deletion or diversion into a regulatory

cell fate, thereby leading to tolerance/suppression. The second signal, termed co-stimulation, is

independent of the antigen receptor and is critical to induce full T-cell activation, sustain cell

proliferation, prevent anergy and/or apoptosis, induce differentiation to effector and memory

status, and allow cell-cell cooperation (Frauwirth and Thompson, 2002). CD28 is a classical co-

stimulatory molecule constitutively expressed on T cells (June et al., 1990). In conjunction with

a TCR signal, ligation of CD28 by its B7 family ligands CD80/CD86 will activate T cells

(Gimmi et al., 1991). Besides CD28, tumor-necrosis factor superfamily (TNFSF) members also

function at various stages of T cell differentiation to enhance T cell proliferation, survival or

effector function. CTLA-4 (cytotoxic T-lymphocyte-associated protein 4), a homologue of the

CD28 molecule that is induced on activated T cells, serves as a negative regulator of T-cell

activation and proliferation (Linsley et al., 1991). Thus, the net result of costimulation is

31

composed of a fine balance between positive and negative signals emanating from many

receptors.

1.4.6 Signal 3: Determining T-cell differentiation into an effector cell

Signal 3 refers to the APC-derived cytokines needed for a T cell to make a productive response

and avoid death and/or tolerance induction. The creation of a particular cytokine environment by

APCs during immunity is critical for the determination of the appropriate type of immune

response. IL-12 and type I IFN are the major types of signal 3 for naïve CD8+ T cells

differentiating into effector T cells (Curtsinger and Mescher, 2010). IL-12, IL-4, IL-6 and TGF

are signal 3 for Th1, Th2, Th17 and Treg cells, respectively (Zhu et al., 2010). The downstream

effects of these cytokines are mediated by transcription factor STAT family members (Zhu et al.,

2010). The upstream pathways regulating the production of IL-12 and type I IFN from DCs are

discussed below.

What signaling pathways regulate the production of IL-12 and type I IFN by activated DCs?

In the setting of helper T cell-dependent cytotoxic T lymphocyte (CTL) activation, the

CD40:CD40L signaling pathway is crucial for the production of IL-12 by DCs (DC licensing)

(Bennett et al., 1998; Koch et al., 1996; Ridge et al., 1998), whereas LTR signaling pathway

involves in the secretion of type I IFN by DCs (Summers-DeLuca et al., 2007; Summers deLuca

et al., 2011). It has been reported that IL-12 (downstream of CD40) and type I IFN induce

complex gene regulation programs that involve, at least in part, chromatin remodeling to allow

sustained expression of a large number of genes critical for CD8+ T cell function and memory

(Agarwal et al., 2009). However, it is unclear how CD40-induced IL-12 and LTR-induced type

I IFNs can distinctly and complementarily program short- and long-term gene expression in

CD8+ T cells.

In the setting of direct activation of CD8+ T cells by DCs that cross-present antigens, IL-12 can

be produced by DC whose TLR signaling is activated. Indeed, CD103+ DCs can express a

various pattern of TLRs and they can produce IL-12 upon TLR ligation (Fujimoto et al., 2011;

Uematsu et al., 2008). The source of type I IFN can be pDC, which is well-known for its

capacity to produce large amount of type I IFN upon viral infection (Cella et al., 1999). Whether

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DC-intrinsic LTR signaling plays a role in a small intestinal viral infection will be

explored in Chapter 4.

1.4.6.1 Expression of Lymphotoxin and LTR signaling

1) Lymphotoxin expression

Lymphotoxin (LT) and lymphotoxin (LT) are TNFSF members TNFSF1 and TNFSF3

respectively, and these two proteins form a membrane-bound heterotrimer LT12 and signals

through LTR. LT can also form soluble homotrimer LT3 which signals via TNFRI/II and

herpesvirus entry mediator (HVEM) (Figure 1-4). LT12 is expressed by cells of the

lymphocyte lineage including B cells, T cells, natural killer (NK) cells, lymphoid tissue inducer

(LTi) cells and retinoic acid- related orphan receptor (ROR) t+ ILC3 (Tumanov et al., 2011;

Ware et al., 1992). The LTR receptor is expressed by radio-sensitive cells such as macrophages

and DCs, and by radio-resistant cells including intestinal epithelial cells, FDCs, endothelial and

stromal cells (Figure 1-4) (van de Pavert and Mebius, 2010; Ware et al., 1995). LIGHT, another

member of TNFSF can also bind to LTR and HVEM as well as DcR3 (decoy receptor 3), a

TNF receptor family member lacking a transmembrane region competes with LTR and HVEM

for LIGHT engagement (Yu et al., 1999)(Figure 1-4).

2) LTR signaling

Upon LTR ligation, two NF-B activating pathways are engaged: the first involves rapid

initiation of classical NF-B signaling followed by more gradual activation of the alternative

pathway, and the second involves exclusive activation of alternative NF-B (Dejardin et al.,

2002). The first pathway leads to activation of p50 and RelA, which control expression of genes

such as vascular cell adhesion molecule 1 (VCAM-1), macrophage inflammatory protein (MIP)

1 and MIP-2. In addition, this pathway leads to an increase in protein levels of the NF-

B2/p100 precursor. The processing of the latter is controlled by a second pathway that involves

the activation of NF-κB-inducing kinase (NIK), which in turn activates IkB kinase (IKK for

the generation of active p52 (derived from the p100 precursor). p52 in association with its

partner (e.g. RelB) translocates to the nucleus and activates the transcription of genes implicated

33

in secondary lymphoid organogenesis and homeostasis such as CCL19, CCL21, CXCL13, and

B-cell activating factor (BAFF) (Figure 1-4) (Dejardin et al., 2002; Yilmaz et al., 2003).

Figure 1-4 The Lymphotoxin system and the LTR signaling pathway.

LT is expressed either as a soluble homotrimer that bind TNFRI and TNFRII and HVEM, or as

a membrane bound heterotrimer when co-expressed with LT. Both LT12 and LIGHT can

bind LTR, and LIGHT can additionally bind to HVEM and DcR3, a decoy receptor expressed

in humans. HVEM also binds two Ig superfamily members BTLA and CD160. Upon LTR

ligation, two NF-B activating pathways are engaged.

1.4.6.2 LTβR signaling is required for the organogenesis of secondary lymphoid tissues and the maintenance of lymphoid tissue microarchitecture

Primary immune responses are initiated in secondary lymphoid organs, including spleen, LNs,

and mucosal associated lymphoid tissues (MALTs). These tissues are situated throughout the

body, draining and sampling sites where antigens from pathogens are most likely to be

encountered. While spleen serves to sample blood-borne antigens, draining LNs collect antigens

34

from nonlymphoid organs via lymphatic vessels. MALTs, such as PPs, ILFs and nasal associated

lymphoid tissues (NALTs), lack afferent lymphatics and acquire antigen directly from the lumen.

All of these secondary lymphoid organs have specialized architecture and microenvironments

that promote the controlled interactions of immune cells in order to elicit a rapid and appropriate

immune responses to infectious agents (Fu and Chaplin, 1999; Randall et al., 2008).

Secondary lymphoid organs develop during embryogenesis or, as in the case of ILFs and NALT,

in the very early post-natal period. This process occurs at pre-determined sites throughout the

body independently of antigen or pathogen recognition, and involves complex interactions

between various hematopoietic, mesenchymal and endothelial cells (Mebius, 2003; Randall et

al., 2008; Ruddle and Akirav, 2009). The very first step of secondary LN formation involves the

production of retinoic acid (RA) from nerve fibers which induces the expression of CXCL13 by

neighboring mesenchymal cells. CXCL13 then attract LTi precursors from the blood to form

initial clusters. Clustering of pre-LTi facilitates signaling through receptor activator of NF-B

(RANK) expressed on LTi. This leads to the induction of LT12-expression by pre-LTi cells

which differentiate into mature LTi cells. Interaction of LT12-expressing LTi cells and LTR-

expressing mesenchymal cells results in their differentiation into stromal lymphoid tissue

organizing (LTo) cells. Upon interactions with LTi, LTo cells express chemokines (CXCL13,

CCL21 and CCL19), adhesion molecules VCAM-1, intercellular adhesion molecule 1 (ICAM-

1), MAdCAM-1 and cytokines (IL-7 and RANKL). These factors support the attraction and

retention of more hematopoietic cells, leading to LN development (van de Pavert and Mebius,

2010). In the absence LTR signaling, mice lack all LNs and PPs (Fütterer et al., 1998).

Constitutive LTR signaling regulates many aspects of immune tissue organization in adult

animals (Gommerman and Browning, 2003). LTR-deficient mice have disorganized splenic B

and T cell zones and lack a mature FDC network (Fütterer et al., 1998; Rennert et al., 1996).

Follicular B cell expression of LT12 maintains differentiation of FDC and induces their

expression of CXCL13 and adhesion molecules, thereby enabling recruitment and retention of B

cells into the follicle (Endres et al., 1999; Fu et al., 1998; Ngo et al., 1999). CXCL13 binding to

CXCR5 can promote up-regulation of LT12 on homing B cells to establish an LTR-

CXCL13 feedback loop for the maintenance of FDC networks and intact B cell follicles (Ansel

35

et al., 2000). Moreover, in mice lacking LTR signaling due to genetic deficiency or

pharmacologic inhibition, MAdCAM-1 and VCAM-1 expression is absent and the MZ is devoid

of metallophilic macrophages, B cells and MZ macrophages (Mackay et al., 1997). In addition,

LTR signaling on HEV regulates expression and localization of MAdCAM-1 and PNAd

(Drayton et al., 2003). Thus, mice in which LTR signaling is blocked have reduced peripheral

LN cellularity due to impaired B cell and T cell homing to LNs (Browning et al., 2005).

1.4.6.3 LTβR signaling in regulating immune responses

The role of LTR signaling in regulating adaptive immune responses is summarized below:

1) LTR signaling in DC homeostasis

LT-deficient mice exhibit a marked reduction in DC numbers in the steady state (Wu et al.,

1999), specifically the CD8-CD11b+ DC subset in lymphoid tissues (Kabashima et al., 2005).

The requirement of LTR in maintaining CD8-CD11b+ DCs is DC intrinsic, and signaling via

LTR is important for homeostatic DC proliferation (Kabashima et al., 2005). The splenic

CD8-CD11b+ DCs can be further divided into ESAMhi and ESAMlo subsets, and the ESAMhi

population is selectively lost when LTR or Notch2 signaling is specifically ablated in DCs

(Lewis et al., 2011). In nonlymphoid tissues, LTR is likely to play a non-redundant role in

homeostasis of intestinal LP CD103+CD11b+ cDCs (Satpathy et al., 2013). Moreover, it has been

reported that LTR is important for maintaining total DCs and CD8-CD11b+/- DCs in the PPs

(Reboldi et al., 2016) (Figure 1-5).

2) LTR signaling in T-cell responses

Consistent with an essential role for lymphoid organs in primary immune responses, LTR

deficient mice display a diminished antigen-specific CD8+ T cell response against some viruses

and intracellular bacteria. For example, both Lta-/- and Ltb-/- mice, whose splenic architecture is

abnormal, display an impaired CD8+ T-cell response (cytotoxicity and IFN production) in the

spleen and a delay in viral clearance upon LCMV Armstrong strain infection (Berger et al.,

1999; Suresh et al., 2002). Transient LTR blockade in New Zealand Black (NZB) mice also

diminishes the LCMV clone 13-specific CD8+ T-cell response; however, since activation of

36

CD8+ T cells in this model is lethal for the susceptible NZB mice, administration of LTR-Ig can

reverse LCMV clone 13 strain-induced disease mediated by CD8+ T cells (Puglielli et al., 1999).

In addition, Lta-/- mice infected with HSV-1 develop similar numbers of antigen-specific CD8+ T

cells, however, their cytotoxicity and cytokine-mediated effector functions are impaired resulting

in enhanced susceptibility to HSV-induced encephalitis (Kumaraguru et al., 2001). These results

suggest that the beneficial or deleterious roles of LTR signaling are context-dependent.

Since the initiation of a CD8+ T-cell response is regulated by DCs, it is reasonable to determine

whether LTR signaling affects the following parameters: 1) migration/retention of antigen-

bearing DCs to/within the draining LN, because signaling of LTR on DCs is important for DC

homeostasis in the spleen, LNs and PPs (Kabashima et al., 2005; Reboldi et al., 2016; Wang et

al., 2005; Wu et al., 1999); and 2) the licensing of LTR-expressing DCs by LT12-expressing

T cells. Previous work from our lab has shown that DC-intrinsic LTR signaling is required for

optimal expansion of antigen-specific CD8+ T cells specific for a model protein antigen via the

production of type I IFN (Summers deLuca et al., 2011), suggesting that signal 3 is fine-tuning

the CD8+ T-cell immune response (Figure 1-5). This LTR/type I IFN axis has been further

demonstrated to play a role in shaping the early CD8+ T-cell response to a nonreplicating self-

antigen in a diabetic mouse model (Ng et al., 2015). However, the type of DC that is primarily

responsible for LTR signaling in these models of T-cell activation is unclear. Moreover, we do

not know what role DC-intrinsic LTR signaling plays in clearance of intestinal viral

infection. Chapter 4 of my thesis addresses the latter question.

3) LTR signaling and B-cell responses

Loss of LTR signaling affects B cell response as well. As mentioned before, Lta-/- (Banks et al.,

1995; De Togni et al., 1994), Ltb-/- (Alimzhanov et al., 1997; Koni et al., 1997), Ltbr-/- (Fütterer

et al., 1998) or LTR-Ig treated (Mackay et al., 1997; Rennert et al., 1996) mice lack FDCs, and

exhibit abnormal secondary lymphoid organ architecture. Thus, it is not surprising that the

humoral immune response would be altered in these mice. Indeed, B cell affinity maturation and

class switching are impaired in the absence of LTR signaling (Banks et al., 1995; Fütterer et al.,

1998; Mackay et al., 1997; Reboldi et al., 2016). However, exceptions do exist. For example,

administration of a high dose of model antigen hapten 4-hydroxy-3-nitrophenyl acetyl (NP) to

37

Lta-/- mice results in a high-affinity anti-NP IgG1 response similar to WT mice (Matsumoto et

al., 1996), suggesting other signals might be involve in the affinity maturation of the NP

response. Secondly, Lta-/- mice generate comparable systemic humoral responses against murine

gammaherpesvirus 68 (MHV-68) compared to WT mice (Lee et al., 2000). Finally, a delayed but

almost fully induced anti-viral IgG response is observed in Lta-/- and Lta-/- WT chimeric mice

upon influenza challenge (Lund et al., 2002; Moyron-Quiroz et al., 2004). Our lab has shown

that in the LN of R- phycoerythrin-immunized Ltb-/- WT BM chimeras, GL7+Fas+PNA+

antigen-specific GC B cells form clusters situated in the follicle, suggesting GCs form normally

in the absence of LTR signaling. However, these GCs do not persist and affinity maturation of

the antigen-specific B cell response is ultimately impaired (Boulianne et al., 2013). Thus, the

reliance on the LT pathway for a productive GC response likely depends on many factors

including the nature and persistence of the antigen and the timing of readouts.

4) LTR signaling in mucosal IgA responses

Banks et al. first reported that in contrast to the similar levels of total serum IgG and IgM in Lt-

/- vs control littermates, the levels of both serum and fecal IgA is dramatically decreased in

unimmunized Lt-/- mice compared with those in WT littermates (Banks et al., 1995). At first it

was assumed that the defect in IgA in these mice was due to the absence of PP and/or MLNs.

However, Yamamoto et al. reported that PP-null mice (offspring from LTR-Ig treated pregnant

WT mice during gestation) possess significant numbers of IgA+ plasma cells in the intestinal LP

(Yamamoto et al., 2000), indicating PPs are not absolutely required for homeostatic intestinal

IgA responses. Moreover, WT BM reconstituted Lta-/- mice or Lta-/-Tnfa-/- mice with display

similar levels of serum IgA and normal numbers of intestinal IgA-producing cells compared with

WT control chimeras (Kang et al., 2002; Ryffel et al., 1998), suggesting even MLNs are not

absolutely needed for homeostatic IgA generation. In response to OVA plus cholera toxin (oral

immunization), PP-null mice mount comparable anti-OVA and anti-cholera toxin IgA response

compared to PP-sufficient mice, whereas Lta-/-Tnfa-/- mice fail to do so (Yamamoto et al., 2000).

In response to Salmonella infection, PP-null mice fail to induce antigen-specific intestinal IgA

antibodies (Hashizume et al., 2008). These results indicate that the requirement of PP in

generating antigen-specific intestinal IgA is context dependent. Additionally, the entry and

residence of B cell/plasma cell into the SILP is disturbed in Lta-/- mice, since the SILP of these

38

mice displays lower levels of the chemokines CXCL13 and CCL21 and vascular addressin

molecules MAdCAM-1 compared to WT controls (Kang et al., 2002; Newberry et al., 2002).

Thus, B cell/plasma cell migration from induction sites to effector sites is impaired in Lta-/- mice.

Taken together, multiple factors in Lta-/- mice contribute to the control of IgA levels.

However, in the presence of PPs, LTR-dependent DCs are found to be required for PP B cell

IgA class switch (Reboldi et al., 2016). RORt+ ILCs produce LT12, which is required for

CD11b+ DC maintenance in the SED. Deficiency in LTR-dependent DCs or RORt+ ILCs

results in reduced IgA+ B cell frequencies in PPs, suggesting LTR signaling is important for

homeostatic IgA responses. Furthermore, SED CD11b+ DCs are found to augment IgA switching

and express av8, an integrin that has an established role in converting TGF from its latent to

its active state and promoting B cell responses (Figure 1-5)(Reboldi et al., 2016).

1.4.6.4 LTR signaling in intestinal disease

The role of LTR signaling in the immune response against Citrobacter rodentium has been

extensively studied. C. rodentium is a murine-adapted mucosal pathogen that shares several

pathogenic attributes with the attaching and effacing enteropathogenic Escherichia coli and

enterohaemorrhagic E. coli, two clinically important human gastrointestinal pathogens (Collins

et al., 2014). C. rodentium infection is used to model several important human intestinal

disorders, including Crohn’s disease (CD) and ulcerative colitis (UC) (Higgins et al., 1999).

Following C. rodentium infection mice develop colitis, and this causes a pronounced dysbiosis

that is characterized by an overgrowth of C. rodentium and a consequent reduction in the

abundance and overall diversity of the resident microbiota (Lupp et al., 2007). Both innate and

adaptive immune responses are important in host defense against C. rodentium infection. Spahn

et al. first reported that mice lacking LTR signaling display increased severity of C. rodentium-

induced colitis, more severe weight loss and a higher burden of systemic C. rodentium when

compared to WT controls (Spahn et al., 2004). Subsequently, Wang et al. demonstrated that

RORt+ ILCs provide the relevant source of LT12 for signaling of LTR on intestinal

epithelial cells, resulting in neutrophil recruitment via the chemokines CXCL1 and CXCL2

during early C. rodentium infection (Wang et al., 2010). LTR in the radio-sensitive

compartment is also involved in the control of C. rodentium infection. Indeed, it has been

39

reported that C. rodentium infection triggers IL-23 production by Notch2-dependent colonic

DCs, which form a positive feedback loop between with RORt+ ILC3 through LTR signaling

(Figure 1-5) (Satpathy et al., 2013; Tumanov et al., 2011). IL-23 subsequently drives RORt+

ILCs to produce IL-22, which is required for the direct induction of AMPs, including RegIII

and RegIII, in colonic epithelial cells (Ota et al., 2011; Tumanov et al., 2011; Zheng et al.,

2008).

Figure 1-5 The role of LTR signaling in the periphery and the intestine

In the periphery, LTR signaling maintains DC homeostasis and promotes an optimal CD8+ T

cell response vis type I IFN. In the PPs, LTR signaling maintains DC homeostasis, which

impacts on IgA class switch. In the colon, LTR expression on both IEC and DCs are important

in host defense against C. rodentium infection.

The role of LTR signaling in host response against bacteria in the colon has been extensively

studied, however, its role in antiviral immunity in the gastrointestinal tract is not well known.

Therefore, Chapter 4 of this thesis will study the role of DC-intrinsic LTR signaling in host

defense against rotavirus, which is a small intestinal tropic virus. The background of this

specific viral infection will be discussed later.

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1.4.7 Fates of mucosal immune responses - Fate 1: Oral Tolerance

After priming by DCs, naïve T cells are activated and differentiate into tolerogenic (oral

tolerance) or immunogenic T cells, depending on the context. B cells are activated with or

without CD4+ T-cell help and differentiate into antibody-secreting cells. In this section, I discuss

the different fates of mucosal immune responses.

The intestinal immune system must discriminate between pathogens and harmless antigens such

as commensal microorganisms and dietary constituents. In the case of pathogens and other

harmful antigens, it is necessary to induce a strong and protective response, resulting in the

elimination of the threat. However, the usual response to harmless antigens or nutrients is to

induce tolerance which prevents unnecessary inflammation and hypersensitivity. The state of

hyporesponsiveness to fed antigen is known as oral tolerance (Scott et al., 2011).Intestinal DCs

are likely integral in ensuring that pathological immune responses to harmless antigens do not

develop. DCs that constitutively traffic out of the intestinal LP have been shown to deliver

antigen from both commensal bacteria and apoptotic epithelial cells to the MLNs (Huang et al.,

2000; Macpherson and Uhr, 2004).

The type of oral tolerance induced is related to the dose of antigen fed: clonal anergy/deletion

(high dose of antigen) or Treg induction (low dose of antigen) (Chen et al., 1995; Faria and

Weiner, 2005). It has been demonstrated that the GALT is a preferential site for the peripheral

induction of Foxp3+ Treg (Sun et al., 2007). Moreover, DCs, especially CD103+ DCs, from the

SILP and MLNs are significantly better than splenic DCs at inducing the expression of Foxp3 in

naïve T cells in the presence of exogenous TGF and RA (Table 1-1)(Coombes et al., 2007;

Mucida et al., 2007; Sun et al., 2007). Indeed, CD103+ DCs can metabolize RA and express

IDO, both features are important for the generation of inducible Treg (Agace and Persson, 2012;

Matteoli et al., 2010). Furthermore, it seems that -catenin and mitogen-activated protein kinase

(MAPK) p38 are required for intestinal DCs to express the RA-metabolizing enzymes, IL-10 and

TGFβ, and to stimulate Treg induction while suppressing inflammatory T effector cells (Huang

et al., 2013; Manicassamy et al., 2010). Signaling pathways within DCs, such as TNF receptor-

associated factor 6 (TRAF6) and TGFR pathways, serve the mechanisms of DC-mediated

coupling of T cell differentiation and Treg induction (Han et al., 2013; Ramalingam et al., 2012).

41

It has been proposed that the tolerogenic properties of intestinal CD103+ DCs might be

conditioned by enterocytes via thymic stromal lymphopoietin (TSLP) and other factors (Iliev et

al., 2009).

In summary, intestinal DCs can provoke immune tolerance to self and innocuous environmental

antigens in the steady state (Steinman et al., 2003). This is accomplished in part by promoting

the differentiation of Tregs and suppressing the induction of effector T cells.

1.4.8 Fate 2: Immune response against harmful pathogens

Overcoming the tolerogenic milieu of the gut is a prerequisite to the generation of an effector T

cell response. In the case of highly virulent pathogens, protective responses must occur rapidly to

contain and control the infection. The intestinal DC compartment is uniquely adapted to perform

this function.

One effector population of DC-driven immunity against pathogens is Th17 cells. The steady state

induction of Th17 cells is dependent on signals from the microbiota, with segmented filamentous

bacteria or SFB being a prominent example (Ivanov et al., 2009). Th17 responses are important

for protection against oral challenge with the fungus Candida albicans or the bacterium

Salmonella (Conti et al., 2009; Raffatellu et al., 2008). It has been demonstrated that intestinal

CD103+CD11b+ DCs play a central role in Th17 homeostasis (Denning et al., 2007). Indeed, this

DC subset is a potent producer of IL-6 and IL-23, cytokines that are critical for the

differentiation and maintenance of Th17 cells (Kinnebrew et al., 2012; Persson et al., 2013b;

Schlitzer et al., 2013).

Besides Th17 responses, Th1 and CTL responses are crucial for protection against intracellular

pathogen challenges, such as bacteria L. monocytogenes (Yamazaki et al., 2013) and C.

rodentium (Simmons et al., 2002), and parasites, including Toxoplasma gondii (Denkers and

Gazzinelli, 1998). Uncontrolled Th1 responses can be harmful or even lethal, which can lead to

Th1-mediated colitis. It has been demonstrated that the intestinal CD103+CD11b- DC subset

controls Th1 and CD8+ T-cell homeostasis and induction (Hildner et al., 2008; Luda et al., 2016).

Indeed, this DC subset specializes in secreting IL-12, which is critical for the differentiation of

Th1 and CD8+ T cells that produce IFN (Mashayekhi et al., 2011; Naik et al., 2005).

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1.4.9 Fate 3: Immune response against self-antigens

Autoimmune diseases are caused by an interplay of a person's genotype and environment

exposures, with DC playing a key role in presenting self-peptides to self-specific lymphocytes.

In the case of the gut, a good example of an inappropriate DC-driven response is the T-cell

response to gliadin peptides in celiac disease. Celiac disease is a chronic enteropathy induced by

ingestion of dietary gluten (as a “trigger”) in genetically predisposed people who also have a

“leaky” gut (increased intestinal permeability) (Meresse et al., 2012). In the context of celiac

disease, gluten is digested by luminal and enterocyte brush-border enzymes into amino acids and

peptides. During infections or as a result of intestinal permeability changes, gliadin peptides (a

composition of gluten) enter the SILP, where they are deamidated by tissue transglutaminase 2

(TG2), allowing interaction with human leukocyte antigen (HLA)-DQ2 (or HLA-DQ8) on the

surface of APCs (Green and Cellier, 2007). Instead of inducing a tolerogenic response (e.g.,

generation of Treg), intestinal DCs present gliadin to CD4+ T cells, resulting in the production of

inflammatory cytokines (IL-21 and IFN) that cause tissue damage (Sollid and Jabri, 2013). This

leads to villous atrophy and crypt hyperplasia as well as the activation and expansion of B cells

that produce antibodies against TG2 and gliadin (Green and Cellier, 2007). In addition to celiac

disease, IBD is also considered a gastrointestinal autoimmune disease, although the etiology and

causative antigen(s) are still unclear and will not be discussed further in this thesis.

Our understanding of anti-viral responses in the gut is relatively limited compared to our

knowledge of anti-bacterial responses. This may be because the intestinal virome was considered

much later than the intestinal bacterial microbiome due to limitations in bioinformatic tools and

sequencing techniques (Virgin, 2014). We now know that the mammalian virome incudes

viruses that infect eukaryotic cells (eukaryotic virome); bacteriophages that infect bacteria

(bacterial virome); viruses that infect archaea (archaeal virome); and virus-derived genetic

elements in host chromosomes that can change host-gene expression, express proteins, or even

generate infectious virus (Virgin, 2014). Since intestinal immunity against viruses is poorly

understood, in Chapters 3 and 4, I take advantage of the rotavirus infection model in mice

to study how intestinal DC impact the local and systemic antiviral responses.

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1.4.10 Rotavirus infection model

RV is a leading cause of acute, often dehydrating gastroenteritis in infants and young children

worldwide. Irrespective of socioeconomic status, by 3 years of age virtually all children will be

infected with RV, with the age of first infection generally being lower and mortality being

greater in resource-limited countries (Clarke and Desselberger, 2015). Conveniently, RV

infection in mice parallels human RV infection reasonably well. In humans, RV-induced diarrhea

is seen primarily in children between 6 months and 2 years of age while mice are most

susceptible to RV-induced diarrhea from 4-14 days of age (Burns et al., 1995). In adult humans

and mice, RV infection is asymptomatic and cleared within a week.

1.4.10.1 Epidemiology and RV vaccines

Human RV was first isolated in epithelial cells of the small intestine from children with diarrhea

in 1973 (Bishop et al., 1973). Since then, RV has been recognized as a leading cause of severe

gastroenteritis among young children worldwide. In the pre-vaccine era, RV was estimated to

account for one-third of the estimated 578,000 deaths from childhood gastroenteritis and more

than 2 million hospitalizations and 25 million outpatient clinic visits among children under 5

years of age each year (Liu et al., 2015; Tate et al., 2016). Because of this tremendous health

burden, prevention of RV is a priority for global health agencies. In 1999, a tetravalent rhesus

reassortant RV vaccine (Rotashield, Wyeth) was withdrawn from the United States market

within a year of its implementation because it caused intussusception, a form of bowel

obstruction (Murphy et al., 2001). Then came the next generation oral RV vaccines - a

pentavalent bovine-human reassortant vaccine (RotaTeq, Merck and Co.) and a monovalent

human vaccine (Rotarix, GlaxoSmithKline/GSK Biologicals), both of which are live attenuated

vaccines (Ruiz-Palacios et al., 2006; Vesikari et al., 2006). Both vaccines were shown to be safe,

were not associated with intussusception, and provided more than 70% and 90% protection

against any RV diarrhea and severe RV diarrhea, respectively. Therefore, the World Health

Organization recommended global implementation of RV vaccines in 2009. The two licensed

vaccines were introduced into more than 60 countries between 2006 and 2013, and have led to

significant reductions in the global burden of RV diarrhea, with a halving of the number of RV-

associated deaths from an estimated 528,000 in 2000 to 215,000 in 2013 (Tate et al., 2016).

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Although the two licensed RV vaccines have an excellent efficacy record in western countries,

their capacity to prevent RV mortality in resource-limited countries, particularly in Africa and

Asia, is still unclear (Angel et al., 2007). Some variables/ differences between developed and

developing countries may contribute to the uncertainty of the vaccine efficiency in resource-

limited countries. First of all, viral transmission frequency, which depends on population density

and climates (highly seasonal in temperate zones versus year-round in tropical zones) may be

different between countries (Minor, 2004). Second, higher infectious doses and/or co-infection

with multiple strains seems to occur in developing countries (Santos and Hoshino, 2005). Third,

the genotypes and/or serotypes of strains circulating in developing countries frequently differ

from the common strains circulating in developed countries (Santos and Hoshino, 2005). Fourth,

bacterial overgrowth and/or helminth, malaria or human immunodeficiency virus (HIV) co-

infection might lower the immunogenicity of vaccines (Grassly et al., 2006). Fifth, higher levels

of pre-immune (maternal) antibodies and/or breastfeeding at the time of vaccination in children

in developing countries may also reduce the immunogenicity of the RV vaccine (Hanlon et al.,

1987). Lastly, other biological factors such as micronutrient malnutrition and altered microbiota

may affect the development of the newborn immune system, which may also reduce vaccine

efficiency (Glass et al., 2006; Harris et al., 2017). Overall, the generation of an RV vaccine

suitable for the resource-limited countries will need to take biological, geographical, financial

and social factors into account.

1.4.10.2 Rotavirus

Rotaviruses are members of the Rotavirus genus of the Reoviridae family, which contains

viruses with segmented dsRNA genomes. RV particles are large and complex, with 3 concentric

protein layers that surround the viral genome of 11 segments of dsRNA. The RV genome

segments encode 6 structural proteins that make up virus particles (viral proteins or VPs) and 6

non-structural proteins (NSPs). VP7 (a glycoprotein or G-type antigen) makes up the outer

capsid shell and VP4 (a protease-sensitive protein or P-type antigen) forms spikes that emanate

through the shell; these induce neutralizing antibody responses and are the basis of a binary

classification system for viral serotypes (Figure 1-6). The intermediate layer is made up of the

major structural protein VP6, whereas the core is composed of VP2 (the scaffolding protein)

with VP1 (the viral RNA-dependent RNA polymerase) and VP3 attached on the inside (Figure

45

1-6). The RV genus is divided into serological groups (A to E). Group A RV causes significant

diarrhea disease in infants and in the young of various mammalian and avian species (Fields et

al., 2007).

Figure 1-6 Rotavirus structure

The figure shows a schematic representation of a rotavirus virion.

RV infects and replicates within mature epithelial cells at the apex of the villus, and viral

progeny are liberated from infected cells by cell lysis or by a non-classical vesicular transport

mechanism in polarized epithelial cells (Starkey et al., 1986). The natural cell tropism for RV is

the differentiated enterocytes in the small intestine, suggesting that differentiated enterocytes

express a specific receptor for viral attachment or they express factors required for efficient

infection and replication (Ramig, 2004). However, recent recognition that extraintestinal spread

of RV occurs (Blutt et al., 2003; Crawford et al., 2006; Fischer et al., 2005) suggests a wider

range of target host cells than previously thought. After attachment, RV penetrates enterocytes

and undergoes uncoating, synthesis of viral transcripts, mRNA translation, replication of

genomic RNA, RNA encapsidation, virion assembly, and lastly virus release into the gut lumen.

1.4.10.3 Pathogenesis

After RV infection, the pathological changes are almost exclusively limited to the small

intestine. Across various animal models, RV infection is associated with virtually no visible

lesions; slight lesions, such as enterocyte vacuolization and loss; or larger changes such as villus

blunting and crypt hyperplasia. Inflammation is generally mild compared to that for other

intestinal pathogens.

46

RV infection alters the function of the small intestinal epithelium, resulting in diarrhea. RV-

induced diarrhea has been attributed to a number of different mechanisms. Firstly, malabsorption

secondary to enterocyte destruction is thought to contribute to diarrhea. Specifically, absorption

of Na+, water, and mucosal disaccharidases are decreased during infection and malabsorption

results in the transit of undigested mono- and disaccharides, carbohydrates, fats, and proteins into

the colon. The undigested bolus is osmotically active, and the colon is unable to absorb sufficient

water, leading to an osmotic diarrhea. The reason for enterocyte destruction under conditions of

malabsorption is thought to be villus ischemia. Secondly, a secreted fragment of NSP4, or certain

NSP4 peptides have been identified as enterotoxins that induce diarrhea when inoculated into

mice. Finally, the enteric nervous system (ENS) is also implicated in RV-associated diarrhea.

Indeed, several drugs that block the action of the ENS attenuate RV-induced diarrhea and ~67%

of the fluid and the electrolyte secretion in RV-induced diarrhea in mice is due to the activation

of the ENS (Lundgren et al., 2000). Thus, pathogenesis of RV infection is multifactorial and

induced by both host and viral factors, which ultimately affect the outcome of the disease.

Additionally, the age of inoculation of animals is important for the manifestation of pathology -

ranging from biliary atresia (newborn mice), diarrhea and some extra-intestinal replication of

virus (7-14 day old mice) or asymptomatic infection (adult mice). Analysis of different viral

reassortment identified several viral proteins as being involved in virulence and include

VP3/NSP2/VP6/NSP3 in the efficiency of virus replication, NSP3 in shut-off of host protein

synthesis, NSP3/VP6 in extra-intestinal spread of virus, VP4/VP7 in viral entry into epithelial

cells, NSP1 in antagonizing type I IFN signaling (Barro and Patton, 2005), and NSP4 in the

induction of diarrhea (Fields et al., 2007).

1.4.10.4 Host immunity to RV infection

Both innate and adaptive immune responses are elicited after RV infection. However, when

comparing the results generated by different groups, some points need to be kept in mind:

1) The genetic background of experimental mice: mice with a C57BL/6 (H2-b) background seem

to be relatively more resistant to RV infection compared to BALB/c (H2-d) or 129 (H2-b) mice.

Thus, protective mechanisms identified in C57BL/6 could differ from those found in BALB/c or

129 mice, and vice versa (Franco and Greenberg, 2000);

47

2) The strain of RV: ECw and EDIM are commonly used wild-type non-cell-culture-adapted

murine homologous strains, whereas rhesus rotavirus (RRV) and simian rotavirus (SA11) are

commonly used as tissue culture adapted heterologous strains. Different viral strains result in

different host immune responses, such as intestinal restriction infection vs

extraintestinal/systemic infection, different kinetics of immune responses and different

requirements for host defense (Allen et al., 2007; Feng et al., 1994; Jaimes et al., 2005).

3) Microbiome influence and littermate controls: The gut microbiota exert a tremendous impact

on health and disease. Since a variety of environmental factors, in addition to mouse genetic

background, can impact intestinal microbiome, it is hard to compare results generated from

different animal facilities. A good example would be WT mice purchased from Taconic that

harbor abundant SFB whereas WT mice from Jackson lab are devoid of SFB (Ivanov et al.,

2009). Moreover, within the same animal facility, separately bred or purchased WT mice often

harbour distinct microbiota compared with separately bred mutant mice. Divergent microbiota

can be vertically transmitted resulting in changes in the immunological baseline (Escalante et al.,

2016; Moon et al., 2015). Therefore, the gold standard for experimentation is to use littermate

controls to determine the relative role of host genetics versus microbiota in conferring a

particular phenotype (Stappenbeck and Virgin, 2016).

i) Innate immune responses to RV infection

Viral infection triggers a cascade of cellular events culminating in the expression and secretion

of immunomodulatory proteins, such as IFNs, which can induce the establishment of an antiviral

state in neighboring cells. The capacity of the antiviral state to suppress viral replication is a

critical mechanism used by the host to control the dissemination of the virus.

Following RV entry into cells, melanoma differentiation-associated protein 5 (MDA5) and RIG-I

detect viral ssRNA and dsRNA, and subsequently signal through mitochondrial antiviral

signaling protein (MAVS) to stimulate the activation of IRF3/7 and IFN/ expression (Broquet

et al., 2011; Pott et al., 2011; Sen et al., 2011). NF-B and AP-1 (via JNK/p38) also are activated

by RV infection, but a role for RIG-1/MDA5/MAVS in this process has not been experimentally

verified. An alternative pathway of RV detection by TLR3 and its adaptor TIR-domain-

containing adaptor-inducing IFN (TRIF) has been demonstrated in vitro and in vivo, also

48

leading to IRF3/7 activation and IFN/ expression (Pott et al., 2012). In the absence of MyD88,

adult mice shed more virus in the feces while neonatal mice display an increase in incidence and

duration of diarrhea, suggesting MyD88-dependent TLR signaling also plays a role in host

control of RV infection (Uchiyama et al., 2015). The protective effects of secreted IFN// are

mediated by autocrine/paracrine stimulation of the IFN/ and IFNreceptor, leading to

STAT1/2 activation, IFN stimulated gene (ISG) expression and inhibition of RV replication by

an unidentified mechanism (Holloway and Coulson, 2013).

In addition to IFNs, cytokine/chemokine expression is likely to result in attraction and activation

of immune cells and resolving the infection. For example, it has been reported that treatment of

mice with bacterial flagellin cured RV infection via a mechanism involving TLR signaling and

induction of the cytokine IL-22 and IL-18 (Zhang et al., 2014a). Recently, it has been suggested

that IFNand IL-22 act synergistically for the induction of ISGs and provide innate immune

responses against RV infection (Hernández et al., 2015). Additionally, NLRP9b-mediated

inflammasome activation is reported to restrict RV replication in intestinal epithelial cells (IECs)

via promoting maturation of IL-18 and gasdermin D-induced pyroptosis (Zhu et al., 2017). In

summary, various innate responses against RV are elicited mainly in RV-infected IECs, which

produce inflammatory mediators or induce cell death to restrict viral replication. Although innate

responses are activated during RV infection, mice lacking an adaptive immune system are unable

to clear the virus and develop chronic disease.

ii) Adaptive immune responses to RV infection

Severe combined immunodeficiency (SCID) mice (BALB/c and C57BL/6 background),

recombinase-activating gene (Rag)2-/- mice (129/C57BL/6) and Rag1-/- mice (C57BL/6), all of

which lack T cells and B cells, develop chronic disease after murine homologous RV infection

(Franco and Greenberg, 1995, 1997; Riepenhoff-Talty et al., 1987; Zhang et al., 2014a),

suggesting that adaptive immune responses are essential for RV clearance.

B-cell responses

After 3-4 days post homologous RV infection, there is a massive induction of B cells in the PPs

and the MLNs (Blutt et al., 2002), suggesting a GC response is activated by RV. At ~5 d.p.i.,

49

anti-RV IgA can be detected in the intestinal wash or fecal pellet (Burns et al., 1995). The IgA

level peaks around 12 d.p.i. and fecal IgA persists for up to 1year following primary infection

(Burns et al., 1995; McNeal and Ward, 1995). The role of B cells and antiviral IgA in primary

RV infection is controversial and inconsistent. Indeed, it has been found that the B-cell response

is not absolutely required for the resolution of primary RV infection, since Jh-/- mice (both adult

and suckling mice) and IgA-/- mice (129/C57BL/6) can clear RV with the same kinetics as WT

controls (Franco et al., 1997; McNeal et al., 1995; O'Neal et al., 2000); whereas other groups

found that MT mice (C57BL/6), IgA-/- mice (C57BL/6 or BALB/c) and J-chain-/- (BALB/c)

mice developed chronic disease (Blutt et al., 2012; McNeal et al., 1995; Schwartz-Cornil et al.,

2002). However, the humoral response does play a role in protecting mice from re-infection,

since Jh-/- mice, MT mice, IgA-/- mice and J-chain-/- mice are susceptible to a secondary RV

infection (Blutt et al., 2012; Franco and Greenberg, 1995; McNeal et al., 1995; Schwartz-Cornil

et al., 2002). In humans, the data are not consistent in terms of whether IgA levels are a good

correlate of protection against RV (Angel et al., 2012).

Is the RV-specific IgA response T-dependent or T-independent?

Intestinal T cell-dependent IgA responses are generated in the PP or MLNs. In the GC,

underpinned by FDCs, the interaction between B cells and follicular helper T cells (Tfh) is

promoted by costimulatory molecules and cytokines. This facilitates B cell proliferation,

induction of activation-induced deaminase (AID), and subsequent class switch recombination

(CSR), somatic hypermutation (SHM) and affinity maturation of the B cell response (Fagarasan

et al., 2010).

T cell-independent of IgA is generated in the SILTs (CPs and ILFs) or the intestinal LP. It is

likely that ILF-resident B cells are activated either after antigen presentation by TNF-

expressing macrophage-DCs or directly by microbial components (e.g., LPS, peptidoglycan).

Consequently, ILF-resident B cells undergo preferential class switching to IgA in the absence of

T cells, under the influence of TGF, BAFF and A proliferation-inducing ligand (APRIL). IgA+

B cells or IgA plasmablasts generated within ILFs undergo differentiation to IgA plasma cells in

the intestinal LP, with the help of IL-6, IL-10, BAFF and APRIL secreted by stromal cells or

DCs (Fagarasan et al., 2010). In the absence of PPs or SILTs, IgA can be generated directly in

50

the intestinal LP (Tsuji et al., 2008b; Uematsu et al., 2008), which is dependent on the TLR-

MyD88 signaling pathway and bacterial load in the gut.

Upon murine RV infection, TCR-/- mice can develop detectable RV-specific IgA and antibody

secreting cells in the intestinal LP, but their level is 4-60 times lower than control WT mice

(Franco and Greenberg, 1997), suggesting that the majority of the anti-RV IgA response is T

cell-dependent. Indeed, anti-CD4 monoclonal antibody (mAb)- treated WT mice display a

similar IgA reduction phenotype compared with TCR-/- mice (Franco and Greenberg, 1997).

Although the IgA level is dramatically decreased, these mice are nevertheless protected from re-

infection, suggesting T-independent IgA responses are sufficient to prevent re-infection.

T-cell responses

RV-specific CD8+ T cells can be detected after RV infection in the small intestine (Offit and

Dudzik, 1989). Several lines of evidence implicate CD8+ T cell response in RV clearance. First

of all, mice deficient in CD8+ T cells (WT mice treated with anti-CD8 mAb or beta 2-

microglobulin (2m)-/- mice) exhibit a 2-3 day delay in RV clearance (Franco and Greenberg,

1995, 1997). Secondly, chronically infected Rag2-/- mice and SCID mice can clear RV upon

adoptive transfer of CD8+ T cells (Dharakul et al., 1990; Franco et al., 1997). Therefore, RV-

specific CD8+ T cells are important in clearance of primary RV infection in WT mice. In terms

of conventional versus T cells, it has been reported that T cells are dispensable whereas

T cells are required for RV clearance (Franco and Greenberg, 1997). Additionally, CD4+ T

cells are dispensable for RV clearance since WT C57BL/6 mice treated with anti-CD4 mAb

cleared RV normally (Franco and Greenberg, 1997). To overcome the caveat that the humoral

immune responses may compensate for the loss of cellular responses (due to the functional

redundancies of immune components), it has been shown that while most Jh-/- mice clear primary

RV infection, Jh-/- mice treated with anti-CD8 mAb develop chronic disease (Franco and

Greenberg, 1995; Franco et al., 1997; McNeal et al., 1995). However, these experiments are

performed on adult mice, leaving a gap in understanding of the role of CD8+ T cells in neonatal

anti-RV responses.

51

The immunodominant epitopes of murine homologous RV recognized by CD8+ T cells have

been mapped by Greenberg’s group (Jaimes et al., 2005). They reported that in homologous

murine RV infections, the majority of intestinal CD8+ T cells recognize the H-2b-restricted

VP6357-366 and VP733-42 epitopes, making these two epitopes good candidates for examining RV-

specific CD8+ T cell responses via flow cytometry (Jiang et al., 2008). It is still unclear which

intestinal DC subset is important for shaping anti-RV adaptive immune responses. Therefore,

Chapter 3 of this thesis will dissect the role of intestinal DCs in modulating host antiviral

responses in both adult and neonatal mice.

1.4.10.5 Contribution of maternal effects on RV immune responses

Lactating mothers nourish neonatal mammals with breast milk rich in factors that compensate for

the virgin intestinal immune system of their offspring. Breast milk contains cytokines such as

TGF and IL-10 that facilitate the tolerogenic response to the microbiota in the newborn

(Garofalo et al., 1995; Letterio et al., 1994). Moreover, suckling mammals ingest large quantities

of immunoglobulins contained in maternal milk. Acquisition of maternal IgG via breast milk

helps protect neonates against pathogens (Niewiesk, 2014). Also, maternal IgG can deliver

microbial molecules to offspring, which increases the number of intestinal ILC3 and reinforces

barrier integrity (Gomez de Agüero et al., 2016). Recently, it has been demonstrated that

maternal-derived, T-cell independent anti-commensal IgG antibodies, which display a broad,

anti-commensal capacity, help restrain microbes in newborn mice. The limited translocation of

microbes reduces the effector Th cell differentiation, which protects the newborns from

developing overwhelming anti-commensal immune responses (Koch et al., 2016). Likewise,

ingestion of IgA can mediate passive immunity to enteric infections and reinforce appropriate

anti-commensal immune responses in offspring (Macpherson et al., 2008; Rogier et al., 2014).

Do maternal antibodies protect neonates from RV infection?

It is reasonable to hypothesize that maternal anti-RV antibodies can protect newborns from RV

infection. Early studies performed on mice revealed a good correlation between the titers of the

RV-specific serum/lacteal IgG (but not IgM or IgA) in the mouse dam and those in her progeny

(Sheridan et al., 1983; Sheridan et al., 1984). Furthermore, neonatal mice positive for RV-

specific intestinal IgG before homologous infection did not develop diarrheal disease (Sheridan

52

et al., 1983). In humans, instead of IgG, RV-specific IgA is present in the breast milk, with

antibody levels at their highest in colostrum and falling significantly as breast-feeding becomes

established (Asensi et al., 2006; Hjelt et al., 1985). Interestingly, it appears that antibody-

mediated passive protection against RV challenge is dependent on serotype, titer of antibody as

well as geographic region (developing versus developed countries) (Clarke and Desselberger,

2015; Offit and Clark, 1985). Additionally, it has been suggested that components other than

antibodies from breast milk (such as lactoferrin and lactadherin) have RV-neutralizing capacity

and may be responsible for some of the protection conferred by breast feeding under certain

conditions (Asensi et al., 2006; Moon et al., 2013; Newburg et al., 1998).

1.5 Human cDC

Three putative cDC subsets have been identified in the human small intestine that can be

distinguished based on surface expression of CD141, CD103 and SIRP, together with a lack of

the monocyte-macrophage markers CD64 and/or CD14 (Watchmaker et al., 2014). It has been

shown that human small intestinal CD103+SIRP- cDCs resemble CD141+ cDCs in other human

tissues as well as murine CD103+CD11b- cDCs, while human small intestinal CD103+SIRP+

cDCs correspond to human tissue/blood derived CD1c+ cDCs and murine CD103+CD11b+ cDCs.

In addition, human small intestinal CD103-SIRP+ cells are heterogeneous, as majority of which

express IRF4, CD11b and intermediate levels of CX3CR1(Watchmaker et al., 2014).

Functionally, as in mice, human intestinal CD103+ cDCs express CCR7 (Mann et al., 2016),

indicating they can migrate to draining LNs. Indeed, CD103+ cDCs can be found in the

migratory compartment of human MLN biopsies (Magnusson et al., 2016). Mouse small

intestinal CD103+CD11b- and CD103+CD11b+ cDCs express high levels of aldehyde

dehydrogenase (ALDH) activity are mutually redundant in inducing Tregs. However, human

small intestinal CD103+SIRP+ cDCs display higher ALDH activity and induce higher Foxp3

expression on CD4+ T cells in vitro. Moreover, this cDC subset induces CCR9 on responding T-

cells more efficiently than CD103+SIPR- cDCs (Watchmaker et al., 2014). The role of human

cDC subsets in regulating intestinal homeostasis, inflammation and infection remains to be

determined.

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1.6 Summary

The work presented in this thesis is divided into two result chapters. The first addresses whether

DCs are required for anti-RV adaptive immune responses in the intestinal mucosa, and if so,

which DC subset(s) are needed. To answer this, I challenged DTx-treated Zbtb46-DTRWT

chimeric mice (lacking all ZBTB46-dependent cDCs), Batf3-/- mice (lacking CD103+CD11b-

DCs) and huLangerin-DTA mice (lacking CD103+CD11b+ DCs) with murine RV and found that

CD103+CD11b- DCs, but not CD103+CD11b+ DCs, are needed for generating an optimal

antiviral CD8+ T-cell response. Moreover, I observed that neonatal mice have a more stringent

requirement for BATF3-dependent DCs in generating antiviral CD8+ T-cell responses.

Furthermore, I observed dysregulated polyclonal CD4+ T cell skewing from a Th1 to Th17

response in both adult and neonatal Batf3-/- mice. Finally, in spite of a considerable deficiency in

DC, I found that the anti-RV IgA response is not impaired in these cDC deficient mice. My

results suggest an age-dependent requirement for DCs in the RV immune response.

In the second data chapter presented in this thesis, I examined the role of LTR in the radio-

sensitive compartment (mainly DC) in the RV infection model. I found that the Ltbr-/- WT

chimeric mice generate more IFN-secreting T cells in the intestinal LP, while homeostatic IL-17

producing CD4+ T cells are decreased. The humoral anti-RV IgA response is generated normally

in spite of increased Th1 and decreased Th17 responses in the context of LTR deficiency in the

radio-sensitive compartment. My results suggest that the requirement of LTR signaling is

context-dependent.

54

Chapter 2

Methods and Materials for Chapter 3 and Chapter 4

55

Mice

Table 2-1 Mouse strains

Stock Name Also known as Vendors

C57BL/6 WT Charles River Laboratories, Senneville

QC, Canada

B6(Cg)-

Zbtb46tm1(HBEGF)Mnz/J Zbtb46-DTR

Gift of Dr. Kenneth Murphy, Washington

University, St. Louis, MO, USA

B6.129S(C)-

Batf3tm1Kmm/J Batf3-/-

The Jackson Laboratory, Bar Harbor, ME,

USA

B6.FVB-Tg(CD207-

Dta)312Dhka/J huLangerin-DTA


USA

B6.FVB-Tg(Itgax-

DTR/EGFP)57Lan/J CD11c-DTR


USA

Ltbr-/- Gift of Dr. Rodney Newberry, Washington

University, St. Louis, MO, USA

Mice indicated above (Table 2-1) were housed in the University of Toronto Division of

Comparative Medicine (DCM) under a specific pathogen-free but not a barrier condition. Mice

were on a standard irradiated chow diet Envigo Teklad (2918). Water was reverse-osmosis and

UV-sterilized and acidified to pH 3. 12 hr light cycle with lights on at 100% intensity from 7 AM

- noon and 50% intensity from 1 PM to 7 PM. Lights were turned off (0%) from 8 PM to 6 AM.

Light intensity was gradually increased or decreased over the course of one hour each transition.

Lighting was not subject to daylight savings time adjustments. Standard bedding for mice was

the Bed-o'Cobs combo bedding. All experiments were approved by the University Animal Care

Committee.

The purchased Batf3-/- mice were first bred with WT mice (Batf3+/+) to generate Batf3+/-

heterozygous F1. Subsequently Batf3+/- F1 mice were back-crossed with Batf3-/- mice to generate

Batf3+/- and Batf3-/- littermates for experimental use. The purchased huLangerin-DTA+/+ mice

were first bred with WT mice (huLangerin-DTA-/-) to generate huLangerin-DTA+/- heterozygous

56

F1. Subsequently huLangerin-DTA+/- F1 mice were back-crossed with WT to generate

huLangerin-DTA and huLangerin-DTA- littermates for experimental use.

Antibodies and Flow cytometry

Anti-mouse CD3 APC eFluor780 (17A2), CD103 APC (2E7), CD4 APC (RM4-5), CD4 FITC

(RM4-5), CD8 FITC (53-6.7), CD11c FITC (N418), CD44 Percp-Cyanine 5.5 (1M7), CD45R

(B220) Percp-Cyanine 5.5 (RA3-6B2), IFN PE-Cyanine 7 (XMG1.2), CD11c PE-Cyanine 7

(N418), MHCII (I-A/I-E) eFluor 450 (M5/114.15.2), anti-mouse/rat Ki-67 eFluor 450 (SolA15),

and anti-mouse IL-22 PE (1H8PWSR), RORt APC (B2D) were purchased from eBioscience

(San Diego, CA). Anti-mouse PE/Dazzle 594 F4/80 (BM8), Brilliant Violet 711 CD8 (53-6.7),

Brilliant Violet 605 IL-17A (TC 11-18H10.1), and anti-mouse/human Brilliant Violet 605

CD11b (M1/70) were purchased from Biolegend (San Diego, CA). Live/Dead fixable Aqua was

purchased from Life Technologies (Carlsbad, CA). After Live/Dead Aqua staining, cells were

washed and then blocked with purified anti-FcRII/III monoclonal antibody (2.4G2). All surface

stains were performed in PBS with 2% fetal bovine serum. Intracellular staining was performed

using a Cytofix/Cytoperm Kit (BD Biosciences, Baltimore, MD). All stained samples were

acquired on a BD FACSCanto, LSR II or LSR Fortessa as appropriate. FlowJo software (Tree

Star, Ashland, OR) was used for fluorescence-activated cell sorting data analysis.

BM chimeras

BM cells (2-4x106) collected from femurs and tibia of Zbtb46-DTR, CD11c-DTR or Ltbr-/- mice

were injected intravenously into WT C57BL/6 mice that had been lethally irradiated (2 x 550

cGy). Recipient mice were left for 8-10 weeks to reconstitute, and were given water

supplemented with neomycin sulfate (2 g/L; BioShop, Canada) for the first 2 weeks.

DTx injection

20 ng per gram body weight of DTx (List Biological Laboratories, USA) was injected

intraperitoneally into Zbtb46-DTRWT chimeric mice 1 day prior to RV inoculation and DTx

injections were then repeated every day for short-term experiments (mice were sacrificed at 7

d.p.i for T-cell assay) or every other day for long-term experiments (mice were sacrificed at 28

57

d.p.i. for ELISA assays). Both treatment strategies resulted in the transient delay of RV antigen

clearance at 7 d.p.i., indirectly confirming the sufficient DC depletion. To exclude any changes

caused by gut microbiome, DTx-treated and PBS-treated Zbtb46-DTRWT chimeric mice were

housed in the same cage for both RV-infected and uninfected conditions.

7 ng per gram body weight of DTx was injected i.p. into CD11c-DTRWT chimeric mice 1 day

prior to RV inoculation and DTx injections were then repeated every 2 days to maintain DC

ablation.

RV mouse infections

The virulent wild type, non-cell-culture-adapted murine RV strain ECw was used to infect mice.

One virus stock was used in these studies. Stocks of RV were prepared as intestinal

homogenates, and the 50% diarrhea dose (DD50) of the ECw virus stock was determined for WT

neonatal mice as previously described (Burns et al., 1995).

The day prior to oral gavage with RV, adult and neonatal mice were transferred to biosafety level

2 (BSL2) facilities for the duration of all studies. Uninfected mice were housed separately in

BSL1 facilities in DCM.

Adult mice (6-8 wk) were orally gavaged with 104 DD50 ECw in 100 l HBSS containing 1 mM

CaCl2 and 0.5 mM MgCl2 after oral administration of 100 μl of 1.33% sodium bicarbonate to

neutralize stomach acidity. Fecal pellets were collected from each mouse on the day of challenge

and for the following days. Serum samples were collected with Microvette Capillary blood

collection tubes (Sarstedt, Germany) according to manufactory instructions. Fecal and serum

samples were stored frozen at -20°C until assayed. For use in the enzyme-linked immunosorbent

assays (ELISAs), 10% (wt/vol) stool suspensions were prepared with PBS containing 0.1%

sodium azide (Merck Millipore, USA).

Neonatal (3-5 days post-natal) Batf3+/- and Batf3-/- littermates were fostered with lactating CD1

dam 1-2 days prior to RV inoculation. The pups stayed with CD1 dam for the duration of the

experiment. Neonatal mice were orally given 5x103 DD50 ECw in 5 l HBSS containing 1 mM

CaCl2 and 0.5 mM MgCl2. On the day of harvest, neonatal mice were euthanized by decapitation

58

(7 d.p.i.) or CO2 asphyxiation (12, 14, 16 d.p.i.). Fecal pellets or colon contents were collected

and were stored frozen at -20 °C until assayed. Serum was collected by intra-cardiac puncture.

Detection of RV antigen and anti-RV IgA by ELISA

ELISA was performed as previously described to detect RV antigen and RV-specific fecal/serum

IgA (Burns et al., 1995), with modifications: using anti-RV mAb (AbD Serotec, Raleigh, NC,

USA), followed by HRP-conjugated anti-mouse IgG2b antibody (SouthernBiotech, Birmingham,

AL, USA) to detect RV antigen and HRP-conjugated anti-mouse IgA (SouthernBiotech) to

detect anti-RV IgA. The OD was read at 450 nm.

Measurement of anti-RV IgA titer by ELISA

For measurement of RV-specific IgA titer we adapted an assay previously described (Gonzalez

et al., 2003). Briefly, ELISA plates were coated overnight with sheep-anti-RV Ab (AbD Serotec,

USA). The plates were then blocked and incubated with inactivated Simian Rotavirus SA11

antigen (Microbix, Canada). After washing, the plates were incubated with 2-fold serial dilutions

of serum samples or fecal supernatant. HRP-conjugated goat anti-mouse IgA (SouthernBiotec,

USA) was applied to capture IgA and then the plates were developed by TMB solution

(BioShop, Canada). The titer of IgA in a serum or fecal sample was defined as log2 transformed

reciprocal of the last dilution exceeding an optical density of the value which was twice the

optical density of blank wells (blanks were those wells without added serum or fecal samples)

(IgA titer= log2(1/last positive dilution)). To be accepted for analysis, the titer of an internal

positive control in a plate could not differ by more than one dilution from plate to plate.

Cell Isolation

For analysis of MLN cells, organs were mashed through a 70 m cell strainer followed by PBS

washing.

For SILP cells, small intestines were dissected and cleaned in situ of mesenteric fat and PPs were

removed. Small pieces of the intestine then were thoroughly washed with washing buffer (Table

2-2) and EDTA solution (Table 2-2) was used to remove IELs. The remaining SILP fraction was

59

then digested with collagenase IV (Sigma-Aldrich, USA) in digestion buffer (Table 2-2) and

lymphocytes were enriched by Percoll gradient (GE Healthcare, Sweden).

For IELs, after washing with an EDTA solution, IELs were enriched by Percoll gradient. Given

the inherent variability and an underestimation in true cell yield in gut preparations, for the most

part we enumerated cellular compartments based on both frequency and absolute numbers.

Table 2-2 List of buffers and solutions used for cell isolation and culture

Buffer/solution name Ingredients

Intestinal cell

isolation buffers

Washing buffer HBSS+ 2%FBS+ 15mM HEPES

EDTA solution HBSS+ 10%FBS+ 15mM HEPES+ 5mM EDTA

Digestion buffer RPMI1640+ 10%FBS+ 15mM HEPES

Complete medium RPMI1640+ 1%Penicillin-Streptamycin+ 1% L-

Glutamine+ 1% HEPES+ 1% Sodium Pyruvate+

0.05mM 2-Mercaptoethanol+ 10% FBS

Intracellular staining (ICS)

To enumerate the number of cytokine-secreting T cells, ICS was performed, as described

previously (Jaimes et al., 2005). In brief, lymphocytes were incubated for 6 hr at 37°C in

complete medium (Table 2-2), supplemented with recombinant human IL-2 (100 U/ml; R&D

Systems, Minneapolis, MN, USA) and GolgiPlug (1 ml/ml; BD Biosciences). Cells were

stimulated with PMA (20 ng/ml; Sigma-Aldrich), ionomycin (500 ng/ml; Sigma-Aldrich),

VP6357–366 peptide (VGPVFPPGM; 2 mg/ml; Genemed Synthesis, San Antonio, TX, USA), or

VP733–40 peptide (IVYRFLFV; 2 mg/ml; Genemed Synthesis) (Jaimes et al., 2005).

Tetramer staining

VP6357-366-biotin was synthesized by NIH tetramer facility and then conjugated with PE-

streptavidin (Life technologies, USA) according manufactory’s instruction. After Live/dead

Aqua staining, cells were incubated with VP6-PE tetramer for 1 hr at 4°C, followed by other

surface staining.

60

RNA isolation, primer sets and qPCR

SILP tissues (stored in RNAlater (QIAGEN, Germany) in -20°C) or fresh cell pellets were used

for RNA extraction. RNA was isolated with Trizol reagent according to the manufacturer’s

instructions (Thermo Fisher Scientific, USA). Genomic DNA was removed with TURBO DNA-

free™ Kit (Thermo Fisher Scientific, USA). RNA was reverse-transcribed into cDNA with

SuperScript™ IV Reverse Transcriptase kit (Thermo Fisher Scientific, USA). Real-time PCR

was performed with SYBR Green Master Mix (Thermo Fisher Scientific, USA) and was run on

an CFX384 Touch™ Real-Time PCR Detection System (Bio-rad). The relative expression of

genes was calculated with the formula 2-∆∆Ct. murine ribosomal protein L19 (mRPL19) was used

as endogenous control housekeeping gene. The primer sets are listed below (Table 2-3).

Table 2-3 Primer sets

Gene Forward sequence Reverse sequence

IFN2/3 5’-AGCTGCAGGCCTTCAAAAAG-3’ 5’-TGGGAGTGAATGTGGCTCAG-3’

IL-22 5’-CATGCAGGAGGTGGTACCTT-3’ 5’-CAGACGCAAGCATTTCTCAG-3’

mRPL19 5’-GCATCCTCATGGAGCACAT-3’ 5’-CTGGTCAGC CAGGAGCTT-3’

Statistics

Comparisons of data were analyzed by student’s t-test (normal distribution) or Mann-Whitney

non-parametric test (non-normal distribution) with GraphPad Prism 6.0 program. Data were

presented as mean values ± SEM. p< 0.05 was considered significant.

61

Chapter 3

Intestinal BATF3-dependent dendritic cells are required for optimal antiviral T-cell responses in adult and neonatal mice

62

3.1 Abstract

Although we know a great deal about which types of dendritic cells (DCs) promote T-cell

priming in the periphery, less is known about which DC subset(s) provoke antiviral responses

within the gut. Here we report that conventional ZBTB46-dependent DCs were critically

required for antiviral CD8+ T-cell responses against rotavirus (RV), the major cause of childhood

gastroenteritis worldwide. Furthermore, we found that in adult mice, BATF3-dependent DCs

were required for generating optimal RV-specific CD8+ T-cell responses. However, in contrast to

mice that lack ZBTB46-dependent DCs, a significant amount of interferon gamma-producing

RV-specific CD8+ T cells were still detected in the small intestine of RV-infected adult Batf3-/-

mice, suggesting the existence of compensatory cross-presentation mechanisms in the absence of

BATF3-dependent DCs. In contrast to adult mice, we found that BATF3-dependent DCs were

absolutely required for generating RV-specific CD8+ T-cell responses in neonates. Loss of

BATF3-dependent DCs also resulted in a skewed polyclonalCD4+ T-cell response in both adult

and neonatal mice upon RV infection, although local and systemic RV-specific immunoglobulin

A production kinetics and titers were unimpaired. Our results provide insights that inform early-

life vaccination strategies against RV infection.

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3.2 Introduction

Dendritic cells (DCs) are the major antigen-presenting cells (APCs) responsible for first-line

defense against pathogens and are principal coordinators of the innate and adaptive immune

response to pathogens. DCs arise from common myeloid progenitors in the bone marrow (BM).

After progressing through developmental stages as macrophage DC precursors and common DC

progenitors, pre-DCs exit the BM and seed lymphoid and non-lymphoid tissues (Murphy, 2013).

ZBTB46 (BTBD4), a transcription factor belonging to the BTB-ZF (Broad complex, Tramtrack,

Bric-a`-brac, and Zinc finger) family, is induced from the pre-DC stage, and its expression is

maintained in fully differentiated conventional DCs (cDCs) but not plasmacytoid DCs (pDCs) or

macrophages (Satpathy et al., 2012). cDCs can be further separated based on the surface

expression of CD8 and CD11b in lymphoid tissues or CD103 and CD11b in non-lymphoid

tissues. Within peripheral lymphoid and non-lymphoid tissues, these DC subsets exert distinct

functions: the CD8+CD11b- or CD103+CD11b- DC subset is specialized in cross-presenting

intracellular pathogens or tumor antigens to CD8+ T cells and producing interleukin (IL)-12

(Hildner et al., 2008; Mashayekhi et al., 2011), whereas the CD8-CD11b+ or CD103+CD11b+

DC subsets, which produce IL-23 and IL-6, are thought to be specialized in the induction of

CD4+ T-cell responses (Persson et al., 2013b; Schlitzer et al., 2013). In the gut-associated

lymphoid tissues, emerging data have shown that specific DC subsets are required for controlling

certain types of pathogen-specific responses, particularly in the large bowel (e.g., C. rodentium

infection (Satpathy et al., 2013)). However, very little is known about which DC subset(s)

mediate clearance of small intestinal-tropic viral infections.

Lineage specification of CD8+CD11b- and CD103+CD11b- cDCs requires basic leucine zipper

transcription factor ATF-like 3 (BATF3) and interferon (IFN) regulatory factor 8, IRF8

(Grajales-Reyes et al., 2015). Mice with BATF3 deficiency (Batf3-/-) exhibit a selective loss of

CD8+CD11b- cDCs within lymphoid tissues and CD103+CD11b- cDCs within non-lymphoid

tissues (Edelson et al., 2010), without apparent abnormalities in other hematopoietic cell types

(Hildner et al., 2008). It has been reported that BATF3-dependent DCs are involved in mediating

adaptive immune responses against various pathogens such as West Nile virus (WNV) (Hildner

et al., 2008) and cytomegalovirus (Krueger et al., 2015; Torti et al., 2011) via antiviral CD8+ T-

cell responses, as well as T. gondii (Mashayekhi et al., 2011) and Leishmania major (Martínez-

64

López et al., 2015) through DC-secreted IL-12. In the respiratory mucosa, Waithman et al.

(Waithman et al., 2013) have shown that BATF3-dependent DCs mediate CD8+ T-cell responses

to influenza virus. However, it is not known whether BATF3-dependent DCs contribute to the

clearance of virus within the intestines, especially the small bowel. Furthermore, it has been

reported that during respiratory syncytial virus infection, neonatal CD103+ DCs in the

mediastinal lymph nodes provoke a fundamentally different CD8+ T-cell response profile than

CD103+ DCs from adult mice (Ruckwardt et al., 2014), suggesting that mucosal DCs exhibit

age-dependent properties. Investigating how intestinal DCs differ between adults and neonates in

initiating adaptive antiviral responses may provide us with better strategies for vaccine design.

Rotavirus (RV) is a double-stranded RNA virus belonging to the Reoviridae family and is a

leading cause of severe diarrhea in children aged <5 years. Although RV infections in adults are

typically asymptomatic or mild, immunosuppressed organ transplantation recipients are

susceptible to RV infection, and these patients can develop significant gastroenteritis (Lee and

Ison, 2014). Similar to children, neonatal/suckling mice also develop diarrhea after oral

infection, while adult mice remain asymptomatic upon infection, although viral shedding can still

be detected in the feces (an indicator of viral presence). RV infection of mice is a well-defined

model system for studying viral infection in the small intestine as RV predominantly infects and

replicates within mature epithelial cells on the tip of the small intestinal villi (Ramig, 2004). In

adult mice, CD8+ T cells have a role in the timely resolution of primary RV infection, while RV-

specific immunoglobulin A (IgA) is important for viral clearance after primary infection and for

preventing re-infections (Blutt et al., 2012; Franco and Greenberg, 1995).

In adult RV-infected mice, it has been reported that pDCs have an important role in promoting

the differentiation of activated B cells into plasma cells via type I IFN secretion (Deal et al.,

2013). In terms of the role of cDC, CD11c+ cells in the subepithelial dome of Peyer’s patches co-

localize with RV (Lopatin et al., 2013). These CD11c+ cells also upregulate co-stimulatory

molecules (CD80, CD86, and CD40) and increase the expression of proinflammatory cytokines

(IL-12/23p40 and tumor necrosis factor ) at early time points post infection (Lopez-Guerrero et

al., 2010). However, it is not clear what specific DC subtype primes RV-specific T cells in adults

or neonates. Furthermore, while the innate response to RV has been studied in neonatal mice

65

(Hernández et al., 2015; Pott et al., 2012), a comprehensive study of the neonatal anti-RV

adaptive immune response contrasted with adult anti-RV responses has not been performed.

In the current study, we generated Zbtb46-diphtheria toxin receptor (DTR)wild-type (WT) BM

chimeric mice and treated reconstituted chimeric mice with diphtheria toxin (DTx) to deplete

cDCs without altering macrophages or pDCs (Meredith et al., 2012a; Satpathy et al., 2012).

DTx-treated Zbtb46-DTRWT chimeric mice were able to clear RV, albeit with prolonged viral

shedding compared with phosphate-buffered saline (PBS)-treated control chimeras. However, in

the absence of cDCs, the antigen-specific CD8+ T-cell response in the small intestinal lamina

propria (SILP) was largely lacking at 7 days post infection (d.p.i.). Likewise, RV-infected Batf3-

/- mice exhibited prolonged viral shedding and decreased RV-specific CD8+ T-cell responses at 7

d.p.i. Unlike the DTx-treated Zbtb46-DTRWT chimeric mice, however, residual antigen-

specific CD8+ T-cell responses were readily detected in Batf3-/- mice, suggesting that other APCs

can compensate for the absence of BATF3-dependent cDCs to mediate cross-presentation of RV

antigen to CD8+ T cells. Interestingly, compared with adult mice, neonates exhibited a more

stringent dependency on BATF3-dependent cDCs for the induction of anti-RV CD8+ T-cell

responses, suggesting differential DC plasticity in adults compared with neonates. Local and

systemic anti-RV IgA responses were largely intact in both DTx-treated Zbtb46-DTRWT

chimeras and Batf3-/- mice, suggesting a dispensable role of cDCs in generating antiviral IgA

responses. These results provide important insights into the CD8+ T-cell response to RV in the

small intestine and may shed light on strategies for vaccine design.

66

3.3 Results

3.3.1 Depletion of ZBTB46-dependent cDCs is not affected by RV infection

ZBTB46 is a transcription factor expressed by cDCs (and pre-DCs) as well as by definitive

erythroid precursors and endothelial cells but not in macrophages or pDCs (Meredith et al.,

2012a; Satpathy et al., 2012). The expression of ZBTB46 on radio-resistant compartments makes

Zbtb46-DTR mice vulnerable to DTx treatment owing to DTx-mediated toxicity (Meredith et al.,

2012a). We therefore reconstituted lethally irradiated WT mice with Zbtb46-DTR BM in order to

avoid targeting DTx sensitive non-hematopoietic cells. Eight to 10 weeks after BM

transplantation, we injected chimeras via the intraperitoneal route with DTx 1 day prior and

throughout the period of RV infection. Using flow cytometry, we found that SILP cDCs were

markedly reduced in DTx-treated Zbtb46-DTRWT chimeric mice compared with PBS-treated

chimeric mice and that depletion efficacy was not affected by RV infection (Figure 3-1A,B;

gating strategies are described in Figure 3-2). Other APCs such as macrophages and B cells were

not altered after DTx treatment or upon RV challenge (Figure 3-1C,D). In terms of cDC subsets

in the SILP, CD103+CD11b- DCs and the majority of CD103+CD11b+ DCs were depleted by

DTx treatment (Figure 3-1E-G). Similar results were observed when absolute number of cells

was tabulated (Figure 3-2B-D). A concomitant increase in the frequency, but not absolute

numbers, of CD103-CD11b+ cells was observed with DTx treatment of Zbtb46-DTRWT

chimeric mice (Figure 3-1H and Figure 3-2E).

Mesenteric lymph nodes (MLNs) are the draining lymph nodes of the intestines, and DCs that

have captured antigen can transport antigen to the MLN via lymphatics in order to cross-prime

CD8+ T cells (Cerovic et al., 2015). We therefore also evaluated the DC populations in the

MLNs from DTx- vs. PBS-treated Zbtb46-DTRWT chimeric mice. DTx treatment of Zbtb46-

DTRWT chimeric mice was found to deplete both migratory and resident DCs (mDCs and

rDCs) and their subsets (Figure 3-2F, G-M). In summary, we confirmed that DTx treatment

efficiently depletes cDC in Zbtb46-DTRWT chimeric mice.

67

Figure 3-1 Conventional DC populations in DTx-treated Zbtb46-DTR WT chimeric mice.

A. Representative flow cytometry plots of CD11c+MHC II+ SILP DCs (detailed gating strategies

were shown in Figure 3-2.1) from mice treated with PBS or DTx at 7 d.p.i.. B. Percentage of

SILP DCs as a frequency of mononuclear cells from mice treated with PBS or DTx at 7 d.p.i.. C

and D. Percentage of SILP macrophages and B cells as a frequency of mononuclear cells

(detailed gating strategies were shown in Figure 3-2.1) at 7 d.p.i.. E. Representative flow

cytometry plots showing the gating strategy of SILP DC subsets (pre-gated as in Figure 3.1A)

from mice treated with PBS or DTx at 7 d.p.i.. F, G and H. Frequency of SILP DC subsets as a

frequency of DCs at 7 d.p.i..

UI, uninfected. RV, rotavirus infected. DP, CD103+CD11b+. DTx, diphtheria toxin. Results were

pooled from 4 independent experiments. Each data point represents a single biological replicate

(one mouse). Data are presented as mean ±SEM. Mann-Whitney test. **p<0.01, ****p<0.0001,

NS= not significant.

69

Figure 3-2 Gating strategy for SILP single cell populations.

A. SILP single cell suspensions were subjected to flow cytometry. Macrophages are defined as

F4/80+MHC IIhi cells, B cells are defined as CD11c-B220+(MHC IIhi) cells, cDCs are defined as

B220-F4/80-CD3-CD11c+MHC IIhi cells. B. Absolute numbers of SILP DCs from Zbtb46-

DTRWT chimeric mice treated with PBS or DTx at 7 d.p.i.. C-E. Absolute numbers of SILP

DC subsets from Zbtb46-DTRWT chimeric mice treated with PBS or DTx at 7 d.p.i.. F.

Representative flow cytometric plots of MLN DCs (pre-gated on B220-F4/80-CD3- live singlet

lymphocytes): CD11c+MHC IIhi migratory DCs (mDCs) and CD11chiMHC II+ resident DCs

(rDCs). mDCs can be further divided into 4 subsets based on the expression of CD103 and

CD11b, while rDCs can be further divided into 2 subsets based on the expression of CD8 and

CD11b. G. Absolute numbers of MLN mDCs from Zbtb46-DTRWT chimeric mice treated

with PBS or DTx at 7 d.p.i.. H. and J. Absolute numbers of MLN mDC subsets from Zbtb46-

DTRWT chimeric mice treated with PBS or DTx at 7 d.p.i.. K. Absolute numbers of MLN

DCs from Zbtb46-DTRWT chimeric mice treated with PBS or DTx at 7 d.p.i.. L. and M.

Absolute numbers of MLN DC subsets from Zbtb46-DTRWT chimeric mice treated with PBS

or DTx at 7 d.p.i..

UI, uninfected. RV, rotavirus infected. DTx, diphtheria toxin. Each data point represents an

individual biological replicate (one mouse) pooled from 2 independent experiments. Data are

presented as mean ±SEM. Mann-Whitney test. *p<0.05, **p<0.01, ***p<0.001, NS= not

significant.

70

3.3.2 ZBTB46-dependent cDCs are required for anti-RV CD8+ T-cell responses

To determine whether cDCs are required for anti-RV CD8+ T-cell responses, DTx-treated

Zbtb46-DTRWT chimeric mice and PBS-treated controls were orally infected with RV.

Although the frequency of SILP CD8+ T cells at steady state was not affected by DTx treatment

(Figure 3-3A), following RV challenge, cDC-deficient mice exhibited severely impaired CD8+

T-cell proliferation corresponding with a reduction in CD8+ T-cell frequency after RV challenge

(Figure 3-3A, B). Antigen-specific CD8+ T-cell responses were subsequently evaluated.

Specifically, the CD8+ T-cell response to VP6357–366, one of the immunodominant RV epitopes

recognized by H-2b-restricted CD8+ T cells (Jaimes et al., 2005), was examined via tetramer

staining along withVP6357–366 peptide restimulation to measure IFN production (Figure 3-4A,

B). At 7 d.p.i., the frequency of VP6357–366-specific CD8+ T cells in the SILP was significantly

reduced in the absence of cDCs (Figure 3-3C). In parallel, after in vitro restimulation with

VP6357–366 peptide, the percentage of CD8+ T cells capable of producing IFN was significantly

reduced in the absence of cDCs (Figure 3-3D). Reduced absolute numbers of CD8+ T cells,

VP6357–366-specific CD8+ T cells, and IFN+ CD8+ T cells were also observed in RV-infected

Zbtb46-DTRWT chimeric mice (Figure 3-4C-F). A similar reduction of antigen-specific CD8+

T cells was also observed in the intraepithelial lymphocyte (IEL) compartment, although the

frequency of CD8+ T cells within the IEL compartment was unaffected by DTx treatment (Figure

3-3E).

Finally, in the absence of cDCs, Zbtb46-DTRWT chimeric mice exhibited a continuous

shedding of RV into the gut lumen (measured in the fecal pellet) until 7 d.p.i. compared with

control chimeras (Figure 3-3F), suggesting a requirement for cDC and the downstream antiviral

CD8+ T-cell response in mediating optimal RV clearance. However, this prolonged viral

shedding was only transient, implying that compensatory mechanisms beyond the RV-specific

CD8+ T-cell response exist to mediate RV clearance. Taken together, these results suggest that

cDCs are required for CD8+ T-cell priming to RV.

71

Figure 3-3 ZBTB46-dependent cDCs are required to prime RV-specific CD8+ T cells.

A. Percentage of CD8+ T cells as a frequency of SILP mononuclear cells at 7 d.p.i.. B.

Percentage of Ki-67+ cells as a frequency of SILP CD8+ T cells at 7 d.p.i.. C. Percentage of

VP6357-366+ cells as a frequency of SILP CD8+ T cells at 7 d.p.i.. D. Percentage of IFN+ cells as a

frequency of SILP CD8+T cells after in vitro restimulation with VP6357-366 peptide at 7 d.p.i.. E.

Percentage of intraepithelial CD8+ T cells as a frequency of IEL mononuclear cells and

percentage of intraepithelial VP6357-366+ cells as a frequency of IEL CD8+ T cells at 7 d.p.i.. F.

Level of RV antigen in the feces measured by ELISA.

UI, uninfected. RV, rotavirus infected. DTx, diphtheria toxin. Results were pooled from 3-4

independent experiments. Each data point represents a single biological replicate (one mouse).

Data are presented as mean ±SEM. A, Student’s t-test; B-F, Mann-Whitney test. *p<0.05,

**p<0.01, ****p<0.0001, NS= not significant.

72

Figure 3-4 Tetramer staining and ICS of SILP CD8+ T cells in the SILP of Zbtb46-

DTRWT chimeric mice.

A. Representative flow cytometry plots showing VP6357-366+ SILP CD8+ T cells (pre-gated on

CD8+ T cells) at 7 d.p.i.. B. Representative flow cytometry plots showing IFN-producing SILP

CD8+ T cells after in vitro restimulation with VP6357-366 peptide for 6 h (pre-gated on CD8+ T

cells) at 7 d.p.i.. C. Absolute number of SILP CD8+ T cells from Zbtb46-DTRWT chimeric

mice treated with PBS or DTx at 7 d.p.i.. D. Absolute number of SILP Ki-67+CD8+ T cells from

Zbtb46-DTRWT chimeric mice treated with PBS or DTx at 7 d.p.i.. E. Absolute number of

SILP VP6357-366+ CD8+ T cells from Zbtb46-DTRWT chimeric mice treated with PBS or DTx

at 7 d.p.i.. F. Absolute number of SILP IFN+CD8+T cells from Zbtb46-DTRWT chimeric

mice after in vitro restimulation with VP6357-366 peptide at 7 d.p.i..

UI, uninfected. RV, rotavirus infected. DTx, Diphtheria toxin. Results were pooled from 2


Data are presented as mean ±SEM. Mann-Whitney test. *p<0.05, **p<0.01, ***p<0.001, NS=

not significant.

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3.3.3 Adult and neonatal Batf3-/- mice exhibit similar deficiencies in CD103+CD11b- cDCs both at steady state and during RV infection

As CD103+CD11b- cDCs specialize in the cross-presentation of viral and tumor antigens to

CD8+ T cells, we speculated that these cDCs would be required for priming CD8+ T cells upon

RV challenge. Recent studies have shown that the transcription factor Batf3 is required for the

development of both lymphoid CD8+CD11b– and non-lymphoid CD103+CD11b– DCs in mice

(Edelson et al., 2010; Hildner et al., 2008). At steady state, we confirmed that Batf3-/- mice

displayed a selective loss of CD103+CD11b- cDCs. This loss in CD103+CD11b- cDCs was

accompanied by an increased frequency of CD103-CD11b+ cells in the SILP, compared with

Batf3+/- littermates, whereas CD103+CD11b+ cDCs remained unchanged (Figure 3-5A). Upon

RV infection, while no changes were observed in the cDC frequency (Figure 3-5B) nor in the

CD103+CD11b- cDC population (Figure 3-5A), both Batf3-/- mice and Batf3+/- littermates

exhibited slightly decreased frequencies of CD103+CD11b+ cDCs and increased frequencies of

CD103-CD11b+ cDCs compared with uninfected (UI) genotype matched controls (Figure 3-5A).

Similar results were observed when absolute number of cells was tabulated (Figure 3-6A-D). In

the MLN, we found that absolute numbers of CD103+CD11b- mDCs and CD8+CD11b- rDCs

were significantly decreased, whereas the absolute number of CD103+CD11b+ and CD103-

CD11b+ mDCs was significantly elevated in Batf3-/- mice (Figure 3-6E-K). In terms of other

APCs, we observed no changes in the macrophage population (Figure 3-5C) and a trend toward a

reduction in B cells following RV infection of Batf3-/- mice (Figure 3-5D).

As neonatal mice are highly susceptible to oral RV infection and the specific DC subset(s)

required to initiate neonatal adaptive responses remains to be determined, we also examined DC

populations in Batf3-/- neonates. Accordingly, day 5–6 postnatal Batf3+/- and Batf3-/- littermates

were examined and are referred to hereafter as neonatal mice. At steady state, compared with

littermate controls, neonatal Batf3-/- mice displayed a profound loss of CD103+CD11b- cDCs, as

well as a modest reduction in CD103+CD11b+ cDCs (Figure 3-5E). Similar to adult Batf3-/- mice,

RV infection did not alter the SILP cDC frequency in neonatal mice (Figure 3-5F). Other APCs

such as macrophages and B cells were maintained at similar frequencies at steady state as well as

upon RV infection in Batf3-/- neonates (Figure 3-5G, H, respectively). Together, these results

74

Figure 3-5 Antigen presenting cells in the SILP of adult and neonatal Batf3-/- mice at steady

state and upon RV infection.

A. Percentage of DC subsets as a frequency of DCs in adult mice at 7 d.p.i.. B. Percentage of

DCs as a frequency of mononuclear cells in adult mice at 7 d.p.i.. C. Percentage of macrophages

as a frequency of mononuclear cells in adult mice at 7 d.p.i.. D. Percentage of B cells as a

frequency of mononuclear cells in adult mice at 7 d.p.i.. E. Percentage of DC subsets as a

frequency of DCs in neonatal mice at 7 d.p.i.. F. Percentage of DCs as a frequency of

mononuclear cells in neonatal mice at 7 d.p.i..G. Percentage of macrophages as a frequency of

mononuclear cells in neonatal mice at 7 d.p.i.. H. Percentage of B cells as a frequency of

mononuclear cells in neonatal mice at 7 d.p.i..

UI, uninfected. RV, rotavirus infected. DP, CD103+CD11b+ DC. Results were pooled from 3-4


Data are presented as mean ±SEM. A-C and E-H, Student’s t-test; D, Mann-Whitney test.

*p<0.05, **p<0.01, ***p<0.001, ****p<0.0001, NS= not significant.

75

Figure 3-6 Kinetics of absolute numbers of DCs and DC subsets in the SILP and MLNs of

Batf3+/- and Batf3-/- mice.

A. Kinetics of SILP DC absolute cell number in adult Batf3+/- and Batf3-/- mice. B-D. Kinetics of

SILP DC subset absolute cell number in adult Batf3+/- and Batf3-/- mice. E. Kinetics of MLN

mDC absolute cell number in adult Batf3+/- and Batf3-/- mice. F-H. Kinetics of MLN mDC subset

absolute cell number in adult Batf3+/- and Batf3-/- mice. I. Kinetics of MLN rDC absolute cell

number in adult Batf3+/- and Batf3-/- mice. J. and K. Kinetics of MLN rDC subset cell number in

adult Batf3+/- and Batf3-/- mice.

DP DC, CD103+CD11b+ DCs. Each data point represents a single biological replicate (one

mouse), with 4-7 mice per group. Data are presented as mean ±SEM. Batf3+/- vs. Batf3-/-, Two-

way ANOVA test. RV-infected Batf3-/- vs. uninfected Batf3-/-, Mann-Whitney test. *p<0.05,

**p<0.01, ****p<0.0001, NS= not significant.

76

suggest that adult and neonatal Batf3-/- mice share similarities in SILP APC profiles before and

after RV infection.

3.3.4 Adult and neonatal mice have distinct DC requirements for mounting RV-specific CD8+ T-cell responses

We next examined the anti-RV CD8+ T-cell response in the SILP, the effector site for primed

CD8+ T cells. Both adult and neonatal Batf3-/- mice exhibited a decreased frequency in total

CD8+ T cells compared with Batf3+/- littermates (Figure 3-7A, B, respectively), which was likely

caused by poor proliferation as indicated by a reduced frequency of Ki-67+CD8+ T cells (Figure

3-7C, D, respectively). Moreover, CD8+ T-cell activation (indicated by CD44 staining) was

reduced in RV-infected adult and neonatal Batf3-/- mice compared with Batf3+/- littermates

(Figure 3-7E, F, respectively). Interestingly, although the frequency of VP6357–366-specific CD8+

T cells (Figure 3-7G) and VP6357–366 peptide-induced IFN-producing CD8+ T cells were

decreased in adult Batf3-/- mice, the CD8+ T-cell response to RV was not eliminated (Figure

3-7H). Similar results in adult Batf3-/- mice were observed when absolute number of cells was

tabulated (Figure 3-8). In concordance with these defects in CD8+ T-cell responses, adult Batf3-/-

mice were also found to shed significantly higher levels of RV compared with Batf3+/- littermate

controls, although RV clearance was restored by 8 d.p.i. (Figure 3-7I).

In contrast to what we observed in adult Batf3-/- mice, neonatal Batf3-/- mice were incapable of

mounting antigen-specific CD8+ T-cell responses in the SILP (Figure 3-7J, K). Nevertheless,

neonatal Batf3-/- mice cleared RV with similar kinetics compared to Batf3+/- littermates (Figure

3-7L), suggesting that a RV-specific CD8+ T-cell response is not absolutely required for RV

clearance in neonates. Together, these results suggest that BATF3-dependent DCs are required

for optimal antiviral CD8+ T-cell responses in both adult and neonatal mice, with a more

stringent requirement for BATF3-dependent DCs in neonatal mice.

77

Figure 3-7 CD103+CD11b- DCs are required for optimal anti-RV CD8+ T-cell responses in

Batf3-/- adult and neonatal mice.

A. and B. Percentage of CD8+ T cells as a frequency of SILP mononuclear cells in adult mice A

and in neonatal mice B at 7 d.p.i.. C. and D. Percentage of Ki-67+ cells as a frequency of SILP

CD8+ T cells in adult mice C and in neonatal mice D after in vitro restimulation with VP6357-366

peptide at 7 d.p.i.. E. and F. Percentage of CD44+ cells as a frequency of SILP CD8+ T cells in

adult mice E and in neonatal mice F after in vitro restimulation with VP6357-366 peptide at 7 d.p.i..

G. Percentage of VP6357-366+ cells as a frequency of SILP CD8+ T cells in adult mice at 7 d.p.i..

H. Percentage of IFN+ cells as a frequency of SILP CD8+ T cells in adult mice after in vitro

restimulation with VP6357-366 peptide at 7 d.p.i.. I. Level of RV antigen in the feces of adult mice

measured by ELISA. J. Percentage of VP6357-366+ cells as a frequency of SILP CD8+ T cells

78

neonatal mice at 7 d.p.i.. K. Percentage of IFN+ cells as a frequency of SILP CD8+ T cells in

neonatal mice after in vitro restimulation with VP6357-366 peptide at 7 d.p.i.. L. Level of RV

antigen in the colon contents of neonatal mice measured by ELISA.

UI, uninfected. RV, rotavirus infected. Results were pooled from 3-4 independent experiments.

Each data point represents a single biological replicate (one mouse). Data are presented as mean

±SEM. A, B, D, F and K, Student’s t-test; C, E, G-J, Mann-Whitney test. *p<0.05, **p<0.01,

***p<0.001, ****p<0.0001, NS= not significant.

79

Figure 3-8 SILP CD8+ T-cell responses (absolute numbers) in adult Batf3-/- mice.

A. Absolute number of SILP CD8+ T cells in adult Batf3+/- and Batf3-/- mice at 7 d.p.i.. B.

Absolute number of SILP Ki-67+CD8+ T cells in adult Batf3+/- and Batf3-/- mice at 7 d.p.i.. C.

Absolute number of SILP VP6357-366+ CD8+ T cells in adult Batf3+/- and Batf3-/- mice at 7 d.p.i..

D. Absolute number of SILP IFN+CD8+T cells after in vitro restimulation with VP6357-366

peptide at 7 d.p.i..in adult Batf3+/- and Batf3-/- mice

UI, uninfected. RV, rotavirus infected. Results were pooled from 2 independent experiments.

Each data point represents a single biological replicate (one mouse). Data are presented as mean

±SEM. A-C, student’s t-test.D, Mann-Whitney test. *p<0.05, **p<0.01, ***p<0.001,

****p<0.0001, NS= not significant.

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3.3.5 Polyclonal antiviral Th1 responses in neonatal mice are BATF3-dependent

Although anti-RV CD4+ T-cell responses are dispensable for RV clearance, they do have a role

in providing help for the anti-RV IgA response (Angel et al., 2007). To evaluate the CD4+ T-cell

profile, we used the mitogen phorbol 12-myristate 13-acetate (PMA) and the calcium ionophore

ionomycin for in vitro restimulation of SILP CD4+ T cells following RV infection. Although the

frequency of CD8+ T cells was reduced in the SILP of Batf3-/- mice (Figure 3-7A, B), the

frequency of CD4+ T cells was comparable between Batf3-/- mice vs. Batf3+/- littermates (both

neonatal and adult), with or without RV infection (Figure 3-9A, B). We found that adult mice of

both genotype exhibited an increase in IFN production in response to RV infection (Figure

3-9C). In contrast, in neonatal mice, RV infection provoked CD4+ T cells to produce IFN only

in neonatal Batf3+/- mice but not in neonatal Batf3-/- mice (Figure 3-9D). These results suggest a

requirement for BATF3-dependent DCs to elicit an anti-RV Th1 response in neonatal mice but

not in adult mice. In contrast with the IFN results, adult Batf3-/- mice exhibited increased IL-17-

producing CD4+ T cells (Figure 3-9E), and neonatal Batf3-/- mice exhibited the same trend albeit

less pronounced (Figure 3-9F). Together, these results imply that loss of Batf3 skews the balance

of T helper type 1 (Th1) versus Th17 cells.

3.3.6 Intact local and systemic anti-RV IgA responses in cDC-deficient mice

Th17 cells have been implicated in promoting antigen-specific IgA responses (Hirota et al.,

2013). Given that Batf3-/- mice exhibited a trend toward increased Th17 responses (Figure 3-9E,

F), we speculated that Batf3-/- mice may display an intact or even enhanced RV-specific IgA

response, which could lead to the resolution RV infection. Indeed, at the time points examined,

adult Batf3-/- mice generated RV-specific IgA with similar kinetics as Batf3+/- littermates both

locally and systemically (Figure 3-10A, C). In terms of the magnitude of the RV-specific

response as expressed in terms of a titer (see Methods section), we found that Batf3-/- mice

produced slightly more IgA in the feces at 14 d.p.i. but less IgA in the serum at 28 d.p.i.

compared with Batf3+/- mice (Figure 3-10B, D), but overall there was no obvious defect in the

IgA response in Batf3-/- mice. Moreover, neither local nor systemic anti-RV IgA responses were

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Figure 3-9 Alteration of Th1 and Th17 responses in adult and neonatal Batf3-/- mice.

A. and B. Percentage of CD4+ T cells as a frequency of SILP mononuclear cells in adult mice A

and neonatal mice B at 7 d.p.i.. C. and D. Percentage of IFN+ cells as a frequency of SILP CD4+

T cells in adult mice C and neonatal mice D at 7 d.p.i.. E. and F. Percentage of IL-17A+ cells as

a frequency of SILP CD4+ T cells in adult mice E and neonatal mice F at 7 d.p.i..

UI, uninfected. RV, rotavirus infected. Each data point represents an individual biological

replicate (one mouse) pooled from 2-3 independent experiments. Data are presented as mean

±SEM. A, D and F, Student’s t-test; B, C and E, Mann-Whitney test. *p<0.05, ***p<0.001,

****p<0.0001, NS= not significant.

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affected by loss of cDCs in DTx-treated Zbtb46-DTRWT chimeric mice (Figure 3-10E-H).

These results suggest that, unlike pDCs, which have an important role in activating B cells to

become RV-specific IgA-producing cells in both humans and mice (Deal et al., 2013), cDCs are

dispensable for generating antigen-specific IgA in adult mice in the context of RV infection.

Finally, similar to adult mice, we found that neonatal mice were able to mount anti-RV IgA

responses (Figure 3-10I-L). In terms of raw optical density measurements, Batf3-/- neonates

exhibited a significant increase in the level of anti-RV IgA in the feces (16 d.p.i.) and in the

serum (14 and 16 d.p.i.) compared with Batf3+/- neonatal littermates (Figure 3-10I, K). However,

the RV-specific IgA titer in the feces and serum was comparable between Batf3+/- and Batf3-/-

neonatal mice (Figure 3-10J, L). These data suggest that humoral antiviral responses are intact in

the absence of BATF3-dependent DCs in neonatal mice.

3.3.7 CD103+CD11b+ DCs are not required for mounting anti-RV adaptive immune responses

CD103+CD11b+ DCs have been reported as a heterogeneous mixture of pre-DC-derived cDCs

and monocyte-derived DCs (Satpathy et al., 2012). We hypothesized that CD103+CD11b+ DC

may have a secondary role in cross-presenting RV epitope(s) to CD8+ T cells in order to

compensate for the loss of CD103+CD11b- DCs in Batf3-/- mice. We tested this hypothesis by

examining the RV-specific CD8+ T-cell response in huLangerin-DTA mice, which lack

CD103+CD11b+ DCs in the SILP (Figure 3-11A) and MLNs (Welty et al., 2013). We found that

the antigen-specific CD8+ T-cell response was not impaired in huLangerin-DTA mice (Figure

3-11B-E). In the neonatal setting, the total frequency of SILP CD8+ T cells post-RV infection in

huLangerin-DTA mice was comparable to UI controls (Figure 3-11F). Although Ki-67 staining

revealed limited proliferation of CD8+ T-cells in neonatal huLangerin-DTA mice (Figure

3-11G), we nevertheless observed comparable frequencies RV-specific CD8+T cells between

neonatal control and huLangerin-DTA mice (Figure 3-11H). Moreover, viral clearance and IgA

production were comparable between adult huLangerin-DTA and control mice (Figure 3-11I-K).

Therefore, although this experiment does not rule out a compensatory role of CD103+CD11b+

DCs in Batf3-/- mice, these data imply that CD103+CD11b+ DCs are dispensable for presenting

RV-antigen to CD8+ T cells in huLangerin-DTA mice.

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Figure 3-10 cDCs are dispensable for the induction and maintenance of local and systemic

antiviral IgA.

A. and B. Level of anti-RV IgA in the feces measured by ELISA in adult mice comparing Batf3-/-

vs. Batf3+/- littermates A and RV-specific IgA titers at 7, 14 d.p.i. B. C. and D. Level of anti-RV

IgA in the serum measured by ELISA in adult mice comparing Batf3-/- vs. Batf3+/- littermates C

and RV-specific IgA titer at 28 d.p.i. D. E. and F. Level of anti-RV IgA in the feces measured by

ELISA in adult mice comparing PBS- vs. DTx- treated Zbtb46-DTRWT chimeric mice E and

RV-specific IgA titers at 7, 14 d.p.i. F. G. and H. Level of anti-RV IgA in the serum measured

by ELISA in adult mice comparing PBS- vs. DTx- treated Zbtb46-DTRWT chimeric mice G

and RV-specific IgA titer at 28 d.p.i. H. I. and J. Level of anti-RV IgA in the feces measured by

ELISA in neonatal mice comparing Batf3-/- vs. Batf3+/- littermates I and RV-specific IgA titers at

14, 16 d.p.i. J. K. and L. Level of anti-RV IgA in the serum measured by ELISA in neonatal

mice comparing Batf3-/- vs. Batf3+/- littermates K and RV-specific IgA titers at 14, 16 d.p.i. L.


individual biological replicate (one mouse) pooled from 2-3 independent experiments. Data are

presented as mean ±SEM. B, D, J and L, Student’s t-test; F, H, I and K, Mann-Whitney test.

*p<0.05, **p<0.01, ***p<0.001, NS= not significant.

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Figure 3-11 CD103+CD11b+ cDCs are not required for anti-RV CD8+ T-cell responses in

the SILP of adult and neonatal huLangerin-DTA mice.

A. Percentage of DC subsets as a frequency of DCs in adult huLangerin-DTA mice at 7 d.p.i.. B.

Percentage of CD8+ T cells as a frequency of mononuclear cells in adult huLangerin-DTA mice

at 7 d.p.i.. C. Percentage of Ki-67+ cells as a frequency of CD8+ T cells after in vitro

restimulation with VP6357-366 peptide in adult huLangerin-DTA mice at 7 d.p.i.. D. Percentage of

VP6357-366 + cells as a frequency of CD8+ T cells in adult huLangerin-DTA mice at 7 d.p.i..

E. Percentage of IFN+ cells as a frequency of CD8+ T cells after in vitro restimulation with

VP6357-366 peptide in adult huLangerin-DTA mice at 7 d.p.i.. F. Percentage of CD8+ T cells as a

frequency of mononuclear cells in neonatal huLangerin-DTA mice at 7 d.p.i.. G. Percentage of

Ki-67+ cells as a frequency of CD8+ T cells after in vitro restimulation with VP6357-366 peptide in

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neonatal huLangerin-DTA mice at 7 d.p.i.. H. Percentage of VP6357-366 + cells as a frequency of

CD8+ T cells in neonatal huLangerin-DTA mice at 7 d.p.i.. I. Level of RV antigen in the feces in

adult huLangerin-DTA mice measured by ELISA. J. and K. Level of RV-specific IgA in the

feces J and in the serum K in adult huLangerin-DTA mice measured by ELISA.

UI, uninfected. RV, rotavirus infected. Control, huLangerin-DTA- littermate. Each data point

represents an individual biological replicate (one mouse) pooled from 3 independent

experiments. Data are presented as mean ±SEM. A, B, D and E, Student’s t-test; C and F-H,

Mann-Whitney test. *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001, NS= not significant.

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3.4 Discussion

In this study, we found that cDCs are required for optimal CD8+ T-cell responses to RV infection

in the small intestine. Furthermore, we demonstrated that BATF3-dependent DCs are the major

DC subset responsible for priming CD8+ T cells in both adult and neonatal mice. Interestingly,

we observed residual CD8+ T-cell proliferation and antigen-driven IFN production in adult

Batf3-/- mice. In sharp contrast to adult Batf3-/- mice, neonatal Batf3-/- mice exhibit a complete

absence of a CD8+ T-cell response to RV, indicating that neonatal mice, strictly require BATF3-

dependent DCs for priming mucosal CD8+ T cells compared with adult mice. Additionally, local

and systemic antiviral IgA production were intact in DTx-treated Zbtb46-DTR chimeric mice,

suggesting a dispensable role of cDCs in mounting antiviral IgA responses.

As the expression of Zbtb46 is restricted to cDCs but not pDCs or macrophages, the Zbtb46-DTR

chimeric mouse model is ideal for studying the role of cDC in priming CD8+ T cells to RV. It

has been previously shown that TLR engagement may downregulate Zbtb46 expression,

resulting in incomplete depletion of cDC (Meredith et al., 2012b). To avoid this issue, we treated

chimeric mice with DTx 1 day prior to RV infection to pre-deplete cDCs, and we further

eliminated newly generated cDCs by subsequently treating chimeric mice with DTx every 1–2

days. Moreover, we found that depletion of cDC was equally potent in UI vs. RV-infected

chimeric mice (Figure 3-1B), suggesting that RV infection does not affect the DC depletion

efficiency. To further bolster our findings using the Zbtb46-DTR system, we compared our

results with DTx-treated Zbtb46-DTR chimeric mice with DTx-treated CD11c-DTRWT

chimeric mice (in this system DTx treatment will deplete DCs and other CD11c-expressing

cells). We found that DTx treatment of CD11c-DTRWT chimeric mice resulted in a similar

phenotype as DTx treatment of Zbtb46-DTR chimeric mice in terms of cDC depletion and the

impact on the CD8+ T-cell response (Figure 3-12A-I). Therefore, although CD11c-DTRWT

chimeric mice are not ideal for studying cDC-specific effects, the parallel phenotypes observed

between this system and the Zbtb46-DTR system confirms that the latter chimeras can be reliably

used to deplete cDC in the context of RV infection.

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Figure 3-12 Similar anti-RV CD8+ T cell responses are observed in CD11c-DTRWT and

Zbtb46-DTRWT chimeric mice.

A. Percentage of SILP DCs as a frequency of mononuclear cells from chimeric mice treated with

PBS or DTx at 7 d.p.i.. B-D. Percentage of SILP DC subsets as a frequency of DCs from

chimeric mice treated with PBS or DTx at 7 d.p.i.. E. Percentage of CD8+ T cells as a frequency

of mononuclear cells from chimeric mice treated with PBS or DTx at 7 d.p.i.. F. Percentage of

Ki-67+ cells as a frequency of CD8+ T cells from chimeric mice treated with PBS or DTx at 7

d.p.i.. G. Percentage of VP6357-366+ cells as a frequency of CD8+ T cells from chimeric mice

treated with PBS or DTx at 7 d.p.i.. H. Percentage of IFN+ cells as a frequency of CD8+T cells

from chimeric mice treated with PBS or DTx after in vitro restimulation with VP6357-366 peptide

at 7 d.p.i.. I. Level of RV antigen in the feces from chimeric mice treated with PBS or DTx

measured by ELISA. J. Percentage of SILP granulocytes as a frequency of total cells from

chimeric mice treated with PBS or DTx at 7 d.p.i..

UI, uninfected. RV, rotavirus infected. ZDC, Zbtb46-DTRWT chimeric mice. CD11c, CD11c-

DTRWT chimeric mice. DTx, diphtheria toxin. Results were pooled from 2-3 independent

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experiments. Each data point represents a single biological replicate (one mouse). Data are

presented as mean ±SEM. Student’s t-test. *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001, NS=

not significant.

89

Another concern with DTx-based system is that an increased level of splenic neutrophils and

monocytes has been reported upon DTx treatment of CD11c-DTR and Zbtb46-DTR chimeric

mice (Meredith et al., 2012a; Tittel et al., 2012). We found that the frequency of granulocytes in

the SILP was unaltered after DTx treatment in Zbtb46-DTR chimeric mice with or without RV

infection, although interestingly, an increase in granulocytes was observed in CD11c-DTR

chimeric mice (Figure 3-12J). Therefore, the use of Zbtb46-DTR chimeric mice has provided us

with the opportunity to compare the RV-specific CD8+ T-cell response in mice that lack all cDC

(Zbtb46-DTR chimeric mice), and when coupled with non-DTx systems that lack select subsets

of cDC (Batf3-/- mice), we may discover which cDC are correlated with CD8+ T-cell responses to

RV infection.

We report here that BATF3-dependent DCs are the principal DC responsible for cross-presenting

RV antigen to intestinal CD8+ T cells. Our results are consistent with Cerovic et al. (Cerovic et

al., 2015) who demonstrated that lymph-borne CD103+CD11b-CD8+ DCs can cross-prime

CD8+ T cells against intestinal epithelial cell (IEC)-derived cellular antigens within the MLNs.

As mentioned, some CD8+ T cells can still undergo proliferation in adult Batf3-/- mice upon viral

challenge. This could either be due to restoration of CD103+CD11b- DCs following RV infection

or alternatively a compensatory cell may prime CD8+ T cells in adult Batf3-/- mice. In support of

the former possibility, Tussiwand et al. (Tussiwand et al., 2012) reported that CD103+CD11b-

DCs can be restored upon IL-12 administration or pathogen-induced IL-12. However, we found

no evidence of a restoration of CD103+CD11b- DCs in the SILP or CD103+CD11b- mDCs and

CD8+CD11b- rDCs in the MLNs during the RV infection time course in Batf3-/- adult mice.

Thus the compensatory IL-12-mediated development of Batf3-independent CD103+CD11b- DC

appears not to be a feature of the RV system during the first week of infection, although it could

be the case at earlier time points.

We favor the hypothesis that an alternative APC can induce some proliferation of RV-specific

CD8+ T cells in Batf3-/- mice. Compared with Batf3+/- littermates, the residual CD8+ T-cell

response to RV correlates with significant increases in CD103+CD11b+ and CD103-CD11b+

mDCs in the MLN as well as CD103-CD11b+ DCs in the SILP (Figure 3-6). As these DCs are

not increased in DTx-treated Zbtb46-DTRWT chimeric mice (Figure 3-2), which completely

lack a CD8+ T-cell response to RV, the residual CD8+ T-cell response to RV in adult Batf3-/-

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mice may therefore be due to compensatory increases in CD103-CD11b+ DCs in the SILP and/or

CD103+CD11b+ and CD103-CD11b+ mDCs in the MLN. Finally, another possible explanation

for the residual CD8+ T-cell response in adult Batf3-/- mice is that atypical/non-professional

APCs such as IECs may also present antigens to T cells (Mayer, 2000). As RV replicates in

IECs, it would be interesting to know whether IECs can present viral antigens to CD8+ T cells in

the absence of BATF3-dependent DCs.

Neonates are highly susceptible to infections with pathogens, such as RV. However, the

mechanism(s) responsible for impaired infant immunity remain elusive. It has been reported that

neonatal T cells are prone to a Th2 bias (Adkins and Du, 1998), and this phenomenon could be

explained by a lack of maturity of neonatal DCs (i.e., lower level of major compatibility complex

(MHC) class II, co-stimulatory molecule CD86 and key cytokines such as IL-12, compared with

adult DCs) (Willems et al., 2009). However, we found that neonatal mice are able to elicit a

robust antigen-specific CD8+ T-cell response, including the production of IFN, indicating that

neonatal DCs are fully capable of priming CD8+ T cells during infection. On the other hand,

unlike adult Batf3-/- mice, Batf3-/- neonates were not able to mount antigen-specific CD8+ T-cell

responses. As SILP T cells from neonatal mice are relatively naive compared with adult T cells,

which can be antigen-experienced, we hypothesize that the activation threshold for neonatal T

cells may be higher, and adequate priming would require a particularly specialized DC subtype,

such as BATF3-dependent DCs.

In spite of the absence of an antigen-specific CD8+ T-cell response to RV infection, Batf3-/-

neonatal mice can nevertheless clear RV with similar kinetics as control mice, suggesting other

antiviral responses beyond CD8+ T-cell responses exist to resolve RV infection. It is likely that

innate immunity has a key role in controlling RV infection in neonates. For example, IL-22 and

IFN produced by type 3 innate lymphoid cells can synergistically stimulate IEC antiviral

responses thereby contributing to RV clearance in neonates (Hernández et al., 2015). We

detected IL-22 in neonatal SILP RORt+ cells at the steady state by flow cytometry and by

reverse transcriptase-PCR (RT-PCR), although these levels were not greatly increased after RV

infection (Figure 3-13A-C). As a positive control, IL-22 induction was detected in the colonic

tissues from mice infected with C. rodentium at 6 d.p.i by quantitative PCR (Figure 3-13C). We

also detected a marked upregulation of IFNin intestinal epithelial lymphocytes after RV

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Figure 3-13 Innate immune responses in neonatal Batf3+/- and Batf3-/- mice with and

without RV infection.

A. Representative flow cytometry plots of IL-22-producing ILC3 (pre-gated on CD3-CD11b-

B220- live singlet lymphocytes) in the SILP of neonatal Batf3+/- and Batf3-/- mice at 1 d.p.i.. B.

Percentage of IL-22-producing RORt- and RORt+ cells as a frequency of SILP Lin

(CD3/CD11b/B220)- cells of Batf3+/- and Batf3-/- mice. C. IL-22 message level in the SILP of

neonatal Batf3+/- and Batf3-/- mice at 1 d.p.i. and in the colonic tissues of adult WT mice 6 days

post C. rodentium infection. D. IFN message level in the IECs of neonatal Batf3+/- and Batf3-/-

mice at 1.d.p.i..

UI, uninfected. RV, rotavirus infected. Citro, C. rodentium infected. Each data point represents a

single biological replicate (one mouse), with 4-5 mice per group. Mann-Whitney test. *p<0.05,

**p<0.01, NS= not significant.

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infection that was comparable between Batf3+/- and Batf3-/- neonatal mice (Figure 3-13D).

Therefore, early innate cytokine production is intact in Batf3-/- neonatal mice, possibly

explaining the normal viral clearance in these mice.

In addition to IL-22 and IFN, pDCs from 7-day postnatal mice are capable of producing similar

amounts of type I IFN as their adult counterparts (Willems et al., 2009), making type I IFN

another candidate for defense against RV infection in neonates. This type I IFN production may

promote neonatal RV-specific IgA responses, and indeed, the local and systemic anti-RV IgA

responses in Batf3-/- neonatal mice are not only intact but were in fact slightly more robust than

the Batf3+/- neonatal mice. However, it is important to note that RV-specific IgA is the

predominant antibody species in lacteal secretions from mice naturally infected with RV or

experimentally infected through the oral route (Sheridan et al., 1984); thus passively acquired

IgA may contribute to the presence of RV-specific IgA in neonates and help resolve RV

infection. It would be interesting to determine whether the robust IgA response in neonates is due

to an elevated type I IFN production by pDCs, in the absence of BATF3-dependent cDCs.

In addition to impaired CD8+ T-cell immunity in Batf3-/- mice, polyclonal CD4+ T-cell cytokine

production is also skewed. During L. major infection, it has been reported that the protective Th1

immune response is severely hindered in Batf3-/- mice, correlating with impaired IL-12

production and a reduction in CD103+ DC numbers (Martínez-López et al., 2015). Consistent

with previous studies (Luda et al., 2016; Muzaki et al., 2016), we found that intestinal Th1

responses were diminished in adult Batf3-/- mice in the steady state and neonatal Batf3-/- mice

challenged with RV. On the other hand, consistent with previous results (Luda et al., 2016), we

found that both adult and neonatal Batf3-/- mice exhibit a trend toward increased Th17 cell

frequency compared with Batf3+/- littermates. Aychek et al. (Aychek et al., 2015) have reported

that during C. rodentium infection, colonic CD103-CD11b+ DC- and macrophage-derived IL-23

can suppress IL-12 production by CD103+CD11b- DCs, resulting in the generation of IFN

producing ex-Th17 cells. We hypothesize that, in the absence of BATF3-dependent DCs,

diminished CD103+CD11b- DC derived IL-12 and enhanced CD103-CD11b+ DC-derived IL-23

results in skewing of cytokines produced by Th cells. Further investigations are need to address

this possibility, with the caveat that a role for BATF3-dependent DCs in modulating intestinal Th

93

responses is likely influenced by the choice of stimuli/pathogen and the age of the mice (neonate

vs. adult).

In adults, the antiviral IgA response in cDC-deficient mice as well as Batf3-deficient mice is

comparable to that of littermate controls. This suggests that IgA responses can occur in the

absence of pre-DC derived cDCs. The normal humoral response we observed in Batf3-/- mice is

consistent with the results from Hildner et al. (Hildner et al., 2008) who found that normal

WNV-specific IgM and IgG responses were induced in WNV-challenged Batf3-/- mice. We

speculate that the antiviral IgA response in cDC-deficient mice may compensate for the

abrogated RV-specific CD8+ T-cell response, resulting in only a transient delay in viral

clearance.

Our study mainly focused on characterizing the function of DC subsets in primary RV infection.

However, it would be interesting to expand our study to examine how cDC subsets affect

secondary RV challenge, eventually translating our study toward a vaccine design strategy. The

two current licensed RV vaccines on the market, Rotarix and RotaTeq, reduce RV-related

morbidity and mortality in developed countries; however, they are not as efficient in resource-

poor countries (Angel et al., 2007; Glass et al., 2006). Additionally, vaccine development has

been highly empirical, leaving large gaps in our understanding of how they induce protection

(Angel et al., 2007). Boosting antiviral CD8+ T-cell responses by modulating BATF3-dependent

DCs might help generate long-term T-cell memory in non-responsive individuals.

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Chapter 4

LTR deficiency in radio-sensitive compartments results in skewed T-cell responses during a mucosal viral infection

95

4.1 Abstract

The lymphotoxin signaling pathway plays an important role in the homeostasis and function of

peripheral and mucosal dendritic cells, and dendritic cell-intrinsic lymphotoxin receptor

(LTR) expression is required for optimal responses to opportunistic intestinal bacteria. Mice

deficient in LT display prolonged rotavirus antigen shedding in the feces. However, it is

unknown whether dendritic cell-intrinsic LTR signaling is required for responses to intestinal

viral infections. We explored this question by orally administrating murine rotavirus to chimeric

mice that lack LTR signaling in the radio-sensitive compartment but retain lymphoid tissues.

We found that although clearance of rotavirus was unimpaired in the Ltbr-/-wild-type (WT)

chimeric mice compared with WTWT chimeric mice, IFN- producing CD8+ and CD4+ T

cells were significantly increased in the SILP of Ltbr-/-WT chimeric mice. In contrast, IL-17-

producing CD4+ T cells were reduced in Ltbr-/-WT chimeric mice in the steady state, and this

reduction persisted after rotavirus inoculation. In spite of this altered cytokine profile in the SILP

of Ltbr-/-WT chimeric mice, the local production of rotavirus-specific IgA was unperturbed.

Collectively, our results demonstrate that LTR signaling in radio-sensitive cells regulates the

balance of IFN and IL-17 cytokine production within the SILP; however, these perturbations do

not affect mucosal antiviral IgA responses.

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4.2 Introduction

LT is a TNF family cytokine that can exist as a membrane-bound LT12 heterotrimer or a

soluble LT3 trimer. Whereas LT3 binds to TNFR I and II, LT12 signals exclusively

through the LTR. Another LTR ligand TNF superfamily member (LIGHT) can also deliver

signals through HVEM. Both LT12 and LIGHT are expressed primarily on lymphocytes,

whereas LTR is expressed on radio-resistant epithelial and stromal cells, as well as radio-

sensitive myeloid cells (Browning, 2008; Ware, 2005).

LT12/LTR signaling is critically required for lymphoid tissue organogenesis and the

maintenance of secondary lymphoid structures (De Togni et al., 1994; Fütterer et al., 1998). In

addition, LTR signaling is involved in host responses to infections in mice, including responses

to lymphocytic choriomeningitis virus (Puglielli et al., 1999), L. monocytogenes, and

Mycobacteria tuberculosis (Ehlers et al., 2003). Moreover, LTR signaling has been found to

regulate acute inflammatory reactions, such as dextran sulfate sodium-induced colitis (Jungbeck

et al., 2008; Stopfer et al., 2004), and to mediate tumor cell apoptosis (Rooney et al., 2000).

Hence, LTR signaling is involved in innate and adaptive immune responses.

Recently, LTR signaling is shown to play a protective role in the immune response to a

mucosal bacterial infection, specifically in the clearance of the attaching and effacing bacterium

C. rodentium (Satpathy et al., 2013; Spahn et al., 2004; Tumanov et al., 2011; Wang et al.,

2010), a mouse model used to understand the consequences of enteropathogenic and

enterohemorrhagic Escherichia coli in humans. Within the radio-resistant compartment, LTR

signaling in intestinal epithelial cells (IECs) is required for the recruitment of neutrophils to the

infection site via production of CXCL1 and CXCL2 chemokines (Wang et al., 2010) and for

protection against epithelial injury via a mechanism that depends on IL-23 (Macho-Fernandez et

al., 2015). Within the radio-sensitive compartment, LTR signaling in LP DCs drives the

production of IL-22 from RORt+ ILCs to maintain barrier integrity, thus providing protection

against C. rodentium (Tumanov et al., 2011). Furthermore, Notch2-dependent CD103+CD11b+

cDCs, which play a critical role in producing IL-23 in response to C. rodentium infection, are

also partially LTR dependent when examined in the context of competitive mixed BM chimeras

(Satpathy et al., 2013). Collectively, these findings support the idea that LTR signaling in radio-

97

resistant and -sensitive compartments is crucial for intestinal homeostasis to limit mucosal

damage caused by bacterial invasion.

Whereas the importance of LTR signaling in host defense against C. rodentium infection in the

colon is well characterized, the role of DC-intrinsic LTR signaling in viral clearance within the

small bowel is less clear. RV infection is a well-defined model system for studying viral

infection in the small intestine, as RV predominantly infects and replicates within mature

epithelial cells on the tip of the small intestinal villi (Angel et al., 2007). Before the introduction

of RV vaccines, RV was a major cause of severe dehydrating diarrhea in infants and children <5

y old (Angel et al., 2007). Previous studies have shown that Lta-/- mice have prolonged intestinal

RV infection corresponding with a defect in anti-RV IgA production, and remarkably, these

lymphoid-tissue deficient mice do eventually mount an IgA response and can clear the virus

(Lopatin et al., 2013). However, this study did not dissect a role for LTR versus TNFR

signaling nor whether the key LTresponding cell type was a DC or an epithelial cell.

Although RV infection in adult mice is asymptomatic, viral particle shedding in the feces is

detectable and correlates with the presence and replication of the virus. The clearance of RV in

adult mice is dependent on cellular and humoral responses, as T cell-deficient mice (TCR-/-,

/TCR-/-, 2m-/-, and anti-CD8 mAb-treated C57BL/6) and IgA-/- mice have varying degrees

of delayed viral clearance. However, even these severely immunocompromised mice can

eventually resolve the RV infection (Blutt et al., 2012; Franco and Greenberg, 1995, 1997), with

the exception of recombination activating gene 2 (Rag2)-/- mice that become chronically infected

and continuously shed viral antigen (Franco and Greenberg, 1995). Although most

immunocompromised mice can clear RV, it is nevertheless a very useful model for studying the

dynamics of CD8+ T cell priming to a small intestinal tropic virus. Here, we focus on RV

infection in adult mice to discern a role for DC-intrinsic LTR signaling on CD8+ T cell priming

in the gut. To evaluate this, we generated Ltbr-/- WT BM chimeric mice and monitored SILP T

cell responses as a readout of DC function, as DCs have been shown to be important for priming

naïve T cells in the gut-associated lymphoid tissues (Mora et al., 2003). Collectively, the results

from our study indicate that loss of LTR signaling in the radio-sensitive compartment shifts the

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gut microenvironment from a Th17- to a Th1-dominant state; yet, this alteration in cytokine

production does not affect the local anti-RV IgA response.

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4.3 Results

4.3.1 Ltbr-/- chimeric mice clear RV with normal kinetics

As mentioned, even highly immunocompromised adult mice can clear RV infections as a result

of redundant humoral and cellular mechanisms. To ascertain if clearance of RV infection is

affected by LTR deficiency in radio-sensitive compartments, we generated Ltbr-/-WT BM

chimeras. At 8–10 wk post-BM reconstitution, Ltbr-/-WT and WT WT chimeras were

inoculated with murine RV by oral gavage, respectively. We found that the kinetics of fecal viral

shedding in the LtbrWT chimeras were similar to those in the WTWT chimeras (Figure

4-1). Thus, as expected, LTR signaling in the radio-sensitive compartment is dispensable for

viral clearance in the intestine.

4.3.2 Ltbr-/- chimeric mice generate more IFN-secreting CD8+ T cells during primary RV infection

The RV system is an excellent system for examining CD8+ T cell responses within the small

intestinal environment (Jaimes et al., 2005). Although Ltbr-/-WT chimeric mice can clear RV

with comparable kinetics compared with WTWT chimeric mice, nevertheless, we followed the

RV-specific CD8+ T cell response to determine if a DC-intrinsic LTR signaling pathway is

required for CD8+ T cell priming, expansion, and/or effector function in response to a mucosal

viral infection. In the current study, we examined the anti-RV CD8+ T cell response at 7 d.p.i. in

the SILP, as this was reported previously as the peak of the intestinal RV-specific CD8+ T cell

response (Jaimes et al., 2005). A gating strategy for total CD4+ and CD8+ T cells in the SILP is

depicted in Figure 4-2. At 7 d.p.i., the proportion of CD8+ T cells as a frequency of total

mononuclear cells was significantly higher in the SILP of Ltbr-/-WT chimeras compared with

WTWT control chimeric mice (Figure 4-3A). Likewise, the percentage of SILP CD8+ T cells

as a frequency of total mononuclear cells that were positive for intracellular Ki-67 staining (an

indicator of cell proliferation) was higher in the Ltbr-/-WT chimeras (Figure 4-3B). However,

there was no difference in SILP CD8+ T cell proliferation when measured as a frequency of the

total CD8+ T cell population (Figure 4-3C), suggesting that the observed increase in frequency of

proliferating CD8+ T cells is a result of an over-representation of CD8+ T cells within the SILP

rather than increased proliferation within the SILP. These results suggested that LTR deficiency

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Figure 4-1 LTR deficiency in the radio-sensitive compartment does not affect local viral

clearance.

Level of RV antigen shedding in the feces was measured over time by ELISA. Data show the

OD reading at 450 nm of individual samples pooled from 3 independent experiments ± SEM.

Samples for 0-7 d.p.i, were collected from 17-20 mice, while samples for 9-20 d.p.i. were

collected from 6-7 mice. UI, uninfected; RV, RV-infected. No significant difference in mean OD

values for viral antigen shedding was observed at any time point.

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Figure 4-2 Gating strategy for the SILP CD4+ and CD8+ T cells.

Gating strategy for all flow cytometry experiments. SILP cells gated in the following orders:

mononuclear cells, singlets, live cells, CD3+ cells and CD4+ versus CD8+ T cells.

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Figure 4-3 Frequency and proliferation status of CD8+ T cells in RV-infected Ltbr-/-

chimeric mice.

WTWT and Ltbr-/- WT (KOWT) chimeric mice were sacrificed at 7 d.p.i.. After

PMA/ionomycin in vitro restimulation for 6 h, SILP CD8+ T cells were analyzed by flow

cytometry. A. Percentage of total CD8+ T cells as a frequency of SILP mononuclear cells. B.

Percentage of Ki-67+CD8+ T cells as a frequency of SILP mononuclear cells. C. Percentage of

Ki-67+ as a frequency of total SILP CD8+ T cells.

UI, uninfected; RV, RV-infected. Each point represents individual mouse pooled from 3

independent experiments. Data presents as average ± SEM. Mann-Whitney non parametric test.

NS, not significant, **p< 0.01, ***p<0.001, **** p< 0.0001

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in the radio-sensitive compartment has the capacity to affect CD8+ T cell accumulation but not

expansion in the SILP after viral infection.

We next examined the quality of the CD8+ T cell response in Ltbr-/-WT versus WTWT

chimeric mice. In vitro polyclonal stimulation revealed that IFN secretion was more robust in

SILP CD8+ T cells derived from Ltbr-/-WT chimeras compared with SILP CD8+ T cells

derived from WTWT chimeras when examined as a frequency of total CD8+ T cells or total

mononuclear cells in the SILP (Figure 4-4A-C). To examine SILP CD8+ T cell responses to

specific antigens, we then measured intracellular IFN production from CD8+ T cells stimulated

with VP6357–366, one of the immunodominant RV epitopes recognized by H-2b-restricted CD8+ T

cells (Jaimes et al., 2005). Although the percentage of IFN-producing CD8+ T cells was

comparable between Ltbr-/-WT and WTWT chimeras when expressed as a frequency of the

total CD8+ T cell population, the percentage of IFN-secreting CD8+ T cells was higher in the

Ltbr-/-WT chimeras when expressed as a frequency of total mononuclear cells in the SILP

(Figure 4-4A, D, and E). Similar results were obtained when CD8+ T cells were stimulated with

VP733–40, another immunodominant epitope recognized by H-2b-restricted CD8+ T cells (Figure

4-4F and G) (Franco and Greenberg, 1999; Jaimes et al., 2005). The overall increase in VP6357–

366- and VP733–40-specific, IFN-producing CD8+ T cells is likely a result of the observed

increase in SILP CD8+ T cells present in the Ltbr-/-WT chimeras (Figure 4-3A). Therefore,

LTR signaling in the radio-sensitive compartment is dispensable for VP6357–366- and VP733–40-

specific CD8+ T cell responses after RV challenge. However, LTR signaling in the radio-

sensitive compartment may be required to regulate other RV-specific CD8+ T cell IFN

responses besides those induced by VP6357–366 and VP733–40 epitopes, as reflected by the increase

in CD8+ T cell IFN production induced by polyclonal stimulation (Figure 4-4A-C).

4.3.3 Polyclonal CD4+ T cell cytokine profiles are skewed in the Ltbr-/- chimeric mice at steady state and after RV infection

It has been shown that depletion of CD4+ T cells results in a delay in the generation of RV-

specific intestinal IgA, which is the principal effector of long-term protection against RV re-

infection (Angel et al., 2007, 2012; Franco and Greenberg, 1999). Therefore, we evaluated

polyclonal CD4+ T cell responses to RV infection in Ltbr-/-WT chimeras at steady state and

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Figure 4-4 Assessment of IFN-producing CD8+ T cells in Ltbr-/- chimeric mice induced by

mitogen or viral peptide stimulation.

WTWT and Ltbr-/- WT (KOWT) chimeric mice were sacrificed at 7 d.p.i..

A. Representative flow cytometry data showing IFN-producing SILP CD8+ T cells after

PMA/ionomycin (up row) and VP6357-366 peptide (bottom row) in vitro restimulation for 6 h (pre-

gated on CD8+ T cells). B and C. After PMA/ionomycin in vitro restimulation for 6 h, SILP

CD8+ T cells were analyzed by flow cytometry. B. Percentage of IFN+ as a frequency of SILP

CD8+ T cells; C. percentage of IFN+CD8+ T cells as a frequency of total SILP mononuclear

cells. D and E. After VP6357-366 peptide in vitro restimulation for 6 h, SILP CD8+ T cells were

analyzed by flow cytometry. D. Percentage of IFN+ as a frequency of SILP CD8+ T cells; E.

percentage of IFN+CD8+ T cells as a frequency of total SILP mononuclear cells. F and G. After

VP733-40 peptide in vitro restimulation for 6 h, SILP CD8+ T cells were analyzed by flow

cytometry. F. Percentage of IFN+ as a frequency of SILP CD8+ T cells; G. Percentage of

IFN+CD8+ T cells as a frequency of total SILP mononuclear cells.

UI, uninfected; RV, RV-infected. Each point represents individual mouse pooled from 2-3


NS, not significant, *p<0.05, **p< 0.01, ***p<0.001, **** p< 0.0001

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during RV infection. Unlike CD8+ T cells, the percentage of CD4+ T cells in the SILP was

comparable between Ltbr-/-WT and WTWT chimeras at steady state and at 7 d.p.i. (Figure

4-5A). However, the proliferation of CD4+ T cells was increased significantly in the Ltbr-/-WT

chimeras at 7 d.p.i. compared with WTWT chimeras (Figure 4-5B), indicating that loss of

LTR signaling in the radio-sensitive compartment can increase the expansion of SILP CD4+ T

cells after viral infection.

Whereas CD4+ T cells from WTWT chimeras did not exhibit augmented IFN production

after RV infection, CD4+ T cells from RV-infected Ltbr-/-WT chimeras exhibited elevated

IFN production compared with uninfected Ltbr-/-WT controls (Figure 4-5C and D). On the

other hand, IL-17 production by CD4+ T cells was significantly reduced in Ltbr-/-WT chimeras

compared with WTWT chimeras, whether at steady state or after RV infection (Figure 4-5C

and E). These results suggest that although primary RV infection does not augment IL-17

production by CD4+ T cells, the frequency of IL-17-producing T cells in the SILP of resting and

infected mice was partially dependent on LTR signaling in radio-sensitive compartments.

4.3.4 Ltbr-/- chimeric mice generate a normal intestinal IgA response to RV

Given that we observed a reduction in SILP Th17 cells in the Ltbr-/-WT chimeras (Figure

4-5E), and Th17 cells have been implicated in promoting antigen-specific IgA responses (Hirota

et al., 2013), we speculated that there might be a delay and/or reduced production of antigen-

specific IgA in response to RV infection in Ltbr-/-WT chimeric mice. Although the initiation of

systemic RV-IgA in the Ltbr-/-WT chimeras was slightly delayed, IgA levels increased

quickly, achieving levels comparable with the WTWT chimeras (Figure 4-6A). Moreover,

intestinal RV-specific IgA production was comparable between the Ltbr-/-WT and WTWT

chimeras at all of the time points examined (Figure 4-6B). These results suggest that antigen-

specific IgA, in response to mucosal viral infection, can be generated in mice lacking LTR

signaling in the radio-sensitive compartment.

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Figure 4-5 Expansion of polyclonal CD4+ T cells in response to RV infection is increased,

and CD4+ T cell cytokine profiles are skewed in Ltbr-/- chimeric mice.

WTWT and Ltbr-/- WT (KOWT) chimeric mice were sacrificed at 7 d.p.i.. SILP

lymphocytes were isolated and restimulated with PMA/ionomycin in vitro for 6 h. Polyclonal

CD4+ T cells were analyzed by flow cytometry.

A. Percentage of SILP CD4+ T cells as a frequency of the total SILP mononuclear cells. B.

Percentage of Ki-67+ as a frequency of SILP CD4+ T cells. C. Representative flow cytometry

data showing IL-17A- and IFN-producing SILP CD4+ T cells (pre-gated on CD4+ T cells). D.

Percentage of IFN+ as a frequency of SILP CD4+ T cells. E. Percentage of IL-17A+ as a

frequency of SILP CD4+ T cells.

UI, uninfected; RV, RV-infected. Each point represents individual mouse pooled from 3


NS, not significant, *p<0.05, **p< 0.01, ***p<0.001, **** p< 0.0001

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Figure 4-6 LTR signaling pathway in radio-sensitive compartments is dispensable for

local antiviral IgA production.

WTWT and Ltbr-/- WT (KOWT) chimeric mice were infected with RV at day 0 and fecal

pellets and serum samples were collected over time.

A. Levels of RV-specific IgA in the serum at 0, 7, 16 d.p.i. were measured by ELISA. B. Levels

of RV-specific IgA in the feces at various time points were measured by ELISA.

Each point represents the individual mouse pooled from 3 independent experiments (samples of

0-7 d.p.i, were collected from 17-20 mice, while samples of 9-20 d.p.i. were collected from 6-7

mice). Data presents as average OD reading at 450 nm of samples ± SEM. Mann-Whitney

nonparametric test. NS, not significant, *p<0.05, **p<0.01

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4.4 Discussion

While it is well known what types of DC and co-stimulatory requirements are required for

priming T-cell response to viral infections in the periphery, including the lung (Ballesteros-Tato

et al., 2010; Belz et al., 2004), less is known about the mechanisms of priming T-cell responses

to gastrointestinal viruses. With respect to RV infection, Lopatin et al. have reported that

CD11c-expressing cells in the SED of PPs colocalized with RV antigen at 24 h post infection

(Lopatin et al., 2013). In addition, it has been demonstrated that DCs from PPs exhibit increased

expression of surface activation markers (CD40, CD80 and CD86), as well as increased mRNA

levels of proinflammatory cytokines such as IL-12/23p40, TNFand IFN shortly after RV

infection (Lopez-Guerrero et al., 2010). However, the molecular mechanisms required for the

effective priming of the naïve T cells by mature SILP-resident DCs have not been well

investigated. Here, we show that LTR signaling in the radio-sensitive compartment dampens

antiviral T cell IFN responses; however, non-specific “homeostatic” IL-17 production by CD4+

T cells in part requires LTR signaling. Nevertheless, this altered cytokine milieu had no

noticeable impact on intestinal virus-specific IgA production nor on viral clearance in Ltbr-/-

chimeric mice.

We have previously shown that DC-intrinsic LTR signaling is critical for CD8+T cell optimal

expansion via type I IFN-dependent mechanism (Summers deLuca et al., 2011). However, our

prior studies measured CD8+ T cell responses to a soluble protein antigen (Ovalbumin, OVA) in

a non-infectious setting with minimal inflammatory stimulus. We found that the collaboration of

DC-intrinsic toll-like receptor (TLR) 4 and LTR signals was required for maximal expression

of type I IFN by DC, and this augmented type I IFN response was needed for optimal expansion

of OVA-specific T cells, but not OVA-driven IFN production (Summers-DeLuca et al., 2007;

Summers deLuca et al., 2011). In our current study, we did not observe a reduction of CD8+ T

cell expansion after mucosal viral challenge in LTR-deficient chimeric mice compared with

WT chimeric mice. We speculate that during a viral infection such as RV, type I IFN can also be

produced by other cell types, such as pDCs (Deal et al., 2013; Mesa et al., 2007), and this may

either over-ride a requirement for LTR signaling in DC, or alternatively TLR4 and RV-

triggered pattern recognition receptors (PRRs) may elicit differential requirements for LTR co-

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signaling. Indeed, responses to influenza virus in the LT-deficient setting are relatively normal

(Lund et al., 2002; Moyron-Quiroz et al., 2004).

While we didn’t observe a defect of RV clearance in adult Ltbr-/- chimeric mice, it is possible

that neonatal mice, which do in fact exhibit diarrhea following RV infection (Little and

Shadduck, 1982), may be more susceptible to the absence of LTR in the radio-sensitive

compartment in terms of their ability to clear an RV infection. However, the Ltbr-/-WT

chimera approach does not allow us to answer this question, and intact Ltbr-/- mice lack

secondary lymphoid tissues (Fütterer et al., 1998), thus introducing a major confounding

variable.

Although the expansion of antigen-specific SILP CD8+ T cells in Ltbr-/-WT chimeric mice was

normal, we observed a significant increase in the percentage of total CD8+ T cells in the SILP of

Ltbr-/-WT chimeric mice after viral challenge. Given that competitive BM chimeras have

revealed that LTR signaling plays a role in maintaining SILP-resident CD103+CD11b+ cDCs

(Satpathy et al., 2013), it is possible that LTR-dependent CD103+CD11b+ cDCs have the

capacity to constrain the CD8+ T cell population within the SILP (or alternatively, a possible

compensatory increase of CD103+CD11b- cDC subset could lead to an increase in gut-homing

CD8+ T cells after viral inoculation). Future studies examining the role of specific SILP DC

subsets and macrophages in the context of mucosal viral infection would shed further light on

mechanisms of T-cell priming in the gut.

The reason(s) for increased IFN production by CD4+ T cells in the LT deficient setting is

unclear but may be related to the complexity of the LT network. Specifically, the LT12-LTR

and the LIGHT-HVEM-BTLA (B- and T-lymphocyte attenuator) systems form an integrated

circuit, controlling intercellular communication between T cells and DCs (Ware, 2005), with

LIGHT serving as a key factor controlling the HVEM BTLA switch between positive and

inhibitory signaling. It has been proposed that the induction of LIGHT during T cell activation

and its occupancy of HVEM displaces BTLA and alleviates inhibitory signaling (Ware, 2005).

Therefore, the loss of LTR expression within the radio-sensitive compartment could promote

preferential binding of LIGHT (expressed by T cells) with HVEM, thus maintaining T cell

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activation. The complex relationship between LT12-LTR and the LIGHT-HVEM-BTLA

systems in the context of SILP resident DC:T cell interactions requires further examination.

Th17 cells have been shown to be responsible for inducing the switch of germinal center B cells

toward the production of high-affinity T cell-dependent IgA (Hirota et al., 2013). Moreover, IL-

17 produced by Th17 cells increases polymeric Ig receptor expression on intestinal epithelial

cells (IECs) and increases the rate of secretory IgA production into the lumen (Cao et al., 2012).

Herein, our results demonstrated that in spite of abrogated SILP Th17 cell homeostasis in Ltbr-/-

WT chimeric mice, decreased production of IL-17 by Th17 cells did not alter local antiviral

IgA production. It is possible that the residual IL-17 production was sufficient to induce the

antiviral IgA response in a T cell-dependent manner. Alternatively, cytokines and growth factors,

such as APRIL and BAFF, which play an important role in T cell-independent IgA class-switch

recombination within isolated lymphoid follicles of the SILP, may provide an independent

mechanism for promoting RV-specific mucosal IgA (Kruglov et al., 2013; Tsuji et al., 2008b).

This T cell-independent IgA induction can be maintained by regulatory T cells (Cong et al.,

2009), LT12-expressing RORt+ ILCs (Kruglov et al., 2013), and APRIL- and BAFF-

expressing plasmacytoid DCs in the mesenteric lymph nodes (Tezuka et al., 2011), which we did

not evaluate in this study. Lastly, soluble LT3, derived from RORt+ ILCs, could promote T

cell-dependent IgA production via TNFRI/TNFRII signaling (Kruglov et al., 2013).

In summary, we show that Ltbr-/-WT chimeric mice are capable of mounting a primary CD8+

T cell response against RV infection. Unlike its critical role in C. rodentium infection, LTR

signaling in the radio-sensitive compartment is not absolutely required for RV clearance in the

small bowel, suggesting that the role of LTR signaling in radio-sensitive compartments varies

with the type of mucosal challenge as a result of factors, such as the site of infection (large bowel

vs. small bowel), the types of pathogen-associated molecular patterns (bacterial vs. viral), and

the severity of disease after challenge (fatal vs. asymptomatic). We previously showed that

LTR signaling in radio-sensitive compartments is required for CD8+ T cell responses to both

self and foreign protein antigens (Ng et al., 2015; Summers deLuca et al., 2011). Presumably the

presence of viral-derived innate signals over-ride a requirement for LTR signaling in radio-

sensitive compartments to prime CD8+ T cells, as has been noted in the case of Influenza virus

infection (Lund et al., 2002).

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The reduction of IL-17-producing CD4+ T cells and the comparable local RV-specific IgA level

in Ltbr-/-WT chimeras suggest that the local humoral anti-RV response does not require

optimal levels of IL-17. Recently, it has been shown that IL-22, a member of the IL-10 family of

cytokines, is essential for protection against RV (Zhang et al., 2014a). Moreover, the main

source of IL-22 production after these challenges is intestinal ILC3 (Hernández et al., 2015;

Tumanov et al., 2011). It is possible that IL-22 may play a more important role than IL-17 in

responses to RV, and it would be of interest to determine if RV-induced IL-22 production by

ILC3 is influenced by the LT pathway. Further studies could focus on the role of LTR signaling

within the radio-resistant compartment (IECs and stromal cells) during intestinal viral responses,

vis-a-vis IL-22 production.

Although the current two licensed, live oral RV vaccines, RotaTeq (Merck, West Point, PA,

USA) and Rotarix (GlaxoSmithKline, Research Triangle Park, NC, USA), prevent up to 74% of

severe RV episodes (Fischer Walker and Black, 2011), the lower vaccine efficacy in resource-

poor countries, as well as the risk of intussusception after vaccination are significant problems

currently without a solution (Parashar et al., 2015). One reason for the low vaccine efficacy in

the low-income countries is a result of reduced immune responses in infants because of

comorbidities or malnutrition, including micronutrient deficiency (Glass et al., 2006). An

understanding of how different dietary conditions affect the SILP cytokine milieu during the

priming phase of RV infection, and how such cytokines impact the formation and potency of

CD8+ effector/memory T cells, may provide a better RV vaccine design.

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Chapter 5

Discussion and Future Directions

114

In this thesis, I have reviewed the biology of DC in the context of rotavirus infection in the small

intestine. I have also provided original data showing that in adult mice, cDCs and LTR

signaling in radio-sensitive compartments are not required for mucosal and systemic RV-specific

IgA responses. In parallel, I found that intestinal anti-RV CD8+ T-cell responses rely on BATF3-

dependent DCs. Interestingly, compared to neonatal mice, adult Batf3-/- mice exhibit a residual

RV-specific CD8+ T-cell response, suggesting compensatory antigen presentation from other DC

subsets or non-professional APCs can substitute for BATF3-dependent DC. In the following

sections, I focus on caveats of data presented in Chapters 3 and 4 as well as four main questions

derived from the data chapters. First, what is the role of DCs and LTR signaling in radio-

sensitive compartments in modulating IgA responses? Next, what cell type present viral antigens

to CD8+ T cells in the absence of BATF3-dependent DCs? Third, what is different between adult

and neonatal intestinal environments and how does this impact immune responses in the

intestine? Last, what is the interplay between enteric viruses and intestinal microbiota (or other

relevant trans-kingdom interactions)?

Caveats arising from Chapter 3

In this Chapter, I used both germline KO mice (Batf3-/- mice and huLangerin-DTA mice) as well

as DTx-inducible KO mice (Zbtb46-DTRWT chimeric mice). Lack of DC and DC subsets

from birth may change the intestinal immunological baseline. Indeed, in the huLangerin-DTA

mice, homeostatic Th17 cells are reduced while the Th1 cells are unaltered compared to WT

littermates (Welty et al., 2013). Although the microbiome composition of colon and cecum is

largely comparable between huLangerin-DTA mice with their littermate controls (Welty et al.,

2013), they may have different levels of SFB in their terminal ileum, which the authors did not

depict in their manuscript. Whether the diminished Th17 cells in huLangerin-DTA mice affect

the steady state IgA response is not clear. It could be that a decreased Th17 response leads to a

decreased expression of pIgR on IEC and less transportation of SIgA into the gut lumen (Cao et

al., 2012). It is not clear whether these changes affect the RV clearance or host anti-RV

responses. In IRF8fl/fl CD11c-cre mice (which is similar to Batf3-/- mice), small intestinal

intraepithelial CD8+ and CD4+CD8+ T cells are almost complete absent (Luda et al., 2016).

Moreover, these mice also lack Th1 cells and fail to support Th1 cells differentiation in MLNs.

Despite these defects in mucosal T cell homeostasis, the composition of the cecal and colonic

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microbiota do not differ between Irf8fl/fl CD11c-cre mice and control Irf8fl/fl mice (Luda et al.,

2016). Again, microbiota composition in the terminal ileum is not depicted in the manuscript.

To study the role of total cDC, I used an inducible depletion model rather than a germline KO

mice. With a transient depletion of cDC, homeostatic Th1 and Th17 cells are not altered (Figure

5-1). By crossing Batf3-/- mice with huLangerin-DTA mice, Welty et al. has reported that these

double KO mice are devoid of both CD103+CD11b- and CD103+CD11b+ cDCs in the intestinal

LP and the MLNs (Welty et al., 2013). Moreover, these double KO mice display reduced Th17

cells and Tregs in the intestinal LP, but unimpaired Tregs in the MLNs. It is currently not known

whether the lack of intestinal CD103+ DCs from birth can affect the development of GALTs and

the intestinal IgA response. For a direct comparison between cDC-deficient mice with DC-subset

deficient mice, I can take advantage of Clec4a4-DTR mice and Clec9a-DTR mice, which allow

for inducible ablation of CD103+CD11b+ cDC and CD103+CD11b- cDC, respectively (Muzaki et

al., 2016).

Figure 5-1 Th1 and Th17 responses in Zbtb46-DTRWT chimeric mice.

A. Percentage of CD4+ T cells as a frequency of SILP mononuclear cells in Zbtb46-DTRWT

chimeric mice at 7 d.p.i.. B. Parentage of IFN+ cells as a frequency of SILP CD4+ T cells in

Zbtb46-DTRWT chimeric mice at 7 d.p.i.. C. Parentage of IL-17a+ cells as a frequency of

SILP CD4+ T cells in Zbtb46-DTRWT chimeric mice at 7 d.p.i..


individual biological replicate (one mouse) pooled from 3 independent experiments. Data are

presented as mean± SEM. Mann-Whitney test. *p<0.05, NS, not significant.

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Caveats arising from Chapter 4

In Chapter 4, I observed that LTR signaling in radio-sensitive compartments is dispensable for

generating a viral specific IgA response systemically or locally at the intestinal mucosa. There

are several caveats by using Ltbr-/- chimeras. First of all, I did not know whether the Ltbr-/- BM

reconstituted WT mice have the same myeloid cell population in the intestine compared to the

WT BM reconstituted WT mice. Since DC-intrinsic LTR signaling is required for the

proliferation of splenic CD8-CD11b+ DC (Kabashima et al., 2005), it is possible that the

corresponding CD103+CD11b+ DCs in the intestine share the similar requirement of LTR

signaling. To test this possibility, I will compare the frequency of SILP DC as well as DC

subsets in Ltbr-/- WT with that in WTWT chimeric mice. A decreased proportion of

CD103+CD11b+ DCs in the intestine may explain the decreased homeostatic Th17 cells found in

Ltbr-/- chimeras. Secondly, the Ltbr-/- WT BM chimeras does not allow me to distinguish

whether my observations are due to a lack of LTR signaling in macrophages or DCs or both. To

test the possibility that lack of DC-intrinsic LTR can recapitulate what I observed in Ltbr-/-

WT BM chimeras, I can either cross Zbtb46-cre with LTRfl/fl mice or reconstitute lethally

irradiated WT mice with 50% Zbtb46-DTR BM and 50% Ltbr-/- BM. Both methods can ensure a

deletion of LTR in cDC compartments, and will provide insights on the contribution of

macrophage-intrinsic LTR versus cDC-intrinsic LTR on controlling anti-RV responses.

Revisiting the role of DC in regulating intestinal IgA responses

It has been demonstrated that the majority of RV-specific IgA is generated in a T cell-dependent

manner (Franco and Greenberg, 1997). Theoretically, DCs loaded with RV antigens migrate to

T-cell zones in the PPs and the MLNs where they activate T cells. Upon induction of the

transcription factor BCL-6, activated CD4+ T cells (Tfh) express CXCR5 and migrate toward

CXCL13 into the FDC-rich environments, where Tfh provide help for activated B cells. Through

CD40:CD40L ligation and cognate TCR-MHCII recognition, activated B cells undergo SHM

and CSR in the GCs of the PPs and the MLNs. In the presence of activated TGF, IL-21 and RA,

class switch to the IgA isotype (rather than IgM or IgG) is preferred. Since DCs initiate this T-

cell dependent IgA response, it is logical to predict that the IgA response against RV would be

impaired in the absence of DCs. However, in Chapter 3, I did not see an impaired RV-specific

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IgA response in DTx-treated Zbtb46-DTR chimeric mice, suggesting that cDCs are dispensable

for IgA induction. To understand why I did not observe the predicted IgA impairment, some

main questions remained to be answered: Are Tfh and GC responses impaired as a consequence

of cDC depletion? Which cells initiate antigen-specific IgA responses in the absence of cDCs? Is

T cell-independent IgA response compensating for the dominant T cell-dependent IgA response?

Are Tfh and GC responses impaired in the absence of cDCs?

Several studies have demonstrated that priming by DCs induces BCL-6 expression in CD4+ T

cells, thus promoting CD4+ T cells to differentiate to a Tfh fate (Goenka et al., 2011; Nurieva et

al., 2009). Moreover, CD8-CD11b+ DCs localized within the interfollicular zone play a pivotal

role in the induction of antigen-specific Tfh cells by upregulating the expression of inducible

costimulator-ligand (ICOS-L) and OX40 ligand through the non-canonical NF-B signaling

pathway (Shin et al., 2015). Based on these observations, it is likely that the induction of Tfh is

disrupted in the absence of cDCs. To test this hypothesis, we can infect BCL-6-YFP reporter

mice (Kitano et al., 2011) with RV and monitor YFP expression by CD4+ T cells and B cells in

the PPs and MLNs at various time points. After the kinetics are defined, we can challenge DTx-

treated Zbtb46-DTR chimeric mice (or huLangerin-DTA mice which lack CD103+CD11b+ DCs)

with RV, and phenotype Tfh and GC B cells by flow cytometry and immunofluorescence

microscopy. In addition, the frequency of RV-specific IgA-producing plasma cells could be

assessed by an Enzyme-Linked ImmunoSpot (ELISPOT) assay to understand whether

differentiation to plasma cells is impaired in the absence of cDCs (or CD103+CD11b+ DCs). If

there is no defect in the generation of Tfh and RV-specific IgA+ plasma cells, other cells may

compensate for cDCs in cDC-deficient mice. Moreover, it has been reported that Th17 (Hirota et

al., 2013) and Foxp3+ Tregs (Tsuji et al., 2009) convert into Tfh cells in the PPs with help from B

cells or DCs. These experiments will shed light on our understanding on cDC-independent Tfh

induction and GC reactions in mucosal tissues.

It could be that we do not observe a Tfh response in mice deficient of cDCs. In this scenario, T

cell-independent IgA class switch is generated after RV infection. To test this possibility, I will

generate mice with MHCII-deficiency on B cells via reconstituting lethally irradiated B6 WT

mice with Jh-/- BM and MHCII-/- BM, and mice with CD40-deficiency on B cells via

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reconstituting lethally irradiated B6 WT mice with Jh-/- BM and CD40-/- BM (Sangster et al.,

2003). By challenging these chimeric mice with RV and evaluate the anti-RV IgA responses in

both fecal and serum samples, I will better understand the nature of the anti-RV IgA responses.

B1 B cells located in the peritoneal and pleural cavities and splenic MZ B cells may also

contribute to the T cell-independent IgA class switch (Fagarasan and Honjo, 2000). It has been

reported that adoptively transferred peritoneal B1 B cells cannot do not ablate RV shedding in

SCID mice, suggesting that B1 B cells do not generate T cell-independent IgA (Kushnir et al.,

2001). To evaluate the contribution of MZ B cells in generating anti-RV IgA, I can cross Ztb46-

DTR mice with Pyk2-/- mice (which lack MZ B cells (Guinamard et al., 2000)) and make BM

chimeric mice (Zbtb46-DTR·Pyk2-/- WT). DTx-treated BM chimeras will lack MZ B cells on

top of cDCs. Alternatively, I can isolate MZ B cells from B6 WT mice and adoptively transfer

these cells into SCID or Rag2-/- mice followed by RV infection. By comparing to SCID or Rag2-

/- mice that do not receive any cells, I will be able to test if MZ B cells are capable to class switch

to T cell-independent IgA+ B cells and produce RV-specific IgA.

Do moDCs promote IgA response in the absence of cDCs?

Although cDCs are depleted in the Zbtb46-DTR chimeric mice treated with DTx, SILP CD103-

CD11b- and CD103-CD11b+ DC subsets are still present (Figure 3-1). Given that they are

resistant to DTx depletion, they could be moDCs, which have been previously implicated in IgA

class-switching (Tezuka et al., 2007). Whether moDCs promote anti-RV IgA responses has not

been addressed yet. Therefore, to directly test this possibility in vivo, we can cross Zbtb46-DTR

mice with Ccr2-/- mice, followed by setting up BM chimeras (donor: Zbtb46-DTR·Ccr2-/- mice)

and challenging these mice with RV. On the other hand, PPs are the primary site for B cell

priming. Given that the CD8+CD11b-, the CD8-CD11b+ as well as the CD8-CD11blo/- DN

DCs are pre-cDC derived (or ZBTB46-dependent) (Bonnardel et al., 2017), all these DC subsets

should be depleted in the DTx-treated Zbtb46-DTR chimeric mice (although I have not looked at

the PP DCs). Interestingly, another DC subset located in the PPs, namely LysoDC

(CD11c+MHCII+SIRP+BST2+CD4-), is monocyte-derived (or ZBTB46-independent)

(Bonnardel et al., 2017). Whether this monocyte-derived LysoDC can promote IgA response is

not known and the Zbtb46-DTR·Ccr2-/- chimeric mice may help us to address this question.

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Do increased pDCs promote IgA response in DTx-treated Zbtb46-DTR chimeric mice?

Tezuka et al reported that, pDCs from the MLNs can promote T cell-independent IgA response

via production of APRIL and BAFF in vitro (Tezuka et al., 2011), suggesting pDCs may play a

prominent role in promoting IgA responses. In DTx-treated Zbtb46-DTR chimeric mice, the loss

cDCs triggers a rapid increase of the FLT3L levels in the serum (Meredith et al., 2012a). FLT3L

is a cytokine promoting the expansion of both cDC and pDC (Maraskovsky et al., 1996; Onai et

al., 2006). Although we did not measure the level of serum FLT3L in our experiments, we do

have data regarding the pDC population in the SILP - I found that the frequency of pDC

increased after DTx treatment in the Zbtb46-DTR chimeric mice (Figure 5-2).

pD

C %

of

liv

e c

ell

s

0 .0

0 .5

1 .0

1 .5

2 .0

+ +++ --R V

D T x

W T z D C C D 1 1 cB M d o n o r

+ ++- +-

**

0 .0 6 8

**

Figure 5-2 Frequency of pDC in the SILP at 7 d.p.i.

Percentage of SILP pDCs as a frequency of mononuclear cells from chimeric mice treated with

PBS or DTx at 7 d.p.i.. Gating strategies for pDC in the SILP can be found in Figure 3-2A. Each

data point represents an individual biological replicate (one mouse) pooled from 3 independent

experiments. Data are presented as mean± SEM. Mann-Whitney test. **p<0.01. zDC, Zbtb46-

DTR. CD11c, CD11c-DTR. RV, rotavirus. DTx, diphtheria toxin. BM, bone marrow.

Deal and colleagues have reported that pDCs can promote optimal B-cell response and viral

specific antibody secretion after RV infection due to pDC-derived type I IFN (Deal et al., 2013).

Therefore, the increased pDC frequency may potentially promote the IgA response in the DTx-

treated Zbtb46-DTR chimeric mice, with a caveat that we didn’t measure the level of type I IFN

in our experiment.

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Interestingly, pDC frequency in the SILP is found to be increased in DTx-treated CD11c-DTR

chimeric mice, compared with WT chimeric mice (Figure 5-2). This piece of data confirmed that

pDCs are not DTx-sensitive in CD11c-DTR mice (Sapoznikov et al., 2007). However, the DTx-

treated CD11c-DTR chimeric mice displayed an impaired (but not ablated) RV-specific IgA

response in both serum and fecal levels (Figure 5-3), which seems to contradict my hypothesis

that pDC can promote IgA response in the absence of cDCs. One possibility could be that since

plasmablasts are also DTx-sensitive in CD11c-DTR mice (Hebel et al., 2006), this may have

resulted in decreased IgA levels. To directly test whether pDCs promote IgA responses in the

absence of cDC, GmAb (Asselin-Paturel et al., 2003) can be administered to DTx-treated

F e c a l R V -Ig A

d .p .i .

OD

45

0n

m

0

1

2

3

P B S

D T x

0 3 5 7 8 9 10 12 14 17

**

*

*

*

0 .0 5 7 1

N S

S e ru m R V -Ig A

d .p .i .

OD

45

0n

m

0 .0

0 .5

1 .0

1 .5

0 7 14 17

*N S

P B S

D T x

Figure 5-3 Fecal and serum RV-specific IgA responses in CD11c-DTR chimeric mice.

CD11c-DTR chimeric mice were infected with RV at day 0 and fecal pellets and serum samples

were collected over time and measured by ELISA.

Each point represents the individual mouse pooled from 3 independent experiments. Data

presents as average OD reading at 450 nm of samples ± SEM. Mann-Whitney test. *p<0.05,

**p<0.01, NS, not significant. DTx, diphtheria toxin

Zbtb46-DTR chimeric mice to deplete pDCs (on top of cDC depletion) and then RV-specific IgA

responses can be examined. The putative function of different DC subtypes in promoting an RV-

specific IgA response is summarized in Table 5-1. The experiments proposed in this section will

help us to uncover the interplay between different DC types and antigen-specific IgA responses

at the intestinal mucosa.

121

Table 5-1 Alterations of different DC types, plasma cells and IgA in different chimeras

Upon RV

infection

cDC pDC moDC Plasma cell RV-specific

IgA

WT

chimeras

- - ? ↑ Normal

DTx-treated

Zbtb46-DTR

chimeras

↓ ↑ ? ? Same as WT

chimeras

DTx-treated

CD11c-DTR

chimeras

↓ ↑ ?

The steady state

equivalent cells are

depleted.

(Rivollier et al.,

2012)

?

The ones express high

level of CD11c may be

depleted at steady

state.

(Hebel et al., 2006)

Decreased and

delayed

compared to

WT chimeras

Is LTR signaling in DCs required for intestinal IgA responses?

After we published our study in the Journal of Leukocyte biology (Sun et al., 2015), another

study conducted by Jason Cyster’s group was published in Science demonstrating that DC-

intrinsic LTR signaling is required for IgA class switch in the PP (Figure 5-4)(Reboldi et al.,

2016). At first glance, there seems to be discrepancies between these two data sets. However, our

studies may not absolutely contrast with each other. First, Reboldi et al. examined the role of

DC-intrinsic LTR in the homeostatic IgA response, on a per cell basis (by flow cytometry).

They found that B cell class switch is skewed from IgA to IgG1 in the PP of WT mice

reconstituted with Ltbr-/- BM, suggesting a requirement of LTR signaling from the radio-

sensitive compartment (mainly DCs) for IgA class switching. In our study, instead of checking

the homeostatic polyclonal IgA/IgG1 expression on the surface of B cells/plasma cells in the PPs

by flow cytometry, we measured RV-specific bulk IgA levels in the fecal pellet by ELISA.

Moreover, we did not measure the level of RV-specific IgG1 given that the IgA response is the

dominant humoral response within the intestinal mucosa after RV challenge (Sheridan et al.,

1983). Thus, it is difficult to compare our data directly with that of Reboldi et al. because of the

differences in readouts. Second, Reboldi et al. checked the IgA class switch at steady state

whereas our study focused on a viral infection setting. It could be true that the Ltbr-/- WT

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chimeric mice display a skewed IgA to IgG1 homeostatic humoral response within PP (although

we did not check this), it nevertheless may not hold true after an intestinal viral infection.

Furthermore, it is known that PPs are not the only place for B cell class switching in the

intestine: MLNs (Mora et al., 2006), ILFs (Tsuji et al., 2008a) and intestinal LP (Fagarasan et al.,

2001) are all able to support B cell class switch to IgA-producing cells. In addition, the priming

of B cells in the setting of RV infection may not be in PPs, and our lab is currently investigating

this possibility. In summary, although Reboldi et al observed a skewed IgA to IgG1 humoral

response in the gut in mice that lack LTR expression on DC, our results are not completely

contradictory to their findings.

Kruglov et al. have proposed that soluble LT3 produced by ILC3 controls T cell-dependent IgA

induction in the small intestinal LP via stromal cells, while membrane-bound LT12 produced

by ILC3 modulates T cell-independent IgA induction in the LP via control of DC function

(Figure 5-4)(Kruglov et al., 2013). As mentioned in the introduction, soluble LT3 does not

signal through LTR, but signals via TNFRI/II (Figure 1-4). Thus, it is tempting to think that

LTR-dependent IgA class switch described by Reboldi et al is primarily influencing T cell-

independent IgA responses. However, Kruglov et al. used RORt-Lta-/- mice in their study, and

these mice lack PPs and other peripheral LNs thus preventing any assessment of the role of ILC3

in PP IgA responses. Since CD40-derived signals are required for the events leading up to IgA

class switch in the Reboldi study, the IgA response in this context is presumably T cell-

dependent. Whether LTR-dependent IgA class switch is T cell-dependent or T cell-independent

needs further investigation.

As a future direction, it will be interesting to dissect the role of LT3 and LT12 on ILC3 in

promoting RV-specific IgA responses in the small intestine. As part of the innate control of RV

infection, ILC3 have been found to produce IL-22, which synergistically acts with IEC-produced

IFN to induce expression of ISGs (Hernández et al., 2015), suggesting ILC3 are activated

during RV infection. I demonstrated that LTR signaling in radio-sensitive compartments is

dispensable for an RV-specific IgA response in Chapter 4, suggesting that a hematopoietic

source of LT12 during adulthood may not play a role. Our next step is to define if ILC3-

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derived LT3 or LT12 signals on stromal cells to induce a T cell-dependent antigen-specific

IgA response upon RV infection.

Figure 5-4 Contrasting views on the role of LT12 in IgA class switch.

LTR signaling in PP DCs is required for T cell-dependent IgA class switch, however the role of

its primary ligand, LT12, is somewhat controversial. Whereas Kruglov et al demonstrated that

LT12 signaling via DC-intrinsic LTR regulates T cell-independent IgA class switch, Reboldi

et al show that LT12 signaling via DC-intrinsic LTR primarily mediates T cell-dependent

IgA class switch.

What cells prime antigen specific CD8+ T cell response in Batf3-/- adult mice?

In Chapter 3, I observed that a residual anti-RV CD8+ T cell response persists in the SILP of

Batf3-/- mice, which are deficient of CD103+CD11b- DCs. This suggests that while BATF3-

dependent cDC are the primary cross-presenting APC subset in adult mice, there may also be

redundancy in the induction of CD8+ T-cell responses in a setting of infection. This kind of

redundancy is not rare in the intestinal tract. For example, intestinal Treg are found in normal

numbers in mice lacking only CD103+CD11b- DC (Welty et al., 2013) or CD103+CD11b+ DC

(Persson et al., 2013b), yet mice lacking both DC subsets have decreased Treg numbers (Welty

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et al., 2013). On the other hand, there is also evidence of antagonism between intestinal

CD103+CD11b- DC and CD103+CD11b+ DC. IL-12 production by CD103+CD11b- DC appears

to reduce the Th2 response to Heligmosomoides polygyrus infection (Everts et al., 2016),

whereas IL-23 produced by CD103+CD11b+ DC and macrophages reduces IL-12 production by

CD103+CD11b- DC during C. rodentium infection (Aychek et al., 2015). These results suggest

that cDC subsets can mutually dampen each other’s responses, probably to help prevent

excessive immune activation and to maintain a balanced immune response (Joeris et al., 2017).

Macrophages and DCs have distinct, yet complementary roles in maintaining gut homeostasis

and immune defense. Although macrophages are the most abundant mononuclear phagocytes in

the steady-state gut LP (Gross et al., 2015), they have been shown to be poor stimulators for T

cells in vitro (Steinman and Cohn, 1973). However, recent evidence suggests that macrophages

can prime naïve CD8+ T cells in vivo (Bernhard et al., 2015; Pozzi et al., 2005). In our study, we

observed that in RV-infected DTx-treated Zbtb46-DTRWT chimeric mice, which lack cDCs

but conserve macrophages (Figure 3-1), the antigen-specific CD8+ T-cell response was abolished

(Figure 3-3). This result suggests that macrophages are not sufficient to induce antiviral CD8+ T-

cell responses in the gut. Whether macrophages help DCs to activate naïve T cells and whether

the memory T-cell response is dependent on macrophages needs to be further determined. Given

that macrophages exceed DCs in quantity and are normally the first immune cells in the body

that come into contact with invading pathogens, they are able to digest pathogens and present a

variety of T-cell priming epitopes to T cells (a broad immune response). On the contrary, DCs

may be more efficient at presenting immunodominant epitopes to cognate T cells and thus elicit

a “narrow and specific” immune response (Bernhard et al., 2015). One caveat in our study is we

only tested one immunodominant epitope of RV, therefore it will be interesting to screen other

epitopes whose affinity is weaker than VP6357-366 to determine whether intestinal macrophages

contribute to induce antiviral CD8+ T-cell immune responses to other epitopes.

What do we know about the role of mucosal DC in the neonatal period?

Historically, the neonatal immune system has been considered to be poorly competent in

generating immune responses and instead is polarized towards the induction of immune tolerance

(Streilein, 1979). A conceptual switch occurred in the late 1990s through some pioneering

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studies demonstrating that adult-like B- and T-cell responses could be achieved in early life

using appropriate dosage and immunization strategies (Adkins et al., 2004). Most studies

regarding neonatal DC biology study the spleen or peripheral LN. Here I show in Chapter 3 that

the requirement of BATF3-dependent DC in inducing the antiviral T-cell response at the

intestinal mucosa is more stringent in neonatal mice than then in adult mice. Although we are

currently not certain why there is an age-associated DC-stringency, I would like to briefly

discuss the differences between adult and neonatal intestinal environments vis-a-vis the immune

system, which may help to solve this puzzle.

(1) Microbiome: It is generally accepted that the healthy fetus is devoid of colonizing viable

microorganisms. During the passage through the birth canal, the fetus first encounters bacteria

derived from the maternal vaginal microbiota (Torow and Hornef, 2017). High inter-individual

variation, but low diversity and density, characterize the neonatal microbial colonization phase,

which is dominated by bacteria specialized in milk fermentation. The microbiota at this stage is

highly sensitive to exogenous perturbations that delay the development of a mature diverse

microbial community (Bokulich et al., 2016). Microbiota alterations, in turn, render the bacterial

ecosystem less resilient to further perturbation (Nobel et al., 2015). With weaning, an

increasingly diverse microbiota is established that is highly individual and remains relatively

stable throughout life. To rule out the impact of microbiome in affecting DC functions, germ-free

(GF) or Batf3-/- mice (adult and neonate) treated with antibiotics can be challenged with RV.

(2) Intestinal barrier: The small intestinal epithelium of newborns exhibits enhanced permeability

to soluble antigens and it is devoid of crypts that harbor intestinal stem cells and AMP-producing

Paneth cells, which emerge at weaning (Bry et al., 1994). The lack of Paneth cell-derived AMPs

may be compensated by the cathelin-related AMPs that are produced by murine enterocytes in

the neonatal small intestine (Ménard et al., 2008). Furthermore, expression of mucin (a building

block of the intestinal mucus layer) is low in the neonate and rises at weaning (Zhang et al.,

2014b). All of these factors might facilitate epithelial invasion, prolong the lifetime of infected

epithelial cells and thereby ultimately allow intraepithelial proliferation and microcolony

formation.

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(3) Immune components: Epithelial innate immune recognition varies in an age-dependent

manner. In mice, epithelial expression levels of the PRRs (e.g. TLR3) are expressed at lowest

levels after birth and increase when mice reach adulthood (Pott et al., 2012). For the adaptive

immune system, TCR+ T cells and B cells are localized exclusively to the PPs and exhibit a

naïve phenotype until weaning (Torow et al., 2015). This is in stark contrast to adult intestinal

tissues, with antigen-experienced IELs and lymphocytes located in PPs and LP, even at steady

state. This age-dependent difference in T-cell populations is relevant to my study, and I did not

test whether the fully reduced antigen-specific CD8+ T cells in the Batf3-/- mice is due to a defect

in neonatal T cells rather than neonatal DCs. To test the possibility, intestinal cDCs from the

neonatal and adult Batf3-/- mice can be sorted out and adoptively transferred to DTx-treated

Zbtb46-DTR chimeric mice. In this way, all the T cells are adult-derived, whereas the DCs are

either from neonatal or adult Batf3-/- mice. Alternatively, CD45.1+ T cells from Batf3-/- adult

mice together with CD45.2+ T cells from Batf3-/- neonatal mice can be adoptively transferred into

Batf3-/-Rag2-/- adult mice. In this way, all DCs are adult-derived, whereas the T cells are either

from neonates or adults. By conducting these experiments, we will achieve a better

understanding of why antigen-specific CD8+ T cell response in neonatal Batf3-/- mice are fully

ablated (as compared to adults).

All these differences between adult and neonate may represent confounders in interpreting the

differences we observed in our experiments. More work is needed to fully understand age-

associated DC phenotypes and this may help us better understand how the neonatal immune

system works in order to ultimately design better pediatric vaccines.

How does the intestinal microbiota interact with enteric viruses and how does this

interaction affect the host immunity?

The enteric virome defined as the collection of retroviruses, noroviruses, rotaviruses,

astroviruses, picornaviruses, adenoviruses, herpesviruses, etc. harbored by the host, regulates,

and are in turn regulated by other microbes (including bacteria, helminths, fungi and protozoa)

through a series of processes termed “trans-kingdom interactions” (Pfeiffer and Virgin, 2016).

Although RV has long been a model virus to study intestinal immune function, few studies

explore its interplay with the resident commensal microbiota. Another enteric virus, murine

127

norovirus (MNV), has been frequently used to study trans-kingdom interactions, thus enhancing

our understanding of intestinal host-microbiota interactions as well as the pathogenesis of IBD.

A link between enteric viral infection and IBD pathogenesis

The first interesting finding with regards to viral infection in intestinal inflammation is a link

between MNV and Paneth cell defects in mice deficient in the CD gene, Atg16l1 (Cadwell et al.,

2010). Atg16l1 is an essential gene mediating autophagy and the CD variant in Atg16l1 is linked

to altered xenophagy and enhanced inflammation (Murthy et al., 2014; Sorbara et al., 2013). In

Cadwell’s study, persistent infection of MNV in the absence of ATG16L1 is able to trigger

increased susceptibility to intestinal inflammation, such as CD. This example demonstrates how

gene-environment interactions are necessary to propagate disease, underscoring the limitation of

mouse studies where mice carrying mutations in human disease susceptibility genes do not

always spontaneously reproduce human pathology. To further this concept, we are testing the

virus-plus-susceptibility gene theory in a different way. Our ongoing studies stem from the

hypothesis that a viral infection in the neonatal period in mice with an IBD susceptible gene

(Nod2) will develop worse colitis in adulthood compared to uninfected mice. RV is a candidate

infectious agent since neonatal mice are highly susceptible to this virus.

Transkingdom interactions between intestinal microbiota and enteric virus

It is well known that intestinal commensals foster host health and limit pathogen colonization.

Recently, it has been reported that the intestinal microbiota can facilitate enteric viral infection

and promote systemic pathogenesis. Antibiotic-treated mice are less susceptible to poliovirus and

reovirus disease (Kuss et al., 2011). Furthermore, poliovirus binds LPS, and exposure of

poliovirus to bacteria enhanced its infectivity. Interestingly, RV infection is also diminished in

both GF and antibiotic-treated mice (Uchiyama et al., 2014). These antibiotic-treated mice

generate higher levels of intestinal and systemic RV-specific IgA and maintain the level of RV-

specific plasma cells in the intestine for a longer time period. The mechanisms underlining the

experimental observations are currently not clear. Along the same lines, Baldridge et al. found

that antibiotics prevented persistent MNV infection, an effect that was reversed by replenishment

of the bacterial microbiota (Baldridge et al., 2015). The receptor for the antiviral cytokine IFN,

as well as the transcription factors STAT1 and IRF3, are required for antibiotics to prevent viral

128

persistence. Lastly, vertical transmission of mouse mammary tumor virus (MMTV, a retrovirus)

also requires the presence of commensal bacteria (Kane et al., 2011). MMTV can bind to

commensal-derived LPS and trigger IL-10 production and thus induce an immune evasion

pathway. In summary, intestinal commensal bacteria facilitate enteric viral infections.

Our understanding towards this complex network is still in its infancy. Basic questions like

whether intestinal DCs modulate such trans-kingdom networks is not known. RV infection,

together with other infections in GF and antibiotic-treated conventional mice, in combination

with mice that lack specific DC subsets, may shed light on this complex regulation.

Overall, results from this thesis demonstrate a role for CD103+CD11b- DC in generating optimal

anti-RV CD8+ T-cell response in the small intestine in both adult and neonatal mice. However, in

the absence of CD103+CD11b- DCs, a compensatory mechanism for presenting antigen to CD8+

T cells exists in adult but not neonatal mice, suggesting that age plays a role in modulating

adaptive immune responses. Local and systemic anti-RV IgA responses are intact in mice

lacking all cDCs and LTR signaling pathway in radio-sensitive compartments, implying either

that T cell-independent IgA responses may take over and compensate the loss of T cell-

dependent IgA response, or non-DC cells can step in to promote an RV-specific IgA response.

Taken together, DCs orchestrate different arms of adaptive antiviral immunity at the intestinal

mucosa in both the neonatal period and adulthood.

129

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