VECTOR SPECIFIC TOLERANCE INDUCTION FOR AIRWAY GENE THERAPY

by

Rahul Kushwah

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

Department of Laboratory Medicine and Pathobiology University of Toronto

© Copyright by Rahul Kushwah 2011

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Vector specific tolerance induction for airway gene therapy

Rahul Kushwah

Doctor of Philosophy

Department of Laboratory Medicine and Pathobiology University of Toronto

2011

Abstract

The success of adenoviral mediated airway gene therapy is hindered by host immune responses

against adenoviral vectors. Helper-dependent adenoviral vectors (HD-Ad) are devoid of viral

coding sequences and have an improved safety profile compared to earlier generation adenoviral

vectors. However, intranasal delivery of HD-Ad vectors potentiates a pulmonary adaptive

immune response, described in chapter 2, which is a barrier to gene therapy. One of the ways to

reduce the immunogenicity of HD-Ad vectors is to increase the efficiency of HD-Ad mediated

gene transfer to the airways, which would lessen the immunogen availability, limiting immune

response against HD-Ad vectors. In chapter 3, a viral formulation strategy using Nacystelyn and

DEAE-Dextran to substantially increase the efficacy of adenoviral mediated gene transfer to the

airways is described. To further reduce the immune response to HD-Ad vectors, I have

developed two novel strategies to induce vector-specific tolerance. The first strategy, described

in chapter 4, involves the use of dendritic cells (DCs) differentiated in presence of IL-10, which

are refractory to HD-Ad induced maturation and instead prime generation of regulatory T cells

which suppress HD-Ad induced T cell proliferation. Delivery of these DCs pulsed with HD-Ad

vectors to mice results in induction of immunological tolerance along with sustained gene

expression following multiple rounds of HD-Ad readministrations. The second strategy,

described in chapter 5, involves delivery of apoptotic DCs followed by delivery of antigen

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towards which tolerance needs to be generated. Apoptotic DCs are readily taken up by viable

DCs, which suppresses DC maturation and induces TGF-β1 secretion, driving generation of

regulatory T cells towards the delivered antigen. This strategy has shown remarkable success in

achieving tolerance towards ovalbumin. Therefore, these strategies can be used to induce

immunological tolerance towards gene therapy vectors which will likely allow for sustained and

long term therapeutic transgene expression.

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In loving memories of

Late Thakur Deep Singh (Baba) and Late Shiv Chandra Singh (Nana)

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Acknowledgments I am grateful to my supervisor Dr. Jim Hu, for all his support and guidance throughout the course

of my graduate studies. I am also grateful to Drs. Li Zhang and Shaf Keshavjee, members of my

thesis advisory committee who have provided me with valuable insights, assistance and resource

throughout my PhD program. Special thanks to our collaborators Drs. Jinyi Zhang and Katherine

A Siminovitch for providing us with transgenic mice for studies using apoptotic dendritic cells.

Also, thanks to Jing Wu and Jordan Oliver for assistance with few experiments and thanks to

Cathleen Duan and Dr. Huibi Cao for production of viral vectors used in this study. A huge thank

you to the Canadian Cystic Fibrosis Foundation for doctoral studentship award.

I am forever grateful to my mom (Nandini Kushwah) and dad (Rajendra Kushwah) for their

unconditional love and support. I am also forever grateful to my grandparents, Late Thakur Deep

Singh, Late Shiv Chandra Singh, Leela Devi, Kamla Devi (Mithai amma) and Urmila Devi

(Kadvi mummy), who always motivated me and were always there whenever I needed them the

most. I am forever grateful to my high school mathematics teacher, Rabindra Singh (Raj), who

was the best mentor I could have ever had and was instrumental in making me achieve my career

goals. I am also grateful to my uncles and aunts: Suman Singh, Padma Singh, Pushpa Singh,

Yogendra Singh, Chandra Kumar Singh (Chanda se pyara Chandamama), Shyama Singh, Ajit

Kumar Singh (Tuntun mama), Ravindra Bala Singh (Rani mami), Anil Pratap Singh (Chhote

mama), Sonia Singh and Arun Pratap Singh (Jhunjhun mama) for their love and support. I am

also grateful to my elder cousin, Shashank Kumar Singh, for all his love and support and to

whom I always looked up to for being a good human being, and also to my sister-in-law Geeta

Singh for her love and support. Last but not least, I am also grateful for all the love and affection

from my choti mom Mahru Sultana (Mona) and the love and support of all my friends and family

members.

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Table of Contents Abstract ii

Acknowledgements v

Table of Contents vi

List of Tables x

List of Figures xi

List of Abbreviations xiv

Dissemination of Thesis Contents xviii

Chapter 1: Introduction

1.1 Gene therapy for airway diseases 1.1.1 Cystic Fibrosis 1 1.1.2 Lung Transplantation 2 1.1.3 Asthma 4 1.1.4 Chronic Obstructive Pulmonary Disease 5

1.2 Vectors for gene therapy 1.2.1 Non viral vectors for gene delivery 6

1.2.1.1 Cationic liposomes 6 1.2.1.2 Glycoconjugates 8 1.2.1.3 Transposon based integrating plasmids 9

1.2.2. Viral vectors for gene delivery 10 1.2.2.1 Retroviral vectors 10 1.2.2.2 Sendai virus based vectors 14 1.2.2.3 Adeno-associated viral vectors 17 1.2.2.4 Adenoviral vectors 20 1.2.2.4.1 Early generation adenoviral vectors 20 1.2.2.4.2 Third generation – Helper dependent adenoviral vectors 21

1.2.3 Challenges to lung gene therapy 23

1.3 Potential for generating vector specific tolerance 1.3.1 Differentiation/Origin of DCs 28

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1.3.2 Dendritic cell subtypes 34 1.3.2.1 Conventional steady state DCs 36

1.3.2.1.1 Migratory DCs 36 1.3.2.1.1.1 Skin 36 1.3.2.1.1.2 Lung 37 1.3.2.1.1.3 Intestinal Tract 37 1.3.2.1.1.4 Liver 39 1.3.2.1.1.5 Kidney 39

1.3.2.1.2 Lymphoid DCs 40 1.3.2.2 Non-conventional DCs 42

1.3.2.2.1 Plasmacytoid DCs 42 1.3.2.2.1.1 pDCs in the liver and the lung 43

1.3.2.2.2Monocyte derived DCs 45 1.3.2.2.2.1 Intestinal monocyte derived DCs 45 1.3.2.2.2.2 Pulmonary monocyte derived DCs 45 1.3.2.2.2.3 Monocyte derived DCs in the skin 46 1.3.2.2.2.4 Monocyte derived DCs in the kidney 47

1.3.3 Regulation of DC apoptosis 48 1.3.3.1TNF superfamily and DC apoptosis 50 1.3.3.2 Nur77 family and DC apoptosis 52 1.3.3.3 HLA-DR and DC apoptosis 52 1.3.3.4 Other factors and DC apoptosis 52

1.3.4 Extrinsic triggers of DC apoptosis 53 1.3.4.2 DC apoptosis is triggered by infections 53 1.3.4.3 DC apoptosis during pathological conditions 54 1.3.4.4 Glucocorticoid induced DC apoptosis 55 1.3.4.5 Tumor induced DC apoptosis 56 1.3.4.6 UV induced DC apoptosis 57 1.3.4.7 T cell induced DC apoptosis 58

1.3.5 Apoptosis and cross priming by DCs 59 1.3.6 Defects in DC apoptosis trigger autoimmune diseases 60 1.3.7 Tolerance induction by DCs 61 1.3.8 Type 1 regulatory T cells 62

1.3.8.1 IL-27 production by DCs drives Tr1 differentiation 64 1.3.8.2 IL-10 production by DCs drives Tr1 differentiation 64 1.3.8.3 ICOSL signalling by DCs drives Tr1 differentiation 66 1.3.8.4 TGF-β1 production by DCs drives Tr1 differentiation 66 1.3.8.5 Other DC driven signals which drive Tr1 differentiation 66

1.3.9 Foxp3+ Regulatory T cells 67 1.3.9.1 Naturally occurring Foxp3+ Tregs 67

1.3.9.1.1 TSLP drives thymic DC mediated nTreg differentiation 67 1.3.9.2 Adaptive Foxp3+ regulatory T cells 68

1.3.9.2.1 IDO in DCs drives aTreg differentiation 70 1.3.9.2.2 TGF-β production by DCs drives aTreg differentiation 71

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1.3.9.2.3 RANKL signalling on DCs drives aTreg differentiation 72 1.3.9.2.4 Retinoic acid producing DCs drive aTreg differentiation 73 1.3.9.2.5 Other DC signals which drive a Treg differentiation 74

1.3.10 Th3 cells 75 1.3.11 Double negative regulatory T cells 75

Hypotheses and Objectives 77

Chapter 2: Characterization of airway adaptive immune response to helper dependent adenoviral vectors 2.1 Abstract 82 2.2 Introduction 83 2.3 Materials and Methods 85 2.4 Results 89 2.5 Discussion 110 Chapter 3: Adenoviral vector formulation in DEAE-Dextran and NAL can increase gene delivery to the airways 3.1 Abstract 114 3.2 Introduction 115 3.3 Materials and Methods 117 3.4 Results 118 3.5 Discussion 128

Chapter 4: Induction of antigen specific tolerance towards adenoviral vectors using immature dendritic cells 4.1 Abstract 130 4.2 Introduction 131 4.3 Materials and Methods 133 4.4 Results 137 4.5 Discussion 158 Chapter 5: Induction of antigen specific tolerance by apoptotic dendritic cells 5.1 Abstract 162 5.2 Introduction 164 5.3 Materials and Methods 167 5.4 Results 173 5.5 Discussion 227

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Chapter 6: Discussion and Future Directions 6.1 Discussion 235 6.2 Future Directions 244 6.3 Conclusions 249 References 250

x

List of Tables

Table 1-1 DC subtypes and their precursors

Table 5-1 Primer sequences for assessment of cytokine induction in viable DCs upon apoptotic

DC uptake

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List of Figures

Figure 1-1 Differentiation of DCs from hematopoietic stem cells.

Figure 1-2 Classification of DC subsets as conventional and non-conventional DCs.

Figure 1-3 Regulation of DC apoptosis.

Figure 1-4 DCs drive differentiation of Tr1 regulatory T cells.

Figure 1-5 DCs drive differentiation of foxp3+ inducible regulatory T cells. (iTregs).

Figure 2-1 Effects of HD-Ad vectors on maturation of bone marrow derived DCs.

Figure 2-2 Induction of CD4+ T cell proliferation in vitro by HD-Ad treated bone marrow derived DCs.

Figure 2-3 Assessment of T cell infiltration in bronchoalveolar lavage fluid upon intranasal delivery of HD-Ad vectors to mice.

Figure 2-4 Assessment of T cell proliferation in the lung and the draining mediastinal lymph node (MLN) in response to pulmonary HD-Ad delivery.

Figure 2-5 Effect of HD-Ad delivery on pulmonary conventional DC (cDC) levels and maturation within the lung.

Figure 2-6 Effect of HD-Ad delivery on DC migration from the lung to the draining MLN.

Figure 2-7 Effect of HDAd delivery on pulmonary plasmacytoid DC (pDC) levels and maturation within the lung.

Figure 2-8 Effects of HD-Ad delivery on CD8α DCs in the draining MLN.

Figure 3-1 X-gal staining of the lung after delivery of FGAd vector particles encoding LacZ under control of CMV promoter (FGAdCMVLacZ) in saline with or without NAL pre-treatment.

Figure 3-2 β-galactosidase activity after delivery of FGAdCMVLacZ vector particles with or without NAL pre-treatment.

Figure 3-3 X-gal staining of the whole lung (left panel) and the trachea (right panel) upon delivery of FGAdCMVLacZ vector particles with or without NAL pre-treatment.

Figure 3-4 Histopathological analysis of the whole lung upon FGAd vector delivery with or without NAL pre-treatment.

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Figure 4-1 HD-Ad mediated transgene expression following readministration is sustained in Rag deficient mice, which are devoid of an adaptive immune response.

Figure 4-2 DCs derived in the presence of IL-10 are refractory to HD-Ad induced maturation.

Figure 4-3 HD-Ad induced maturation of normal DCs is not inhibited by addition of exogenous IL-10.

Figure 4-4 DCs derived in the presence of IL-10 suppress T cell proliferation upon exposure to HD-Ad and instead induce generation of Tr1 regulatory T cells.

Figure 4-5 DCs derived in the presence of IL-10 suppress pulmonary DC maturation and migration in response to HD-Ad vectors.

Figure 4-6 DCs derived in the presence of IL-10 suppress HD-Ad induced pulmonary T cell infiltration in BALF and suppress pulmonary cytotoxic T cell response.

Figure 4-7 DCs derived in the presence of IL-10 suppress HD-Ad induced pulmonary T cell proliferation even after three rounds of HD-Ad delivery.

Figure 4-8 DCs derived in the presence of IL-10 suppress HD-Ad induced antibody production even after three rounds of HD-Ad delivery.

Figure 4-9 Gene expression following three rounds of HD-Ad delivery is sustained in mice immunized with HD-Ad pulsed, IL-10 derived DCs.

Figure 5-1 UV radiation induces apoptosis in DC and splenocytes.

Figure 5-2 Viable DC take up apoptotic DC and this uptake is inhibited by cytochalasin D.

Figure 5-3 Immature/mature apoptotic DC and necrotic DC do not induce maturation of viable DC.

Figure 5-4 Viable DCs fail to upregulate CD86 expression and IL-12 production in response to LPS upon uptake of apoptotic DC.

Figure 5-5 Viable DC become tolerogenic DC upon uptake of apoptotic DC.

Figure 5-6 Viable DC take up apoptotic DC and induce differentiation of naïve T cells into Foxp3+ Treg in vitro.

Figure 5-7 In response to LPS, viable DC that have taken up apoptotic DC, induce secretion of TGF-β1 and upregulate TGF-β2 gene expression, which mediates generation of Foxp3+ Treg.

Figure 5-8 mTOR pathway is involved in induction of TGF-β1 secretion upon uptake of apoptotic DC by viable DC.

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Figure 5-9 Apoptotic or necrotic DCs are taken up by viable DCs in vivo.

Figure 5-10 Uptake of apoptotic DCs by viable DCs in vivo.

Figure 5-11 Delivery of apoptotic DCs to mice results in their uptake by viable DCs, which do not undergo maturation but produce TGF-β1.

Figure 5-12 Apoptotic DCs suppress LPS-induced DC migration from periphery to draining lymph nodes in vivo.

Figure 5-13 Apoptotic DCs suppress LPS-induced DC maturation in vivo.

Figure 5-14 Apoptotic DCs can suppress LPS-induced airway inflammation.

Figure 5-15 Intranasal delivery of apoptotic DCs suppresses LPS induced IL-12 production in DCs in MLN.

Figure 5-16 Intranasal delivery of apoptotic DCs results in the secretion of TGF-β1 and Treg expansion.

Figure 5-17 Apoptotic DCs induce antigen-specific Tregs in vivo.

Figure 5-18 Apoptotic DCs induce expansion of Tregs in vivo.

Figure 5-19 Proposed model for tolerance induction by apoptotic DCs.

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List of Abbreviations

AAV: Adeno associated virus

Ad: Adenovirus

AhR: Aryl hydrocarbon receptor

aTreg: Adaptive Foxp3+ regulatory T cells

BALF: Bronchoalveolar lavage fluid

BMDC: Bone marrow derived dendritic cells

BrdU: Bromodeoxyuridine

cAMP: Cyclic adenosine monophosphate

CAR: Coxsackie and adenoviral Receptor

CCR5: Chemokine (C-C motif) receptor 5

CCR7: Chemokine (C-C motif) receptor 7

CD: Cluster of differentiation

CDP: Common DC progenitor

CF: Cystic Fibrosis

CFA: Complete Freund’s adjuvant

CFSE: Carboxyfluorescein succinimidyl ester

CFTR: Cystic fibrosis transmembrane conductance regulator

CLP: Common lymphoid progenitor

CMP: Common myeloid progenitor

CMV: Cytomegalovirus

COPD: Chronic obstructive pulmonary disease

AAT: Alpha 1- antitrypsin

Cox-2: Cyclooxygenase-2

CTLA4: Cytotoxic T-lymphocyte antigen 4

DCs: Dendritic cells

DEAE-Dextran: Diethylaminoethyl Dextran

DN: Double negative

DNA: Deoxy-ribonucleic acid

CAT: Chloramphenicol acetyltransferase

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DNAzyme: Deoxyribozymes

DOPE: Dioleoyl Phosphatidylethanolamine

DOTMA: N-[1-(2, 3-dioleyloxy) propyl]-N,N,N-trimethylammonium chloride

EAE: Experimental autoimmune encephalomyelitis

EDTA: Ethylenediaminetetraacetic acid

FACS: Fluorescence-activated cell sorting

FG-Ad: First generation adenoviral vectors

FITC: Fluorescein isothiocyanate

FIV: Feline immunodeficiency virus

GALT: Gut associated lymphoid tissue

GFP: Green Fluorescent Protein

HIV: Human immunodeficiency virus

GM-CSF: Granulocyte-macrophage colony stimulating factor

GVHD: Graft versus Host disease

H&E: Haematoxylin and eosin

HAA: 3-hydroxyanthranilate

hAAT: human alpha 1- antitrypsin

HD-Ad: Helper-dependent adenoviral vectors

HGF: Hepatocyte growth factor

HIV: Human immunodeficiency virus

HLA: Human leukocyte antigen

ICOS: Inducible T cell costimulator

ICOSL: Inducible T cell costimulator ligand

IDO: Indoleamine 2,3-dioxygenase

IFN: Interferon

IFN-γ: Interferon gamma

IFNR: Interferon receptor

IgG: Immunoglobulin G

IL: Interleukin

ILT: Ig like transcript

ITR: Inverted terminal repeats

iTreg: Inducible Foxp3+ regulatory T cells

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LC: Langerhans cells

LP: Lamina Propria

LPC: Lipophosphatidylcholine

LPS: Lipopolysaccharide

LTR: Long term repeats

MDP: Macrophage DC Progenitor

MHC: Major histocompatibility complex

MLN: Mediastinal lymph node

MOI: Multiplicity of infection

mTOR: Mammalian target of rapamycin

MUC1: Mucin 1, cell surface associated

NAL: Nacystelyn

nTreg: Naturally occurring Foxp3+ regulatory T cells

OPG: Osteoprotegerin

OVA: Ovalbumin

pDC: Plasmacytoid dendritic cells

PDL-1: Programmed death ligand – 1

PEG:Polyethylene glycol

PFA: Paraformaldehyde

PGD2: Prostaglandin D2

PI: Propidium iodide

Poly I:C: Polyinosinic: Polycytidilic acid

Poly I:C: Polyinosinic:polycytidylic acid

PP: Peyer’s patches

RA: Retinoic acid

RANK: Receptor activator of nuclear factor κB

RANKL: Receptor activator of nuclear factor κB ligand

RAR: Retinoic acid receptors

RNA Ribonucleic acid

RXR: Retinoic X receptors

SeV: Sendai virus

siRNA: small interfering RNA

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STAT: Signal transducer and activator of transcription

STAT6: Signal transducer and activator of Transcription 6

TCDD: 2,3,7,8-Tetrachlorodibenzo-p-dioxin

TGF: Tumor Growth Factor

TLR: Toll like receptor

TNF: Tumor necrosis factor

Tr1: Type 1 regulatory T cells

Tregs: Regulatory T cells

TSLP: Thymic stromal lymphopoetin

UVR: Ultraviolet radiation

VD3: 1α,25-dihydroxyvitamin D3

VDR: 1α,25-dihydroxyvitamin D3 receptor

VIP: Vasoactive intestinal peptide

VSV: Vesicular stomatitis virus

X-Gal: 5-bromo-4-chloro-3-indolyl-beta-D-galacto-pyranoside

X-SCID: X-linked severe combined immunodeficiency

xviii

Dissemination of thesis content

Publications

Kushwah R and Hu J (2011) Complexity of dendritic cell subsets and their function in the host

immune system. Immunology.133(4): 409-19.

Kushwah R and Hu J (2011) Role of dendritic cells in the induction of regulatory T cells. Cell

and Bioscience. 1(1):20.

Kushwah R and Hu J (2010) DC apoptosis: regulation of tolerance versus immunity. Journal of

Immunology 185(2): 795-802.

Kushwah R, Wu J, Oliver JR, Jiang G, Zhang J, Siminovitch KA and Hu J. (2010) Uptake of

apoptotic DC converts immature DC into tolerogenic DC, which induce differentiation of

Foxp3+ regulatory T cells. European Journal of Immunology 40(4):1022-35.

Kushwah R, Oliver JR, Zhang J, Siminovitch KA and Hu J (2009). Apoptotic dendritic cells

induce tolerance in mice through suppression of dendritic cell maturation and induction of

antigen-specific regulatory T cells. Journal of Immunology 183: 7104 -7118.

Kushwah R, Cao H and Hu J (2008) Characterization of pulmonary T cell response to helper

dependent adenoviral vectors following intranasal delivery. Journal of Immunology 180: 4098-

4108.

Kushwah R, Oliver JR, Cao H and Hu J (2007), Nacystelyn enhances adenoviral vector-

mediated gene delivery to mouse airways. Gene Therapy Aug; 14(16):1243-8.

Kushwah R, Cao H and Hu J. (2007) Potential of helper-dependent adenoviral vectors in

modulating airway innate immunity. Cellular and Molecular Immunology Apr; 4(2):81-9.

The contents of the above manuscripts have been used after obtaining copyright permissions.

1

Chapter 1 Introduction

1.1 Gene therapy for airway diseases

Gene therapy involves insertion of therapeutic genes into cells to replace a mutant allele for

treatment of diseases. Gene therapy is particularly attractive for diseases that currently do not

have satisfactory treatment options, and is attractive for monogenic disorders. Gene therapy has

been extensively investigated for airway diseases, largely due to the accessibility of the airways

to gene delivery. The potential of gene therapy for airway diseases has been suggested by animal

studies and the rationale is most apparent for monogenic airway diseases such as cystic fibrosis.

Airway gene transfer efforts have been directed toward recombinant viruses, particularly

adenoviruses, adeno-associated viruses, retroviruses, and complexes of plasmid DNA with

carrier molecules, such as cationic lipids or polymers. However, efficient delivery and expression

of the therapeutic transgene at levels sufficient to result in phenotypic correction of the diseased

state have proven elusive. Adenoviral vectors amongst all the vectors show the highest efficacy

for airway gene delivery, but the drawback has been induction of subsequent immune responses.

1.1.1 Cystic Fibrosis

Cystic Fibrosis (CF) has been the most prominent disease for which lung gene therapy has been

actively investigated (Griesenbach et al., 2004). CF is an autosomal recessive monogenic

disorder that afflicts up to 1 in 3,000 people amongst the Caucasian population and is

characterized by mutations in cystic fibrosis transmembrane conductance regulator (CFTR) gene

which encodes an epithelial chloride channel (Riordan et al., 1989). Clinical manifestations of

CF include incompetent electrolyte transport in several epithelia including airways, pancreas and

sweat glands (Fuller and Benos, 1992). However, the major cause of morbidity and mortality in

CF is due to lung disease (Davis, 2006). The lungs of CF infants are almost normal at birth

except for widening of mouths of submucosal glands indicative of mucus impact (Sturgess and

Imrie, 1982). However, as infants age there is gradual colonization of airways by bacterial

species such as Haemophilus influenzae, Staphylococcus aureaus and eventually Pseudomonas

aeurigonosa (Saiman, 2004), which initially clear with vigorous antibiotic therapy. However,

later on there is permanent establishment of bacterial colonies which are nearly impossible to be

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cleared, probably caused by slower clearance and capture of bacteria due to extremely thick

mucus along with damaged airway epithelium (Davis, 2006). Although the mechanism of Cftr

mutations leading to CF airway disease is not clear, it has been shown that the airway fluid

volume in CF lungs is reduced and this reduction is believed to impair the lung mucociliary

function which is the first line of lung defense. In addition, CF epithelia have exaggerated

responses to proinflammatory stimuli. It is likely that this combination of inefficient airway

clearance and exaggerated inflammatory responses contributes to the excessive neutrophil

infiltration in the CF lung. The chronic infection results in persistence of neutrophil infiltration

along with chronic inflammation leading to destruction of the airways (Tomashefski et al., 1985).

Therefore, the most important aim in terms of CF therapy is treatment of the respiratory system.

The predominant location for CFTR expression in the lung is the submucosal gland serous cells,

found in trachea and bronchi (Koehler et al., 2001). Since, the definite target cell has not been

characterized completely; the CFTR expression using gene therapy is being targeted to both

surface epithelial cells along with submucosal glands. Gene therapy for cystic fibrosis has been

actively investigated for the last 10-20 years, with the tenet being that delivery of the gene

encoding CFTR to the airways would result in functional CFTR and hence a cure for CF defect.

1.1.2 Lung transplantation

For various end-stage lung diseases, lung transplantation is the only option to save the lives of

patients. However, the application of lung transplantation is limited by shortage of donors along

with severe acute rejection and impaired long-term graft function. Studies have shown that in

human lung transplantation, there is elevation of cytokines such as tumor necrosis factor (TNF)-

α, interferon (IFN)-γ, interleukin (IL)-10, IL-12, and IL-18, during the cold ischemic time and

decrease over time after reperfusion (De Perrot et al., 2002). Levels of IL-8 increase after

transplant reperfusion and also correlate with the length of ICU stay along with the loss in lung

function (De Perrot et al., 2001; De Perrot et al., 2002; Fisher et al., 2001). Therefore, gene

therapy is being explored for delivering anti-inflammatory genes to the lung to potentially

increase graft survival and limit lung inflammation. The anatomical structure of the lung along

with the requirement of transient gene expression makes lung transplantation an ideal candidate

for gene therapy. Administration route, timing, vector selection and gene selection are important

aspects that need to be considered for development of effective gene therapy for lung

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transplantation (Sato and Keshavjee, 2006). Studies have shown that adenoviral (Ad) vectors are

by far most highly suited, due to their ability to easily transduce airway cells and being non-

integrating which reduces the risk of integration induced oncogene activation (Kushwah et al.,

2007a). Additionally, helper-dependent Ad vectors, which do not encode for any viral genes can

be employed for lung gene therapy for transplantation as they are far less immunogenic than

earlier used Ad vectors (Kushwah et al., 2007a).The problem of gene therapy vector induced

inflammation is attenuated in lung transplantation since the transplant patients are already under

an immunosuppressive drug regime, which inhibits IFN-γ and anti-Ad IgG production (Cassivi et

al., 2000; Cassivi et al., 1999b; Suga et al., 2002). Initially gene therapy for lung transplantation

was hindered by poor transfection rate obtainable during the period of cold temperature required

to preserve the organ, and by the drawback of unnecessarily transfecting other organs if the gene

was delivered systemically, which has been overcome by delivering through transtracheal route

before retrieving and storing organs at 4C(Cassivi et al., 1999a; Chapelier et al., 1996).

IL-10 has been chosen as a therapeutic gene for lung transplantation due to its effects in

suppressing inflammatory responses. Initial studies in animal model of bronchiolitis obliterans

showed that delivery of the vector encoding for IL-10 within 5 days after lung transplantation

inhibited the development of fibrous airway obliteration (Boehler et al., 1998). Delivery of Ad

vectors encoding IL-10 to the lungs of donor rats ameliorated ischemia-reperfusion injury and

improved early post-transplant graft function, providing evidence that IL-10 gene therapy on

donor lung can improve graft function (Fischer et al., 2001). Additionally, gene delivery with IL-

10 switches the mode of cell death upon ischemia-reperfusion from necrosis to apoptosis, which

also suppresses lung inflammation (Fischer et al., 2003).

An important issue with gene therapy relates to the minimum time necessary to obtain sufficient

gene expression before lung retrieval. A gap of 24 hours between brain death and organ retrieval

is associated with complications including immune response which can lead to loss of the donor

organ. Studies have shown that lungs retrieved 12 hours after Ad mediated IL-10 gene delivery

are associated with significant improvement in post-transplant lung function. In contrast, lungs

retrieved at shorter times show significantly elevated levels of TNF-α and MIP-2, which

increases the extent of injury (de Perrot et al., 2003). This is likely due to the initial immune

response to Ad vectors which is eventually suppressed by IL-10 production from transduced

cells. More recently, this approach has been employed in larger animal models such as pigs and

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studies have shown that IL-10 gene therapy reduced ischemia-reperfusion injury and improved

graft function after lung transplantation (Martins et al., 2004). Furthermore, the approach

prevented the release of inflammatory cytokines such as IL-6 in lung tissue and plasma.

Recently, delivery of short hairpin RNA encoding for caspase 3 in rats has also shown efficacy

in preventing lung-ischemia reperfusion injury by preventing cell death in the transplant (Zhang

et al., 2009b). Studies conducted to date provide proof of principle of using gene therapy in lung

transplantation. However, further improvements in vector design will ultimately improve the

efficacy of the procedure and improve outcomes in lung transplantation.

1.1.3 Asthma

Asthma is a chronic disease which affects both children and adults. Asthma is characterized by

T-helper type 2 (Th2) driven chronic inflammation and narrowing of airways resulting in clinical

symptoms which include cough, shortness of breath, wheeze and chest tightness. Various stimuli

can induce “episodes” or attacks such as allergens, viral infections and even exercise. Gene

therapy has been explored for asthma and most strategies have focused on blocking expression of

pro-inflammatory molecules or transcription factors in the disease pathogenesis using antisense

oligonucleotides, DNAzymes, small interfering RNA, or blocking of microRNAs using

antagomirs (Maes et al., 2011).

Delivery of viral vectors expressing siRNA targeted against Th2 cytokines such as IL-4, and IL-

5 has been shown to result in reduction of airway hyper-responsiveness, reduction of cellular

infiltration in the lung tissue, reduction of airway remodeling along with reduction of other

hallmarks of asthma (Huang et al., 2008; Karras et al., 2007). Activation of the transcription

factor STAT6 is crucial for differentiation of Th2 and also for infiltration of eosinophils and Th2

lymphocytes in the airways. Therefore gene therapy strategies have focused on inhibition of

STAT6 using siRNA. Preclinical studies have demonstrated efficacy of the strategy in

suppressing pathology associated with asthma (Darcan-Nicolaisen et al., 2009). Similarly,

inhibition of GATA-3, the transcription factor that drives lineage commitment towards Th2 cell

type using short hairpin RNA and DNAzyme has shown efficacy in suppressing the cardinal

features of asthma in animal models (Lee et al., 2008; Sel et al., 2008). Dendritic cells are the

key players in asthma by initiating priming of Th2 cells. Therefore gene therapy strategies have

been explored which focus on targeting DC and T cell interactions to suppress Th2

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differentiation. Gene therapy using siRNA targeted against CD86 and CD40, which are

costimulatory markers expressed on mature DCs has been also shown to suppress features of

asthma in animal models (Crosby et al., 2007; Suzuki et al., 2009). Altogether, gene therapy

strategies have shown great promise as a treatment strategy for asthma. Further developments in

vector design will ultimately increase the efficiency of gene therapy strategies targeting asthma.

1.1.4 Chronic obstructive pulmonary disease

COPD is a category of chronic lung diseases characterized by the pathological limitation of

airflow in the airway that is not fully reversible. Individuals with COPD experience shortness of

breath, increased sputum and coughing and are prone to serious conditions such as recurring

chest infections, respiratory failure, pulmonary hypertension and heart failure(Chung and

Adcock, 2008). COPD is common among seniors in Canada and the most common forms of

COPD are chronic bronchitis and emphysema (Cosio Piqueras and Cosio, 2001). Chronic

bronchitis is characterized by persistent cough and sputum production with evidence of

hyperplasia and hypertrophy of goblet cells in the airways resulting in mucus secretion and

airway obstruction (Banning, 2006). Emphysema is characterized by enlarged alveolar sacs

causing a reduction in gas surface exchange area as well as loss in lung elasticity, which burdens

the thoracic cavity and clinically manifests as difficulty in breathing (Banning, 2006). Cigarette

smoking is the principal cause of COPD; however chronic exposure to pollutants can also

contribute to development and exacerbation of COPD (Cosio and Guerassimov, 1999).

Furthermore, individuals with a hereditary disease such as alpha1-antitrypsin (AAT) deficiency

are also predisposed to COPD development. The major function of AAT is to protect tissues

against neutrophil elastase, and pulmonary emphysema associated with AAT deficiency is due to

the unrestrained proteolytic activity of neutrophil elastase on lung connective tissue leading to

alveolar destruction (Lomas and Parfrey, 2004). AAT is primarily synthesized by the liver and is

secreted into the blood where it circulates and diffuses into the lung parenchyma. AAT is also

made by lung epithelial cells and macrophages.

Lung epithelial cells seem to be an ideal target for gene therapy to treat AAT deficiency because

the lung is where AAT is needed most and local production of AAT should be the most effective.

Considerable effort has been invested in attempts to deliver a normal hAAT gene in vivo. Many

of these approaches have been limited by either short-lived gene expression or inability to

6

achieve the very high levels of circulating hAAT (1 mg/ml) presumably required to prevent the

progression of emphysema in hAAT-deficient patients (Stecenko and Brigham, 2003). However,

data from recent studies have provided encouraging results. Adeno-associate viral vector

mediated delivery of AAT to lungs in mice and dogs has been shown to result in therapeutic

levels of AAT expression in the lungs (Halbert et al., 2011). Furthermore, transduction of

alveolar macrophages in the mouse lung by AAT encoding lentiviral vectors has been shown to

result in long term gene expression with amelioration of emphysema (Wilson et al., 2010).

Improvements in vector design along with generation of strategies to prevent the immune

response directed against gene therapy vectors are likely to result in better outcomes for AAT

gene therapy.

1.2 Vectors for lung gene therapy

Broadly, two classes of vectors have been investigated for lung gene therapy, which include viral

and non-viral vectors. Among the viral vectors: lentivirues, adeno-associated viruses and

adenoviruses have been actively investigated. Additionally, owing to its efficient transduction of

airways, sendaivirus is also being intensively investigated. In contrast to viral vectors, non-viral

vector approaches to lung gene therapy have also been developed and these include use of

cationic liposomes, glycoconjugates and transposon based integrating plasmid. In spite of

intensive vector development, there still are challenges to be overcome which include efficient

transgene expression, immunological response against the vectors and physical barriers

preventing vector accessibility to target epithelial cells.

1.2.1 Non-viral vectors for gene delivery

1.2.1.1 Cationic Liposomes / DNA complex

Since the introduction of Lipofectin in 1987, which is a 1:1 mixture of cationic lipid DOTMA

and colipid DOPE (Mahato et al., 1997), many cationic lipid / DNA complexes have been tested

for in vitro and in vivo gene delivery. There are usually two interactions that occur between

DNA and lipids. First being the fusion of lipids with DNA due to charge difference and the

second being collapse whereby due to the small surface size of lipids compared to DNA, they

collapse resulting in formation of DNA coils (Gershon et al., 1993). Upon interacting with cells,

7

the lipids fuse with the cells and deliver the DNA content to the cells. Though liposomes

mediated gene delivery demonstrated efficacy in in vitro and in vivo studies, the clinical trials

did not reveal the same efficacy.

Stribling et al. delivered chloroamphenicol acetyltransferase (CAT) encoding plasmids in

combination with cationic liposomes to mice by aerosol. CAT expression could be observed in

lung up to 21 days especially in airway epithelial and cells of alveolar lining with no treatment

related histopathology (Stribling et al., 1992). Studies were also conducted using CFTR encoding

plasmids in mice models. Hyde et.al. delivered human CFTR encoding plasmids in complex with

cationic liposomes in CFTR knockout mice and CFTR expression could be detected throughout

the airway by in situ hybridization (Hyde et al., 1993). In order to assess correction of CF defect,

the authors measured cAMP stimulated chloride secretion, which was significantly improved in

CF knockout animals receiving CFTR and liposomes complex. This study clearly demonstrated

the efficacy of using cationic liposomes in airway gene therapy. The efficacy of cationic

liposomes for airway gene delivery was also tested in rabbits, however gene expression using

alpha anti –1 antitrypsin as a transgene could only be observed for a period of 7 days in the lung

(Canonico et al., 1994).

BGTC/DOPE cationic liposomes were also tested for fetal airway gene delivery in sheep with

transgene expression observed in airway cells, though at the same time histopathological lesions

could be observed indicating toxicity (Luton et al., 2004). This was particularly due to the large

dose required for sufficient levels of transduction because though sufficient amount of gene

expression can be achieved in lung, the efficiency of gene transfer is extremely low (Lee et al.,

2005). Therefore for sufficient levels of gene expression, a large dose needs to be administered

in humans. Studies looking at effects of airway delivery of higher doses of cationic liposomes

and plasmid complexes did demonstrate significant toxicity. Scheule et.al assessed safety profile

of cationic lipid GL-67 by intranasal delivery in BALB/C mice (Scheule et al., 1997). A dose-

dependent relationship was observed with pulmonary inflammation indicated by neutrophil,

macrophage and even lymphocyte infiltration to the lung. At the same time there were significant

increases in levels of inflammatory cytokines such as IL-6, TNF-α and IFN-γ. Similarly, studies

were conducted with a rat lung transplant model which again indicated a dose-dependent toxicity

with the use of cationic liposomes-DNA complexes (Nagahiro et al., 2000).

8

Eight clinical studies have been conducted so far using cationic liposomes in CF patients (Lee et

al., 2005). In a clinical trial conducted by Alton et.al., CFTR encoding plasmids were complexed

with cationic liposomes and delivered to nasal epithelium of CF patients (Alton et al., 1999).

Seven among eight patients showed flu like symptoms indicating inflammatory response.

Another clinical trial conducted by Noone et.al. in CF patients resulted in no vector specific

mRNA detection in nasal epithelium scrape biopsies after administration of CFTR encoding

plasmid vectors in complex with liposomes intransally (Noone et al., 2000). Therefore, the

current level of transduction efficiency achieved with the use of cationic liposomes is too low to

produce any therapeutic benefit and hence is a major drawback in clinical application of cationic

liposomes in airway gene delivery.

1.2.1.2 Glycoconjugates

The use of glycoconjugates for airway gene delivery is based on the premises that many

receptors present on airway epithelial cells contain covalently linked carbohydrates, which bind

to lectins. Therefore, delivery of vectors in complex with lectins could result in interaction with

the receptors followed by endocytosis. In a study conducted by Yin et.al., lectin-PL/His

conjugates were mixed with plasmids encoding for LacZ and transfected into human airway

epithelial cells, resulting in beta-galactosidase expression and providing the proof of principle of

using glycoconjugates for gene delivery to airway epithelial cells (Yin and Cheng, 1994). Many

studies have looked at the lectin binding patterns of different cells in the human lung indicating

different lectin moieties with varying specificity for different kinds of cells within the lung

(Dorscheid et al., 1999).

Therefore, the knowledge of the specific kinds of lectins that can bind to differentiated human

airway epithelial cells could be a very important tool for development of glycoconjugate based

airway gene delivery therapeutics. Yi et.al., screened 32 lectins and identified 15 lectins that

could bind to the apical membrane of epithelial cells (Yi et al., 2001). Among the lectins that

bound to apical membrane, various lectins demonstrated varying efficiency in being taken up by

cells, with Jacalin lectin binding to PNAS positive cells and Peanut Agglutinin binding to

subpopulations of both ciliated and unciliated cells. Upon looking at endocytosis, lectins such as

Con A were internalized by endocytosis within 1 hour whereas others such as Jacalin took four

hours to be internalized, indicating the variability among lectins in binding to surface versus

9

endocytosis. Currently many different lectins are being investigated; however the best lectin for

gene delivery should bind efficiently to apical surface and be taken up readily via endocytosis.

The efficiency of glycoconjugate mediated gene therapy is determined both by specificity of the

lectins expressed on the cell surface along with lectins present inside cell which mediate

intracellular trafficking (Fajac et al., 1999).

Glycoconjugates have been tested in vitro as well as in vivo models for airway gene delivery.

Modified chitosan oligomer polyplexes by complexed with plasmids encoding for luciferase

have been tested in human epithelial cells along with mice with success, though the persistence

of gene expression has not been addressed (Issa et al., 2006). Vectors encoding for CFTR in

complex with glycosylated polylysines have been administered to tracheal gland serous cells

derived from CF patients with detectable CF expression (Allo et al., 2000). However, no studies

to date have demonstrated efficacy of glycoconjugates in CF gene therapy and neither has been

detailed analysis conducted to study the immunological responses associated with delivery of

glycoconjugates. Though the strategy holds promise, further preclinical studies are needed before

any clinical trials can be initiated.

1.2.1.3 Transposon based integrating plasmids

Transposons are a form of mobile DNA sequences that can move across the genome and

integrate at different sites. The site of integration is mediated by the sequences flanking the

transposable element and integration is mediated by integrase enzyme. Among all the

transposable elements, sleeping beauty transposon system belonging to Tc1/mariner superfamily

has been investigated for airway gene therapy (Izsvak et al., 2000). Sleeping beauty transposon

itself is comprised of two inverted repeats which are 340 bp in size and can efficiently integrate

into human chromosome. The integration is mediated by sleeping beauty transposase which can

be provided to the cell as a gene or mRNA (Hackett et al., 2005). Upon integration the biggest

benefit is that the therapeutic gene can be stably expressed throughout the lifetime of the patient.

Studies demonstrate that using Sleeping beauty system, approximately 10,000 different

integrations can be achieved in human cells (Izsvak et al., 2000).

10

There is an inverse relationship between size of transposon and transposon efficiency. The ability

of sleeping beauty transposon to mediate delivery to the lung has been tested in mouse models.

In a study conducted by Belur et.al., sleeping beauty transposons encoding for luciferase were

injected intravenously in wild type as well as transgenic mice expressing sleeping beauty

transposase (Belur et al., 2003). In wild-type animals expression in the lung could be detected up

to 3 months whereas in transgenic mice the expression was sustained for a much longer time,

indicating importance of transposase. Luciferase expression was widely distributed in the

respiratory zone with no staining observed in conducting airways. Though the expression was

sustained, the efficiency was only 2-3% as assessed by immunohistochemical staining and

integration occurred preferentially at TA rich sites. Liu et.al., combined an endothelial specific

promoter with GFP in sleeping beauty transposase to mediate sustained gene expression in

airway endothelium for greater than 2 months (Liu et al., 2004).

Safety profile associated with the use of transposons for airway gene delivery has not been

assessed. The proponents of transposons argue that the transposable elements have been a part of

human genome for the last 50 million years without any acute toxicity (Essner et al., 2005). In

contrast, a study conducted by Dupuy et.al. demonstrated that sleeping beauty transposons can be

mobilized in mouse somatic cells at very high frequencies which can induce cancer in wild-type

mice (Dupuy et al., 2005).

1.2.2 Viral vectors for gene delivery

1.2.2.1 Retroviral vectors

In 1990, a study conducted by Drumm et.al.(Drumm et al., 1990), was the first ever report of the

use of retroviruses for CFTR gene delivery. Amphotropic retroviruses encoding for CFTR were

used to efficiently transduce a pancreatic adenocarcinoma cell line (CEPAC-1) derived from a

CF patient. Retroviral gene delivery was able to correct the defect as assessed by measurement of

chloride efflux using whole cell patch clamp studies. Subsequently, Olsen et.al.(Olsen et al.,

1992), demonstrated retroviral mediated CFTR delivery in a human epithelial cell line (CFT1),

leading to correction of the defect and partial maintenance of the correction up to six months.

11

Several studies were also conducted to demonstrate the utility of retrovirus in targeting

progenitor cells. Primary cultures of tracheal epithelial cells could be transduced by LacZ

encoding retrovirus and upon transfer of these cells to denuded trachea implanted in nude mice,

these cells differentiated into a variety of epithelial cells, with all cells expressing lacZ

(Engelhardt et al., 1991; Grove et al., 2002). At the same time, studies were conducted to identify

the progenitor cells, targeting of which with retroviruses could theoretically result in permanent

cure of the CF defect (Duan et al., 1998). Thus, the transduction efficiency and the benefits of

retroviral vectors were being realized. However, studies were quick to report that the efficiency

of retroviruses was dependent on the differentiation and mitotic state of the cell, with highest

transduction efficiency achieved with maximum differentiation and almost no transduction in

fully differentiated epithelium (Engelhardt et al., 1992). Studies were also conducted to measure

retroviral transduction in basal as well as secretory airway epithelial cell population(Halbert et

al., 1996) with no differences observed between the two. In vivo studies demonstrated that

transduction of rabbit airway epithelial cells only occurred when the trachea was wounded.

Wang et al.(Wang et al., 1998) reported the use of Keratinocyte growth factor (KGF) to induce

proliferation of airway epithelium resulting in increased retroviral transduction efficiency.

However, there could be multiple drawbacks and side-effects associated with use of KGF such as

malignancy. There were reports also indicating that the age of an animal could play a major role

in retroviral mediated transduction efficiency (Johnson et al., 1998). Due to the reduced

transduction efficiency and the requirement for cellular differentiation, attention gradually

focussed towards the use of lentiviruses as vectors for gene therapy.

Lentiviruses have been investigated for airway gene therapy due to their ability to integrate their

genomes, resulting in the possibility of permanent correction of the genetic disorder through

targeting of progenitor cells. Lentivirus belong to the family of Retroviridae, which are

characterized by single stranded RNA genome which is reverse transcribed upon infection by the

virally packaged RT. Among Lentiviruses, Human Immunodeficiency Virus – 1 (HIV-1) and

Feline Immunodeficiency virus (FIV) have been investigated as vectors for airway gene delivery.

Lentiviral genomes encode for structural (gag, env, pol) as well as regulatory proteins (tat, rev,

vif, nef, vpr and vpu)(Goldman et al., 1997). Gag gives rise to three structural proteins which

include matrix, nucleocapsid and capsid. Pol encodes for protease and reverse transcriptase,

wherease env encodes for surface glycoprotein (gp120) and transmembrane protein

12

(gp41)(Krambovitis and Spandidos, 2006). The gradual development of lentiviral vectors was

initiated with the use of other retroviral vectors such as Moloney murine leukemia virus

(MuMLV) for gene therapy.

HIV-1 is the prototype of lentiviral vectors and over the past 20 years there has been

development of 3 generations of HIV-1 vectors. Naldini et al. (Naldini et al., 1996), were the

first to report replication defective forms of HIV-1 which could be used as vectors for gene

therapy. The first generation vectors used three plasmids: packaging construct, envelope

construct and transgene construct, co-transfected in 293T cells for production of vector particles.

The packaging construct was derived from HIV provirus and was deleted for packaging signal,

env sequences and LTR’s were replaced with CMV promoter. The envelope was provided in a

second plasmid and a third plasmid encoding for transgene with the HIV packaging signal was

used, to be packaged in viral particles produced. In the second generation vectors, all the

virulence factor encoding genes including vif, vpu and nef were deleted from the packaging

construct (Zufferey et al., 1997). Eventually there was development of so called third generation

vectors, where packaging construct only encoded for gag, pol and rev (Dull et al., 1998).

HIV entry into cells is mediated by gp120 and gp41, encoded by env. Gp120 binds to CD4 and

secondary interactions with CXCR4/CCR5 mediate entry (Krambovitis and Spandidos, 2006).

Therefore, the natural tropism of HIV limits their infection to CD4+ T cells and macrophages.

Therefore, pseudotyping of HIV envelope protein with VSV-G was used to broaden the tropism

of HIV derived vectors to efficiently transduce airway epithelial cells. VSV-G uses a common

membrane phospholipid as a receptor, resulting in an extremely broad tropism. Goldman

et.al.(Goldman et al., 1997) showed that in contrast to retroviral vectors, VSV-G pseudotyped

HIV based vectors could efficiently transduce non-dividing epithelial cells in vitro. At the same

time, the vector could be used to efficiently transduce undifferentiated CF derived epithelial cells

in human bronchial xenograft model, resulting in correction of the defect. VSV-G pseudotyped

lentiviral vectors were also able to efficiently transduce human fetal tracheal xenografts (Lim et

al., 2003). At the same time, VSV-G pseudotyped vectors could only transduce airway epithelia

in vivo when access was provided to the basolateral surface by opening the tight junction

(Kobinger et al., 2001). Pretreatment of mouse nasal epithelium with detergent

lipophosphatidylcholine (LPC) followed by instillation of lentiviral vectors encoding for LacZ

resulted in prolonged gene expression up to 92 days and CFTR encoding vectors led to partial

13

correction of CF defect in mice up to a period of 110 days (Limberis et al., 2002). Due to the

need of opening tight junctions associated with VSV-G pseudotyping, a lot of work has been

carried out on identification of proteins that can be used to pseudotype HIV derived vectors for

efficient delivery to airway epithelium. Filoviridae, which include Marburg and Ebola virus have

natural tropism for respiratory epithelium, making their envelope proteins good candidates for

pseudotyping HIV derived vectors. Pseudotyping of HIV vector with envelope protein derived

from Zaire strain of Ebola has been shown to result in efficient transduction of airway epithelial

cells in vitro as well as in vivo without a need of opening tight junctions (Kobinger et al., 2001).

However, the safety concern associated with HIV also led to investigation of FIV derived vectors

for airway gene delivery. Though FIV does not infect human cells, Poeschla et al. (Poeschla et

al., 1998) were the first group to report the use of pseudotyped replication defective FIV to

efficiently transduce dividing, growth-arrested, and postmitotic human targets. Wang et al.

(Wang et al., 1999) developed second generation FIV vectors by deleting trans-acting elements

(vif, orf2) and used them to efficiently transduce airway epithelial cells in vitro as well as in

vivo. FIV vectors encoding for CFTR were able to efficiently transduce cultures of primary

differentiated human airway epithelial cells derived from CF patients and correct the CF defect.

At the same time upon EGTA treatment to open tight junctions, FIV vectors were able to

efficiently transduce dividing as well as non dividing airway epithelial cells in vivo. FIV vectors

have also been pseudotyped with envelope protein of Marburg and Ebola, resulting in efficient

gene transfer when applied to the apical surface of airway epithelium (Sinn et al., 2003). In order

to increase the safety profile associated with the use of Ebola coat proteins, mutant Ebola coat

proteins have also been used to demonstrate efficient transduction of airway epithelium (Medina

et al., 2003).

Along with Ebola, pseudotyping of HIV and FIV vectors with influenza and respiratory syncitial

virus (RSV) has also been investigated. However, the results have been discouraging with

inefficient transduction of human epithelial cells when applied on the apical side (Kobinger et

al., 2001). Overall, so far HIV vectors pseudotyped with Ebola envelope proteins have

demonstrated the highest efficiency in transducing airway epithelium in vitro as well as in vivo

(Wilson, 2004).

14

Most of the clinical trials conducted so far have used retroviral vectors. Most importantly, X-

SCID gene therapy clinical trial, where MFG retroviral vectors were used for ex vivo gene

transfer to bone marrow derived cells, showed the most important drawback associated with

retroviral/lentiviral vectors (Hacein-Bey-Abina et al., 2003b). Three out of eleven patients, went

on to develop leukemia due to integration of retroviral vector near the LMO-2 proto-oncogene

promoter (Hacein-Bey-Abina et al., 2003a). Recently, in 2003, the first clinical trial using

lentiviral vectors was initiated by VIRxSYS (Morris, 2005). Studies have looked at lentiviral

integration in cells and indicate that integration is favoured in transcription units in both dividing

and non-dividing cells, with this trend being significantly stronger in non-dividing cells (Ciuffi et

al., 2006).

1.2.2.2 Sendai virus-based vectors

Sendai virus, also known as hemagglutinating virus of Japan (HVJ), has been investigated as a

vector for airway gene delivery over the last ten years. Sendai virus (SeV) is a negative strand

RNA virus closely related to human parainfluenza virus type I (hPIV1) and belongs to the family

of Paramyxoviridae. SeV causes severe respiratory illness in mice, however the virus is

considered to be non-pathogenic in humans (Bitzer et al., 2003). The SeV genome encodes for

six proteins which include nucleocapsid (N), viral phosphoprotein (P), matrix (M), fusion (F),

hemagglutinin-neuroaminidase (HN) and major polymerase subunit (L)(Bitzer et al., 2003). SeV

replication is independent of the nucleus and occurs in cytoplasm without a DNA phase through

the use of viral RNA polymerase. The RNA genome is tightly bound to N and the RNA-

dependent RNA polymerase activity depends on proteins N, P and L (Sedlmeier and Neubert,

1998).Therefore, SeV is incompetent in transforming cells by integrating its genome (Li et al.,

2000), resulting in a requirement for vector readministration for sustained transgene expression.

Hemagglutinin-neuroaminidase (HN) and the fusion protein (F) determine the tropism of Sendai

virus (Tashiro and Seto, 1997). HN serves two different functions, firstly, it binds to sialic acid

containing gangliosides present on mammalian cell membranes and secondly it induces cleavage

of sialic acid residues to prevent self-aggregation of progeny virus around the same cell. Upon

binding of HN to cell membranes, F protein mediates fusion of the viral particle to the cell

membrane (Tashiro et al., 1990). Though the ability of Sendai virus in mice was characterized

due to its natural tropism, studies were conducted to assess the ability of SeV to replicate in non-

15

human primate cells. In 2002, Kano et.al., assessed the SeV replication in macaques. SeV vectors

encoding for SIV gag were readministered intranasally in macaques and robust gag expression

was observed in nasal mucosa, up to 13 days after administration (Kano et al., 2002).

Additionally, no adverse clinical manifestations were observed in the immunized animals.

Similarly, the ability of SeV to replicate was also demonstrated in other primates such as African

green monkeys and chimpanzees (Skiadopoulos et al., 2002).

The process of SeV vector generation involves transfection of a host cell line with plasmids

encoding for genome RNA and transgene along with plasmids encoding for N, P and L, with

expression of all plasmids driven by a T7 polymerase, supplied by a recombinant virus (Garcin et

al., 1995). Once in the cell, N, P and L proteins are produced and they self-assemble into

nucleocapsids which bind to and replicate the genomic RNA. The replicated RNA is then

packaged to generate SeV particles (Leyrer et al., 1998). Gradually there was development of

second generation SeV vectors, which were completely devoid of viral F protein encoding

sequences. The strategy involves transfection of packaging cell with F encoding plasmid in

addition to all the plasmids used for first generation along with deletion of F sequences on the

genome RNA (Hirata et al., 2002). Upon transduction of the target with second generation

vector, since the vector does not encode for F protein, the progeny is devoid of F protein and

hence cannot infect other cells i.e. no secondary infection is feasible (Waddington et al., 2004).

In 2000, Yonemitsu et al.(Yonemitsu et al., 2000), investigated the use of SeV vectors for gene

transfer to airway epithelium. LacZ encoding SeV vectors were used to achieve 70-80%

transfection efficiency in mice with similar efficiencies observed in ferrets. At the same time,

when freshly isolated primary human nasal epithelial cells were transfected with SeV vectors in

vitro, efficient transfection was achieved which was approximately 2 fold higher than the use of

cationic liposomes. In contrast to liposomes, only a brief exposure time with the cells was

sufficient for efficient transduction, for less than 10% virus could only be recovered, 5 minutes

after treatment of primary cells. Additionally, SeV gene transfer to mucous covered or depleted

sheep trachea was virtually identical. Overall the study indicated efficient transduction of

epithelium in a very short contact time irrespective of the presence of mucous. SeV’s ability to

efficiently transduce airway epithelium has also been exploited to produce therapeutic levels of

circulating proteins such as IL-10 (Griesenbach et al., 2002).

16

SeV has also been used to efficiently transduce differentiated human airway epithelial cells in

vitro. Application of GFP encoding SeV to differentiated human airway epithelial cells has been

shown to result in up to 90% of the cells expressing the transgene followed by a gradual decrease

in the numbers of GFP positive cells after 3 weeks (Pinkenburg et al., 2004). Second generation

SeV vectors have also demonstrated efficient transduction of mice airway epithelial cells in vivo

and human airway epithelial cells in vitro, with levels comparable to the use of F protein

encoding first generation SeV vectors (Ferrari et al., 2004). Second generation SeV-CFTR

vectors have also been shown to correct chloride transport defect in CF knockout mice

(Griesenbach et al., 2005). However, the correction was transient and indicated a requirement for

repeated readministration.

Currently, one of the limitations with the use of SeV is the small size of the packaging construct.

So far, only gene with sizes upto 3.4 kb have been inserted in SeV vector (Bitzer et al., 2003).

However, the upper limits have not been analyzed, though theoretically it is feasible to use larger

genes. At the same time, though there are no infectious particles produced after the use of second

generation SeV vectors, the F protein lacking particles could still be immunogenic and may

potentiate an immune response against SeV vectors. Therefore there is a need to develop SeV

based vectors, which upon infection do not result in production of any particles. At the same

time, the second generation vector encodes for M protein, which has been shown to promote SeV

particle formation (Takimoto et al., 2001) and hence needs to be deleted from the vector

backbone for enhanced safety.

The antibodies generated against surface glycoproteins of hPIV1 have been shown to cross-react

with SeV HN protein(Bitzer et al., 2003). hPIV1 infects infants repeatedly until there is

generation of efficient immunity against the virus and eventually, there is an induction of long

lasting memory immune response which cross reacts with Sendai virus, suggesting SeV to be a

good candidate for hPIV1 vaccine (Smith et al., 1994b). Therefore, the presence of pre-existing

immunity against hPIV1 could be a potential problem in the utilization of SeV as vectors for

human airway gene therapy. Griesenbach et al. (Griesenbach et al., 2006a), used intranasal

delivery of T cell immunodominant epitopes to induce tolerance in mice before delivery of SeV

vectors. The strategy resulted in partial reduction of T cell proliferation in response to SeV

vectors with no reduction seen in levels of antibody production. This indicated the complexity

17

associated with the immune response against SeV vectors and the importance of immune

responses against SeV in developing a therapeutic agent for human gene therapy.

Phase I clinical trials with SeV have been conducted, though not in the context of lung gene

therapy. Intranasal delivery of live SeV in healthy adults demonstrated good tolerance with

evidence of immunogenicity (Slobod et al., 2004).

1.2.2.3 Adeno-associated viral vectors

Adeno-associated virus is a non-pathogenic single stranded DNA parvovirus part of the

dependovirus genus, which requires coinfection with a helper virus such as adenovirus or herpes-

simplex virus to complete its lytic life cycle(Conway et al., 1997). The 4.7kB viral genome is

flanked by inverted terminal repeats (ITRs) and encodes for three structural viral capsid proteins

(VP1, 2 and 3) encoded by cap along with four proteins required for replication (Rep 78, 68, 52

and 40) encoded by rep(Berns and Linden, 1995). The ITRs contain all the cis-acting sequences

required for viral replication and site-specific integration(Flotte, 2005a). Two promoters along

with alternative splicing are used for generation of replication proteins and a third promoter in

combination with alternative splicing and alternate translation codons is used for generation of

structural proteins (Choi et al., 2005). In the absence of helper virus, the virus undergoes a latent

phase through rep proteins mediating integration of viral genome into host cell DNA (Cheung et

al., 1980) without any pathological consequences (Flotte and Berns, 2005).

Among all the serotypes, AAV serotype 2 is the most frequently studied and has been used for

development of gene therapy vectors (Flotte, 2005b). AAV2 vector plasmids, depleted of rep and

cap genes, but with ITRs flanking the therapeutic transgene and appropriate promoter and

polyadenylation signals are transfected into permissive cells along with helper plasmids

encoding for rep and cap (Tratschin et al., 1984). At the same time, cells are infected with helper

virus usually, adenovirus, which promotes the lytic cycle (Tratschin et al., 1985). The AAV

particles produced, package the transgene due to it being flanked by ITRs, which are then

purified and used as vectors for gene therapy. Approximately 100 different variants of primate

AAV have been identified along with avian, canine and bovine strains (Gao et al., 2002). Though

all the strains are non-pathogenic and stable, they differ in their tissue tropism.

18

One of the earliest uses of AAV vectors was for CF gene therapy. Initially, one of the difficulties

with the use of AAV vectors was the packaging limit of 5 kb including the requirement of ITRs

(0.3kb). This limited the capacity to 4.7 kb, with the CFTR gene itself being 4.4 kb in size,

thereby limiting the use of exogenous promoters. The expression was dependent on the use of

AAV p5 promoter. The construct was tested using neomycin resistance gene and transfected into

IB3 (airway epithelial cell line derived from CF patients) cells. Approximately 70% transduction

efficiency could be achieved and 5’-ITR seemed to have an enhancer effect on p5 promoter

(Flotte et al., 1992). The combination of p5 promoter and ITR was also tested with CFTR

transgene in IB3 cells resulting in restoration of cAMP regulation of the chloride efflux (Flotte et

al., 1993b). Eventually, in vivo rabbit models were used to test the efficacy of AAV-CFTR

vectors. The vector was transferred to one lobe of rabbit lung using a fiberoptic bronchoscope

and vector DNA along with recombinant CFTR mRNA as well as protein could be detected for

up to six months (Flotte et al., 1993a), indicating efficient and sustained gene transfer using

AAV vectors. Additionally, analysis of vector DNA in targeted cells indicated the presence of

vector DNA only in low molecular weight episomal form, with no vector DNA detected in high

molecular weight, genomic DNA (Flotte et al., 1994), demonstrating that even without

integration, episomal vector DNA can direct transgene expression.

Although, wild type AAV2 integrates in a site specific fashion on chromosome 19, in vitro

studies have indicated that recombinant AAV2 vectors, though they stay episomal

predominantly, some integration can also occur in a non-specific fashion (Kearns et al., 1996).

Studies were also conducted to assess the efficiency and safety of AAV-CFTR vector in

primates. Conrad et al., delivered AAV-CFTR vectors to the posterior basal segment of the right

lower lung lobe of Rhesus macaques and assessed inflammation as well as vector persistence.

Vector DNA in episomal form along with mRNA could be detected upto a period of 6 months

with no evidence of toxicity or inflammation as assessed by pathological examination along with

cytokine measurements (Conrad et al., 1996).

Preclinical studies demonstrated the efficiency as well as safety of AAV-CFTR vectors.

Eventually phase I and phase II clinical trials were initiated. Phase I trials conducted,

demonstrated dose-dependence in vector gene transfer, with more vector particles resulting in

higher gene transfer. Gene transfer as assessed by the presence of vector DNA indicated the

presence of 0.6 copy/cell at 14 days, which declined to 0.1 copy/cell by 30 days, with no

19

observable detection by 90 days (Aitken et al., 2001). Though the vector was tolerated well with

no adverse reaction, the period of vector prevalence was much lower than that observed in

primate studies. In another clinical study conducted by Flotte et al.(Flotte et al., 2003) to assess

vector safety, 25 CF patients were administered AAV-CFTR vectors to the right lungover a wide

dose range. The study indicated that at high doses, there was significant development of anti-

AAV-neutralizing antibodies, which would interfere with re-administration of the vector. At the

same time, no change could be observed in nasal potential difference measurement, a readout of

CFTR function after vector delivery. Phase II clinical trials to measure safety and efficacy of

readministration were also initiated. Moss et al. (Moss et al., 2004) conducted a phase II clinical

trial, where CF patients were given repeated administrations of AAV-CFTR vectors (3 doses, 1

month apart). This study, noted improvement in lung function, as assessed by forced expiratory

volume. However, the improvement would only be seen after first administration and not after

second or third. Additionally, increase in antibody titer was reported by Flotte et al. (Flotte et al.,

2003). Later, large studies based on vector-readministration were initiated, however the trials did

not meet primary outcome and were discontinued (Griesenbach et al., 2006b). In CF patients,

airway epithelial cells have a significantly higher rate of turnover compared with healthy

controls, which further reduces the persistence of the virus and hence requires increased re-

administrations. Perhaps, in order to reduce the re-administrations, there is a need to develop

AAV vectors that integrate in a site-specific fashion and persist longer in the airway epithelial

cells.

AAV2 has been used in human lung gene therapy clinical trials, however AAV5 and AAV6 have

been showed to mediate a more efficient transduction of mice airways and human airway

epithelial cells (Halbert et al., 2006). Heparan sulphate proteoglycan (HSPG) is the primary

receptor used by AAV2 for attachment into cells (Sanlioglu et al., 2001). The relatively low

abundance of this receptor on the apical surface of epithelial cells limits the transduction

efficiency of AAV2 vectors. Other barriers include extracellular inactivation by neutrophil

derived defensins along with the presence of pre-existing antibodies against AAV2 capsid

proteins. In order to enhance transduction efficiency, studies have been conducted to pseudotype

AAV2 with capsid of other serotypes along with use of stronger promoters. Pseudotyping of

AAV2 with AAV5 or AAV1 capsid demonstrated a significant increase in transduction

20

efficiency (Virella-Lowell et al., 2005). Similarly, use of CMV enhancer/chicken beta-actin

hybrid promoter led to increased transgene expression (Virella-Lowell et al., 2005).

1.2.2.4 Adenoviral vectors

Adenovirus belongs to the family of double stranded DNA viruses with a linear genome of

approximately 34-38 kb in size (Tatsis and Ertl, 2004). They are species-specific and have

different serotypes, with approximately 51 human serotypes (Tatsis and Ertl, 2004) identified so

far. The human serotypes cause mild respiratory or gastrointestinal disease in children and in

immuno-compromised adults. The genome contains five early units (E1A, E1B, E2, E3, E4), two

units (IX, IVa2) expressed after initiation and one late unit (L1-5) along with two species of

RNA, referred to as virus-associated RNA (VA RNA). Among the early genes, E1A and E1B

encode for regulatory proteins that impact on host cell cycle and proteins required for expression

of other early and late genes, E2 encodes proteins for viral DNA replication, E3 for immune

response and E4 for inhibiting host cell apoptosis (Imperiale and Kochanek, 2004). The late

genes encode for structural proteins.

1.2.2.4.1 Early generation adenoviral vectors

The initial vectors derived from adenovirus were referred to as first generation vectors (FGAd).

In these vectors, there was deletion of E1 region or E1 and E3 region. This allowed a maximal

transgene capacity of approximately 5 kb in E1 deleted and 8.2 kb in E1 and E3 deleted vectors

(Danthinne and Imperiale, 2000). Since the E1 product is necessary for viral replication, the

generation process involved transfection of permissive cells with E1 deleted vector, with E1

being provided in trans. However, deletion of E1 could be easily overcome by high multiplicity

of infection and also by E1 like factors present in the cell such as NF-IL6, a member of CEB/P

family (Yang et al., 1995).

Engelhardt et al. (Engelhardt et al., 1993), utilized a human bronchial xenograft model to study

efficiency of LacZ encoding FGAd vectors. The vectors were able to transduce all the cells in

surface epithelium except for basal cells and expression sustained for a period of 3-5 weeks

correlating with E2a expression. In spite of the transduction efficiency observed in vitro, in vivo

studies painted a different picture. Grubb et al. demonstrated the inefficacy of FGAd-CFTR in

correcting sodium transport defect in CF mice, even when high vector dose / repeated doses were

21

administered (Grubb et al., 1994). This could be attributed to the localization of coxsackievirus

and adenovirus receptor (CAR) which is the receptor used by adenovirus. The receptor is

selectively localized onto the basolateral surface of the epithelial cells, which is inaccessible to

the vector particles applied on the apical side due to the presence of tight junctions, thereby

limiting the transduction efficiency (Pickles et al., 2000).

In vivo studies indicated that upon vector administration, there was a gradual development of an

inflammatory immune response. Administration resulted in mild, transient and a dose-dependent

cellular inflammatory responses (Yei et al., 1994b). Repeated readministration resulted in

significantly reduced levels of gene expression compared to naïve animals and this reduction

inversely correlated with the amount of anti-adenovirus neutralizing antibody levels (Yei et al.,

1994a). Clinical trials were also initiated with FG-Ad vectors in CF patients. After lobular

administration of low dose there were reports of cough and fever. However, at high doses 67%

patients developed high fever, transient myalgias, increased sputum production along with

cellular infiltrates (Joseph et al., 2001). Along with epithelial cells, even mononuclear

inflammatory cells were infected by the vector. The local inflammatory response caused by the

expression of viral genes from the FGAd induced a potent immune response to transduced cells,

which was a major factor limiting the efficiency of FGAd vectors (Knowles et al., 1995). The

immune response led to elimination of virally transduced cells, which further limited the

transgene expression (Yang et al., 1994). Due to the presence of serious immunological problems

associated with FGAd, a second generation adenoviral vectors (SGAd) was developed, which in

addition to E1, was also deleted for E2 and E4 sequences (Schaack, 2005), allowing for a cloning

capacity of up to 14 kb (Alba et al., 2005) . Though, the inflammatory immune response was

diminished with the use of SGAd, it was not completely eliminated, probably due to the

continued leaky expression of viral late genes (Schaack, 2005; Yang et al., 1995).

1.2.2.4.2 Third generation – Helper dependent adenoviral vectors

Eventually, third generation adenoviral vectors, also referred to as gutless or helper-dependent

adenoviral vectors were generated. The only adenoviral DNA present in these vectors is the

packaging signal along with 3’ and 5’ ITRs, allowing for a cloning capacity of up to 36 kb.

However, since adenoviruses efficiently package DNA which is only 75-105% of Ad genome,

and transgenes rarely are 36kB in size, stuffer DNA was used for designing vector DNA of

22

appropriate size (Alba et al., 2005). Initially, lambda phage DNA was used as stuffer DNA.

However, upon administration of gutless vectors, there was a cellular immune response, owing to

the immunogenicity of the peptides encoded by stuffer DNA being presented on the cell surface

(Parks and Graham, 1997). Therefore, the choice of stuffer DNA is important and normally it is

the human intronic sequences that are used.

Generation of gutless vectors requires a helper virus, thereby giving the name of helper-

dependent adenoviruses (HD-Ad) to gutless vectors. Permissive cells, usually Cre/293 cells,

which express endogeneous Cre recombinase are transduced with a helper adenovirus which

essentially is the complete adenoviral genome, with the packaging signal spanned by loxp sites

(Parks et al., 1996). At the same time, a gutless vector containing the transgene and stuffer

sequences, flanked by 3’ and 5’ ITRs is also transfected. The helper virus encodes for all the

proteins required to produce the virus, however Cre recombinase produced by the cell cuts at the

loxP site, removing the packaging signal and preventing the helper virus genome to be packaged.

On the other hand, gutless genome with the packaging signal is packaged, resulting in production

of a complete virus encoding only for transgene.

Since one of the biggest concerns with the previous vectors was the immune response, several

studies were conducted to investigate the immune response to HDAd vectors. Studies comparing

HDAd to FGAd report negligible inflammation upon administration of HDAd along with a more

sustained transgene expression up to a period of 28 days with beta-galactosidase and 15 weeks

with human alpha fetal protein (Toietta et al., 2003). O’Neal et.al., compared the toxicity of

FGAd with HDAd and reported that upon administration of FGAd, there was transgene

expression increasing from 1 to 3 days which drastically dropped after 7 days, and at the same

time there was rise in liver enzymes and hepatocyte proliferation indicating toxicity (O'Neal et

al., 2000). In contrast, administration of HDAd led to more stable transgene expression without

any observable hepatic toxicity. Therefore, in contrast to FGAd, HDAd are able to mediate long-

term, high level transgene expression in the absence of chronic toxicity due to the absence of any

viral protein expression in transduced cells.

The efficacy of HDAd vectors encoding for CFTR has been tested in vitro as well as in vivo in

rodents along with primate models. HDAd vectors encoding for CFTR were administered

intranasally in CFTR knockout mice and transgene expression as assessed by RNA and protein

23

expression, could be detected upto a period of 28 days in whole lung and bronchioles with

minimal toxicity (Koehler et al., 2003). The therapeutic potential was assessed by infecting the

animals with Burkholderia cepacia, a bacterium that causes severe lung hisopathology in CFTR

knockout mice but not in wild type animals with functional CFTR. CFTR knockout animals

receiving HDAd CFTR vectors upon Burkholderia infection, showed significant reduction in

lung histopathology compared to CFTR knockout animals not receiving any gene therapy

vectors. This clearly demonstrated the therapeutic potential of HDAd vectors in reducing

susceptibility of CF patients to opportunistic infections. The ability of the vector was also tested

in rabbits, by aerosolizing vector particles using an intracorporeal nebulizing catheter. HDAd

particles encoding for LacZ were aerosolized and delivered with 0.1% lipophosphatidylcholine.

X-gal staining demonstrated transduction of all types of surface epithelial cells with extensive

transduction of airway epithelium. However, fever and mild pneumonia were also observed

which could possibly be due to the inflammatory nature of LPC (Koehler et al., 2005) . The

system has also been tested in primate models with encouraging results. Altogether, Ad vectors

have been shown to be most promising for lung gene therapy applications, but several challenges

need to be overcome prior to their applications in a clinical setting.

1.2.3 Challenges to lung gene therapy

The two biggest challenges associated with lung gene therapy are the physical barriers to gene

therapy vectors and the immunological response against the vectors. Although, correction of 5%

of the epithelial cells results in more than 50% correction of chloride ion transport defect(Dorin

et al., 1996), it still represents an extremely daunting task due to the ability of airways to

neutralize foreign particles including vectors encoding CFTR along with the physiological

features of the airways which acts as a physical barrier.

The prominent physical barriers to lung gene therapy include the production of surfactant

proteins and mucus (Gautam et al., 2002) along with the mucociliary clearance system,

glycocalyceal barrier and slow rate of luminal endocytosis by epithelial cells (Pickles, 2004).

The lumen of the human airways is lined with mucociliary epithelium which is comprised of

ciliated cells along with mucus secreting goblet cells. At the same time, in chronic diseases, such

24

as cystic fibrosis the sub mucosal glands enlarge, there is as increase in the number of mucus

secreting goblet cells and these cells also appear in distal airways where they are normally absent

(Bernacki et al., 1999). In this setting, there is a significant increase in levels of mucus

production along with impairment of mucociliary clearance, which eventually results in chronic

bacterial infection. The build-up of bacterial biofilms along with chronic inflammation further

increases the viscosity of the mucus in the airways and acts as a significant barrier against gene

therapy vectors. Removal of mucus in sheep airway epithelial model has been shown to result in

25 fold enhancement of gene delivery using liposomes mediated gene transfer (Kitson et al.,

1999).

The glycocalyx of the airway epithelium is composed of complex carbohydrates including

glycoproteins and proteoglycan. This is assembled into an organized structure that is bound to

the surface of epithelial cells and constitutes a meshwork that needs to be crossed by gene

therapy vectors to access the cell. In addition to glycocalyx, airway epithelial cells also secrete

mucin and other matrix components such as collagen and fibronectin (Wang et al., 2002). The

barriers erected by glycocalyx have been well documented with most of the vectors. Primary

human airway epithelial cells expressing mucins such as MUC1 are eight fold less refractory to

adenoviral mediated transduction compared to cells not expressing mucin (Arcasoy et al., 1997).

Even after deletion of MUC1 in mice, adenoviral transduction was very inefficient, due to other

components of glycocalyx (Stonebraker et al., 2004). Similar observations have been made with

AAV2 vectors, indicating that treatment of cells with glycosidases and neuraminidases is needed

to increase transduction efficiency (Bals et al., 1999). Additionally, the same drawback has been

observed with retroviral along with lentiviral vectors (Wang et al., 2002).

Another barrier is the localization of receptors used by viruses for entry into the cell. The

primary receptor used by adenovirus for attachment to the cell is the Coxsackie’s and adenoviral

receptor (CAR), which is localized onto the basolateral surface of epithelial cells. Application of

the virus to the apical surface results in very inefficient transduction, though application of

agents such as lipophosphatidylcholine (LPC) which open the tight junctions and allow for viral

entry to basolateral surface, results in significant increase in transduction efficiency. In contrast,

though AAV vectors can enter epithelial cells from the apical surface, the efficiency is higher

when applied from the basolateral side. This is due to endosomal processing which targets AAV

25

capsid proteins through ubiquitination. Application of proteasome inhibitors has been shown to

significantly increase AAV transduction efficiency when virus is applied from the apical side

(Duan et al., 2000). In chapter 3, we devise a new strategy of formulating adenoviral vectors

which can significantly enhance efficiency of gene transfer to the airway epithelial cells upon

intranasal delivery.

The host immune response poses the biggest challenge to lung gene therapy and is composed of

innate and adaptive responses. Epidemiological studies indicate that approximately 80% of the

population have antibodies directed against AAV2, with approximately 30% of the population

maintaining neutralizing antibodies (Peden et al., 2004). Similarly, there is significant prevalence

of anti-Ad5 antibody, ranging from approximately 37% of the population in the USA to 85% of

the population in South Africa (Nwanegbo et al., 2004). Neutralizing antibodies bind to the

vector particles and prevent their entry into host cells, thereby reducing the efficiency of vector

uptake (Hodges et al., 2005). In chapter 2, we have carried extensive studies and identified that

intranasal delivery of empty HD-Ad can in fact induce a potent CD4+ and CD8+ T cell response.

Additionally, the innate immune response which is comprised of natural killer (NK) cells,

macrophages, neutrophils and complement along with many cytokines, is the first barrier

encountered by viral vectors. Adenoviruses can infect vast kinds of cells including macrophages,

which are also responsible for uptake of neutralized viral particles (Jooss and Chirmule, 2003). It

has been shown that administration of toxic compounds to animals results in release of

inflammatory cytokines, such as IL-1β and TNF-α and upon macrophage depletion, up to 95%

reduction in levels of inflammatory cytokines is observed (Salkowski et al., 1995). Application

of macrophage depletion in adenoviral gene therapy has demonstrated significant enhancement

of the transduction efficiency of adenoviral vectors and decrease in immune responses against

the vectors (Kuzmin et al., 1997). However, it needs to be noted that these studies were

conducted using FGAd and with significant improvement in vector design and development of

HD-Ad, the immune responses can be further reduced. Additionally, modification of the viral

capsid with polyethylene-glycol (PEG) has been shown to reduce the ability of the vector to

infect macrophages and dendritic cells, resulting in reduction of inflammatory responses (Croyle

et al., 2001). Similarly, AAV vectors can also potentiate innate immune response, albeit to a

much lower extent that adenoviral vectors (Zaiss et al., 2002).

26

Since adenoviral vector genomes are present episomally in the cell; there is a need for vector

readministration for sustained therapeutic gene expression. In contrast, though lentiviral and to a

lesser extent adeno-associated viral vectors have the ability to integrate their genomes, due to the

rapid rate of epithelial cell turnover, the transduced cells are lost. This again leads to the

requirement of vector readministration. Koehler et al. looked at the feasibility of adenoviral

readministration to the mouse lung using beta-galactosidase as the transgene. Studies indicate

that the immunological responses induced by first dose of HD-Ad were almost negligible

compared to that induced by FG-Ad. However, with re-administrations of HD-Ad, there was a

significant loss in beta-galactosidase expression and a dramatic increase in the levels of anti-

adenoviral antibody titer (Koehler et al., 2006). This therefore indicates that though one dose of a

vector may be non-inflammatory, subsequent rounds of re-administrations can significantly

boost the immune response and decrease vector efficiency. Similar studies using AAV vectors in

the rabbit have demonstrated that with a round of readministration there is significant loss in

transgene expression due to an increase in neutralizing antibody titer (Halbert et al., 1997). In

contrast to viral vectors, though non-viral vectors are not as immuno-stimulatory, their efficiency

in transfecting epithelial cells is too low to achieve a beneficial therapeutic effect.

There has been tremendous development in the field of adenoviral gene therapy over the last

decade, from the initial use of first generation adenoviral vectors to the recent development of

helper-dependent or so called gutless viral vectors. The gutless vectors have demonstrated

significant improvement over first generation vectors in terms of toxicity, efficiency and using

cell specific promoters such as cytokeratin 18, vectors can be targeted specifically to the airway

epithelium, which further reduces the inflammatory response. At the same time, due to reduced

toxicity, helper-dependent systems can also be used for delivery of anti-inflammatory siRNAs,

which would have been a problem with first generation vectors. However, the innate immune

responses elicited against the vector along with priming of the immune response with re-

administration still poses a challenge and work is being undertaken in many laboratories to

circumvent this problem and take HDAd vectors from the laboratory bench to clinical use in

humans. In chapter 4 and 5, we identify two new strategies that can be used to generate vector

specific tolerance and can likely suppress the immune response against HDAd vectors.

27

1.3 Potential for generating vector specific tolerance

Although with HD-Ad vectors the immune response is reduced, subsequent re-administrations

with high dose of vector can increase it to levels seen with FG vectors, thereby limiting

transgene expression. Therefore there is a need to induce tolerance within the host towards the

HD-Ad vectors without compromising the immunity towards other infections. Dendritic cells

play a critical role in the initiation and modulation of immune responses and determine the

balance between tolerance and immunity. Tolerance depends on a specialized subset of T cells

referred to as Regulatory T cells, which suppress the activity of autoreactive T cells against self

antigens and have also been shown to suppress innate immune responses. Dendritic cells are very

important in peripheral development of regulatory T cells and can possibly be used to induce

vector specific tolerance towards HD-Ad vectors.

The hallmark of the vertebrate immune system is adaptive immunity, which in contrast to innate

immunity, provides specific and efficient immunity. This is mediated by random generation of

receptors in developing lymphocytic clones in the thymus along with B cells through somatic

cell rearrangement (Cooper and Alder, 2006). The process of thymic selection results in a

population of autoreactive T cells which has been shown to be present in all individuals (Ohashi,

2003). Therefore there is a peripheral tolerance mechanism to keep these self-reactive pathogenic

T cells in check, which is mediated by a specialized subpopulation of T cells called regulatory T

cells (Treg) (Lohr et al., 2005). Regulatory T cells exert their tolerogenic properties in a contact

dependent fashion along with secretion of anti-inflammatory cytokines such as TGF-beta and IL-

10. Recent data suggest that contact-dependent suppression is perhaps the primary way by which

Tregs exert their effects. Surface expression of CTLA4 and TGFβ-1 has been shown to be

required for suppression. The cells exert their suppressive ability by interacting with the target T

cell and inhibiting expression of IL-2 R alpha chain (Annunziato et al., 2002). Blockade of

surface CTLA4 using Fab fragments has been to result in inhibition of target suppression by

Tregs in vitro (Sakaguchi, 2005). Another surface molecule recently implicated in suppression is

GITR. In vitro studies have indicated that cross-linking of GITR on Tregs along with TCR

stimulation, prevents Treg mediated suppression of reactive T cells (Shimizu et al., 2002).

Another property of regulatory T cells in maintaining tolerance is through induction of infectious

tolerance. It has been shown that Tregs can not only anergize CD4+ cells, but can also induce

28

them to secrete high levels of IL-10. These anergized IL-10 secreting CD4+ cells can then

further suppress T cell proliferation in an IL-10-dependent manner (Dieckmann et al., 2002).

Suppressive effects of Tregs have been shown to be dependent on IL-2 and in vitro studies

implicate the importance of IL-2 in mediating Treg suppressive function (Thornton et al., 2004).

The emerging model is that Tregs do not respond to initial activation of target T cells. However,

when the target T cells begin to proliferate, they secrete IL-2, which activates Tregs and results

in their expansion as well as suppressive function. The expanded Tregs then mediate contact

dependent antigen specific suppression and mediate infectious tolerance with acts as a positive

feedback. Interestingly, the suppressive functions of regulatory T cells are not only restricted to

the adaptive immune response (T and B cells) but can also affect the activation and functioning

of innate immune response cells (monocytes, macrophages and dendritic cells) (Taams and

Akbar, 2005). In vitro culture of monocytes and regulatory T cells derived from healthy donors

has been shown to suppress pro-inflammatory cytokine production and activation of

macrophages (Taams et al., 2005). At the same time, incubation of monocytes with Tregs

resulted in decreased secretion of IL-6 and TNF-α by macrophages following LPS stimulation.

Therefore, along with adaptive immunity, Tregs can also exert their effects on innate immunity.

Hence, induction of vector specific tolerance towards HD-Ad vectors can limit the immune

response against vectors and result in sustained gene expression in the lungs. Dendritic cells

(DCs) are professional antigen presenting cells and are essential mediators of immunity and

tolerance. The tolerogenic function of DCs can potentially be harnessed to generate tolerance

towards HD-Ad vectors.

1.3.1 Differentiation/Origin of dendritic cells

DCs were first discovered in 1973 as a novel cell population in mouse spleen which was clearly

distinct from macrophages (Steinman et al., 1975; Steinman and Cohn, 1973; Steinman and

Cohn, 1974; Steinman et al., 1979; Steinman et al., 1974). Similar to other leukocytes, DCs are

derived from hematopoietic stem cells. Although the pathways leading to generation of DCs are

not completely understood, recent findings have shed light on DC ontogeny (Fogg et al., 2006;

Liu and Nussenzweig, 2010; Liu et al., 2009). During haematopoiesis, hematopoietic stem cells

give rise to common myeloid (CMP) and common lymphoid progenitors (CLP), with monocytes,

macrophages, megakaryocytes, granulocytes and erythrocytes originating from CMP and T cells,

B cells and Natural Killer cells, originating from CLP (Akashi et al., 2000; Kondo et al., 1997).

29

Studies have documented that injection of purified CLP as well as CMP into irradiated mice

results in their differentiation into DCs (Traver et al., 2000). Recently, it has also been shown

that human multilymphoid progenitors can give rise to all lymphoid cell types along with

monocytes, macrophages and DCs (Doulatov et al., 2010). Although, DCs are traditionally

thought to be of myeloid origin, these studies indicate that even lymphoid progenitors can give

rise to DCs. However, since CMPs are several fold more abundant than CLPs, most of the DCs

are likely derived from the myeloid lineage (Liu and Nussenzweig, 2010).

DCs are comprised of a heterogeneous population of cells with DCs in various organs possessing

unique set of cell surface markers. Moreover, different routes of DC differentiation from

precursors, adds another layer of complexity to the heterogeneity of DC populations. It has

become quite evident that many distinct DC subtypes exist, each with a particular location and

specialized function in the immune system (Shortman and Liu, 2002).

Over the last decade, studies have established the potential of monocytes to differentiate into

DCs. Monocytes are circulating leukocytes that were classically known as precursors to

macrophages. Mouse monocytes have been classified into two subsets which are Ly6Chigh

monocytes, which are CX3CR1low, CCR2+, CD62L+, and CCR5−, and Ly6Clow monocytes, which

are CX3CR1high, CCR2−, CD62L−, and CCR5+ (Geissmann et al., 2003). Previous studies

indicated that it is particularly during inflammation that DCs arise from monocytes. However

recent findings challenge the notion and instead indicate that even during steady state conditions

DCs can arise from monocytes. Ly6Chigh monocytes have been shown to give rise to CD103-

DCs in the intestinal lamina propria under steady state conditions (Bogunovic et al., 2009; Varol

et al., 2009). Ly6Clow monocytes are thought to play a tolerogenic role and recent evidence

indicates that they can be differentiated into DCs in vitro upon culturing with GM-CSF and IL-4

(Peng et al., 2009). Moreover, in vivo studies indicate that injection of apoptotic thymocytes

results in their uptake by Ly6Clow monocytes, which subsequently migrate to the spleen and

differentiate into immunosuppressive DCs (Peng et al., 2009; Swirski et al., 2009). It is

important to note that adoptive transfer of purified monocytes under steady state conditions to

mice has failed to reconstitute the entire DC repertoire, whereas upon induction of inflammation

using complete Freund’s adjuvant (CFA) monocytes have been able to differentiate into certain

DC subsets (Naik et al., 2006). Therefore, monocytes cannot be regarded as the absolute

30

precursors to conventional DCs but likely differentiate into specialized DC subsets under specific

conditions.

The common precursor to macrophages, monocytes and DCs is the macrophage-DC progenitor

(MDP) which is classified as Lin−CX3CR1+CD11b−CD115+cKit+CD135+ (Fogg et al., 2006).

MDP is derived from CMP and only gives rise to monocytes, macrophages and DCs (Fogg et al.,

2006). MDP likely differentiate into a DC-restricted progenitor, called common DC progenitor

(CDP) which gives rise to DCs but not monocytes or macrophages (Liu et al., 2009). Although

both MDP and CDP reside exclusively in the bone marrow, a precursor DC population (Pre-

DCs) derived from CDP has been identified in bone marrow, blood, spleen and lymph nodes,

which comprise less than 0.05% of the leukocytes in respective tissues (Liu and Nussenzweig,

2010; Liu et al., 2009). These pre-DCs have been shown to migrate to lymphoid tissues through

the blood and undergo proliferation and differentiation into DCs (Liu et al., 2009). Therefore

CMPs give rise to MDPs, which give rise to CDPs, which subsequently give rise to Pre-DCs,

which function as immediate precursors to DCs. Figure 1-1 provides a schematic for

differentiation of DCs from precursors.

31

Figure 1-1: Differentiation of DCs from hematopoietic stem cells.

HSCs differentiate into CLPs and CMPs; CMPs subsequently differentiate into monocytes and

pre-DCs in the bone marrow. Subsequently, monocytes and pre-DCs enter the blood and migrate

to lymphoid organs and peripheral tissues, where they give rise to lymphoid DCs and tissue

resident DCs. In addition to CMPs, CLPs also have the potential to give rise to DCs, but their

contribution is not well understood.

32

Although the myeloid origin of DCs has been established, lymphoid origins of DCs from CLPs

cannot be ignored. Recently, studies have identified that TLR9 activation via CpG DNA on

CLPs promotes generation of DCs (Welner et al., 2008). It has also been shown that induction of

TLR4 signaling via LPS treatment of CLPs promotes DC differentiation (Nagai et al., 2006).

Flt3, a receptor tyrosine kinase, is involved in haematopoiesis and although it is not needed for

generation of CDPs in bone marrow, it plays a role in DC development in peripheral tissue along

with DC homeostasis and expansion (Waskow et al., 2008). It is particularly important for

development of plasmacytoid DCs along with CD8+ DCs and CD103+ DCs and functions by

signalling through the mammalian target of rapamycin (mTOR) pathway (Sathaliyawala et al.,

2010). Studies have indicated that adoptive transfer of CLPs followed by injection of Flt3L

drives DC differentiation from CLPs, which indicates that CLPs do have the potential to

differentiate into DCs, but still does not address whether under steady state conditions, CLPs act

as precursors to DC populations (Karsunky et al., 2003). Therefore, it is likely that under certain

conditions, certain sub-types of DCs can be derived from lymphoid progenitors as well. Table 1-

1 provides an overview of the various DC populations and their precursors.

33

Table 1-1: DC subtypes and their precursors

34

1.3.2 Dendritic cell subtypes

DCs were initially broadly classified into two groups, which include the steady state

conventional DCs and non-conventional DCs (Shortman and Naik, 2007). Steady state

conventional DCs were regarded to have a DC form and function, whereas non-conventional

DCs were DCs usually not seen in the steady state but those that arise in response to

inflammatory stimuli. Non-conventional DCs initially included plasmacytoid DCs and monocyte

derived DCs (Liu and Nussenzweig, 2010; Liu et al., 2009; Shortman and Liu, 2002; Shortman

and Naik, 2007). However, the identification of DC subsets that are monocyte-derived but arise

in the absence of inflammation under steady state conditions, further complicates DC

classification. Since DCs have multiple routes of development, DCs which arise from pre-DCs

with a classical DC function can be regarded as conventional DCs, whereas non-conventional

DCs can include monocyte derived DCs along with plasmacytoid DCs, which are distinct in their

function. Figure 1-2 provides a classification of various DC subtypes as conventional or non-

conventional DCs.

35

Figure 1-2: Classification of DC subsets as conventional and non-conventional DCs.

Conventional DCs are derived from common DC progenitor and pre-DC population and are

further divided into migratory and lymphoid DCs. Non-conventional DCs include plasmacytoid

DCs, which are derived from pre-DC population along with monocyte-derived DC subsets

,found in various peripheral organs.

36

1.3.2.1 Conventional steady-state DCs

Conventional DCs are comprised of DC subsets derived from CDP and pre-DCs and can be

further divided into migratory and lymphoid DCs. Migratory DCs have the ability to migrate

from peripheral tissues to lymphoid organs, whereas lymphoid DCs reside in the lymphoid

organs and lack migratory function. Migratory DC subsets include DC subsets found in the skin,

lung, intestinal tract, liver and kidneys. Lymphoid DCs are found in lymphoid organs such as

lymph nodes, spleen and thymus and have been further divided depending on the varied

expression of CD4 and CD8.

1.3.2.1.1 Migratory DCs

Migratory DCs are derived from CDPs and pre-DCs and reside in peripheral tissues such as skin,

lung, intestinal tract, liver and kidney. Migratory DCs are characterized by their unique ability to

acquire antigen and subsequently migrate to the draining lymph nodes, where interaction with T

cells takes place.

1.3.2.1.1.1 Skin

Skin DCs include Langerhans cells (LCs) and dermal DCs which participate in immune

responses against pathogens that gain access to the epidermal and the dermal layers. These DCs

have slow turnover with half-life of greater than 21 days for LCs and approximately 12 days for

dermal DCs. Dermal DCs includes dermal langerin+CD103+ DCs and langerin-CD103-DCs.

CD103 corresponds to integrin αE, which is expressed on a subset of effector CD8+ T cells and

CD4+ and CD8+ Tregs along with DC subsets (del Rio et al., 2010). Dermal langerin+ DCs can

be classified as conventional DCs, derived from bone marrow precursors with pre-DC precursor

as the likely precursor for this dermal DC subset (Ginhoux et al., 2007). However, the origins of

dermal langerin-CD103- DCs are not well understood.

Dermal langerin+CD103+ DCs have been discovered to be the most efficient subset in processing

viral antigens to MHC I pathway likely via cross-presentation (Bedoui et al., 2009b). Recently,

this particular subset has been shown to play a key role in initiating Th1 and Th17 responses

during subcutaneous sensitization and has also been shown to cross present antigens expressed

by keratinocytes (Henri et al., 2010; King et al., 2010). Studies have also reported that dermal

37

langerin+ DCs can play a role in mediating contact hypersensitivity with no evidence of tolerance

induction (Kaplan et al., 2005; Wang et al., 2008; Yoshiki et al., 2009). In addition to dermal

langerin+ DCs, langerin-dermal DCs have also been shown to potentiate CD8+ T cell responses,

with dermal langerin- DCs comprising the major population of migrating DCs following

intradermal injection of lentiviral vectors (Furmanov et al., 2010). However, in models of

leishmaniasis, it has been shown that dermal langerin- DCs play a role in priming CD4+ T cell

responses and dermal langerin+ DCs play a role in priming CD8+ T cell responses (Brewig et al.,

2009). Overall, these studies point towards a role of both dermal DCs and dermal langerin+ DCs

as initiators of immune response.

1.3.2.1.1.2 Lung

Lung DC consist of 3 DC sub-populations which include CD103+CD11chighCD11b- DCs in the

intraepithelial network, CD103-CD11chighCD11b+ DCs in the lamina propria of conducting

airways along with plasmacytoid DCs(Lambrecht and Hammad, 2009). Among the three subsets,

the CD103+CD11chighCD11b- DC population is regarded as a conventional migratory DC

population which is derived from the pre-DC precursor population (Ginhoux et al., 2009).

CD103+ DCs in the lung have the ability to migrate to the draining lymph nodes, produce IL-12

and are specialized in cross-presenting antigens to CD8+ T cells(del Rio et al., 2007; Sung et al.,

2006). Among the resident pulmonary DCs, it appears CD103+ DCs are the major initiators of

CD8+ T cell response to poxvirus infection (Beauchamp et al., 2010). However, it appears that

pulmonary CD103+ DCs may comprise a heterogeneous population with CD8α+ and CD8α- DCs,

with CD8α-CD103+ DCs being the major initiators of CD8+ cytotoxic T cell response

(Beauchamp et al., 2010; Pascual et al., 2008).

1.3.2.1.1.3 Intestinal tract

In the intestinal tract, DCs are found in Peyer’s patches (PP), lamina propria (LP) and mesenteric

lymph nodes (MLN). Intestinal DCs are classified primarily on varied expression of CD103 and

CX3CR1. The two main lamina propria subsets are CD103+CX3CR1-CD11b+ (CD103+ LP) DCs

and CD103-CX3CR1+CD11b+ DCs, with with the latter outnumbering CD103+ LP DCs by 3-4

fold. CD103+ LP DC is a conventional DC population derived from CDPs along with pre-DCs,

under control of Fltl3 and GM-CSFR ligands (Bogunovic et al., 2009; Varol et al., 2009).

38

CD103+ DCs are also found in Peyer’s patches and are also a conventional DC subset derived

from CDPs as well as pre-DCs under control of Fltl3 but not GM-CSF signalling (Bogunovic et

al., 2009). Studies have documented CD103+ DCs to serve both a regulatory and an

immunogenic role in the intestinal tract.

CD103+ LP DCs express high levels of CCR7 and are substantially depleted in the mesenteric

lymph nodes of CCR7-/- mice but not in the LP of CCR7-/- mice (Bogunovic et al., 2009; Jang

et al., 2006; Johansson-Lindbom et al., 2005). This indicates that CD103+ DCs constitute the

major population of migratory DCs. In various experimental models such as salmonella

infection, CD103+ DCs have been shown to be the DC subset responsible for antigen transport to

the MLN (Bogunovic et al., 2009). CD103+ LP DCs have been shown to recognize pathogenic

intestinal bacteria via TLR5 and secrete pro-inflammatory cytokines (Uematsu et al., 2006).

Additionally, CD103+ DCs have also been shown to induce expression of gut homing receptor

CCR9 on T cells and drive induction of gut homing CD8+ T cells (Annacker et al., 2005;

Johansson-Lindbom et al., 2005). TLR5-mediated bacterial recognition by CD103+ DCs

combined with their migratory ability makes CD103+ DCs the major DC subset responsible for

initiating adaptive immune response in the intestinal tract.

Contrary to their role in initiating immune responses, CD103+ DCs have been shown to serve a

regulatory function, for depletion of CD103+ DCs exacerbates colitis in mice (Varol et al., 2009).

CD103+ DCs have also been shown to drive Treg differentiation under steady-state conditions

through a mechanism dependent on TGF-β and retinoic acid (Coombes et al., 2007). DC specific

β-catenin knockout mice display reduced numbers of Tregs in the intestine with normal Treg

frequencies in the spleen, indicating a role of β-catenin signalling in intestinal DCs to promote

tolerance (Manicassamy et al., 2010). β-catenin signalling is essential in intestinal DCs to

promote expression of anti-inflammatory mediators, such as retinoic acid-metabolizing enzymes,

IL-10, TGF-β and suppression of pro-inflammatory cytokines, which cumulatively promotes

tolerance induction (Manicassamy et al., 2010). Furthermore, intestinal epithelial cells are also

known to secrete factors such as TSLP which can promote tolerance induction by CD103+ DCs

under steady-state conditions (Rimoldi et al., 2005). In the absence of inflammation, E-cadherin

from intestinal epithelial cells may interact with CD103 on CD103+ DCs to activate β-catenin

pathway promoting tolerance. However, during the presence of inflammatory stimuli CD103 and

39

E-cadherin interaction may be affected which could inactivate the β-catenin pathway and drive

CD103+ DCs to an inflammatory phenotype to prime immune responses.

1.3.2.1.1.4 Liver

Hepatic DCs were initially identified as CD11c+B220- DCs and CD11c+B220+ plasmacytoid

DCs (Jomantaite et al., 2004). CD11c+B220- hepatic DCs have been shown to play a role in

peripheral tolerance under steady-state conditions and can reduce ischemia/reperfusion induced

liver injury through secretion of IL-10 (Bamboat et al., 2010). However, CD11c+B220- DCs have

been further divided into CD103+CD11b- and CD103-CD11b+ subsets, with both expressing

aldehyde dehydrogenase, an enzyme which controls retinoic acid production and has been

associated with Treg induction(Guilliams et al., 2010). Fltl3 is highly expressed in the CD103+

subset compared to the CD103- subset and Fltl3 knockout mice are substantially depleted of liver

CD103+ DCs (Ginhoux et al., 2009). Flt3 is involved in generation of DCs from pre-DCs

pointing towards pre-DCs being the precursor to CD103+ DCs, which can therefore be classified

as conventional DCs (Ginhoux et al., 2009). In contrast to liver CD103+ DCs, CD103- hepatic

DCs can originate both from MDPs as well as monocytes, indicating that CD103- DCs may

include both pre-DC as well as monocytes derived populations (Liu and Nussenzweig, 2010). In

lieu of the recent findings, although CD103+CD11b- hepatic DCs can be classified as

conventional, CD103-CD11b+ hepatic DCs require further investigation to identify their origins.

1.3.2.1.1.5 Kidney

DC subsets in kidney include the recently-identified interstitial CX3CR1+CD11b+ DCs,

CX3CR1+CD11b- DCs and CD103+ DCs (Soos et al., 2006). The CD103+ subset is largely a

conventional DC subset, arising from pre-DC precursor population (Ginhoux et al., 2009).

CD103+ DCs have been shown to play an important role in mediating tolerance to kidney

allografts (Degauque et al., 2006). Renal DCs have been shown to secrete IL-10 and their

depletion has been associated with aggravated glomerular damage in a model of nephrotoxic

nephritis (Scholz et al., 2008). In contrast, renal DCs have recently been shown to play a role in

progression of kidney disease (Heymann et al., 2009). It is likely that CD103+ kidney DCs may

play a tolerogenic role, with immunological role attributed to other renal DC subsets. The

discrepancy could largely be because of studies relying on CD11c as a marker for renal DCs.

40

Further investigation into subsets of renal DCs along with expression of CD103 on

CX3CR1+CD11b+ and CX3CR1+CD11b- DCs is further required to shed light on role of various

renal DC subsets in tolerance versus immunity.

1.3.2.1.2 Lymphoid DCs

Lymphoid DC subsets are found in lymphoid organs such as thymus, spleen, lymph nodes and

include CD8+CD4+ DCs, CD8-CD4- DCs and CD8-CD4+ DCs. CD8-CD4- and CD8-CD4+ share a

higher degree of similarity than CD8+CD4+ DC and hence are collectively referred to as CD8-

DCs. CD8-CD4- DCs and CD8-CD4+ DC differ significantly in one functional aspect: whereas

CD8-CD4- DC, like CD8+ DC, can make IL-12 p70 when appropriately stimulated, CD4+ DC

appear unable to do so (Edwards et al., 2003). Under steady-state conditions although CD8+ DCs

can be observed in the T-cell areas of the lymphoid tissues, CD8- DCs are found within the

marginal zones of lymphoid tissues and only migrate to T cell areas upon stimulation (Sathe and

Shortman, 2008; Steinman et al., 1997). The recently-identified pre-DC precursors can give rise

to both CD8- as well as CD8+ DCs (Naik et al., 2006). However, pre-DC precursors are further

divided into two subsets based on CD24 expression and these include CD24high and CD24low pre-

DC precursors. CD24high pre-DC, which are DEC205-MHCII- further differentiate into CD8-

CD24+ DEC205+MHCII+ cells which differentiate into CD8+ DCs without dividing (Bedoui et

al., 2009a). In contrast to CD24high pre-DC, the CD24low population gives rise to CD8- DCs

(Sathe and Shortman, 2008).

Although CD8+ DCs have been well characterized, our understanding of CD8- DCs is fairly

limited. Among lymphoid DC subsets, CD8-CD4- DCs have been shown to secrete the highest

amounts of IFN-γ and act as potent initiators of cytotoxic T cell response upon intravenous

immunization with male antigen (Hochrein et al., 2001; McLellan et al., 2002). In contrast to

their role in driving T cell responses, other studies have reported that CD8-CD4- DCs are poor

stimulators of CD8+ T cells in vitro and instead prime regulatory Tr1 cells, which secrete IL-10

and suppress immune response (Zhang et al., 2005). The ability of CD8-CD4- DCs to induce

tolerance by driving Tr1 differentiation is mediated via TGF-β1 secretion. The ability of CD8-

CD4- DCs in driving tolerance versus immunity is dependent on TLR9 signaling. Stimulation of

CD4-CD8- DCs via TLR9 signaling has been shown to convert tolerogenic CD8-CD4- DCs into

immunogenic DCs which could potentiate a T cell response (Zhang et al., 2010). This raises the

41

likelihood that under steady state conditions, CD8-CD4- DCs may play a role in tolerance

induction but upon stimulation by TLR9 ligands, may revert to an immunogenic phenotype

which can prime a cytotoxic T cell response. CD8-CD4+ DCs have been shown to reduce

severity of EAE in murine models, firstly through secretion of IL-10 and secondly through

tolerizing effects on Th1 cells (Legge et al., 2002). Studies have shown that CD8-CD4+ DCs are

unable to drive IFN-γ production in T cells and this ability is independent of IL-10 production

(Legge et al., 2002).

CD8+ DCs are found in the spleen, lymph nodes and the thymus and their turnover ranges from

only 3 days in the spleen to up to 10 days in the thymus (Shortman and Heath, 2010). Moreover,

though CD8+ DCs in the spleen and the lymph nodes likely derive from pre-DCs, the DNA of

thymic CD8+ DCs contains IgH gene D–J arrangements as in T cells, raising the likelihood that

thymic CD8+ DCs may have lymphoid origins (Corcoran et al., 2003). CD8+ DCs play a key role

in viral immunity along with immune responses to intracellular pathogens, particularly in the

spleen and the lymph nodes (Shortman and Heath, 2010). CD8+ DCs are the most potent

producers of IFN-α among lymphoid DCs, which plays a role in increasing cytotoxicity of NK

and T cells and further contributes to viral immunity (Hochrein et al., 2001). Initially, CD8+ DCs

were thought to express the complete CD8 molecules found on T cells. However, later it became

apparent that T cells express CD8αβ heterodimer, whereas CD8+ DCs express CD8αα

homodimer. Although CD8 is used as a marker to classify DCs, no studies to date have been able

to show a functional and/or developmental significance of CD8 on DC surface. CD8+ DCs

express CD36 and Clec9A, which are receptors that give CD8+ DCs the ability to readily

phagocytose dead cells (Iyoda et al., 2002). However, the distinguishing feature of CD8+ DCs is

their ability to cross-present exogenous antigens through MHC class I pathway (den Haan et al.,

2000). Initially the ability of CD8+ DCs to phagocytose dead cells was thought to be responsible

for their ability to cross present. However, later studies identified that CD8+ DCs can even

uptake soluble antigens and cross-present via the MHC I pathway, indicating an intrinsic

difference in their antigen processing machinery compared to other DC subsets (Schnorrer et al.,

2006). This makes CD8+ DCs potent inducers of CD8+ T cell responses to exogenous antigens

and CD8- DCs potent inducers of CD4+ T cell responses (Dudziak et al., 2007). Stimulation of

CD8+ DCs with TLR ligands induces CD40 induction on CD8+ DCs, which drives production of

high levels of IL-12p70, which in combination with MHC class I antigen presentation, drives an

42

effector CD8+ T cell response (Schulz et al., 2000). Furthermore, CD8+ DCs have also been

shown to secrete IFN-λ in response to polyinosinic:polycytidilic acid (poly I:C), which is a

double stranded RNA known to function as an adjuvant (Lauterbach et al., 2010).

In addition to the induction of immune response, CD8+ DCs have also been implicated in

tolerance induction. DEC205 is an endocytic receptor highly expressed on CD8+ DCs, which

mediates the efficient processing and presentation of antigens on MHC class II products in vivo

(Shrimpton et al., 2009). Targeting of antigen to DEC205 by coupling with anti-DEC205

antibodies has been shown to induce CD8+ T cell tolerance (Bonifaz et al., 2002). This was

mainly attributed to deletional tolerance and also to induction of regulatory T cells (Kretschmer

et al., 2005). Furthermore, CD8+ DCs have been shown to induce peripheral self-tolerance by

capturing self antigens and presenting to both naive CD4+ and CD8+ T cells via the cross

presentation pathway (Belz et al., 2002). It is likely that exposure to an antigen in the presence of

TLR ligands which drive CD40 induction results in induction of an immune response, whereas in

the absence of any inflammatory stimuli, the antigen is cross-presented and tolerance is

achieved.

1.3.2.2 Non-conventional DCs

Non-conventional DCs include plasmacytoid DCs, which although are derived from CDP, are

unique in their ability to secrete high amounts of IFN α/type-I which distinguishes them from

conventional DCs, resulting in them being classified as non-conventional DCs. Additionally,

there are several DC subsets derived from monocytes and not CDPs, which results in them being

also classified as non-conventional DCs.

1.3.2.2.1 Plasmacytoid DCs

Plasmacytoid DCs (pDCs) comprise a distinct DC subset found both in lymphoid and non-

lymphoid organs and characterized by rapid production of Type I Interferons in response to viral

infections. pDC differentiation is dependent on Fltl3 and MDP and CDP give rise to pDCs (Liu

et al., 2009) (Auffray et al., 2009). pDCs express TLR7 and TLR9 which recognize viral RNA

and DNA and signal downstream via PI3 kinase, which regulates IRF7, a key transcriptional

regulator of IFN to drive type I IFN production (Guiducci et al., 2008). TLR signalling and IFN

production by pDCs is positively regulated by Ly49Q, which is expressed on all pDCs and binds

43

to MHC I (Tai et al., 2008). Recognition of viral particles by TLRs leading to IFN production by

pDCs does not require viral replication. Instead, TLR recognition of virus leads to IFN

production, which positively feedbacks via IFNR to drive further IFN production by pDCs

(Kumagai et al., 2009). In addition to serving as a source of IFN, pDCs have also been shown to

be critical for differentiation of activated B cells to plasma cells via secretion of Type I

interferons and IL-6 (Jego et al., 2003). Activated pDCs behave differently from conventional

DCs in antigen presentation following stimulation via TLR 9 ligands such as CpG DNA. In

models of influenza infection, conventional DCs undergo maturation and present antigens in

complex with MHC II, with a parallel downregulation of MHC II synthesis. In contrast, although

pDCs also undergo maturation and present antigens, MHC II synthesis is not downregulated,

giving pDCs the ability to continuously present endogenous viral antigens in the activated state

(Young et al., 2008).

pDCs have been associated with maintenance of peripheral tolerance as well as in induction of

autoimmune responses. Studies have shown that in models of experimental autoimmune

encephalitis, pDCs migrate to the lymph nodes and interact with myelin-specific CD4+ T cells

via MHC II and induce Treg expansion, which dampens the autoimmune response (Irla et al.,

2010). Moreover, pDCs have also been shown to upregulate ICOS-L expression upon

undergoing maturation, which drives induction of IL-10 producing Tregs (Ito et al., 2007). In

contrast to tolerance, pDCs have also been associated with autoimmune responses. Antimicrobial

peptide L77 (CAMP) has been shown to bind to self DNA, which is then recognized by TLR9 in

endocytic compartments of pDCs and drives IFN production leading to an autoimmune response

(Lande et al., 2007). Moreover, in the absence of conventional DCs, alloantigen expressing pDCs

have been shown to prime T cell responses in models of GVHD (Koyama et al., 2009). The

differential ability of pDCs to drive immunity versus tolerance is not completely understood and

may in fact be attributed to individual pDC subsets. The ability of pDCs to drive tolerance

induction could be attributed to a CCR9 expressing subset of pDCs, which have been shown to

be potent inducers of Tregs and suppressors of antigen specific immune responses (Hadeiba et

al., 2008).

1.3.2.2.2 pDCs in the liver and the lung

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Although pDCs have been implicated in hepatic immune responses, their role in the liver is not

completely understood. Under steady state conditions, hepatic DCs are poor stimulators of T cell

proliferation but have a pro-inflammatory cytokine profile (Kingham et al., 2007). Hepatic

pDCs, upon TLR9 ligation become potent inducers of NK cells, NKT cells and antigen specific

CD8+ T cells in vitro (Kingham et al., 2007). During hepatitis C infection, hepatitis C infected

cells trigger a robust IFN response in pDCs, which plays a role in inhibiting infection (Takahashi

et al., 2010). Furthermore, during chronic hepatitis C infection, depletion of pDCs is observed,

which may contribute to viral persistence, indicating that hepatic pDCs play a key role in

initiating immune responses against hepatitis C (Lai et al., 2007). However, hepatic pDCs may

also play a role in tolerance. Hepatic pDCs express high levels of NOD2, a pattern recognition

molecule, which binds to its ligand muramyl dipeptide (MuDP) (Castellaneta et al., 2009).

MuDP treatment of pDCs results in reduction of T cell stimulatory capacity along with an

increased expression of IRF-4, which is inhibitory to NFκB pathway (Castellaneta et al., 2009).

Altogether these studies indicate that although pDCs can drive hepatic inflammation, they also

may have a self-regulatory mechanism to control hepatic inflammation.

The role of pulmonary pDCs is highly controversial, with studies pointing towards their role in

immunity as well as tolerance. Adenoviral delivery to mice induces maturation of both pDC and

conventional DCs, with only conventional DCs migrating to the draining lymph nodes, raising

the likelihood that pulmonary pDCs may play an indirect role in potentiating immune responses

by modulating conventional DCs (Kushwah et al., 2008). In contrast to the role of pulmonary

pDCs in immune responses, studies have shown that pulmonary pDCs can suppress generation of

effector T cells in asthma models (de Heer et al., 2004). Moreover, in the model of dust mite

induced allergy, increased frequency of pDCs in the lung is associated with suppression of

airway inflammation (Lewkowich et al., 2008). Studies indicate that programmed death-1/

programmed death ligand 1 pathway is important for pDC-mediated suppression of airway

inflammation and is independent of the pDC maturation status (Kool et al., 2009). Furthermore,

pulmonary pDCs may also suppress conventional DC maturation as increased ratio of pDC to

conventional DCs in the lungs results is associated with reduced inflammation along with

reduction in Th2 driving chemokines (Kohl et al., 2006). pDCs in the lung may affect the

balance between Th1 and Th2 in the lung and shift towards Th1 responses, which may also result

in amelioration of Th2-associated inflammation observed in asthma models.

45

1.3.2.2 Monocyte derived DCs

Monocytes are derived from CMPs and MDPs and can give rise to DCs under inflammatory as

well as steady-state conditions. Monocyte derived DCs are found in peripheral tissues such as the

intestine, lung, skin and kidneys, and also have the ability to take up antigen and subsequently

migrate to draining lymph nodes.

1.3.2.2.1 Intestinal monocyte derived DCs

E-cadherin+ DCs and CD103-CX3CR1+ intestinal DCs are monocytic in origin. Ly6hi monocytes

home to site of inflammation in the intestine and give rise to E-cadherin+ DCs which have a

proinflammatory gene expression profile (Siddiqui et al., 2010). E-cadherin+ DCs secrete very

high amounts of IL-23 and IL-12 upon stimulation and also exacerbate T-cell colitis in mice,

pointing towards their role in intestinal inflammation (Siddiqui et al., 2010). Another monocyte-

derived DC population in the intestine is that of CD103-CX3CR1+ DCs, which are derived from

Ly6Chigh monocytes under the control of GM-CSF (Bogunovic et al., 2009; Varol et al., 2009).

CX3CR1+ DCs are capable of taking up bacteria via transepithelial dendrites from the intestinal

lumen, indicating that this DC subset may act as the first line of defense to mucosal pathogens

(Niess et al., 2005). CD103-CX3CR1+ DCs are longer lived than conventional intestinal DCs and

are poor stimulators of T cell proliferation with poor migration to the draining lymph nodes

(Schulz et al., 2009). During the course of an infection, such as Salmonella, depletion of

CX3CR1+ DCs only during the early course of infection attenuates Salmonella-induced colitis

(Hapfelmeier et al., 2008). Altogether, these findings point towards the role of CX3CR1+ DCs in

mediating the initial innate immune response against pathogens and not in initiating intestinal T

cell responses, thereby playing a role in gut homeostasis.

1.3.2.2.2 Pulmonary monocyte derived DCs

Pulmonary CD103-CD11chiCD11b+ DCs are monocyte-derived and Ly6lo monocytes have been

shown to give rise to this particular subset under steady state conditions (Jakubzick et al., 2008).

However, in response to a pulmonary fungal infection, largely Ly6hi monocytes have been shown

to give rise to this particular subset (Osterholzer et al., 2009). In response to allergen challenge,

monocytes are recruited from the blood to the lung where they undergo differentiation into

CD103-CD11chiCD11b+ DCs. These DCs are key producers of CCL17 and CCL22, which are

46

critical for infiltration of Th2 cells and eosinophils into the airways to drive allergen induced

inflammation (Medoff et al., 2009). In house dust-mite induced airway allergy model, CD11b+

DCs uptake antigen, increase in numbers, undergo maturation and secrete cytokines that play a

role in inducing a Th17 immune response (Lewkowich et al., 2008). Increased ratio of CD11b+

DCs compared to pDCs corresponds to an increased pulmonary immune response, indicating that

CD103-CD11chiCD11b+ DCs play a critical role in airway inflammation by regulating Th2 and

Th17 immune responses.

1.3.2.2.3 Monocyte derived DCs in the skin

LCs possess an unique cellular organelle called the Birbeck granule, with langerin (CD207), a C-

type lectin as its main component and plays a role in antigen uptake (Romani et al., 2010).

Among skin DCs, only epithelial resident LCs express E-cadherin, which mediates their

attachment to neighbouring keratinocytes as well as langerin (Jakob and Udey, 1998). In human

LCs, glycolipids from pathogens such as Mycobacterium leprae are endocytosed via langerin

and then loaded onto CD1a antigen without endosomal acidification, which can then

subsequently present antigens to T cells (Hunger et al., 2004). TGF-β1 is crucial to development

of Langerhans cells (LCs) as mice lacking TGF-β1 lack LCs (Borkowski et al., 1996).

Furthermore, it is the TGF-β1 derived from LCs which acts in an autocrine manner for

development/survival of LCs (Kaplan et al., 2007). The receptor for colony stimulating factor -1

(CSF-1) is also important for LC development as under steady state conditions CSF-1 lacking

mice lack LCs (Ginhoux et al., 2006). Ly6Chi monocytes have been shown to give rise to LCs

during inflammation, pointing towards a myeloid lineage of LCs (Ginhoux et al., 2006). Along

with the myeloid ancestry of LCs, studies have also shown that mouse lymphoid progenitors can

give rise to LCs upon transfer, pointing towards a lymphoid ontogeny of LCs (Anjuere et al.,

2000). In addition to the studies supporting myeloid/lymphoid ancestry of LCs, several studies

point to a distinct route of LC development, whereby yolk sac primitive macrophages migrate to

developing skin from E10 to E16.5 and populate the LC compartment (Ginhoux and Merad,

2010). Inspite of the controversy surrounding development of LCs, LCs can be regarded as non-

conventional DCs due to their origins from monocytes and/or macrophages.

Under steady state conditions, LCs do not express surface MHC II and most of MHC II

molecules are intracellular. LCs do express E-cadherin which mediates there adherence to

47

keratinocytes. However, upon encounter with an immunogen, MHC II is rapidly transported to

the cell surface and expression of E-cadherin is downregulated which allows LCs to detach from

keratinocytes and migrate to the skin-draining lymph nodes (Pierre et al., 1997). Activation

induced LCs also elongate their dendrites and penetrate the keratinocyte tight junctions to sample

external antigens below the stratum corneum of the skin (Kubo et al., 2009). LCs carry the

antigen to the draining lymph nodes, where CD8+ DCs take up the antigen and cross present to

CD4+ and CD8+ T cells (Allan et al., 2006; Carbone et al., 2004). However, studies have

challenged the immunogenic potential of LCs and have instead showed that depletion of LCs

leads to increase in ear swelling in contact hypersensitivity models, pointing towards a role of

LCs in mediating immunological tolerance (Kaplan et al., 2005). It has been shown that

disruption of E-cadherin under steady-state conditions leads to upregulation of costimulatory

molecules, MHC II and chemokine receptors on LCs, triggered via the β-catenin pathway (Jiang

et al., 2007). However, these LCs fail to release immunostimulatory cytokines and instead

promote tolerance induction via generation of regulatory T cells. Since β-catenin signalling in

DCs has been association with a tolerogenic phenotype; it is likely that under steady state

conditions, the β-catenin pathway promotes LC induced tolerance induction (Manicassamy et al.,

2010). However, signalling induced during the presence of inflammatory stimuli may override

the β-catenin pathway and promote induction of an immune response instead of tolerance.

1.3.2.2.4 Monocyte derived DCs in kidney

CX3CR1+ interstitial DCs which comprise CD11b+ and CD11b- DCs found in the kidney also

express F4/80, a macrophage marker, indicating that this DC subset could be monocyte derived

(Soos et al., 2006). CD11b+ (CX3CR1+CD11b+) DCs in the kidney express the monocytic

marker Gr1, indicating monocytic origin and they have been shown to increase in numbers in a

mouse model of glomerular disease and are critical for infiltration of T cells (Heymann et al.,

2009). In contrast to CD11b+ DCs, the function and the origins of CX3CR1+CD11b- DCs are not

well understood, although the presence of CX3CR1 and F4/80 indicates that CX3CR1+CD11b-

may also be monocytic in origin.

Although DCs comprise a heterogenous population, they are mostly derived from pre-DCs or

monocytes. However, lymphoid progenitors can also give rise to DCs and the contribution of

lymphoid progenitors to DC development is not completely understood. Further studies are

48

needed to identify whether lymphoid derived DCs arise normally during steady state conditions

or under the influence of inflammatory stimuli. Studies have tried to associate various DC

subsets with an ability to drive tolerance verus immunity. It is likely that the local environment

and the presence of extrinsic signals can drive a DC subset to behave as tolerogenic or

inflammatory. Recent studies have shed light on signals such as β-catenin pathway, but further

studies are needed to better understand the signals that can drive DC from behaving as

tolerogenic to immunostimulatory. Identification of such signals can then subsequently be

employed to develop new strategies to induce tolerance induction.

1.3.3 Regulation of DC apoptosis

DC apoptosis regulates the magnitude of immune responses by limiting antigen availability to T

cells and is regulated both by extrinsic and T cell mediated signals. Environments with

significant DC apoptosis are immunosuppressive, promote regulatory T cells (Tregs) generation,

and display functional impairment of remaining DCs, indicating that DC apoptosis may

contribute to tolerance. The importance of DC apoptosis is further highlighted by studies

identifying defects in DC apoptosis as triggers of autoimmune diseases (Bouillet et al., 1999;

Chen et al., 2007; Chen et al., 2006; Cohen and Eisenberg, 1992).

Though DCs play a critical role in the immune system, nevertheless, their life span is fairly

limited. BrdU labeling experiments indicate that under steady state conditions, splenic DCs have

a very rapid rate of turnover, with half life ranging between 1.5-2.9 days (Kamath et al., 2000).

Furthermore, peripheral DCs such as LCs upon arrival in the lymphoid organs have a life span

ranging from 1-9 days (Kamath et al., 2002). Upon activation, DCs regulate a certain set of genes

which allows for induction of maximal immune response, but a consequence of that is the

signaling that initiates cell death. LPS induced DC maturation initiates DC apoptosis through

CD14 mediated NFAT activation (Zanoni et al., 2009).Although, the specific genes that regulate

DC lifespan have not been completely identified, studies have shown involvement of multiple

pathways that regulate DC apoptosis (Figure 1-3).

49

Figure 1-3: Regulation of DC apoptosis.

DC apoptosis is regulated by T cell-dependent signals, such as TRANCE, CD154, and FASL and

T cell-independent signals, such as amyloid peptides, TRAIL, LPS, Type I IFN, Leptin, and

CCR7. Immature DCs are susceptible to apoptosis induced by FasL (CD95L), the susceptibility

to which is lost in mature DCs because of expression of cFLIP, which inhibits caspase 8. LPS

induces apoptosis of mature DCs through CD14-mediated NFAT activation, independent of

TLR4, which is normally required for LPS-induced DC maturation. MINOR has been identified

recently as an inducer of DC apoptosis; however, the signal for MINOR induced apoptosis is not

known.

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Two major pathways regulate apoptosis and these are the extrinsic and the intrinsic pathways

respectively. The extrinsic pathway involves binding of a death inducing ligand to a receptor,

which results in formation of death-inducing signaling complex (DISC) (Riedl and Salvesen,

2007). DISC in turn cleaves and activates activator caspases 8 or 10, which in turn cleave and

activate effector caspases. Effector caspases in turn cleave important molecules within the cell,

resulting in apoptosis. The TNF superfamily in particular induces DC apoptosis through the

extrinsic pathway. The intrinsic apoptosis pathway involves signals from within the cell which

induce permeabilization of mitochondrial outer membrane resulting in activation of caspases

which induce apoptosis (Hengartner, 2000; Hengartner, 2001). Anti-apoptotic members of the

Bcl-2 superfamily normally suppress mitochondrial membrane permeabilization, by inhibiting

pro-apoptotic proteins Bax and Bak, which are known to induce mitochondrial membrane

depolarization. Bcl-2 is normally highly expressed in immature DCs but as DCs undergo

maturation, its expression is downregulated, which makes mature DCs more susceptible to

apoptosis.

1.3.3.1 TNF superfamily and DC apoptosis

Members of the TNF superfamily are known to regulate DC apoptosis by interacting with

adaptor molecules that are upstream of caspases or by modulating expression of anti-apoptotic

proteins such as Bcl-2 and Bcl-2–related proteins (Smith et al., 1994a). Several members of the

TNF superfamily are known to regulate DC apoptosis/survival and these include CD154,

TRANCE, CD95L and TRAIL (Bjorck et al., 1997; Blum et al., 2006; Chino et al., 2009;

Cremer et al., 2002; Josien et al., 2000; Koppi et al., 1997).

CD40, a member of TNF receptor superfamily, is a co-stimulatory molecule expressed on DCs,

which is required for DC activation. Triggering of CD40 signaling via CD154 (CD40L, a

member of TNF superfamily), which is expressed on activated T cells, increases the persistence

and survival of DCs, highlighting the importance of CD40 signaling in regulation of DC

apoptosis. CD40-/- DCs have poor rates of persistence and survival upon adoptive transfer (Miga

et al., 2001). Triggering of CD40 via CD154 acts as a survival signal from T cells to increase

survival of mature DCs, which largely occurs through modulation of anti-apoptotic Bcl-2(De

Smedt et al., 1998) (Park et al., 2006). Upon interaction of T cells with DCs, there is formation

of the immunological synapse, which is known to induce anti-apoptotic signaling in DCs via

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induction of Akt1 activation, which largely occurs through induction of CD40 signaling (Riol-

Blanco et al., 2009).

TRANCE (TNF related activation induced cytokine) is a member of the TNF superfamily and is

expressed by osteoblasts and fibroblasts, activated T cells, subcapsular sinus macrophages,

metallophilic macrophages and certain myeloma (Cremer et al., 2002). It binds to its receptor

RANK (receptor activator of NFkB), which appears to be upregulated on DCs upon activation

(Wong et al., 1997). TRANCE is a survival factor only for DCs and also enhances DC mediated

T cell proliferation, with no pro-survival effects on other APCs. Interstitial DCs in human skin

co-express TRANCE and RANK, constitutive interaction of which is necessary for DC longevity

(Cremer et al., 2002). Upon binding of TRANCE to RANK on DCs, there is activation of NFkB

and JNK pathways which play a role in initiating anti-apoptotic signaling. NFkB is particularly

essential for TRANCE induced DC survival since survival of p50(-/-)cRel(-/-) DCs is not

affected by TRANCE (Ouaaz et al., 2002). Signaling via TRANCE also results in upregulation

of Bcl-xl which suppresses apoptosis. Preclinical studies demonstrate the utility of TRANCE in

enhancing the efficacy of DC immunotherapy by prolonging DC survival upon delivery to mice

(Josien et al., 2000).

Another receptor to which TRANCE binds is osteoprotegerin (OPG). OPG is a member of the

TNFR superfamily and functions as a decoy receptor to limit TRANCE-RANK interactions.

OPG-/- DCs have better survival than wild type DCs, which is likely due to absence of OPG in

limiting TRANCE-RANK interactions (Chino et al., 2009).

DC maturation also results in upregulation of CD95(Fas), a member of the TNF receptor

superfamily, which is a known inducer of pro-apoptotic signaling (Leverkus et al., 2000). In

contrast to induction of apoptotic cascade, ligation of CD95 on mature DC surface with its ligand

CD95L(FasL), a member of TNF superfamily, results in upregulation of c-FLIPL (Willems et

al., 2000). C-FLIPL is a homologue of caspase 8, and inhibits apoptosis by preventing

recruitment of caspase 8 to the death inducing signaling complex. Inhibition of c-FLIPL by

treatment of DCs with protein synthesis inhibitors results in loss of resistance to CD95 induced

apoptosis (Willems et al., 2000). In addition to FLIPL, studies indicate that anti-apoptotic

signaling induced via CD40-CD40L interaction in mature DCs can counteract CD95 induced

pro-apoptotic signaling (Bjorck et al., 1997; Koppi et al., 1997). In contrast to mature DCs,

52

immature DC express very low levels of c-FLIP and CD40, which likely mediates their

susceptibility to CD95 induced apoptosis.

TRAIL (TNF related apoptosis inducing ligand) is a transmembrane protein expressed on

effector cells and induces apoptosis upon binding to DR4 and DR5 receptors. It can also bind to

DcR1 and DcR2 which are truncated decoy receptors. TRAIL dependent apoptosis involves the

caspase dependent non-mitochondrial extrinsic pathway with caspase 8 and 10 acting as initiator

caspases. Leukemic plasmacytoid DCs express DR4 and DR5 and therefore are susceptible to

TRAIL induced apoptosis (Blum et al., 2006).

1.3.3.2 Nur77 family and DC apoptosis

Nur77 family is a group of zinc finger transcription factors belonging to the steroid nuclear

receptor superfamily (Cheng et al., 1997). The Nur77 family consists of 3 orphan nuclear

receptors which include Nurr77, Nurr1 and MINOR. Nurr77 and MINOR have been implicated

in regulation of T, B cell apoptosis and differentiation (Winoto and Littman, 2002). Although

Nurr77 is constitutively expressed in DCs, it does not regulate DC apoptosis. In contrast to

Nur77, MINOR is upregulated in DCs upon activation and forced expression of MINOR leads to

induction of apoptosis and its suppression results in inhibition of DC apoptosis (Chen et al.,

2009).

1.3.3.3 HLA-DR and DC apoptosis

Immature DCs are sensitive to HLA-DR induced apoptosis and this sensitivity further increases

as DCs undergo maturation (Bertho et al., 2000). The mechanism of how HLA-DR signaling

triggers DC apoptosis is not well understood, though it appears to be independent of caspase

activation or tyrosine kinase/phosphatases. The hallmarks of HLA-DR induced DC apoptosis is

the rapid depolarization of mitochondria, independent of the Bcl-2 pathway (Leverkus et al.,

2003). HLA-DR signaling triggers re-localization of protein kinase C δ (PKC δ) in the nucleus

and inhibition of PKC suppresses HLA-DR-induced DC apoptosis (Bertho et al., 2002).

1.3.3.4 Others

The chemokine receptor CCR7 is upregulated on the DC surface upon maturation and plays a

role in regulating DC survival. Stimulation of DCs with the CCR7 ligands CCL19 and CCL21

53

results in inhibition of apoptosis, via activation of the Akt1 pathway, which results in inhibition

of pro-apoptotic GSK3β and FOXO1, along with translocation of prosurvival NFkB to the

nucleus (Escribano et al., 2009; Sanchez-Sanchez et al., 2004). Leptin, an adipocyte-derived

hormone also promotes DC survival by inducing Akt1 and NFkB activation along with

upregulation of Bcl-2 and Bcl-xl gene expression (Mattioli et al., 2009; Mattioli et al., 2005).

Type I Inteferon has also been shown to regulate survival of splenic DC by promoting DC

apoptosis through regulation of apoptosis-related genes, in particular through downregulation of

Bcl-2 and Bcl-xl gene and protein expression (Mattei et al., 2009). Furthermore, even amyloid

peptides have been shown to induce DC apoptosis, which occurs via activation of acid

sphingomyelinase, resulting in production of ceramide, which results in autocatalysis of caspase

8 resulting in activation of apoptotic signaling (Xuan et al.).

1.3.4 Extrinsic triggers of DC apoptosis

1.3.4.1 DC apoptosis triggered by infections

Virus, parasitic as well as bacterial infections are known to induce DC apoptosis which results in

a bystander effect of induced immunosuppression. Measles virus (MV) infection is usually

associated with secondary infections due to immunosuppression. MV infects resting as well as

mature DCs and induces DC apoptosis via the Fas pathway and likely also through the TRAIL

pathway (Servet-Delprat et al., 2000; Zilliox et al., 2006). MV pulsed DC-T cell cultures show

enhanced rates of DC apoptosis along with an impairment of existing DCs to induce T cell

proliferation and undergo maturation (Fugier-Vivier et al., 1997). Other viruses such as foot and

mouth disease virus have also been shown to induce apoptosis of immature DCs(Jin et al., 2007).

Brugia malayi, a nematode and causative agent of lymphatic filariasis (elephantiasis) is known to

induce T cell hyporesponsiveness upon infection. Recently, it has been shown that it can induce

DC apoptosis through the extrinsic apoptosis pathway via induction of the TRAIL pathway,

raising the possibility that T cell hyporesponsiveness may be a consequence of DC apoptosis

(Semnani et al., 2008). It remains to be investigated whether infection-induced DC apoptosis has

a direct effect on viable DCs, resulting in their functional impairment.

Several bacterial infections are also known to trigger DC apoptosis. Gram positive bacteria such

as Streptococcus pneumoniae, certain group A streptococci strains and Listeria monocytogenes

54

induce apoptosis of DCs via bacterial encoded virulence factors, pneumolysin, streptolysin O and

listeriolysin respectively (Colino and Snapper, 2003; Cortes and Wessels, 2009; Guzman et al.,

1996). Similarly many gram negative bacteria also induce DC apoptosis and these include

Legionella pneumophila, Yersini enterocolitica and Pseudomonas aeruginosa. Legionella

pneumophila infection of DCs results in apoptotic death mediated by caspase-3 activation

(Nogueira et al., 2009). Yersinia enterocolitica induces DC apoptosis by injecting Yersinia outer

protein P, which induces degradation of cFLIPL, thereby rendering DCs sensitive to apoptosis

and also induces pro-caspase 8 cleavage to active form (Grobner et al., 2007). Treatment of DCs

with Pseudomonas aeruginosa results in induction of apoptosis through caspase dependent

pathway with no effect on apoptosis of other cell types such as epithelial cells (Worgall et al.,

2002). Studies need to be conducted to address whether there is some extent of impairment of

DC function upon bacterial infections which are known to induce DC apoptosis.

1.3.4.2 DC apoptosis during pathological conditions

DC apoptosis has been observed in several pathologies such as breast cancer, sepsis and trauma,

which parallels induction of immunosuppression. Patients with advanced breast cancer have

defective cellular immunity in mounting immune response to different pathogens. Studies have

identified that DCs isolated from breast cancer patients have a defect in inducing T cell

proliferation, and display an immature phenotype, with a defect in undergoing maturation in

response to an inflammatory stimuli (Gabrilovich et al., 1997; Satthaporn et al., 2004).

Furthermore, recently it has also been shown that breast cancer patients have proportionately

higher levels of apoptotic DCs than controls (Pinzon-Charry et al., 2006). Studies are needed to

identify whether DC apoptosis in breast cancer has a direct effect on the observed suppression of

DC function.

One of the hallmarks of the septic syndrome is the induced immunosuppression which is

responsible for sepsis-induced mortality. Sepsis induced immunosuppression is associated with

rapid and extensive caspase-3 mediated apoptosis of over 50% of DCs, observed both in humans

as well as mouse models of sepsis (Tinsley et al., 2003). Concomitant to DC apoptosis, there is

also an increase in the levels of circulating Tregs, both due to expansion and differentiation,

along with an increase in their suppressive function (Cavassani et al., 2010; Kessel et al., 2009;

Scumpia et al., 2006). However, the mechanism of observed increase in Tregs is not well

55

understood, though recently a study has shown that post-septic splenic DCs are good inducers of

Foxp3+ Tregs in vitro (Cavassani et al., 2010). Studies have shown that rescuing DC apoptosis

via generation of transgenic mice overexpressing anti-apoptotic protein Bcl-2 in DCs results in

resistance to endotoxin-induced sepsis and immunosuppression, clearly highlighting the role of

DC apoptosis in mediating sepsis pathology (Gautier et al., 2008). Cumulatively, these findings

indicate that DC apoptosis during sepsis has an effect on splenic DCs, which renders them

tolerogenic with a potential to prime Treg differentiation.

Severe acute trauma is often followed by complications involving organ dysfunction and

septicemia. This usually involves over activation of immune response followed by

immunosuppression. DCs isolated from multiple trauma patients, show a differential pattern of

expression of apoptosis related genes with reduced expression of anti-apoptotic genes and higher

expression of pro-apoptotic genes, identifying enhanced sensitivity of DC post traumatic stress to

undergo apoptosis (Maier et al., 2009). Studies have revealed that 3-5 days post trauma in

patients with multiple trauma, there is significant depletion of myeloid DCs, with approximately

a 3 fold decrease in the ratio of myeloid DCs to plasmacytoid DCs and a parallel increase in the

plasma levels of IL-10 (Henrich et al., 2009). However, there is also an impairment of monocyte

conversion to immature DC which likely further contributes to the state of immunosuppression

post trauma (De et al., 2003).

Studies have also shown that post trauma-hemorrhage, there is increased apoptosis of splenic DC

and surviving splenic DCs have a significant reduction in their antigen presentation ability as

well as MHC II expression (Kawasaki et al., 2006). Furthermore, the surviving splenic DC show

impaired IL-12 as well as IFN-γ production. Overall, the studies indicate that in

immunosuppressive environments induced in different pathologies with a high incidence of DC

apoptosis, there is a diminished ability of remaining viable DCs to undergo activation. This

raises the possibility that perhaps apoptotic DCs could be mediating the immunosuppressive

effect in certain pathologies by acting on viable DCs.

1.3.4.3 Glucocorticoid- induced DC apoptosis

Studies indicate that after liver, heart and kidney transplantation, there is a strong decline in the

circulating numbers of DCs. However, gradually as the dosage of immunosuppressive

56

glucocorticoids such as prednisolone is tapered, the DC numbers gradually recover. Furthermore,

patients and even healthy volunteers treated with corticosteroids show a decrease in the levels of

circulating plasmacytoid DCs. It has been shown that prednisolone as well as dexamethasone

treatment of plasmacytoid DCs can induce apoptosis via the extrinsic pathway (Abe and

Thomson, 2006; Boor et al., 2006).

1.3.4.4 Tumor- induced DC apoptosis

Studies indicate that tumor progression is usually associated with an impairment of DC function.

Tumors secrete factors which impair the function of both circulating as well as tumor-infiltrating

DCs, thereby suppressing the adaptive immune response and resulting in tumor progression.

Apoptotic DCs have been observed in many different tumors and studies have shown that co-

culture of DCs with media derived from tumor cell cultures, results in DC apoptosis, indicating

that tumors secrete factors that can induce DC apoptosis. Triggering of DC apoptosis by tumor

cells appears to occur via the extrinsic pathway as signaling through CD154(CD40L) is able to

suppress tumor-induced DC apoptosis (Esche et al., 1999).

Lymphatic drainage from the tumor occurs in the tumor draining lymph node, also called

sentinel lymph node (SLN). SLN has been observed in multiple cancers including non-small cell

lung cancer, breast cancer, melanoma, gastric and colon cancer (Cochran et al., 2006). Studies

have looked at the cellular composition of SLN and have identified a drastic reduction of DC

number in SLN compared to other lymph nodes (Botella-Estrada et al., 2005; Poindexter et al.,

2004). It appears that tumor-derived soluble factors play a major role in depletion of DCs. In

non-small cell lung cancer, there is increased apoptosis of DCs in the SLN, caused by the release

of TGF-β1 from the tumors (Ito et al., 2006). In cancers such as hepatocellular carcinoma

(HPCC), patients show a functional impairment of DCs. At the same time, high levels of α-

fetoprotein (AFP) are present in sera as well as in tumors, which has been shown to induce DC

apoptosis, indicating that DC apoptosis may somehow regulate functional impairment of

remaining viable DCs (Um et al., 2004). Human melanoma tumors have also been shown to

secrete factors such as gangliosides that induce DC apoptosis. Gangliosides are

glycosphingolipids which contain sialic acid and are found on plasma membrane. Normal

melanocytes only express GM3 ganglioside, whereas melanoma cells express other gangliosides

such as GD2 and GM3, and also secrete them into the circulation, which induce DC apoptosis

57

and results in immunosuppression (Peguet-Navarro et al., 2003). In addition, many melanoma

and fibrosarcoma tumor lines are known to secrete factors that mediate accumulation of C16 and

C24 ceramide in DCs, which induces DC apoptosis via activation of caspase 8 (Kanto et al.,

2001).

1.3.4.5 UV-induced DC apoptosis

Langerhans cells form an extensive network in the epidermis to capture antigens and

subsequently potentiate an adaptive immune response. UV radiation alters the function of

Langerhans cells and results in induction of tolerance towards antigen rather than induction of an

immune response (Elmets et al., 1983). Furthermore, Langerhans cells isolated from mice treated

with UV light fail to prime immune responses upon transfer to naïve mice but rather induce long

lasting tolerance which is dependent on generation of Tregs, indicating that Langerhans cells are

a major target of UV radiation and UV radiation converts them into tolerogenic DCs. However,

it is interesting to note that UV-induced immunosuppression is absent in transgenic mice which

overexpress anti-apoptotic molecules or lack pro-apoptotic molecules (Ullrich, 2005). UV

radiation has been shown to induce DC apoptosis both in vitro and in vivo. UVB radiation

induces apoptosis in DCs by activation of caspases 3, 8 and 9, loss of mitochondrial membrane

potential along with cellular and nuclear degradation. Immature DCs are highly susceptible to

UVB-induced radiation, whereas mature DCs seem to upregulate c-FLIP and Bcl-2 which makes

them less susceptible to UV-induced apoptosis (Nicolo et al., 2001). In response to UVB

exposure, there is DNA damage in Langerhans cells; however, the cells do retain their ability to

migrate to the draining lymph nodes, where they undergo apoptosis (Pradhan et al., 2008).

Prevention of apoptosis of Langerhans cells results in lack of UV-induced immunosuppression,

raising the likelihood that as UV-treated DCs undergo apoptosis in draining lymph nodes, they

somehow interact with the neighboring viable DCs which may have an immunosuppressive

effect. It is still not understood whether it is the UV-induced apoptotic DCs which directly

impact existing DCs to become tolerogenic DCs with a potential to induce Tregs.

UV radiation is also used as a therapeutic agent in extracorporeal photopheresis (ECP). During

ECP, peripheral blood mononuclear cells are treated with photoactive 8-methoxypsoralen (8-

MOP), and then exposed to UV light, which results in activation of 8-MOP, which then

covalently binds to DNA and induces apoptosis. ECP has been used for treatment of many

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autoimmune disorders which are normally not responsive to traditional immunosuppressive

regimens. The mechanism of ECP-induced immunosuppression was thought to be directly

dependent on induction of lymphocyte apoptosis. However ECP results in modulation of APC

function, which plays a key role in ECP induced immunosuppression. Recently, studies have

shown that ECP induces apoptosis of DCs (Holtick et al., 2008; Rao et al., 2008). It has been

reported that immature DCs have a higher susceptibility to ECP-induced apoptosis with upto

50% DCs undergoing apoptosis within 24 hours of treatment and over 90% undergoing apoptosis

within 72 hours of ECP treatment (Rao et al., 2008).Furthermore, studies have revealed that ECP

induces DC apoptosis and remaining viable DCs after ECP become tolerogenic with a severely

diminished ability to undergo maturation in response to LPS and also a defect in the ability to

induce T cell proliferation (Legitimo et al., 2007; Rao et al., 2008). At the same time, studies

have shown that there is a parallel increase in the levels of Tregs (Xia et al., 2009). These

findings point towards a role of UV-induced apoptotic DCs in mediating ECP-induced

immunosuppression by affecting the function of remaining viable DCs.

1.3.4.6 T cell-induced DC apoptosis

NKT cells are a distinct population of T cells conserved between mice and humans and are

stimulated by α-glycosylceramides and α-glucosylceramide in a CD1d- and TCR- dependent

manner. Among the human subpopulations of Vα24NKT cells, there are CD4– CD8– (double-

negative, DN) Vα24NKT and CD4+ Vα24NKT cells and both have been shown to have

cytotoxic activity against DCs from normal donors (Nicol et al., 2000; Nieda et al., 2001).

Activated CD4+ Vα24NKT cells upregulate CD40L and induce DC apoptosis via CD40/CD40L

signaling through the extrinsic apoptosis pathway. Studies indicate that NKT cells prevent

autoimmunity and induction of DC apoptosis could be one of the mechanisms used by these cells

to prevent hyperactivation of the immune system.

In tumor lymph nodes, Foxp3+ Tregs can directly interact with tumor antigen-bearing DCs and

induce their apoptosis through perforin-dependent pathways (Boissonnas et al., 2010). CD8+ T

cells can also play a role in inducing DC apoptosis. Antigen-loaded mDCs rapidly undergo

depletion in mice with an ongoing immune response and the rate of DC depletion is accelerated

in transgenic animals with increased levels of CD8+ T cells (Hermans et al., 2000).

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1.3.5 Apoptosis and cross priming by DCs

One of the unique abilities of DCs is to take up apoptotic cells and to present antigens derived

from apoptotic bodies to naïve CD8+ T cells. Though macrophages are more efficient than DCs

at taking up apoptotic cells, they fail to generate effective levels of peptide-MHC class I

complexes and instead degrade the antigen derived from apoptotic cells rather than cross-present

(Albert et al., 1998a). Studies have demonstrated that DCs can uptake apoptotic macrophages

infected by influenza or salmonella and induce a potent CTL response against the virus proteins,

indicating cross-presentation of antigens derived from apoptotic cells to naïve CD8+ T cells

(Albert et al., 1998b). CD8αα+TCRαβ+ T cells recognize self-peptides presented by MHC class

1b molecule Qa-1 and play a role in controlling experimental autoimmune encephalitis by killing

pathogenic CD4+ T cells and also in controlling colitis disease. The activation of this particular

T cell subset is mediated by DCs which capture apoptotic CD4+ T cells and cross present self-

peptides via MHC I to the CD8αα+TCRαβ+ T cell subset (Smith et al., 2009).

Lox-1 is a scavenger receptor expressed on certain DC subsets and has affinity for apoptotic cells

and heat shock proteins, and thereby is also involved in cross-presentation. IFN-α conditioned

DCs, which have been shown to resemble naturally occurring DCs upregulate Lox-1, which

plays a key role in the uptake of apoptotic lymphocytes and mediates cross presentation of

apoptotic cell derived antigen to CD8+ T cells (Parlato et al., 2009). Other scavenger receptors

expressed on immature DC surface are αvβ5 integrin and CD36, which make immature DC

highly efficient at taking up apoptotic cells and cross-presenting apoptotic cell-derived peptides

to CD8+ T cells compared to mature DCs (Albert et al., 1998a). Upon phagocytosis, apoptotic

cells can be found in vesicles containing MHC class I and class II molecules for up to 6-8 hours,

though thereafter antigens derived from apoptotic cells are not observed in the cytoplasm. It is

feasible that scavenger receptors trigger cytoskeletal rearrangement in DCs promoting apoptotic

cell uptake and then directing apoptotic cells to specialized pathways which mediate cross-

presentation. Recently, T-cell immunoglobulin mucins (TIMs) have been shown to mediate

cross-presentation upon apoptotic cell uptake by DCs (Kobayashi et al., 2007; Nakayama et al.,

2009).

DCs have limited lifespan and rapidly undergo apoptosis upon interaction with antigen-specific

T cells. However, inspite of rapid DC apoptosis, studies have shown that antigen presentation

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persists beyond 7 days, which is the peak time point for experimental CTL response. The

persistence of antigen presentation is mediated by exosomes which are released as DC undergo

apoptosis (Luketic et al., 2007). These are small vesicles which contain many surface markers

along with costimulatory molecules found on the DC surface and possess the ability to activate

naïve T cells. Cumulatively, studies conducted to date indicate that cross-presentation by DCs

plays an important role in the induction of immune responses. Since DCs have a limited lifespan,

apoptotic DCs must therefore interact with viable DCs which could likely lead to cross-

presentation. However, the effects of this interaction are not well-understood. Moreover, the role

of cross-presentation by DCs in induction and/or maintenance of immunological tolerance

warrant further investigation.

1.3.6 Defects in DC apoptosis trigger autoimmune diseases

Mice homozygous for gld and lpr alleles, which are genetic defects in FasL and Fas, go on to

develop autoimmune diseases, highlighting the importance of apoptosis in the maintenance of

immunological tolerance (Cohen and Eisenberg, 1992). However, when mice were generated,

where Fas was selectively ablated in T and/or B cells, it was insufficient for induction of

lymphoproliferative disease observed in lpr mice; also, induction of autoantibody titer was also

diminished compared to lpr mice (Hao et al., 2004). Furthermore, selective expression of

apoptosis inhibitory enzymes such as cytokine response modifier A (crmA), a serpin-like

protease inhibitor encoded by cowpox virus, in T cells failed to induce autoimmunity in mice

(Smith et al., 1996; Walsh et al., 1998). Overall these findings pointed towards apoptosis defects

in non-lymphoid cells resulting in induction of autoimmunity.

Chen et al generated transgenic mice expressing baculovirus p35, an inhibitor of apoptosis which

functions by inhibiting caspase 8, under control of the CD11c promoter (Chen et al., 2006).

These transgenic mice had DC-specific apoptosis defects and displayed hyperactivation of T and

B cells, along with autoimmune manifestations characterized by severe lymphocytic infiltration.

Furthermore, upon adoptive transfer, transgenic DCs were able to induce autoantibody

formation, indicating a direct role of DC apoptosis in the maintenance of peripheral tolerance.

Bim is a proapoptotic BH3-only protein in the Bcl-2 family and Bim-/- mice develop

autoimmunity (Bouillet et al., 1999). This is mainly due to defective negative selection of T and

61

B lymphocytes. However, studies indicate that Bim-/- DCs are defective in undergoing apoptosis

and are highly potent in induction of T cell activation and autoantibody formation, indicating that

defects in DC apoptosis could also be contributing towards the autoimmunity observed in Bim-/-

mice (Chen et al., 2007).

DCs have a high rate of turnover, and their apoptosis is regulated by many different pathways.

DC apoptosis is observed in several pathologies paralleled with induced immunosuppression.

These findings raise the likelihood that DC apoptosis could be one of the mechanisms of

inducing peripheral tolerance, which can be exploited for induction of tolerance in gene therapy,

transplantation along with other pathologies. In Chapter 5, we explore the tolerogenic potential

of apoptotic DCs and identify apoptotic DCs as tools which can be used for induction of antigen-

specific tolerance through generation of Tregs.

1.3.7 Tolerance induction by DCs

DCs are the key players in maintaining immune tolerance, for their ablation has been shown to

result in autoimmunity, highlighting the active role that DCs play under steady state conditions

in maintaining immune tolerance (Ohnmacht et al., 2009). In order to prevent autoimmune

reactions, self-reactive lymphocytes need to be deleted or their function needs to be suppressed.

The generation of normal lymphocyte repertoire which is largely self-tolerant depends on

positive and negative selection, which occurs in the thymus, process referred to as central

tolerance. However, some self-reactive lymphocytes that escape thymic deletion enter peripheral

tissues and the suppression of their function is needed to prevent autoimmune reactions, this is

referred to as peripheral tolerance. Central tolerance in the thymus is largely mediated by cortical

epithelial cells, medullary epithelial cells and thymic DCs and involves deletion of self reactive

thymocytes along with induction of naturally-occurring regulatory T cells (Tregs), which play a

key role in maintaining self tolerance and suppressing a variety of pathological immune

responses (Sakaguchi et al., 2006). In contrast to central tolerance, peripheral tolerance is

mediated by DCs through generation of Tregs and clonal deletion of self reactive T cells. Tregs

generated in the periphery are thought to be important in controlling immune response to non-

self antigens. Peripheral Tregs include IL-10-secreting Tr1 Tregs, adaptive foxp3+ Tregs, Th3

cells and double negative Tregs. DC-induced generation of these Treg subsets is largely

mediated by IL-27, TGF-β, IL-10, retinoic acid, indoleamine-2,3-dioxygenase and vitamin D.

62

The generation of these Tregs is either mediated by tissue resident specific DC subsets with a

specialized Treg-inducing function or by the action of mediators present in the local tissue

microenvironment, which act on DCs and drive them to behave as tolerogenic DCs and induce

Treg differentiation.

1.3.8 Type-1 regulatory T cells

Type-1 regulatory T cells (Tr1) cells are a group of Tregs characterized by production of IL-10.

Although, initially, studies pointed towards a central role of IL-10 in mediating Tr1 generation,

recent studies indicate that Tr1 generation could also be dependent on IL-27. Both IL-10 and IL-

27 are produced by DCs. Aryl hydrocarbon receptor (AhR), which is a ligand-activated

transcription factor belonging to the basic helix-loop-helix-PER-ARNT-SIM family, is induced

in Tr1 cells and during Tr1 differentiation, physically associates with c-maf, a transcription

factor belonging to the family of basic region leucine zipper domain transcription factors, and

activates IL-10 and IL-21 promoters (Apetoh et al., 2010; Marshall and Kerkvliet, 2010). Studies

to date have pointed towards a role of DC derived IL-27, IL-10, and TGF-β1 along with a role of

ICOSL signalling by DCs in induction of Tr1 cells. Figure 1-4 provides an overview of various

DC-derived signals which can drive Tr1 differentiation.

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Figure 1-4 DCs drive differentiation of Tr1 regulatory T cells.

DCs secrete IL-27, IL-10 and TGF-β1, which induce AhR and c-maf in T cells. AhR and c-maf

physically associate with each other and activate IL-10 and IL-21 promoters, driving Tr1

differentiation. IL-27 suppresses production of Th17-inducing cytokines, such as IL-1β, IL-6 and

IL-23 and drives Tr1 differentiation. IFN-γ suppresses Th17 inducing osteoprotegerin (OPG) and

drives IL-27 production, thereby promoting Tr1 differentiation. Furthermore, PD-1/PDL-1

signaling and bacterial peptides along with vasoactive intestinal peptide (VIP) drive IL-10

production, which also induces Tr1 differentiation. Moreover, ICOS/ICOSL signalling as well as

TGF-β production by DCs has also been implicated in driving Tr1 differentiation.

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1.3.8.1 IL-27 production by DCs drives Tr1 differentiation

DCs cultured with Foxp3+ Tregs secrete elevated levels of IL-10, IL-27 and TGF-β1, among

which TGF- β1 and IL-27 are important for driving differentiation of Tr1 cells (Awasthi et al.,

2007). IL-27 suppresses production of Th17 polarizing cytokines IL-1β, IL-6 and IL-23 from

DCs and acts on naive T cells to drive expression of the transcription factor c-maf, IL-21 and

ICOS, which collectively drive differentiation of Tr1 cells (Murugaiyan et al., 2009; Pot et al.,

2009). Furthermore, IL-27 production by DCs also drives IL-10 transcription in T cells by

activation of STAT1 and STAT3, which are recruited to the IL-10 promoter, further promoting

differentiation of Tr1 cells (Iyer et al., 2010). Recently, IFN-γ has also been identified to

promote DC-induced Tr1 generation. Studies have shown that IFN-γ inhibits Th17-inducing

osteoprotegerin (OPG) in DCs and instead promotes IL-27, which drives induction of Tr1 cells

(Murugaiyan et al., 2010; Pot et al., 2009). Hepatic DCs preferentially secrete IL-27 instead of

IL-12 upon LPS stimulation, indicating that hepatic DCs may also act as inducers of Tr1 cells

(Chen et al., 2009).

1.3.8.2 IL-10 production by DCs drives Tr1 differentiation

Differentiation of DCs from bone marrow in the presence of IL-10 leads to generation of a

CD11clowCD45RBhigh DC subset with a plasmacytoid morphology and an immature phenotype

(Wakkach et al., 2003). These DCs secrete high amounts of IL-10 upon stimulation and drive

differentiation of naive T cells into IL-10 secreting Tr1 cells. Similarly, skin-derived Langerhans

cells have also been shown to produce IL-10 which can also contribute towards generation of

Tr1 cells (Igyarto et al., 2009). Similar to murine Tr1 cells, DCs secreting IL-10 also drive

differentiation of human Tr1 cells, which is dependent on human leukocyte antigen (HLA)-G

and (Ig-like transcript) ILT4 molecules (Gregori et al., 2010). HLA-G, which is a non classical

MHC I molecule, plays a central role in maintaining fetal-maternal tolerance during pregnancy

and is expressed on IL-10-producing tolerogenic human DCs (Hunt et al., 2005). HLA-G binds

to inhibitory immunoglobulin-like transcript (ILT)-2 and ILT-4 receptors which has been shown

to suppress DC maturation (Allan et al., 2002; Ristich et al., 2005). IL-10 produced by DCs acts

in a positive feedback manner, sustaining HLA-G and ILT4 expression on DCs along with an

induction of HLA-G expression on T cells. ILT4 on tolerogenic DCs interacts with HLA-G on T

cells and HLA-G on DCs interacts with ILT2 on T cells, which drives differentiation of T cells

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into IL-10 producing Tr1 cells (Gregori et al., 2009). Monocyte derived human DCs cultured in

the presence of 1α,25-dihydroxyvitamin D3 (VD3) show a semi-mature phenotype characterized

by low levels of MHC Class II and costimulatory molecule expression along with production of

IL-10 and impairment of IL-12 production, driving Tr1 differentiation. Moreover, DCs cultured

with VD3 upregulate programmed death ligand -1 (PDL-1) upon activation, inhibition of which

suppresses Tr1 differentiation (Unger et al., 2009). PDL-1 signalling on DCs likely promotes IL-

10 production, since triggering PDL-1 on DCs by soluble PD-1 has been shown to suppress DC

maturation and promote IL-10 production (Kuipers et al., 2006). Altogether, these studies

indicate that VD3 treatment of DCs drives PDL-1 upregulation which acts as an inducer of IL-10

production by DCs, thereby driving Tr1 differentiation. However, it remains to be investigated

whether HLA-G and ILT4/ILT2 signalling is involved in VD3-driven induction of Tr1 cells.

Repetitive stimulation of peripheral CD4+ T cells by immature allogeneic DCs can also drive Tr1

generation (Levings et al., 2005). T cells cultured under stimulation by immature DCs selectively

upregulate cytotoxic T-lymphocyte antigen 4 (CTLA4) and lose their ability to produce IFN-γ,

IL-2, IL-4 and subsequently differentiate into Tr1 cells (Jonuleit et al., 2000). This is dependent

on the ability of immature DCs to secrete IL-10, which is severely diminished as DCs undergo

maturation. IL-10 production by pulmonary DCs also appears to be critical for induction of Tr1

induced tolerance. Pulmonary DCs in mice exposed to respiratory antigens undergo maturation

but secrete IL-10 which drives induction of Tr1 cells. Moreover, these Tr1 cells can suppress

airway responsiveness and their development is dependent on IL-10 production by DCs, since

adoptive transfer of DCs from IL-10-deficient mice fails to induce Tr1-mediated tolerance

(Akbari et al., 2001). In a model of food-induced anaphylaxis, tolerance induction is mediated by

gastrointestinal lamina propria DCs, whereby sugar modified antigens are targeted to C-type

lectin receptor SIGNR-1, resulting in preferential production of IL-10 but not IL-6 or IL-12 p70,

which ends up driving differentiation of naive T cells into Tr1 cells (Zhou et al., 2010b).

Several strategies that can induce DC production of IL-10 and prevent maturation have been

employed for generation of Tr1 cells. Neuropeptide vasoactive intestinal peptide is released

during inflammatory/autoimmune conditions, which induces generation of DCs with a capacity

to produce IL-10 and an inability to undergo complete maturation, which drives the generation of

Tr1 cells (Gonzalez-Rey et al., 2006). Exposure of DCs to cyclo-oxygenaase-2 overexpressing

gliomas results in IL-10 production by DCs which also drives Tr1 responses, which may be

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dependent on robust secretion of Prostaglandin E2 from the glioma (Akasaki et al., 2004).

Additionally, bacterial peptides such as filamentous hemagglutinin from Bortadella pertussis can

directly affect DCs by suppressing their ability to produce IL-12 and instead induce production

of IL-10, which drives generation of Tr1 cells (McGuirk et al., 2002). Individuals infected with

Plasmodium vivax have elevated levels of Tr1 cells and culture of mononuclear cells from

healthy individuals with Plasmodium vivax extracts can drive generation of Tr1 cells, indicating

that certain peptides produced from Plasmodium vivax may affect DC function which leads to

Tr1 generation (Jangpatarapongsa et al., 2008).

1.3.8.3 ICOSL signalling by DCs drives Tr1 differentiation

In addition to IL-10 production by pulmonary DCs, ICOS-ICOSL signalling is also critical for

Tr1 induction. Pulmonary DCs upregulate ICOSL upon maturation which is also critical for Tr1

induction since inhibition of ICOSL on pulmonary DCs suppresses Tr1 induction (Akbari et al.,

2002). Plasmacytoid DCs also upregulate ICOS ligand upon maturation, which has been shown

to drive differentiation of T cells into Tr1 cells (Ito et al., 2007).

1.3.8.4 TGF-β1 production by DCs drives Tr1 differentiation

In addition to IL-10 and IL-27, TGF-β may also have a role in priming Tr1 differentiation, since

addition of neutralizing antibodies against TGF-β to a coculture of CD4-CD8- splenic DCs and T

cells drastically reduces the production of IL-10 by T cells (Zhang et al., 2005). CD4-CD8-

splenic DC subsets secrete elevated levels of TGF-β upon stimulation with lipopolysaccharide

(LPS) and subsequently prime differentiation of IL-10-producing Tr1 cells (Zhang et al., 2005).

1.3.8.5 Other DC-driven signals which drive Tr1 differentiation

Along with TGF-β, CD40 signaling also plays a role in the ability of CD4-CD8- DCs to prime

Tr1 generation, since CD40 ligation abrogates the ability of CD4-CD8- DCs to prime Tr1 cells

(Zhang et al., 2009a). Furthermore, TLR9 stimulation via CpG DNA and TLR4 stimulation via

LPS can convert these Tr1-inducing DCs into Th1- and Th1/Th17-inducing DCs respectively

(Zhou et al., 2010b). Mature pDCs isolated from peripheral blood of rheumatoid arthritis patients

express high levels of indolamine-2,3-dioxygenase, which has also been implicated in promoting

differentiation of T cells into Tr1 phenotype (Kavousanaki et al., 2010).

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1.3.9 Foxp3+ Regulatory T cells

Foxp3+ regulatory T cells (Foxp3+ Tregs) are critical for maintaining immune tolerance and

preventing autoimmune reactions (Sakaguchi et al., 2008). Foxp3+ Tregs are identified by their

expression of CD25 (IL-2 receptor) and Foxp3, a transcription factor critical for Treg

differentiation (Zheng and Rudensky, 2007). The Foxp3+ Treg population can be divided into the

naturally-occurring Foxp3+ Treg population (nTreg), generally found in the thymus and the

adaptive Treg population (aTreg), which is derived in the peripheral tissues from CD4+CD25-

precursors upon activation in presence of TGF-β (Curotto de Lafaille and Lafaille, 2009). DCs

are critical for Treg induction and in this section I offer an insight in the recent advances in our

understanding of how DCs can drive nTreg and aTreg differentiation.

1.3.9.1 Naturally-occurring Foxp3+ Tregs

Natural Foxp3+ Tregs (nTregs) comprise a distinct lineage pathway determined at the double

positive (CD4+CD8+) stage of thymocyte development due in part to co-stimulatory signals

initiating Foxp3 expression. nTregs develop in the thymus during thymic development upon

recognition of self antigens. Although the role of DCs in thymic selection is documented, the

role of DCs in generation of nTregs is highly controversial (Wirnsberger et al., 2011). Several

studies have shown that DCs are dispensable for nTreg generation, whereby antigens specifically

expressed in thymic epithelial cells are sufficient to drive differentiation of nTregs

(Aschenbrenner et al., 2007). Conversely, there are studies identifying the contribution of thymic

DCs to generation of nTregs (Ziegler and Liu, 2006).

1.3.9.1.1 TSLP drives thymic DC mediated nTreg differentiation

Epithelial cells in Hassall’s corpuscles in the thymus produce thymic stromal lymphopoetin

(TSLP) which acts on thymic DCs by binding to TSLPR and IL-7R alpha complex and drives

induction of CD80 and CD86 (Ziegler and Liu, 2006). These DCs subsequently induce

differentiation of CD4+CD8-CD25- thymocytes into nTregs, which is dependent on IL-2 and

CD28 signaling (Watanabe et al., 2005). Therefore, TSLP-activated myeloid DCs in the thymus

are likely critical for positive selection of medium to high affinity self reactive thymocytes to

develop into nTregs (Ziegler and Liu, 2006). In addition to myeloid DCs, plasmacytoid DCs

(pDCs) residing in the thymus can also induce differentiation of CD69hiTCRhiCD4+CD8+

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thymocytes into nTregs and this is dependent on CD40L crosstalk (Martin-Gayo et al., 2010).

Thymic pDCs also express TSLP receptor along with IL-7 receptor complex and become

responsive to TSLP produced by thymic epithelial cells of Hassall’s corpuscles. TSLP-activated

pDCs can then drive differentiation of nTregs from CD4+CD8-CD25- thymocytes, which can be

inhibited by Th1- and Th2-polarizing chemokines IL-12 and IL-4 respectively (Hanabuchi et al.,

2010).

The role of TGF-β in driving nTreg differentiation is highly controversial. Previous studies have

shown normal nTreg numbers in TGFβ-R1-deficient mice (Curotto de Lafaille and Lafaille,

2009). However, recently it has been shown that mice with TGFβ-R1 deletion have nTreg

deficiency between postnatal day 3 and 5, but subsequently there is a surge in nTreg generation

due to increased responsiveness of the cells to IL-2 (Liu et al., 2008). It remains to be

investigated whether thymic DCs produce TGF-β1 which can affect nTreg generation.

1.3.9.2 Adaptive Foxp3+ Regulatory T cells

Adaptive Foxp3+ regulatory T cells (aTregs) are generated in the periphery by DCs and their

generation appears to be dependent on IDO, retinoic acid, Vitamin D and TGF-β. aTregs cells

play essential roles in immune tolerance and in the control of severe chronic allergic

inflammation (Curotto de Lafaille et al., 2008). Furthermore, since aTregs are induced in the

periphery, they also act as barriers in preventing the clearance of microorganisms and tumors,

whereby both are known to generate conditions that can drive aTreg differentiation (Curiel,

2008; Wohlfert and Belkaid, 2008). Figure 1-5 provides an overview of the DC-derived signals

which can drive aTreg differentiation.

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Figure 1-5: DCs drive differentiation of foxp3+ adaptive regulatory T cells. (aTregs).

DCs secrete TGF-β, which induces foxp3 in naive T cells, driving differentiation of naive T cells

into iTregs. Activation of AhR and TLR9 drives induction of IDO, which catalyzes tryptophan

metabolism. Tryptophan metabolites promote aTreg generation through induction of TGF-β

production and suppression of the Th17-inducing cytokine, IL-6. Furthermore, uptake of

apoptotic DCs by viable DCs along with exposure to haptens, glucocorticoids and UV radiation

also induces TGF-β production, which drives aTreg differentiation. Other signals such as

RANK/RANKL signalling by vitamin D-treated keratinocytes and treatment of DCs with

vasoactive intestinal peptide (VIP), hepatocyte growth factor (HGF) and prostaglandin-D2

(PGD2) also promote aTreg differentiation. Moreover, retinoic acid promotes aTreg

differentiation by suppressing cytokines which are inhibitory to aTreg differentiation and

targeting of antigen to DEC205 drives aTreg differentiation through a TGF-β dependent

mechanism.

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1.3.9.2.1 Indoleamine 2,3-dioxygenase in DCs drives aTreg differentiation

DC populations expressing indoleamine 2,3-dioxygenase (IDO) can play a critical role in

immune tolerance by promoting aTreg induction (Munn and Mellor, 2007). IDO catalyzes

tryptophan metabolism via the kynurenine pathway and therefore depletes the local environment

of tryptophan. Tryptophan catabolism likely plays an important role in suppressing T cell

proliferation by arresting T cells in the G1 phase of cell cycle (Brandacher et al., 2008).

However, recent studies have highlighted an important role of tryptophan catabolites in

mediating aTreg induction by exerting their effects directly on DCs. During HIV infection, IDO

activity is critical in regulating the Treg/Th17 balance with increased IDO levels produced by

DCs, associated with a chronic inflammatory state in progressive HIV disease due to a

breakdown of the mucosal barrier (Favre et al., 2010). Certain DC subsets such as pDCs, certain

splenic DCs populations such as CD19+ DCs and nasal DCs have been identified to upregulate

IDO upon stimulation (Mellor et al., 2005). Induction of IDO in DCs appears to be dependent on

aryl hydrocarbon receptor (AhR), for DCs lacking Ahr fail to upregulate IDO and prime T cell

response instead of tolerance induction (Nguyen et al., 2010). Activation of Ahr in mice by

2,3,7,8-Tetrachlorodibenzo-p-dioxin(TCDD), commonly referred to as dioxin, for 10 days results

in IDO induction both in the lungs and spleen along with upregulation of Foxp3 in the spleen,

which could be suppressed by inhibiting IDO (Vogel et al., 2008).

DCs residing in the nasal lymph nodes play an important role in inducing tolerance to inhaled

antigens. Studies have identified selective induction of IDO in non-plasmacytoid DCs in the

nasal lymph nodes, which is critical for inducing tolerance, for abrogation of IDO induction

results in elimination of tolerance induction towards the inhaled antigen (van der Marel et al.,

2007). In a murine model of experimental autoimmune encephalitis, IDO-deficient mice show

exacerbation of encephalitis, which can be inhibited by treatment with the tryptophan metabolite

3-hydroxyanthranilate (3-HAA), generated during IDO-mediated tryptophan catabolism.

Treatment with 3-HAA drives TGF-β production from DCs and also suppresses IL-6 production,

which ends up driving aTreg induction (Yan et al., 2010). Kynurenine, the first metabolite of

IDO-driven tryptophan metabolism activates the AhR on T cells, thereby driving aTreg

differentiation (Mezrich et al., 2010). TLR9 ligation drives induction of IDO in pDCs which

suppresses IL-6 production, suppressing conversion of naive T cells into Th17 cells and instead

promoting aTreg induction (Baban et al., 2009). IDO-driven kynurenine generation also appears

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to be important for pDC-mediated aTreg differentiation (Chen et al., 2008). Human monocyte

derived DCs, cultured under low tryptophan conditions, selectively upregulate inhibitory

receptors ILT3 and ILT4 and drive aTreg induction (Brenk et al., 2009).

1.3.9.2.2 TGF-β production by DCs drives aTreg differentiation

Skin DCs include Langerhans cells (LCs) and dermal DCs, with LCs being frequently associated

in maintenance of immune tolerance for acute depletion of LCs has been associated with an

enhancement of dermal immune responses (Bobr et al., 2010). Patients with Langerhans cell

histiocytosis, a condition with uncontrolled proliferation of LCs, show expansion of Foxp3+ Treg

populations, indicating a role of LCs in Foxp3+ Treg expansion (Senechal et al., 2007). Exposure

of UVR-exposed skin to haptens results in induction of aTregs, which is not observed upon LC

depletion, supporting the role of LCs in inducing aTregs and suppressing immune reaction in the

skin (Schwarz et al., 2010). Mice with LC-specific TGF-β depletion, show signs of autoimmune

disease in the skin and fail to develop LCs, indicating a role for LC-derived TGF-β in LC

development and also pointing towards a role of LC in maintenance of immune tolerance

(Kaplan et al., 2007). TGF-β production by LCs could in fact be a potential mechanism of how

LCs can prime differentiation of Tregs. In nickel allergy patients, administration of oral

glucocorticoids leads to TGF-β production by LCs, which expands aTregs and results in

reduction of clinical symptoms (Stary et al., 2010).

CD8α+ DCs were initially identified in the mouse spleen with a propensity to drive aTreg

induction (Stock et al., 2004). A unique CD8+ splenic DC subset, which expresses DEC205, a

type I transmembrane protein with multiple C-type lectin domains, has been identified in the

mouse spleen, and preferentially drives differentiation of aTregs (Yamazaki et al., 2008).

CD8+DEC205+ DCs can drive aTreg differentiation both in vitro and in vivo in the presence of

low dose of the antigen without addition of any exogenous TGF-β. However, aTreg induction

mediated by CD8+DEC205+ DCs is dependent on TGF-β for addition of TGF-β-neutralizing

antibody suppresses aTreg differentiation (Yamazaki et al., 2008). Furthermore, polyinosinic:

polycytidylic acid (poly I:C)-induced maturation of CD8+DEC205+ DCs reduces their ability to

drive aTregs, pointing towards their role in maintaining peripheral tolerance under steady state

conditions. Targeting of small amounts of antigen to DCs by using antigen fused to DEC205

antibody under conditions of suboptimal DC activation has been shown to drive aTreg induction

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(Kretschmer et al., 2005). Another CD8- splenic subset, which expresses DCIR-2, a type II

transmenbrane protein with a single external C-type lectin domain, drives aTreg differentiation

when exogenous TGF-β is added. However, in the absence of exogenous TGF-β, CD8- splenic

DCs are better at stimulating nTregs rather than driving aTreg differentiation (Yamazaki et al.,

2008).

Pulmonary DCs mediate inhalational tolerance which occurs during non-inflammatory settings

through CCR7-dependent migration of pulmonary DCs to the draining bronchial lymph node

(Hintzen et al., 2006). In the absence of inflammatory signal, pulmonary DCs acquire antigens

and subsequently acquire a semi-mature phenotype characterized by intermediate expression of

costimulatory molecules and high levels of MHC II expression, followed by subsequent

migration to bronchial lymph nodes, where tolerance is induced (Lambrecht and Hammad,

2003). Additionally, the local microenvironment in the lung also plays a role in driving aTregs.

Pulmonary stromal cells can produce cytokines, such as TGF-β, which can drive differentiation

of pulmonary DC into IL-10- and TGF-β-producing DCs which can subsequently drive aTreg

differentiation (Li et al., 2008).

Uptake of apoptotic DCs by viable DCs suppresses DC maturation and instead induces

production of TGF-β1 via the mTOR signalling pathway (Kushwah and Hu, 2010; Kushwah et

al., 2009; Kushwah et al., 2010). TGF-β1-producing DCs subsequently interact with naive T

cells and drive Foxp3 induction, thereby driving aTreg differentiation.

1.3.9.2.3 RANKL signalling on DCs drives aTreg differentiation

The local environment within skin could also contribute towards maintenance of tolerance by

DCs. The interplay between vitamin D and RANKL-RANK signalling in the skin plays a role in

inducing an environment which promotes DC-induced aTreg induction. The activated metabolite

of vitamin D (1,25-dihydroxyvitamin D3, VD3) exerts actions through its nuclear receptor, the

VD3 receptor (VDR) (Carlberg et al., 1993). VDR is expressed on immune cells such as DCs and

Vitamin D treatment of DC inhibits maturation along with their ability to prime alloreactive T

cell responses (Penna and Adorini, 2000). Keratinocytes in inflamed skin overexpress RANKL,

which through RANKL-RANK signalling modulates the function of DCs in the epidermis to

expand aTregs (Loser et al., 2006). Application of the topical vitamin D analog, calcipotriol,

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followed by transcutaneous immunization with a protein agent results in induction of aTregs,

primarily due to induction of RANKL on keratinocytes which likely modulates DC function to

drive aTreg differentiation (Ghoreishi et al., 2009). aTreg induction upon topical application of

vitamin D is absent in mice lacking vitamin D receptor, indicating vitamin D-driven RANKL as

the likely mechanism of how vitamin D can mediate skin tolerance by inducing aTregs

(Ghoreishi et al., 2009).

1.3.9.2.4 Retinoic acid-producing DCs drive aTreg differentiation

Oral intake of protein antigens leads to induction of oral tolerance, which is largely mediated

through generation of aTregs in the mesenteric lymph nodes. DCs in the mesenteric lymph nodes

express cyclooxygenase-2 (cox-2), which plays a role in aTreg induction. Suppression of cox-2

in mesenteric DCs results in induction of GATA-3 along with IL-4 in T cells and suppresses

aTreg induction (Broere et al., 2009). In addition to cox-2, mesenteric DCs express high levels of

B7-H1 and B7-DC, which are B7 family costimulatory molecules and are also essential for

mesenteric DC driven induction of aTregs (Fukaya et al., 2010). The most well-studied

mechanism of how oral tolerance induces aTregs is through retinoic acid. Retinoic acid (RA) is

an active metabolite of vitamin A which regulates multiple cellular processes such as cell death,

proliferation and differentiation through the retinoic acid receptors (RAR, including α, β and γ

subtypes) and the retinoic X receptors (RXR, also including α, β and γ subtypes). RA has been

shown to suppress inflammatory responses in animal models of multiple diseases, such as

inflammatory bowel disease and experimental autoimmune encephomyelitis. A unique

population of DCs, characterized by the expression of alpha E integrin, CD103 has been

identified in the gut-associated lymphoid tissue (GALT) as well as in the mesenteric lymph

nodes with a specialized function of inducing Tregs and maintaining immune tolerance

(Coombes et al., 2007; Sun et al., 2007). CD103+ DCs selectively drive aTreg differentiation

through RA- and TGF-β-dependent process, since addition of inhibitors of RA production or

TGF-β neutralizing antibody suppresses aTreg induction (Coombes et al., 2007; Sun et al.,

2007). CD103+ DC-derived RA also drives induction of α4β7 integrin and CCR9 on newly-

generated aTregs, which makes them home to GALT (Benson et al., 2007). Furthermore, RA

also sustains the stability and function of aTregs, even under inflammatory settings which further

results in tolerance induction (Zhou et al., 2010a). The mechanism of how RA can drive aTreg

induction is not completely understood. Initially, RA was thought to inhibit effects of IL-6

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signaling, which promoted aTreg induction rather than induction of Th17 in the presence of

TGF-β (Mucida et al., 2007). Later studies indicated that RA suppresses the generation of

CD44hi effector memory T cells, which secrete IL-4, IL-21 and IFN-γ, and suppress TGF-β-

mediated aTreg differentiation (Hill et al., 2008). However, RA can also interfere with the effects

of inhibitory cytokines on aTreg differentiation and can promote aTreg induction in the absence

of inhibitory cytokines, which is dependent on RAR-α (Nolting et al., 2009). Immune deficient

mice lacking CD103 fail to suppress T cell-mediated colitis upon transfer of Tregs, pointing

towards a role of CD103+ DCs in maintaining intestinal immune homoestasis (Annacker et al.,

2005). Curcumin treatment of bone marrow-derived DCs drives expression of Aldh1a, an

enzyme involved in RA production, which makes DCs behave similarly to mucosal CD103+ DCs

and drive RA mediated induction of aTregs (Cong et al., 2009). The local intestinal environment

also modulates DC function. Intestinal epithelial cells produce TGF-β, and retinoic acid which

drive a tolerogenic DC phenotype, which can then subsequently drive aTreg differentiation (Iliev

et al., 2009a; Iliev et al., 2009b). Additionally, lamina propria macrophages also suppress

intestinal DC-induced Th17 responses, which could inadvertently prime aTreg differentiation

(Denning et al., 2007).

A CD103-CD11b+ DC subset has been identified in the skin and expresses 3 aldehyde

dehydrogenases, which catalyze conversion of retinal to RA, giving this subset the unique

property of priming aTreg differentiation (Guilliams et al., 2010). Furthermore, treatment of

mice with AhR ligand 2-(1'H-indole-3'-carbonyl)-thiazole-4-carboxylic acid methyl ester (ITE)

has been shown to result in generation of tolerogenic DCs which promote aTreg differentiation

through a RA-dependent mechanism (Quintana et al., 2010).

1.3.9.2.5 Other DC-derived signals which drive aTreg differentiation

Mast cell-derived Prostaglandin-D2 (PGD2) is a mediator of inflammation which promotes

infiltration of eosinophils and Th2 cells into the lung during asthma. PGD2 can act through DP1

or DP2 receptor. Studies have shown that treatment of asthmatic mice with a DP1 agonist can in

fact suppress features of asthma by acting on pulmonary DCs and inducing cAMP-dependent

protein kinase A activation, which suppresses the ability of DCs to drive Th2 responses and

instead promotes induction of aTregs (Hammad et al., 2007).

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Additionally, treatment of DCs with immunosuppressive peptides has also been shown to drive

aTreg differentiation. DCs treated with immunosuppressive neuropeptide, vasoactive intestinal

peptide (VIP) drive aTreg differentiation, which is likely mediated by the ability of VIP to

suppress DC maturation and proinflammatory cytokine production (Chorny et al., 2006a; Chorny

et al., 2006b). Moreover, upon treatment with hepatocyte growth factor (HGF), DCs also drive

aTreg differentiation, which is abrogated if DCs are treated with antibodies against HGF receptor

(Benkhoucha et al., 2010). However, the mechanisms of how HGF receptor signalling in DC

drives aTreg differentiation is not understood.

1.3.10 Th3 cells

Th3 cells were first identified as a novel population of T cells induced upon induction of

peripheral tolerance upon oral delivery of myelin basic protein, which suppressed experimental

autoimmune encephalitis in mice (Chen et al., 1994). These Th3 cells are class II-restricted T

cells with identical αβ TCR as Th1 and Th2 cells. Moreover, they are characterized by

production of high levels of TGF-β, along with low amounts of IL-4 and IL-10 with no

production of IFN-γ or IL-2. The ability of these cells to suppress EAE is largely TGF-β-

dependent. Secretion of TGF-β by Th3 cells drives induction of Foxp3 in activated T cells,

driving them towards an aTreg phenotype (Carrier et al., 2007a). Furthermore, Foxp3 can also be

induced in Th3 cells for studies have shown that transient induction of TGF-β1 in T cells during

activation in absence of IL-2 can drive generation of Foxp3+ Th3 cells which comprise a distinct

Treg phenotype, which is CD25- and can control a hyperproliferative T cell response (Carrier et

al., 2007b).

1.3.11 Double negative regulatory T cells

TCR+CD3+CD4-CD8- double negative (DN) regulatory T cells inhibit immune responses by

Fas/FasL destruction of effector cells in an antigen-specific fashion (Zhang et al., 2000).

Although the mechanism(s) of how DCs can prime differentiation of DN Tregs is not

understood, syngeneic DCs have been successfully utilized for expansion of antigen-specific DN

Tregs (Thomson et al., 2007).

DCs play a critical role in the induction of tolerance. One of the active mechanisms whereby

DCs induce/maintain tolerance is through induction of Tregs. Over the last decade, significant

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progress has been made in understanding the DC-specific signals that can drive induction of

Tregs. These findings can potentially be employed to generate tolerogenic DCs which can be

used to generate vector-specific tolerance for airway gene therapy. In chapter 4 and chapter 5, I

describe two novel strategies that can be used for generation of vector-specific tolerance by using

tolerogenic DCs.

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Hypothesis and Objectives

Introduction

Adenoviral vectors have been investigated extensively for pulmonary gene therapy due to their

ability to efficiently transduce a wide variety of cell types and also due to their large carrying

capacity (Cao et al., 2004). Initially, FG-Ad vectors were used for gene therapy but their success

was hampered by the host immune response against viral antigens which led to destruction of

transduced cells (Yang et al., 1996a). Gradually, there was development of HD-Ad vectors,

which had a much improved safety profile, compared to FG-Ad vectors and could mediate long-

term gene expression in the absence of the chronic toxicity observed with FG-Ad vectors (Ng et

al., 2001; Ng et al., 1999; Parks, 2000). Moreover, HD-Ad vectors have also been shown to

mediate unprecedented levels of transgene expression in the lungs of large animals such as

rabbits and baboons (Koehler et al., 2005). However, because Ad vectors do not integrate into

the host cell and airway epithelial cells have a rapid rate of turnover, vector readministrations are

needed to maintain sustained and long-term expression of the therapeutic gene. It turns out that

readministration of HD-Ad vectors is associated with induction of an adaptive immune response

against these vectors. Our group has previously shown that following HD-Ad vector

readministration to the mouse lung, there is a significant decrease in the levels of transgene

expression with a concomitant increase in the antibody levels against these vectors (Koehler et

al., 2006). This indicates that even HD-Ad vectors can potentiate an adaptive immune response.

Therefore, it is extremely important to understand the pulmonary adaptive immune response

against HD-Ad vectors. In chapter 2, I have investigated the pulmonary immune response

against HD-Ad vectors by performing intranasal delivery of empty HD-Ad vectors (not encoding

for any transgene) and the findings indicate that delivery of a high dose of HD-Ad vectors is

associated with infiltration of both CD4+ and CD8+ T cells in the lungs along with T cell

proliferation, which peaks around day 6-7 post vector delivery. Although delivery of a low dose

also potentiates T cell proliferation, no T cell infiltration is observed in the BALF and also the

proportions of CD8+ T cells are reduced compared to those observed with a high dose of HD-Ad

vector delivery. Furthermore, although both conventional and plasmacytoid DCs undergo

maturation upon HD-Ad vector delivery to the lungs, only the conventional DCs migrate to the

draining lymph nodes and likely potentiate a CD4+ T cell response. Moreover, CD8α DCs in the

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draining lymph nodes undergo maturation and this DC subset is associated with cross-

presentation. Therefore, it is likely that in the absence of viral transcription, CD8α DCs cross-

present HD-Ad derived epitopes via MHC class I pathway to drive a cytotoxic T cell response.

Since the findings indicate reduced pulmonary immune responses with a low dose of HD-Ad

vectors, it is perhaps important to develop novel strategies which can be used to enhance the

efficiency of HD-Ad mediated gene delivery to the airways to reduce the viral dose and thereby

reduce the immune response against HD-Ad vectors. In chapter 3, I have identified a novel

strategy, whereby delivery of a mucolytic agent, Nacystelyn (NAL) to the airways of mice

followed by vector delivery in DEAE-Dextran can mediate up to 60-70 fold enhancement

compared to delivery of Ad vectors in saline. Moreover, airway histology indicates that this

strategy can also suppress pulmonary inflammation induced by Ad vectors. Therefore, this

strategy can likely be used to further reduce the vector dose which can further limit the immune

response against HD-Ad vectors. The drawback of the strategy is that in cases where several

rounds of HD-Ad vector readministration to the lung are needed, eventually the immune

response will likely be amplified which will substantially limit long-term transgene expression.

In order to mediate long term stable gene expression in the lungs following HD-Ad vector

readministration, perhaps it is most desirable to abrogate immune responses against HD-Ad

vectors in an antigen-specific manner. Therefore in chapter 4, I devise a novel strategy to induce

antigen specific tolerance towards HD-Ad vectors. Delivery of DCs derived in presence of high

concentrations of IL-10 upon being pulsed with HD-Ad vectors to mice mediates long term

tolerance induction towards HD-Ad vectors, whereby pulmonary DC maturation, T cell response

along with antibody response to HD-Ad vectors is suppressed even after three rounds of

pulmonary HD-Ad vector readministration and there is concomitant induction of Tr1 regulatory

T cells. Moreover, sustained gene expression is also observed in the lungs of mice immunized

with HD-Ad pulsed, IL-10 derived DCs following three rounds of HD-Ad administrations to the

lung, highlighting the efficacy of this strategy in mediating long-lasting tolerance. Furthermore,

induction of tolerance is specific towards HD-Ad vectors since delivery of ovalbumin-pulsed IL-

10 derived DCs fails to mediate long lasting gene expression following HD-Ad vector

readministration.

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In addition to the use of IL-10- derived DCs to drive HD-Ad specific tolerance; I have developed

another novel strategy based on the use of apoptotic DCs to induce antigen specific tolerance,

described in chapter 5. Apoptotic DCs are rapidly taken by viable DCs, which subsequently

suppress maturation of viable DCs and instead drives secretion of TGF-β1, which mediates

differentiation of naïve T cells into Foxp3+ Tregs. The ability to mediate Treg induction is

specific to apoptotic DCs since apoptotic splenocytes cannot mediate the same effects.

Furthermore, delivery of apoptotic DCs to mice followed by delivery of antigen in CFA leads to

uptake of apoptotic DCs by viable DCs in the draining lymph nodes and spleens, which

suppresses maturation of viable DCs and instead drives induction of antigen-specific tolerance

through induction of antigen-specific Foxp3+ Tregs. Further work is needed to employ this

strategy for induction of antigen-specific tolerance towards HD-Ad vectors, to allow for

sustained transgene expression in the lungs following vector readministration.

Altogether, this thesis describes strategies to enhance adenoviral-mediated gene delivery to the

airways. The goal of the research described in this thesis is to identify novel formulations that

can enhance adenoviral mediated gene transfer to the airways and to develop novel strategies that

can be used to induce antigen-specific tolerance towards adenoviral vectors, which can allow for

repeated vector administration to sustain long term gene expression.

Hypothesis

1. An adaptive immune response is initiated upon intranasal delivery of HD-Ad vectors.

2. Efficiency of Ad mediated pulmonary gene transfer can be increased by mucolytic

agents.

3. Immature DCs can induce Ad specific immunological tolerance.

4. Apoptotic DCs promote immunological tolerance.

Objectives

1. To study the pulmonary immune response initiated upon intranasal delivery of HD-Ad

vectors and to identifying underlying mechanisms.

2. To increase the efficiency of Ad mediated gene transfer to the airways.

3. To study whether DCs derived in the presence of IL-10 can generate Tregs towards HD-

Ad particles.

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4. To study the effects of apoptotic DCs on viable DCs and the adaptive immune response

and to identify whether apoptotic DCs can be used to induce antigen-specific tolerance.

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Chapter 2

Characterization of airway adaptive immune responses to helper

dependent adenoviral vectors

The contents of this chapter have been published in the Journal of Immunology: Rahul

Kushwah, Huibi Cao and Jim Hu. Characterization of pulmonary T cell response to helper-

dependent adenoviral vectors following intranasal delivery. Journal of Immunology. 2008 Mar

15;180(6):4098-108.

Acknowledgements: I would like to acknowledge Huibi Cao for generation of adenoviral

vectors used in this study.

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2.1 Abstract

In spite of extensive research in the field of gene therapy, host immune responses continue to be

the major barrier in translating basic research to clinical practice. Helper-dependent adenoviral

(HD-Ad) vectors show great potential for pulmonary gene therapy, but the knowledge of

pulmonary immune responses toward these vectors is very limited.

Objective of Study: In this study I explored the pulmonary adaptive immune response upon

delivery of HD-Ad vectors, lacking a transgene.

Summary of Results: In this study, I show that HD-Ad vectors are potent stimulators of dendritic

cell (DC) maturation, thus leading to stimulation of T cell proliferation with approximately 6%

of naive CD4(+) T cells from pulmonary mediastinal lymph nodes responding to HD-Ad-treated

DCs. Furthermore, through in vivo pulmonary studies in mice, I show that HD-Ad vectors can

prime CD4(+) and CD8(+) T cell responses in the lung at high and substantially low doses. This

indicates cross-presentation of HD-Ad-derived epitopes by DCs to prime CD8(+) T cell

responses. To assess the basis of the pulmonary T cell response against HD-Ad vectors, I

examined the response of conventional DCs (cDCs) and plasmacytoid DCs (pDCs) in the lung.

In response to HD-Ad delivery, there is induction of maturation in both cDC and pDC subsets,

but it is the cDCs, not pDCs, that migrate rapidly to draining lymph nodes within the first 2 days

after vector delivery to prime adaptive immune response against these vectors.

Conclusion: These findings demonstrate that empty HD-Ad particles are potent inducers of

pulmonary adaptive immune response and therefore strategies are needed to develop tolerance

against these vectors to improve and sustain HD-Ad mediated gene transfer.

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2.2 Introduction

Inspite of the extensive research in the field of gene therapy, host immune responses continue to

be the major barrier in translating basic research to clinical practice. Helper-dependent

adenoviral (HD-Ad) vectors show great potential for pulmonary gene therapy, but the knowledge

of pulmonary immune responses towards these vectors is very limited. The understanding of the

pulmonary adaptive immune response to HD-Ad can hold the key towards development of

strategies to prevent immune responses against vectors and may hold the key to understanding

the reason behind the sudden appearance of unwanted immune responses in many gene therapy

clinical trials.

Due to the lack of viral coding sequences, HD-Ad vectors are thought to be less immunogenic

than the first generation vectors and it is widely believed that HD-Ad vectors do not induce an

adaptive immune response (Alba et al., 2005; Liu and Muruve, 2003; Muruve, 2004). However,

other studies have shown that with readministration of HD-Ad vector particles, there is a

decrease in transgene expression which correlates with an increasing antibody titer against the

virus, indicating that there may be an adaptive response being mounted against HD-Ad particles

(Koehler et al., 2006). Nevertheless, none of the studies to date have assessed the ability of HD-

Ad particles to induce T cell immune responses upon airway delivery (Koehler et al., 2006;

Morral et al., 1999). Furthermore, adeno-associated virus (AAV) derived vectors, which are

thought as being the least immunogenic may also induce a cytotoxic CD8+ T cell response, for

in recent AAV clinical trials, there has been the appearance of cytotoxic CD8+ T cells upon

vector delivery (Wilson, 2007; Zaiss and Muruve, 2005). These observations clearly highlight

the importance of studying adaptive immune responses to the so-called “less-immunogenic”

vectors such as HD-Ad vectors. The ability to develop safe and effective gene therapy

therapeutics depends on better understanding of the immunological processes in the lung, for

targeting/inhibition of these processes will help in increasing the efficacy and persistence of gene

therapy vectors.

The ability of adenoviral-derived vectors to transduce a wide variety of cells, both dividing and

non-dividing, has led to their development as an efficient system for pulmonary gene transfer

(Campos and Barry, 2007). However, problems of host adaptive and innate immune responses

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have limited the use of adenoviral vectors for gene therapy because of the concern of their safety

and efficacy in vivo. Over the last decade, immense research has been conducted on improving

the vector design, which has resulted in development of helper-dependent adenoviral (HD-Ad)

vectors, which do not encode any viral genes and the only viral sequences present are the

terminal repeats along with the packaging signal (Flotte et al., 2007). This has resulted in HD-Ad

vectors as being the vector of choice for pulmonary gene therapy for diseases such as Cystic

Fibrosis (Brunetti-Pierri and Ng, 2006; Kushwah et al., 2007a; Kushwah et al., 2007b).

Dendritic cells are potent antigen presenting cells and lung dendritic cells are ideally positioned

in the airway epithelium network to perform surveillance of the inhaled antigens (Banchereau

and Steinman, 1998; Lambrecht et al., 1998; Upham, 2003). Upon uptake of foreign antigens,

airway DCs migrate to the T-cell zone of the draining lymph nodes, in particular the mediastinal

lymph node (MLN), where they interact with naïve T cells and prime adaptive immune

responses (Lambrecht et al., 2000a; Lambrecht et al., 2000b). Despite the critical role of airway

DCs in modulating immune responses to inhaled antigens, to our best knowledge, studies have

not examined the effects of HD-Ad vectors on pulmonary DCs. Moreover, pulmonary DCs

include both the conventional (cDC) and plasmacytoid DC (pDC) subsets and the relative

contribution of the two subsets and their response to HD-Ad vectors has not been investigated to

date.

In this study, we assessed the pulmonary adaptive immune responses to HD-Ad vectors. Since

incorporation of a transgene can further elicit/potentiate immune responses; we chose to assess

immune responses to empty HD-Ad vector which did not encode any transgene. We show that

HD-Ad vectors do have the ability to potentiate CD4+ as well as CD8+ T cell response upon

pulmonary delivery at high as well as substantially low dose. Furthermore, we also show that

HD-Ad vectors are potent stimulators of DC activation in vitro as well as in vivo with cDCs

playing the major role in priming T cells in the draining MLN. Moreover, HD-Ad vector

delivery also resulted in maturation of CD8α DCs within the draining MLN, indicating that this

particular subset of DC may play a role in cross-presentation of HD-Ad-derived antigens and

inducing CD8+ T cell proliferation. In contrast, though pulmonary pDCs do mature upon HD-Ad

delivery, they do not migrate to draining MLN, perhaps acting as a local source of IFN-α to

prime innate and adaptive immune response against HD-Ad vectors.

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2.3 Materials and Methods

Antibodies and other reagents

All the antibodies used were directed against mouse antigens. The following antibodies were

purchased from eBioscience (San Diego, CA): CD86 PECy5, CD80 PE, MHC II PE, CCR7 PE,

BrdU FITC, CD11c PE, mPDCA1 PE and the following from BD Biosciences (Missisauga,

ON): CD11c FITC, CD4 PECy7, CD8PECy7, CD8αPECy7, CD3 PE. CD11b antibody was a

kind gift from Dr. Jim Xiang (Saskatoon Cancer Center, SK). Isotype control IgGs were obtained

from eBioscience and/or Serotec (Raleigh, NC). CFSE was obtained from Molecular Probes

(Burlington, ON) and BrdU, DEAE-Dextran, heparin as well as FITC-Dextran from Sigma-

Aldrich (Oakville, ON). GM-CSF was obtained from R&D Systems (Minneapolis, MN). Cell

proliferation ELISA based on BrdU incorporation and chemiluminescent detection and

Collagenase D was obtained from Roche (Laval, QC). Aerrane was obtained from Baxter

(Missisauga, ON).

Mice and vector delivery

C57BL/6 mice were purchased from Charles River and maintained as per guidelines of SickKids

animal facilities. All the animal studies were reviewed and approved by the SickKids

Institutional committee for humane use of laboratory animals. Mice that were 8-11 wks of age,

were lightly anesthetized by Aerrane inhalation and a highly purified batch of HD-Ad vector

particles, purified via caesium chloride density gradient centrifugation were delivered

intranasally in a volume of 50ul with 5X109 or 1X1011 particles in complex with DEAE-Dextran

for efficient delivery, as described previously (Kushwah et al., 2007b).

Generation of bone marrow derived dendritic cells (BMDC) and HD-Ad induced

maturation

Bone marrow cells were isolated from tibias and femurs of adult mice and cultured in the

presence of GM-CSF as described previously (Lutz et al., 1999). On day 7, weakly adherent cells

were isolated and 85-90% of the cells were confirmed to be CD11c+ DCs via FACS analysis.

Preparation of “empty” HD-Ad vector particles was performed in our laboratory as described

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(Koehler et al., 2003).CD11c+ DCs were treated with HD-Ad vectors at different multiplicities

of infection and after 24 hours, marker expression was assessed via FACS analysis.

In vitro T cell proliferation and DC irradiation

HD-Ad treated CD11c+ DCs were exposed to single dose of 20 Gray of 60Co γ irradiation

(Princess Margaret Hospital) and then cultured with CD4+CD25- T cells, isolated from draining

MLN via cell sorting, for a period of 72 hours and BrdU solution was added during the last 12

hour to label the cells, after which proliferation was measured using BrdU cell proliferation,

chemiluminescent ELISA according to the manufacturer’s instructions (Roche, QC). In order to

calculate precursor frequencies, HD-Ad treated DCs were cultured with CFSE labelled naïve

CD4+ T cells isolated from draining MLN, for a period of 7 days, after which FACS analysis

was performed to assess CFSE dilution. T cell precursor frequencies were calculated from CFSE

dilution data as described (Chen et al., 2003).

Preparation of tissue lymphocytes

At different time-points after HD-Ad vector delivery, mice were sacrificed by i.p. injection of

Euthanyl (Bimeda-MTC, QC). In order to collect bronchoalveolar lavage fluid (BALF), mouse

lungs were lavaged as described previously(Koehler et al., 2006). After performing lavage, lungs

were perfused with 10 ml of PBS containing 10U/ml heparin via the right ventricle of the heart

in order to remove blood cells from the lung vasculature. Perfusion was performed until the

lungs turned completely white in color. Lungs were dissected out and after removal of draining

MLN; lungs were minced and digested for 25 minutes at 37 C using 250U/ml Collagenase D

solution, with addition of EDTA (10 mM final) during the last 5 minutes of incubation.

Fragments of digested lungs were passed through a 100 micron cell strainer (BD Biosciences)

and hypotonic lysis was used to remove erythrocytes. Cells were then counted and resuspended

at appropriate concentration for different experiments. Similarly, draining MLN were digested,

followed by suspension at appropriate concentrations.

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BrdU labelling

At different time points after vector delivery, mice were anaesthetized using Aerrane inhalation

and then BrdU was administered intranasally in a volume of 50ul at 16mg/ml concentration. 24

hours after BrdU delivery, mice were killed and lungs/draining MLN were isolated.

Ex vivo assessment of T cell specificity upon HD-Ad delivery

6 days after delivery of HD-Ad vector particles, mice were sacrificed and MLN were isolated. T

cells isolated from MLN by nylon-wool enrichment were co-cultured with saline / HD-Ad

treated, irradiated BMDCs for a period of 72 hours BrdU solution was added during the last 12

hour to label the cells, after which proliferation was measured using BrdU cell proliferation

ELISA according to the manufacturer’s instructions.

In vivo labelling of DCs

Mice were anaesthetized using Aerrane inhalation and 50ul of 1mg/ml FITC-Dextran or 50ul of

5mM CFSE was delivered intranasally, 2 hours prior to viral delivery. At indicated time points,

mice were killed and draining MLN were isolated.

Measurement of absolute DC counts

Lung or draining MLN were isolated and single cell suspensions were prepared as described

above. Cell suspensions were stained for DC specific markers and absolute counts were assessed

in the lung or draining MLN for each mouse separately.

Ab labelling and flow Cytometry

For in vitro BMDC experiments, DCs were labelled at 4 C for CD11c and maturation markers. In

order to assess pulmonary T cell proliferation, cells from BALF / lung or MLN single cell

suspensions were stained for CD3 and CD4/CD8 with/without BrdU staining as described

(Carayon and Bord, 1992). Migration of FITC-Dextran labelled DCs was assessed by labelling

of MLN single cell suspensions with CD11c. DCs in the airways were identified by staining

single cell lung suspensions with CD11c and CD11b antibodies for cDCs and with CD11c and

mPDCA1 antibody for pDCs. Maturation of DCs in vivo was assessed by staining with CD86

antibody as marker for DC maturation. Maturation of CD8α DCs was assessed by gating on

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CD11c+ CD8α+ cells from the draining lymph nodes and staining with CD86 antibody as a

maturation marker. Alveolar macrophages are unique amongst various macrophage

subpopulations in being very similar to DCs in the expression of surface markers (Guth et al.,

2009). Therefore, cells were first gated according to their FSC versus SSC characteristics to

discriminate highly autofluorescent macrophages from DCs and then gated on CD11c+CD11b+

populations to identify DCs (Grayson et al., 2007). Flow Cytometry data were acquired for each

of the experiments using a BD FACSCalibur (BD Immunocytometry Systems) at the SickKids-

UHN Flow Cytometry Facility and was analyzed using FlowJo flow Cytometry analysis software

(Treestar, Oregon).

Statistical analysis

Student’s t test was used to assess statistical significance between means. Significance was set at

p<0.05. All the data are presented as means +/- standard deviation.

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2.4 Results

HD-Ad vectors induce maturation of BMDCs

BMDCs generated in the presence of GM-CSF showed classical characteristics of immature

myeloid DCs, characterized as CD11c+ with low expression of MHC II, CD80, CCR7 and CD86

(Figure 2-1A). The effect of HD-Ad vector on the surface phenotype of BMDCs (day 7) was

assessed 24 hours after incubation with HD-Ad vector by Flow Cytometry. Incubation with HD-

Ad caused an increase in the intensity of expression along with the percent of DCs positive for

CD86, CD80 and MHC II expression (Figure 2-1A, B). Similarly, the chemokine receptor

CCR7, which is also a hallmark of DC activation, was highly upregulated upon treatment with

HD-Ad vectors (Figure 2-1A, B). As a control, the intensity of CD11c, which is expressed at

similar levels by both mature and immature DCs stayed stable (Figure 2-1B).

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Figure 2-1: Effects of HD-Ad vectors on maturation of bone marrow derived DCs.

(A) Representative histograms showing expression of CD86 (i), CD80 (ii), MHC II (iii) and

CCR7 (iv) on CD11c+ bone marrow derived DCs, 24 hours after treatment with HD-Ad vector

particles or saline. (B) Frequency of different subsets of CD11c+ bone marrow derived DCs after

HD-Ad or saline treatment. Using Flow Cytometry, expression of different markers was assessed

on CD11c+ DCs. Isotype matched IgGs were used as isotype controls and all values represent

means of 4-5 independent experiments; *p<0.05 compared to the DCs treated with saline (-Ad).

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HD-Ad vector treated DCs are potent inducers of T cell proliferation in vitro

Since HD-Ad treated DCs had the surface phenotype of mature myeloid DCs, we next assessed

induction of T cell proliferation by HD-Ad treated DCs in vitro. Lymphoid drainage from the

lung is predominantly into MLN; thereby, we isolated naïve CD4+ T cells from MLN as

representative of T cells that encounter HD-Ad presenting DCs in lymph node upon pulmonary

delivery of the vector. In order to determine the potency of adenoviral matured DCs to induce T

cell proliferation, BMDCs were matured overnight in the presence of different multiplicities of

infection (MOI) of HD-Ad vectors, ranging from MOI of 40 to 400. HD-Ad matured DCs were

irradiated and co-cultured with naïve CD4+ T cells from draining MLN for 3 days to assess T

cell proliferation via BrdU incorporation. Increasing MOI of HD-Ad used to treat DCs led to a

parallel increase in levels of T cell proliferation, which plateaued around MOI 80-200, indicating

that this particular MOI was inducing maximal T cell proliferation (Figure 2-2A). Therefore,

MOI of 100 was used for all the experiments and T cell proliferation in response to HD-Ad

vectors was also confirmed using CFSE dilution analysis (Figure 2-2B). Furthermore, we wanted

to estimate the frequency of naïve T cells from draining MLN responding to adenoviral vectors.

To do that, we assessed T cell proliferation in a co-culture of DCs with CFSE labelled naïve

CD4+ T cells. As a CFSE labelled cell undergoes division, the CFSE is halved and every round

of division gives a different peak due to varying CFSE intensity upon FACS analysis and

eventually, the frequency of responsive T cells can be calculated. Therefore, naïve CD4+ T cells

from draining MLN, labelled with CFSE were cultured with HD-Ad matured DCs for a period of

7 days, after which CFSE dye dilution was assessed via Flow Cytometry (Figure 2-2B). The

results indicated approximately 9% of naïve CD4+ T cells from draining MLN to be responding

to HD-Ad epitopes presented by DCs (Figure 2-2C). As a parallel control, we used co-culture of

CFSE labelled naïve T cells with DCs without addition of any HD-Ad vectors for maturation, for

this would indicate the percent of non-specific T cell proliferation. Non-specific proliferation

was determined to be around 3% by CFSE dilution analysis (Figure 2-2C). Therefore, upon

accounting for non-specific proliferation, results indicate that approximately 6% of naïve T cells

underwent proliferation in response to HD-Ad derived epitopes presented by HD-Ad-treated

DCs.

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Figure 2-2: Induction of CD4+ T cell proliferation in vitro by HD-Ad treated bone marrow

derived DCs.

(A) Assessment of CD4+ T cell proliferation using BrdU incorporation 3 days after co-culture of

naïve CD4+ T cells isolated from draining MLN with DCs treated with HD-Ad at different MOI

(RLU – relative light units, was a chemiluminescent measurement of BrdU incorporation by

proliferating cells). (B) Representative histogram depicting CFSE dilution profile of CFSE+

naïve CD4+ T cells which were cultured with HDAd treated DCs or media treated DCs for a

period of 7 days. (C) Frequency of T cells proliferating in co-culture of naïve CD4+ T cells

(CFSE+) with HD-Ad/media treated DCs, calculated from CFSE dilution analysis. *p<0.05,

compared to precursor frequency of T cells cultured in presence of media treated DCs. Results

for panel A were normalized to control measuring T cell proliferation in naïve CD4+ T cell and

DC co-culture, where DCs were not treated with HD-Ad vectors Results are representative of 3

independent experiments.

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HD-Ad vectors induce T cell response upon pulmonary delivery

HD-Ad vectors are regularly employed for pulmonary gene therapy studies under the assumption

that they do not induce an adaptive immune response in vivo. We next wanted to explore the

ability of HD-Ad vectors to induce T cell responses upon pulmonary delivery. Analysis of

immunological responses was performed at two different doses. The first dose being 1X1011

vector particles/mouse as a high dose, which has been shown to result in therapeutic effects in

CFTR knockout mice (Koehler et al., 2003). The second dose was a low dose of 5X109 vector

particles/mouse and is 3 fold lower than the low dose normally used for HD-Ad pulmonary

delivery (Koehler et al., 2006; Koehler et al., 2003).At first, we looked at the infiltration of T

cells in the BALF of mice at different time points after vector delivery. Under basal conditions,

in the absence of any inflammatory response, prior to delivery of HD-Ad vectors, BALF is

primarily composed of macrophages and only 2-3% of the cells can be identified as T

lymphocytes (Figure 2-3A). However, upon delivery of a high dose of HD-Ad vector (1 X 1011

vector particles), there was a gradual recruitment of T lymphocytes in BALF, which peaked

around day 7, when approximately half of the cells in BALF were T lymphocytes. The

infiltration gradually receded and was close to basal levels by day 14 (Figure 2-3A). Further

assessment of the phenotype of T cells indicated that there was infiltration of both CD4+ as well

as CD8+ T cells, which followed a similar trend as overall T cell responses, peaking around day

7, with approximately 10-15% of the cells in BALF being CD8+ T cells and 30-40% being

CD4+ T cells (Figure 2-3B). In contrast to high dose, delivery of a low dose of HD-Ad vector

(5X109 particles), which is even lower than the low dose used in gene therapy experiments, did

not result in any significant changes to the composition of the BALF (Figure 2-3A).

Since the presence of T cells in BALF is indicative of extensive inflammation and absence of T

cells in BALF does not indicate absence of T cell proliferation in response to HD-Ad, we also

assessed T cell proliferation within the airways along with the draining MLN. In order to directly

estimate the frequency of proliferating T cells responding to intranasal delivery, we used the

strategy of in vivo uptake of the thymidine analog BrdU with a flow-cytometry based analysis to

identify BrdU+ proliferating cells within the airways and draining lymph nodes. BrdU was

delivered 24 hour prior to sacrificing mice and single cell suspensions of lung and lymph node

cells were analyzed via Flow Cytometry to look at BrdU+ T cells. Figure 2-4A shows

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representative histograms indicating BrdU staining on CD3+ T cells on day 0 and day 6 post

HD-Ad delivery compared to control receiving saline intranasally.

As expected, mice that received high dose (1 X 1011 particles) of HD-Ad vector showed a

gradual increase in frequency of BrdU+ T cells in the lung, which followed a similar trend to that

of T cell infiltration in BAL, for it peaked around day 6 where approximately 45% of the T cells

in the airways were BrdU+ and receded to basal levels by day 14 (Figure 2-4B). Assessment of

composition of proliferating BrdU+ T cells in the lung on day 6 post HD-Ad delivery indicated

that at a high dose, approximately 35% of proliferating T cells in the lung were CD8+ T cells and

65% being CD4+ T cells (Figure 2-4C). Thus, overall indicating that at high dose, there is

potentiation of CD8+ T cell responses along with CD4+ T cell response within the lung. A

similar trend was also observed in draining MLN, where proliferating T cells were not detectable

in the draining MLN at a significant frequency before 3-4 days after HD-Ad delivery. The rate of

T cell proliferation in the draining MLN upon delivery of high vector dose (1 X 1011 particles)

reached a maximum during the day 6-8 interval, where approximately 25% of the T cells were

proliferating (BrdU+), beyond which the frequency eventually decreased after day 8 and returned

to basal levels by day 14 (Figure 2-4D). In contrast, though delivery of low dose of HD-Ad

vector did not result in T cell infiltration in BALF, there was indeed T cell proliferation observed

within the lung along with draining MLN (Figure 2-4B, D). The trend observed was similar to

that seen at a high dose, with an increase in the frequency of proliferating T cells both in the lung

and draining MLN during day 3-4 and eventually reaching maximum around day 6-8 interval,

with approximately 10% of the T cells in the MLN and 30% of the T cells in the lung to be

proliferating T cells (Figure 2-4B, D). Among proliferating BrdU+ T cells in the lung,

approximately 20% of proliferating T cells were CD8+ T cells and 80% were CD4+ T cells

(Figure 2-4C). Although the absolute frequencies of proliferating T cells were lower at the low

dose than those observed at a high dose, the overall trend stayed the same, with frequencies

returning to basal levels by day 14. In order to confirm the specificity of T cell proliferation

being observed in the draining MLN, T cells isolated from draining MLN of HD-Ad treated

animals on day 6 were cultured with HD-Ad treated BMDCs and T cell proliferation was

assessed by BrdU incorporation. As a control, T cells isolated from MLN of mice treated with

saline were co-cultured with saline treated BMDCs, which resulted in basal levels of

proliferation, perhaps indicating some non-specific proliferation. In contrast, co-culture of T cells

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from animals treated with saline with HD-Ad treated BMDCs resulted in higher levels of

proliferation, perhaps indicating the proliferation of naïve T cells in response to HD-Ad (Figure

2-4E). However, co-culture of T cells isolated from MLN of mice treated with high or a low dose

of HD-Ad resulted in significantly higher levels of T cell proliferation, indicating that HD-Ad

treated animals had significantly higher levels of proliferating T cells responding to HD-Ad

derived epitopes (Figure 2-4E). Therefore, the results clearly indicated that HD-Ad vectors also

induce T cell proliferation in vivo even at a dose below the so called low-dose used in gene

therapy experiments(Koehler et al., 2006).

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Figure 2-3: Assessment of T cell infiltration in bronchoalveolar lavage fluid upon

intranasal delivery of HD-Ad vectors to mice.

Mice were delivered a high dose (1X1011 particles) or a low dose (5X109 particles) of HD-Ad

vectors intranasally and at different time points, the presence of T cells in BAL fluid was

assessed via Flow Cytometry. (A) Percent of total T cells in BAL at different time points after

HD-Ad delivery. (B) Percent of CD4+ and CD8+ T cells in BAL at different time points after

delivery of a high dose of HD-Ad particles. All values represent n=3-4 mice per time-point.

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BrdU BrdU

Freq

uen

cy

Freq

uen

cy

Saline

HD-Ad

i ii

A

98

Figure 2-4. Assessment of T cell proliferation in the lung and the draining mediastinal

lymph node (MLN) in response to pulmonary HD-Ad delivery.

(A) Representative histogram showing BrdU incorporation among CD3+ T cells in the mouse

lung upon HDAd delivery (low dose - 5X109 particles) on day 0 (i) and day 6 (ii) compared to

mice receiving saline. (B) BrdU incorporation by CD3+ T cells in the lung at different time

points after delivery of high (1X1011 particles) or low dose (5X109 particles) of HD-Ad. (C)

Proportion of CD4+ and CD8+ T cells among proliferating BrdU+ T cells on day 6, upon

delivery of high (1X1011 particles) or low dose (5X109 particles)of HD-Ad. (D) BrdU

incorporation by CD3+ T cells in the draining MLN at different time points after delivery of high

(1X1011 particles) or low dose (5X109 particles) of HD-Ad. (E) Proliferation of draining MLN

derived T cells from HD-Ad or saline treated mice upon co-culture with HD-Ad treated DCs

(Control refers to co-culture of T cells derived from saline treated animals with saline treated

DCs, RLU – relative light units, was a chemiluminescent measurement of BrdU incorporation by

proliferating cells). *p<0.05 compared to control and saline groups. The percentage of BrdU

expression (B, D) is shown for CD3+ gated T cells at the indicated number of days after HD-Ad

delivery. All values represent n=3-4 mice per time point and n=8 per group (E).

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HD-Ad vectors induce maturation of cDC and alter their levels within the lung

In order to assess the response of DCs to HD-Ad in vivo, we delivered HD-Ad intranasally and

looked at effects on cDCs at different time-points. cDCs are usually identified via high

expression of CD11c along with CD11b (Smit et al., 2006). In order to clearly identify lung

cDCs, at first we performed bronchoalveolar lavage to remove most of alveolar macrophages

prior to isolation of cells from the lung. Moreover, gating characteristics were further used to

clearly identify lung the cDC subset.

In order to assess maturation of cDCs, we looked at the expression of CD86 costimulatory

marker on lung cDCs at different time-points after HD-Ad delivery. Figure 2-5A shows

representative histogram depicting that in response to HD-Ad there is an increase in CD86

expression on day 1, which returns to basal levels by day 5. The percent increase in mature lung

cDCs was calculated relative to the levels in control mice receiving saline instead of HD-Ad

vectors. Maximum increase in DC maturation was observed 1 day after HD-Ad delivery, which

decreased on day 2 and eventually returned to basal levels by day 5 (Figure 2-5B). At the same

time, we also looked at the absolute numbers of lung cDCs in response to HD-Ad delivery and

found that around day 2 after HD-Ad delivery there was a marked decrease in absolute numbers

of lung cDCs that eventually returned to normal levels by day 5 (Figure 2-5C). The decrease in

absolute numbers of cDCs mirrored the increase in maturation of cDCs in response to HD-Ad

vectors, with increased maturation correlating with reduced numbers of cDCs in the lung.

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CD86 CD86

Freq

uen

cy

Freq

uen

cy

+ Saline+ HD-Ad

i iiA

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Figure 2-5: Effect of HD-Ad delivery on pulmonary conventional DC (cDC) levels and

maturation within the lung.

(A) Representative histograms showing expression of CD86 on CD11c+ CD11b+ pulmonary

cDCs, which is upregulated on day 1 (i) after HD-Ad delivery and returns to normal by day 5

(ii). (B) Increase in relative levels of pulmonary cDCs in the lung expressing CD86 in mice

receiving HD-Ad vector compared to the levels of CD86 expressing pulmonary cDCs in mice

receiving saline. (C) Absolute numbers of pulmonary DCs in the lung at different time points

after HD-Ad delivery to mice. All values represent n=3-4 mice per time point.

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HD-Ad vectors induce migration of pulmonary DCs to draining MLN

To better understand the basis for the decrease in lung cDC levels after HD-Ad delivery, we next

used FITC-dextran to track the location of lung cDCs by assessing their migration. FITC-dextran

is a carbohydrate with a very high molecular mass, which is easily uptaken by phagocytic cells

including DCs and hence allows for tracking the location of phagocytic cells. In order to assess

DC migration after HD-Ad delivery, we delivered FITC-Dextran, 2 hours prior to HD-Ad

delivery to label phagocytic cells, including DCs within the airways. The efficacy of using FITC-

Dextran is demonstrated in Figure 3-6A which clearly shows that 1 day after FITC-dextran and

HD-Ad delivery, FITC-dextran+ cells can be observed within the draining MLN. In general,

cDCs rapidly migrate from peripheral tissues to the draining MLN in response to infection.

Using FITC-dextran, we were able to identify the subset of DCs that migrated from the lung to

the draining lymph node in response to HD-Ad vectors. In response to HD-Ad, there was a

dramatic increase in the percentage of FITC+ DCs (CD11c+ DCs) in the lymph node, 24 hours

after HD-Ad delivery (Figure 2-6B). In contrast, controls receiving only saline did not show this

dramatic increase; instead a small increase on day 1 was observed, indicating basal level of DC

migration (Figure 2-6B). The dramatic increase seen in response to HD-Ad delivery compared to

delivery of saline was thus an event specific to HD-Ad delivery. At different time-points, we also

measured the absolute count of DCs in the draining MLN; by looking at total DC count along

with the count of FITC+ as well as FITC- DCs. The trend observed was similar to that seen with

percent of FITC+ DCs, for both the FITC+ as well as FITC- DCs increased in levels after day 0

and peaked around day 2, after which they receded to basal levels by day 5 (Figure 2-6C). This is

consistent with the knowledge that mature DCs have a very short lifespan, so probably mature

DCs are entering LN, where they are priming T cell responses and then dying within 24-48

hours.

The peak in DC count within the MLN occurred around day 2 (Figure 2-6B), which correlates

with the observed reduction in absolute number of mature cDCs within the lung on day 2 (Figure

2-5C). Similarly, the percentage of mature DCs was also reduced on day 2 (Figure 2-5B),

indicating that probably migration of mature cDCs from the lung to the draining MLN resulted in

decrease of cDCs within the lung and a dramatic increase of FITC+ mature DCs within the

MLN.

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Figure 2-6: Effect of HD-Ad delivery on DC migration from the lung to the draining MLN.

(A) Representative histogram, 1 day after HD-Ad delivery showing that FITC+ DCs can be

identified in draining MLN (FITC expression by CD11c+ DCs in MLN). (B) Relative levels of

FITC+ CD11c+ DCs in MLN at different days after intranasal delivery of HDAd or saline, 2

hours after FITC-Dextran delivery. (C) Absolute number of DCs, both FITC- and FITC+ along

with total DCs in MLN were quantified at different time points after HD-Ad delivery via

expression of CD11c. All values represent n=3-4 mice per condition.

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HD-Ad vectors induce maturation of plasmacytoid DCs but do not affect their levels in the

lung

Since pDCs may play a key role in priming immune response to viral particles; we also looked at

the effects of HD-Ad vectors on pDCs within the airways (Santini et al., 2002). First we assessed

the effects of HD-Ad on maturation of pDC. Figure 2-7A shows representative histograms

showing CD86 expression on lung pDCs, at day 0 and day 1 after HD-Ad vector delivery. We

observed that upon HD-Ad delivery, pDCs matured rapidly, with maximum maturation being

observed around day 1 and eventually receding by day 2 and returning to basal levels by day 5

(Figure 2-7A, B). Thus, the trend was similar to that observed with cDCs. Since, the trend

observed with cDCs was associated with migration to the draining LN; we next assessed the

migration of pDCs from the lung to draining LN in response to HD-Ad. It has been reported that

FITC-Dextran cannot be used for identifying pDCs for pDCs do not readily take up FITC-

Dextran. CFSE has been used before to track pDCs and hence, we used intranasal delivery of

CFSE to track movement of pDCs from the lung to the draining LN(Legge and Braciale, 2003).

CFSE was delivered 2 hours prior to viral delivery to label all the cells in the airways. However,

we did not detect the presence of mPDCA1+ cells among the CFSE+ population within the

lymph nodes, indicating that there was no migration of pDCs induced in response to HD-Ad

vectors (Figure 2-7C). At the same time, we also measured the absolute numbers of pDC within

the lung at different time points and observed that upon HD-Ad vector delivery, the number of

pDC in the lung stayed very consistent, further confirming the lack of pDC migration from the

lung (Figure 2-7D).

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CD86 CD86

Freq

uen

cy

Freq

uen

cy

A

Saline

HD-Ad

i ii

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Figure 2-7: Effect of HDAd delivery on pulmonary plasmacytoid DC (pDC) levels and

maturation within the lung.

(A) Representative histograms showing expression of CD86 on mPDCA1+ pDCs prior to HDAd

delivery (day 0) (i) and 1 day after delivery (ii). (B) Increase in mPDCA1+CD86+ DC

population in the lungs of mice at different time points after HDAd delivery, relative to

mPDCA1+CD86+ pDC population in the lungs of mice receiving saline. (C) Representative

histogram showing lack of mPDCA1 expression on CFSE+ cells from the draining MLN, 24

hours after delivery of CFSE and HD-Ad vector particles. (D) Absolute numbers of pulmonary

pDCs in the lung at different time points after HD-Ad delivery to mice. All values represent n=3-

4 mice per time point.

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HD-Ad vectors induce maturation of CD8α DCs in the draining MLN

CD8α DCs are thought to play a key role in cross-presentation of antigens and since upon HD-

Ad delivery, we observed induction of a CD8+ T cell proliferation even without viral

transcription, we assessed the effects of HD-Ad delivery on CD8α DCs to identify if CD8α DCs

can play a potential role in induction of CD8+ T cell proliferation in response to intranasal

delivery of HD-Ad vectors. Since CD8α DCs are known to be lymphoid-resident; we did not

observe recruitment of CD8α DCs in the lungs. However, upon assessing for maturation of

draining MLN resident CD8α DCs by assessing for expression levels of CD86, we observed a

dramatic increase in the levels of CD86 expression by CD8α DCs in response to HD-Ad

delivery, indicating that intranasal delivery of HD-Ad was resulting in maturation of CD8α DCs

within the draining MLN (Figure 2-8).

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Figure 2-8: Effects of HD-Ad delivery on CD8α DCs in the draining MLN.

Flow cytometric analysis of CD86 expression was performed on CD8α DCs, 24 hours after HD-

Ad delivery. Isotype matched IgG was used as isotype control and results are representative of

n=8 animals.

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2.5 Discussion

Several clinical trials with first generation adenoviral vectors were initiated and ultimately

discontinued due to an induction of immune response, attributed to leaky expression of viral

genes from this vector in airway epithelium (Kushwah et al., 2007a). In contrast, development of

HD-Ad vectors with no viral coding sequences led to the assumption that adaptive immune

responses will be substantially reduced, since there is no leaky expression of any viral genes.

Due to this assumption, adaptive immune responses to HD-Ad vectors were never extensively

studied and hence the knowledge of pulmonary immune responses to these vectors is practically

absent. To the best of our knowledge, our study is the first study that demonstrates induction of

pulmonary adaptive immune response to intranasal delivery of HD-Ad vectors, clearly

illustrating the ability of HD-Ad vectors to induce both CD4+ as well as CD8+ T cell response at

high as well as substantially low dose. However, it is also important to note that though an

adaptive immune response was initiated at a low vector dose, the extent of pulmonary T cell

infiltration and T cell proliferation was substantially reduced. CD4+ T cells responses can

possibly be induced via uptake of exogenous HD-Ad vector particles by DCs and presentation

via MHC II pathway; however, induction of CD8+ T cell response in the absence of any viral

transcription was unexpected, since viral gene expression is usually needed for antigens to be

shuttled through the MHC I pathway.

In two recent gene therapy clinical trials using AAV serotype 2 vectors, which are thought to be

the least immunogenic; there were reports of apparent immune responses with appearance of

CD8+ effector T cells (Manno et al., 2006). This gave rise to the so-called capsid T-cell

hypothesis, which postulates that exogenous vector proteins may be shuttled into the MHC class

I pathway for T-cell priming (Wilson, 2007). Our results demonstrate similar phenomena

occurring upon pulmonary delivery of HD-Ad vectors, whereby we are seeing proliferation of

CD8+ T cells along with CD4+ T cells. This observation indicates that HD-Ad derived peptides

are being shuttled to MHC I pathway and cross-presented in context with MHC class I to induce

CD8+ T cell response along with presentation with MHC class II to induce CD4+ T cell

response. Five dominant CD8+ T cell epitopes on the capsid hexon protein of adenovirus have

been identified and since HD-Ad vector does indeed have the capsid proteins, it is possible that

cross-presentation of these epitopes may play a role in inducing CD8+ T cell responses (Leen et

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al., 2004). Our results indicate that perhaps CD8α DCs may play a role in cross-presenting HD-

Ad derived epitopes, since this subset of DC undergoes maturation within the draining MLN

upon HD-Ad delivery. We did not observe recruitment of CD8α DCs in the lungs as expected,

since these DCs are thought to be non-migratory. However, maturation of CD8α DCs indicates

that perhaps cDCs may be migrating to the draining MLN and transferring the antigens to CD8α

DCs, which thereby results in maturation of CD8α DCs. Mature CD8α DCs may then go on to

induce CD8+ T cell response, as has been observed with lung and sub-cutaneous infection using

HSV(Allan et al., 2006; Belz et al., 2004). Moreover, CD103+ DCs with the ability to cross-

present inhaled antigens and migrate to draining MLN have also been identified in the mouse

lung (del Rio et al., 2007). Therefore, there may be a specialized population of DCs that may be

initiating CD8+ T cell responses upon HD-Ad delivery via cross-presentation, which may

include CD103+ DCs along with CD8α DCs, which we have shown to mature in draining MLN

in response to HD-Ad. MHC:peptide complexes of immunodominant epitopes can have a half-

life over 7 days (Lazarski et al., 2005; Sant et al., 2005). Therefore, HD-Ad transduced epithelial

cells may present vector derived epitopes for a substantial amount of time to allow for

responding T cells to migrate to the airways and gradually target/eliminate some transduced

cells. Furthermore, upon readministration, memory T cells are probably activated which are

recruited to the site within the first 3 days and mediate gradual clearing of transduced cells and

hence may account for observed loss in transgene expression upon vector readministration

(Hikono et al., 2006; Koehler et al., 2006). Moreover, the extent of both CD4+ as well as CD8+

proliferation in the lung as well as draining MLN, peaked around day 6-7, which is similar to the

timeline of pulmonary T cell responses associated with other pulmonary infections such as

influenza (Lawrence and Braciale, 2004; Lawrence et al., 2005).

We assessed the response of pulmonary DCs to HD-Ad to gain insight into the priming of T cell

response to HD-Ad vectors upon pulmonary delivery. Our results demonstrated that early after

HD-Ad delivery, there is rapid maturation of both cDC along with pDC, which peaks around day

1. As DCs mature, there is migration of only cDCs to the lymph nodes, resulting in reduction of

percent of mature cells as well as reduction in absolute count of cDC in the lung by day 2 after

HD-Ad vector delivery. During the first 48 hours after delivery, there is massive increase in the

numbers of lung-derived DCs in the draining MLN, which eventually recedes to basal levels by

day 5. In contrast to cDCs, pDCs do not migrate and probably act as local sites of IFN-α

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secretion to prime innate as well as adaptive immune responses. Ex vivo studies have further

demonstrated that adenovirus can induce IFN-α secretion by pDCs via the TLR-MyD88 pathway

(Zhu et al., 2007). The observation of priming of DC maturation and migration within the first 48

hours is consistent with previous findings involving other respiratory viruses such as

paramoxyviruses (Grayson et al., 2007). Thus, cDCs seem to play the central role in priming

pulmonary immune responses against HD-Ad vectors and hence intervention within the first 24

hours of vector delivery to prevent DC maturation/migration may help in mitigating the adaptive

immune responses induced by HD-Ad vectors. In contrast to our findings that HD-Ad vectors act

as potent stimulators of pulmonary DC maturation and pulmonary T cell response, it was

recently reported that infection with adenovirus can suppress lung DC function, and such

suppression was related to adenovirus transcription (Thiele et al., 2006). We reason that the

report’s conclusion may be instead attributed to gene products encoded by the E3 region which

function to modulate host immune response (Rawle et al., 1989). In particular, E3gp19K protein

encoded by E3 region of the adenoviruses genome has been shown to sequester MHC I

complexes in endoplasmic reticulum, thereby suppressing CD8+ T cell responses (Sparer et al.,

1996). Since HD-Ad particles do not encode any viral genes including the E3 region, these

vectors cannot suppress pulmonary DC responses and are rather initiators of adaptive immune

response.

In conclusion, our data demonstrates that pulmonary delivery of HD-Ad results in induction of

CD4+ as well as CD8+ T cell responses that peak around day 5-7. These responses are primarily

orchestrated by cDCs which mature and migrate to draining MLN within the first 48 hours after

HD-Ad delivery. At the same time, pDCs also mature but do not migrate to MLN and hence may

act as local source of IFN-α to prime innate and adaptive immune response. Our findings

indicate that with a lower dose of HD-Ad vectors, the pulmonary adaptive immune response can

be reduced. Therefore, it is likely that by further increasing the efficacy of HD-Ad mediated gene

transfer, the vector dose can be reduced even further, which will likely limit the immune

response against HD-Ad vectors. Identification of adaptive immune responses initiated upon

pulmonary delivery of HD-Ad vectors indicates that strategies targeted to improve vector

delivery and persistence need to target both adaptive and innate immune responses. Moreover,

identification of the time-line of these responses may help in identifying the time-point for

intervention to prevent subsequent unwanted immunological responses.

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Chapter 3

Adenoviral vector formulation in DEAE-Dextran and NAL can

increase gene delivery to the airways

The contents of this chapter have been published in Gene Therapy: Rahul Kushwah, Jordan

Oliver, Huibi Cao and Jim Hu. Nacystelyn enhances adenoviral vector-mediated gene delivery to

mouse airways. Gene Therapy. 2007 Aug; 14: 1243-1248.

Acknowledgements: I would like to acknowledge Jordan Oliver for assistance with analyzing

airway histology and Huibi Cao for generation of adenoviral vectors used in this study.

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3.1 Abstract:

Adenoviral vector-mediated gene delivery has been extensively investigated for cystic fibrosis

(CF) gene therapy; however, one of its drawbacks is the immune responses to viral vectors as

well as the presence of a thick mucus layer in the airways of CF patients. Nacystelyn (NAL) is a

mucolytic agent with anti-inflammatory and antioxidant properties, and has been clinically tested

in CF patients to reduce mucus viscosity in the airways (Vanderbist et al., 2001). In chapter 2, I

have identified that pulmonary adaptive immune response can be reduced by reducing the vector

dose of adenoviral particles. Therefore, I explored the potential of NAL in increasing the

efficiency of pulmonary gene transfer, which could potentially reduce the vector dose, resulting

in reduced immune response against adenoviral vectors.

Objective of Study: In this study I explored the potential of NAL in enhancing adenoviral

mediated gene delivery to the airways.

Summary of Results: In this study, I show that pretreatment of the airways with NAL followed

by administration of adenoviral vectors in complex with DEAE-Dextran can significantly

enhance gene delivery to the airways of mice without any harmful effects. Moreover, NAL

pretreatment can reduce airway inflammation, which is normally observed after delivery of

adenoviral particles.

Conclusion: Taken together, these results indicate that NAL pretreatment followed by

adenoviral vector-mediated gene delivery can be beneficial to CF patients by increasing the

efficiency of gene transfer to the airways. This could potentially limit the immune response

against HD-Ad vectors by reducing the vector dose required to achieve therapeutic levels of gene

expression in the lungs.

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3.2 Introduction:

Over the last decade, adenoviruses have been widely used as vectors for gene therapy

applications, including treatment of cystic fibrosis (CF). Adenoviral entry into the cells requires

an initial binding of viral fiber to the CAR (Coxsackie and adenoviral receptor) receptor

followed by internalization. Unfortunately, the localization of CAR receptors to the basolateral

side of epithelial cells creates a new obstacle by reducing the efficiency of gene transfer.(Grubb

et al., 1994) Additionally, the airways of CF patients are covered with a thick mucus layer due to

chloride channel defects, which can further reduce the efficiency of gene delivery to the

epithelial cells.(Sanders et al., 2000) Furthermore, the host innate and adaptive immune

responses against adenoviral vectors also hinder their application in gene therapy protocols.

Development of helper-dependent adenoviral (HDAd) vectors has reduced the adaptive immune

response; however, due to the presence of capsid protein, the host innate immune response still

remains a problem. Therefore, enhancement of gene transfer can lead to reduction in the viral

dosage, which could further reduce the acute toxicity associated with the use of adenoviral

vectors. CF is an autosomal recessive monogenic disorder caused by mutations in the cystic

fibrosis transmembrane conductance regulator (CFTR) gene, which codes for an epithelial

chloride channel.(Koehler et al., 2004) The major cause of morbidity and mortality in CF

patients is lung disease, and hence the most important aim of CF gene therapy is to treat the

respiratory system. By far, CF gene therapy has garnered most research due to the feasibility of

theoretically correcting the disease by providing a single copy of the CFTR gene to airway

epithelial cells. Among all the vectors available for gene delivery, vectors derived from adeno-

associated virus (AAV) and adenoviruses have been most widely used. Multiple clinical trials

were initiated with AAV vectors; however, due to unsatisfactory primary outcomes, most of

them were discontinued. Similarly, adenoviral vector-mediated gene therapy suffered from the

drawbacks of vector toxicity and low efficiency of gene transfer.

Nacystelyn (NAL) is a derivative of N-acetyl-cysteine with an added L-lysine residue, and has

been used for treatment of impaired mucociliary clearance and chronic mucus retention in CF

patients.(Vanderbist et al., 2001; Vanderbist et al., 1999) NAL has been shown to possess

antioxidant (Gillissen et al., 1997), mucolytic and anti-inflammatory properties, thus indicating

that it may be beneficial for CF patients.(Tomkiewicz et al., 1994; Tomkiewicz et al., 1995)

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Clinical trials conducted with NAL in CF patients have demonstrated an improvement in

symptoms, with a dose-dependent decrease in sputum viscoelasticity along with a decrease in

sputum solids content, and an increase in chloride and sodium concentrations, thus indicating

improved mucus clearance.(Dasgupta and King, 1996). At the same time, no adverse effects

were observed in CF patients, even at a high dose of 24 mg given via inhaler (App et al., 2002).

Presently, multiple phase I and phase II clinical trials are being undertaken in Europe and the

USA to assess the efficacy of NAL treatment in CF patients. Studies conducted in humans have

also indicated that NAL prevents the respiratory burst of peripheral blood polymorphonuclear

cells, indicating that it may have potential in reducing damage to lungs caused by an active

inflammatory response (Nagy et al., 1997), which could be of a potential benefit in the context of

adenoviral vector-mediated CF gene therapy. Furthermore, NAL can inhibit the maturation of

human dendritic cells (Vosters et al., 2003), which in turn may limit the host adaptive immune

response against adenoviral vectors. NAL has also been shown to enhance cationic liposome

mediated gene transfer to the airways (Ferrari et al., 2001; Stern et al., 1998). In spite of this

wealth of knowledge, no studies have assessed the efficacy of NAL in enhancing adenoviral

vector-mediated gene delivery to the lung.

Complexing adenoviral particles with polycations has been shown to enhance transduction of

cell lines which are normally resistant to adenoviral infection in vitro (Kaplan et al.,

1998).DEAE-Dextran is a positively charged derivative of dextran sugar, which has been shown

to enhance the efficiency of adenoviral-mediated gene transfer to the airways (Gregory et al.,

2003).

To further explore the potential properties of NAL in gene therapy, we investigated whether

NAL can enhance adenoviral vector-mediated gene delivery to the airways of mice, and

simultaneously reduce the acute toxicity induced by adenoviral vector particles. Our data suggest

that pre-treatment of the airways with NAL prior to adenoviral vector administration can

enhance the efficiency of gene delivery. We propose a two-step delivery approach, where

adenoviral vectors complexed with DEAE-Dextran are delivered after NAL pre-treatment,

resulting in maximum enhancement of mouse airway epithelial cell transduction by adenoviral

vectors, along with a reduction in airway inflammation.

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3.3 Materials and Methods

Viral delivery

Solutions of DEAE-Dextran and viral particles were heated separately to 30oC, followed by

addition of DEAE-Dextran to the virus at the desired concentration. The mixture was then heated

for 20 minutes at 30oC to allow complex formation and delivery was done intranasally with or

without 100mM NAL pre-treatment, 30 minutes prior to vector administration. 6-10 weeks old,

female C57BL/6 mice (Charles River Laboratories, Wilmington, MA, USA) were used for the

experiment. Mice were briefly anaesthetized by isofluorane inhalation and viral or NAL solution

was placed in small drops on the nares, from where it was aspirated into the lungs.

X-gal staining

3 days after vector delivery, lungs were isolated and X-gal staining of the whole lung was

performed as described (Chow et al., 2000). After staining, tissues were washed and fixed in 4%

paraformaldehyde (PFA) for 4 hours, followed by transfer in 70% ethanol and cleared using

methyl salicylate prior to being photographed.

Beta galactosidase activity assay

3 days after vector delivery, mouse airways, including lungs and trachea were isolated,

homogenized and assayed for beta-galactosidase activity using a chemiluminescence kit

(Galacto-Light, Applied Biosystems, Foster City, CA, USA) and a microplate luminometer

(E.G&G. Berthold LB96V, Bad Wildbad, Germany), as described (Koehler et al., 2006), and the

values were normalized to the total protein concentration .

Histological analyses

2 days after vector delivery, lungs were infused and fixed overnight in 10% formalin. Formalin

fixed, paraffin-embedded mouse lung tissue samples were sectioned at 4 μm and stained with

haematoxylin and eosin (H&E) for histological examination under a light microscope.

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3.4 Results

Pretreatment of airways with NAL enhances adenoviral mediated gene delivery

We assessed the ability of NAL to enhance adenoviral vector-mediated gene delivery in vivo. We

administered adenoviral vectors encoding LacZ in saline to C57BL/6 mice, with or without NAL

pre-treatment. LacZ was chosen as a transgene due to a relative ease of quantification of

expression at the protein level, along with the ease to assess localization of gene

delivery/expression throughout the lung by performing X-gal staining. Moreover, first generation

adenoviral (FGAd) vectors were used for most of the experiments, since these vectors are much

easier to produce than the helper dependent adenoviral (HDAd) vectors. We delivered 5X109

FGAd vector particles per animal, which we identified to be the optimum dose based on our

preliminary studies. One of the aims of gene therapy is to use the lowest number of viral particles

possible while obtaining maximal transduction, in order to reduce any toxicity associated with

the viral vector itself. Furthermore, pre-treatment with NAL did result in an enhancement of

adenoviral gene delivery (Figure 3-1B) compared to control, which received adenoviral vectors

in saline without NAL pre-treatment (Figure 3-1A); however, the level of enhancement was not

very high. Thus, we hypothesized that delivery of adenoviral vectors in other formulations, such

as DEAE-Dextran, after NAL pre-treatment may result in a significant enhancement of

adenoviral vector-mediated airway epithelial cell transduction.

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Figure 3-1: X-gal staining of the lung after delivery of FGAd vector particles encoding

LacZ under control of CMV promoter (FGAdCMVLacZ) in saline with or without NAL

pre-treatment.

(A) Viral delivery in saline. (B) NAL pre-treatment followed by viral delivery in saline 30

minutes later. 5X109 adenoviral particles encoding for LacZ were delivered intranasally to 6-10

weeks old, female C57BL/6 mice in a volume of 50 μl with or without pre-treatment with 100

mM NAL. After 3 days, lungs were isolated and X-gal staining of the whole lung was

performed. After staining, tissues were washed and fixed in 4% paraformaldehyde (PFA) for 4

hours, followed by transfer in 70% ethanol and cleared using methyl salicylate prior to being

photographed. Images are representative of one mouse per group (n=3 mice per group).

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NAL treatment of airways followed by delivery of adenoviral vector formulation in DEAE-

Dextran results in highest levels of transgene expression

Adenoviral vectors encoding LacZ were complexed with different concentrations of DEAE-

Dextran and delivered to C57BL/6 mice with or without NAL pre-treatment. NAL pre-treatment

led to a significant enhancement of LacZ transgene expression in the mouse airways upon

adenoviral vector delivery (Figure 3-2A). NAL-mediated enhancement of transgene expression

could be observed both in mice receiving viral formulation in 10 μg/ml as well as 20 μg/ml

DEAE-Dextran, indicating that a combinatorial approach involving NAL pre-treatment and

DEAE-Dextran can be used for significant enhancement of adenoviral vector-mediated gene

delivery to the airways. In contrast, viral delivery in saline after NAL pre-treatment only led to a

minor enhancement, indicating that NAL by itself may not open tight junctions to allow vector

particles to reach the basolateral surface, and therefore, DEAE-Dextran is needed to partially

fulfill this requirement. Pre-treatment with NAL followed by adenoviral vector delivery in 20

μg/ml DEAE-Dextran resulted in approximately 64 fold enhancement in gene transfer compared

to viral delivery in saline without NAL pre-treatment (Figure 3-2A), whereas viral delivery in 20

μg/ml DEAE-Dextran without NAL pre-treatment resulted in approximately 20 fold

enhancement (Figure 3-2A), indicating a further 3-4 fold enhancement by NAL pre-treatment

over DEAE-Dextran without NAL pre-treatment (Figure 3-2B). Similar 3-4 fold enhancement of

LacZ transgene expression was observed upon comparing mice receiving NAL pre-treatment

followed by viral delivery in 10 μg/ml DEAE-Dextran with mice receiving viral delivery in 10

μg/ml DEAE-Dextran without NAL pre-treatment (Figure 3-2A).

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Figure 3-2: Beta-galactosidase activity after delivery of FGAdCMVLacZ vector particles

with or without NAL pre-treatment.

(A) Normalized beta-galactosidase activity and fold enhancement in beta-galactosidase activity

by viral delivery in saline or in DEAE-Dextran with or without NAL pre-treatment (pre-NAL),

where n=8 mice in all the groups except pre-NAL-Saline, where n=4. (B) Combination of NAL

together with FGAd vector particles in 10 μg/ml DEAE-Dextran versus delivery in 10 μg/ml

DEAE-Dextran without any NAL, where n=5 mice per group. (C) Fold enhancement in beta-

galactosidase activity by NAL pre-treatment over delivery of viral particles (FGAd or HDAd) in

20 μg/ml DEAE-Dextran without NAL pre-treatment, where n=5 mice per group. 5X109

FGCMVLacZ or HDK18LacZ vectors were delivered in a volume of 50μl with or without NAL

pre-treatment (Noveon, USA), in a formulation of saline or DEAE-Dextran. Error bars represent

SEM. *Significantly different compared to all other groups (P≤0.05), RLU – Relative light unit.

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Effects of NAL in enhancing gene transfer are not observed if it is combined in the

formulation with adenoviral vectors and DEAE-Dextran

The simplest approach for gene delivery would be a combination of NAL together with DEAE-

Dextran and virus rather than any form of NAL pre-treatment. To test this idea, we assessed the

efficacy of NAL mediated enhancement, by combining it with DEAE-Dextran and virus instead

of pre-treating with NAL. The results indicate that a combination of NAL administered together

with virus in DEAE-Dextran ameliorated the enhancement effects seen with NAL pre-treatment

(Figure 3-2C). Hence, NAL can only enhance adenoviral vector-mediated gene delivery if it is

administered prior to vector delivery.

Effects of NAL in enhancing gene transfer are not vector dependent

We also assessed whether NAL-mediated enhancement of gene delivery was specific for first

generation adenoviral vectors or whether it could be applied to third generation of viral vectors

(HDAd vectors). We assessed the ability of NAL pre-treatment to enhance gene delivery of

HDAd vectors using LacZ construct driven by K18 promoter (HDAdK18LacZ). The K18

promoter is derived from the cytokeratin K18 gene, and it has been previously shown to mediate

airway epithelium specific transgene expression.(Koehler et al., 2003) Analysis of beta-

galactosidase activity confirmed that NAL pre-treatment can also be used to enhance gene

delivery mediated by HDAd vectors with airway epithelium specificity to similar levels as seen

with FGAd vectors (Figure 3-2D).

Effects of NAL and DEAE-Dextran in enhancing adenoviral mediated gene transfer are

observed throughout the lungs

Since the beta-galactosidase assay relies on protein expression in the whole lung and cannot

differentiate between localized or uniform expression, we performed staining of the whole lung

to assess localization of LacZ expression after delivery of adenoviral vectors to the mouse

airways in saline without NAL pre-treatment, DEAE-Dextran without NAL pre-treatment or

DEAE-Dextran with NAL pre-treatment (Figure 3-3). X-gal staining of the lungs isolated from

mice that received saline without any virus did not show any positive LacZ staining throughout

the airways as expected (Figure 3-3A). Delivery of FGAd vector particles in saline led to weak

transduction of bronchioles in the lung (Figure 3-3B, left panel) whereas delivery of virus in

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DEAE-Dextran without NAL pre-treatment led to transduction throughout the lung except for

the lower extremities of the airways (Figure 3-3C, left panel). In contrast, pre-treatment with

NAL followed by viral delivery in DEAE-Dextran led to uniform transgene expression

throughout the lung, including the lower extremities of the airways (Figure 3-3D, left panel). We

also took a closer look at staining within the trachea, and in contrast to viral delivery in saline or

DEAE-Dextran without NAL pre-treatment, pre-treatment with NAL followed by viral delivery

in DEAE-Dextran resulted in maximum transduction of the trachea as indicated by an increase in

X-gal staining intensity (Figure 3-3B-D, right panels). In contrast to the previous reports of

DEAE-Dextran mediated-enhancement of gene transfer only in the lower lung (Kaplan et al.,

1998), our observations indicated enhancement throughout the airways including the mouse

trachea which could be due to higher DEAE-Dextran concentration used in our experiments

(Figure 3-3C, right panel). The whole lung X-gal staining data in combination with results from

beta-galactosidase activity assay (Figure 3-2) confirmed that NAL pre-treatment followed by

viral delivery in DEAE-Dextran can significantly enhance gene delivery throughout the airway

epithelium.

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Figure 3-3: X-gal staining of the whole lung (left panel) and the trachea (right panel) upon

delivery of FGAdCMVLacZ vector particles with or without NAL pre-treatment.

(A) Control with saline, no virus. (B) Virus in saline. (C) Virus in 20 μg/ml DEAE-Dextran. (D)

NAL pre-treatment (100mM) followed by viral delivery in 20 μg/ml DEAE-Dextran. Briefly,

after 3 days lungs were isolated followed by fixation, washing and overnight staining in X-gal

solution. Images are representative of one mouse per group (n=3 mice per group).

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NAL reduces the adenoviral vector induced acute inflammation in the airways

Having established the efficacy of NAL pre-treatment in enhancing adenoviral vector-mediated

gene transfer to the airways, we next assessed whether the use of NAL can potentially reduce the

acute toxicity of adenoviral particles. Histological analysis of the mouse airways was conducted

by a veterinary pathologist. Mild perivascular and peribronchiolar inflammation characterized by

an infiltration of neutrophils, lymphocytes, and macrophages was observed in the lungs of mice

in all groups except for the control group which appeared to have normal lung histology (Figure

3-4). The data showed that inflammation was more prominent in mice receiving adenoviral

vector particles in saline or DEAE-Dextran without NAL pre-treatment (Figure 3-4B, C) as

compared to mice which received pre-treatment with NAL (Figure 3-4D). This indicates that

NAL pre-treatment can reduce the acute toxicity of adenoviral vector particles. Further analysis

could have been carried out to quantify inflammation by grading various parameters such as the

site and cell composition of lung inflammation as well as alveolitis. Additionally, other

histological features such as subpleural inflammation and interstitial inflammation of the

parenchyma are also taken into account (Khelef et al., 1994).

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Figure 3-4: Histopathological analysis of the whole lung upon FGAd vector delivery with or

without NAL pre-treatment.

(A) Control – saline with no virus. (B) Viral delivery in saline without NAL pre-treatment. (C)

Viral delivery in 20 μg/ml DEAE-Dextran without NAL pre-treatment. (D) NAL pre-treatment

(100mM) followed by viral delivery in 20 μg/ml DEAE-Dextran. Images were taken at 100x

magnification (left panels) and right panels refer to a 400x magnified region indicated by a box

within the left panels. 5X109 FGCMVLacZ particles were delivered to mice in saline or DEAE-

Dextran with or without NAL pre-treatment, and after two days, lungs were infused and fixed

overnight in 10% formalin. Images are representative of one mouse per group (n=3 mice per

group).

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3.5 Discussion

In summary, our results demonstrate the utilization of NAL, a clinically used mucolytic agent in

CF patients, in enhancing adenoviral vector-mediated gene delivery to the mouse airways.

Additionally, NAL is able to reduce the acute toxicity associated with delivery of adenoviral

particles in saline, indicating its importance as an anti-inflammatory agent with the potential of

enhancing adenoviral gene delivery in a clinical setting for CF patients. However, it needs to be

noted that murine airways are different from human airways for they secrete low amounts of

mucus due to limited number of submucosal glands (Fehrenbach, 2002). Therefore, enhancement

of gene delivery seen in mice airways may be mostly due to a combination of antioxidant,

mucolytic and anti-inflammatory effects of NAL. In contrast, human airways both in normal and

particularly in CF patients secrete a lot of mucus; therefore, we can speculate that pre-treatment

with NAL may result in even higher levels of enhancement of adenoviral vector-mediated gene

delivery than seen in mice, mostly due to its mucolytic activity in combination with anti-

inflammatory effects. Taken together, our findings establish the combination of NAL and

DEAE-Dextran as a novel and a highly effective formulation for enhancing adenoviral vector

mediated gene delivery.

In chapter 2, we identified that although HD-Ad vectors are devoid of viral coding sequences,

intransal delivery of HD-Ad vectors does in fact elicit an adaptive immune response

characterized by induction of both CD4+ along with CD8+ T cells. One of the strategies to limit

the immune response is to limit the dose of vector delivered, which will limit the availability of

HD-Ad immunogen. Therefore, by formulating HD-Ad vectors in NAL and DEAE-Dextran, we

can further reduce the vector dose needed for therapeutic levels of gene expression, which likely

will limit the immune response to HD-Ad vectors.

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Chapter 4

Induction of immunological tolerance towards HD-Ad vectors using

immature dendritic cells

Acknowledgements: I would like to acknowledge Cathleen Duan for assistance in performing

some of β-galactosidase activity assays (few samples for Figures 4-1 and 4-9B) and would like to

acknowledge Drs. Li Zhang and Shaf Keshavjee for reading the manuscript.

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4.1 Abstract:

HD-Ad vectors are able to mediate high-level transgene expression in the absence of chronic

toxicity observed with FG-Ad due to the absence of viral coding sequences. Although with HD-

Ad vectors the immune response is reduced, subsequent re-administrations with high dose of

vector can increase it to levels seen with FG-Ad vectors, thereby limiting transgene expression.

Therefore there is a need to induce tolerance within the host towards the HD-Ad vectors without

compromising the immunity towards other infections.

Objective of Study: In this study we generated dendritic cells refractory to Ad induced

maturation and assessed their potential in inducing tolerance to HD-Ad vectors.

Summary of Results: DCs generated in presence of IL-10 were refractory to HD-Ad induced

maturation and instead of inducing T cell differentiation, primed differentiation of IL-10

producing regulatory T cells which suppressed T cell proliferation. Delivery of HD-Ad pulsed;

IL-10 modified DCs to mice suppressed the adaptive immune response against HD-Ad vectors

following intranasal delivery and also primed differentiation of IL-10 producing Tregs in vivo,

which could suppress HD-Ad induced T cell proliferation.

Conclusion: Taken together, these results indicate that delivery of HD-Ad pulsed DCs generated

in the presence of IL-10 to mice suppresses the immune response to HD-Ad vectors and likely

induces a state of immunological tolerance.

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4.2 Introduction:

Adenoviruses are a family of DNA viruses with a linear, double-stranded genome of about 36

kb. Adenoviral (Ad) vectors have been extensively studied for pulmonary gene therapy due to

their ability to efficiently transduce a wide variety of proliferating and non-proliferating

cells(Cao et al., 2004; Kushwah et al., 2007a; St George, 2003). Initially, the use of First

Generation adenoviral (FG-Ad) vectors has demonstrated substantial host immune responses to

viral antigens leading to destruction of transduced cells and prevention of re-administration

(Yang et al., 1996a). Significant improvement in the safety and efficacy of Ad-based vectors

came with the development of Helper-Dependent adenoviral vectors (HD-Ad), which do not

encode any viral genes (Ng et al., 2001; Ng et al., 1999; Parks, 2000). In contrast to FG-Ad, HD-

Ad vectors are able to mediate long-term, high-level transgene expression in the absence of

chronic toxicity observed with FG-Ad due to the absence of viral coding sequences. We and our

collaborators have previously demonstrated unprecedented levels of transgene expression when

HD-Ad vectors were delivered to the airway of rabbits(Koehler et al., 2005) and baboons(Hiatt et

al., 2005). Although with HD-Ad vectors the immune response is reduced, subsequent vector re-

administrations can increase it to levels seen with FG-Ad vectors, thereby limiting transgene

expression (Koehler et al., 2006). Therefore there is a need to induce tolerance within the host

towards HD-Ad vectors without compromising immunity towards other infections to mediate

stable gene expression following multiple vector readministrations to the lung.

Dendritic cells (DCs) are professional antigen-presenting cells derived from the same bone

marrow precursors as macrophages. DCs reside in the tissues as “immature” DC, which in the

presence of appropriate signals, turn into “mature” DC. Mature DCs are potent stimulators of T

cell proliferation and effector cell development. In contrast to mature DCs, immature DCs have

been implicated in generation of peripheral tolerance through regulatory T cell (Treg)

development. Tregs constitute a suppressor subset of T cells which are critical in suppressing

autoimmunity by suppressing the function of autoreactive T cells (Vignali et al., 2008).

Restimulation of blood cord-derived naïve CD4+ cells with immature DC has been shown to

induce the development of Tregs, whereas restimulation with mature DCs leads to Th1 effector

phenotype (Jonuleit et al., 2000). Therefore, the maturation status of DCs is critical in deciding

between tolerance and immunity. Moreover, adoptive transfer of immature DCs into rats before

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cardiac transplant, has been shown to significantly enhance graft survival through induction of

Tregs (DePaz et al., 2003). Therefore, immature DCs presenting HD-Ad derived epitopes can

likely be used to generate long-lasting tolerance towards HD-Ad vectors, which can prevent loss

in transgene expression associated with multiple rounds of HD-Ad mediated gene delivery to the

airways.

In this study, we assessed the feasibility of inducing immunological tolerance towards HD-Ad

vectors using immature DCs pulsed with HD-Ad vectors. DCs generated in presence of IL-10

were refractory to HD-Ad-induced maturation and instead of inducing T cell differentiation,

primed differentiation of IL-10 producing regulatory T cells which suppressed T cell

proliferation. Delivery of HD-Ad-pulsed IL-10-modified DCs to mice suppressed the adaptive

immune response against HD-Ad vectors following intranasal delivery and also primed

differentiation of IL-10-producing Tregs in vivo, which could suppress HD-Ad induced T cell

proliferation. After three rounds of vector delivery, mice that received HD-Ad pulsed, IL-10-

modified DCs had significantly elevated levels of vector encoded transgene expression compared

to control groups. Altogether, these findings highlight a new DC-based strategy to induce

tolerance towards HD-Ad vectors, which allows for sustained gene expression in the airways

after multiple rounds of vector readministration.

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4.3 Materials and Methods

Viral delivery

Solutions of DEAE-Dextran and viral particles were heated separately to 30oC, followed by

addition of DEAE-Dextran to the virus to a final concentration of 10µg/ml. The mixture was

then heated for 20 minutes at 30oC to allow complex formation and delivery was performed

intranasally as described previously (Kushwah et al., 2007b). 6-10 weeks old, C57BL/6 mice or

Rag deficient mice (Charles River Laboratories, Wilmington, MA, USA) were used for the

experiment. All the animals were housed at SickKids animal facility or Toronto Centre for

Phenogenomics and all the animal studies were conducted followings institutional guidelines.

β-galactosidase activity assay

3 days after vector delivery, mouse airways, including lungs and trachea were isolated,

homogenized and assayed for β-galactosidase activity using a chemiluminescence kit (Galacto-

Light, Applied Biosystems, Foster City, CA, USA) and a microplate luminometer (E.G&G.

Berthold LB96V, Bad Wildbad, Germany), as described (Koehler et al., 2006) and the values

were normalized to the total protein concentration.

X-gal staining

3 days after vector delivery, lungs were isolated and X-gal staining of the whole lung was

performed as described (Chow et al., 2000). After staining, tissues were washed and fixed in 4%

paraformaldehyde (PFA) for 4 hours, followed by transfer in 70% ethanol and cleared using

methyl salicylate prior to being photographed.

Generation of bone marrow derived dendritic cells

C57BL/6 mice were used as a source of bone marrow cells. Briefly, humerus, tibia and femur

bones were isolated and bone marrow (BM) cells were flushed out using saline. BM cells were

cultured in presence of GM-CSF (Peprotech, Rocky Hill, NJ, USA) only to generate normal DCs

or in presence of 70ng/ml IL-10 (Peprotech) to generate IL-10-modified tolerogenic DCs.

Around day 3 of culture, non-adherent cells were removed, for they comprised mostly lymphoid

lineages. DCs were ready for use on day 7, during which either they were used as is or were

incubated with HD-Ad particles.

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Cytokine measurements

Cytokine levels were measured using IL-1β, IL-10, IFN-γ and IL-12 ELISA kits (R&D systems,

Minneapolis, MN, USA).

Flow cytometry

All the antibodies were obtained from eBioscience (San Diego, CA, USA) unless indicated

otherwise. For in vitro DC experiments, DCs were labelled at 4 C with CD11c and maturation

markers. In order to assess pulmonary T cell proliferation, cells from BALF / lung or MLN

single cell suspensions were stained for CD3 and CD4/CD8 with/without BrdU staining as

described (Carayon and Bord, 1992). Migration of FITC-Dextran labelled DCs was assessed by

labelling of MLN single cell suspensions with CD11c. DCs in the airways were identified by

staining single cell lung suspensions with CD11c and CD11b antibodies. Maturation of DCs in

vivo was assessed by staining with CD86 antibody as a marker for DC maturation. Cells were

gated according to their FSC versus SSC characteristics to discriminate highly autofluorescent

macrophages from DCs as described (Grayson et al., 2007). Intracellular cytokine staining of T

cells was performed as described (Kushwah et al., 2009; Kushwah et al., 2010). Intracellular

cytokine staining for adenoviral uptake was performed using anti-hexon antibody (Chemicon,

Temekula, CA, USA) as described (Weaver and Kadan, 2000). Flow Cytometry data were

acquired for each of the experiments using a BD FACSCalibur at the SickKids-UHN Flow

Cytometry Facility and was analyzed using FlowJo flow Cytometry analysis software (Treestar,

Oregon, USA).

T cell proliferation assay

DCs incubated overnight with HD-Ad vectors were irradiated (25 Gray) and co-cultured with

naïve T cells for a period of 3-7 days. During the last 24 hours of culture, BrdU was added to be

incorporated by proliferating cells. T cell proliferation was measured using Cell proliferation

chemiluminescence kit (Roche, Laval, QC, Canada).

ELISA for anti-Ad antibodies

A pan-specific (IgA, IgE, IgGs, IgM) ELISA for mouse anti- Ad5 antibodies was performed as

described (Croyle et al., 2001; Koehler et al., 2006). 96-well ELISA plates (Corning Costar,

Acton, MA, USA) were coated overnight at 4°C with 5 x 1010 particles of Ad5 per well in 100

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mM bicarbonate buffer pH 9.6. The plates were then washed 4 times with TBS-T (TBS with

0.05% Tween-20) and blocked for 3 h at room temperature with 3% BSA in TBS. Mouse BALF

diluted 1:100, or serum diluted 1:2000 in TBS, was added to the wells for overnight incubation at

4°C. The plates then were washed 4 times with TBS-T and incubated with anti-mouse-Ig-biotin

(BD Pharmingen, San Diego, CA, USA), diluted 1:5000 in TBS, for 3 h at room temperature.

The plates were washed 4 times with TBS-T and then incubated with avidin-alkaline

phosphatase (Sigma-Aldrich, USA), diluted 1:50 000 in TBS, for 2 h at room temperature. The

plates were washed 4 times with TBS-T, incubated with 1 mg/ml p-nitrophenyl phosphate

(Sigma-Aldrich) in 100 mM diethanolamine buffer pH 9.8, containing 0.5 mM MgCl2, for 10

min at room temperature. The reaction was stopped by the addition of EDTA (40 mM final), and

optical density was read at 405 nm.

Assessment of regulatory T cell generation

CFSE (Sigma Aldrich) labelled naïve T cells were cultured with adenoviral incubated DCs. After

5 days, CFSE+ cells were isolated and added at different ratios to a fresh co-culture of naïve T

cells and adenoviral incubated DCs derived from bone marrow cells cultured in the presence of

GM-CSF only. T cell proliferation was assessed as described above. To assess for Tr1

generation, experiments were repeated with non labeled naïve T cells and IL-10 production by

CD4+ T cells was assessed by flow cytometry.

BrdU labeling

At different time points after vector delivery, mice were anaesthetized using Aerrane inhalation

and then BrdU was administered intranasally in a volume of 50µl at 16mg/ml concentration. 24

hours after BrdU delivery, mice were killed and lungs/draining MLN were isolated.

In vivo labelling of DCs

Mice were anaesthetized using Aerrane inhalation and 50µl of 1mg/ml FITC-Dextran(Sigma-

Aldrich, USA) was delivered intranasally, 2 hours prior to viral delivery as described

previously(Kushwah et al., 2008).

Proliferation and cytokine production by T cells from lymph nodes

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Six days after challenge with HD-Ad vector particles, mice were sacrificed and MLN were

isolated. T cells isolated from MLN by nylon-wool enrichment were co-cultured with HD-Ad

treated, irradiated DCs for 72 hours and BrdU solution was added during the last 12 hours to

label the cells, after which proliferation was measured using BrdU cell proliferation ELISA

according to the manufacturer’s instructions (Roche, QC). Media was collected and used for

measurement of cytokine production.

In vivo delivery of dendritic cells

5 X 106 DCs suspended in 150µl of saline were injected via pulmonary injections as described

(Onn et al., 2003; Umeoka et al., 2004).

Preparation of single cell suspension from lungs

At different time-points after HD-Ad vector delivery, mice were sacrificed by i.p. injection of

Euthanyl (Bimeda-MTC, QC, Canada). In order to collect bronchoalveolar lavage fluid (BALF),

mouse lungs were lavaged as described (Koehler et al., 2006). After performing lavage, lungs

were perfused with 10 ml of PBS containing 10U/ml heparin via the right ventricle of the heart

in order to remove blood cells from the lung vasculature. Perfusion was performed until the

lungs turned completely white in color. Lungs were dissected out and after removal of draining

MLN; lungs were minced and digested for 25 min at 37 C using 250U/ml Collagenase D

solution, with addition of EDTA (10 mM final) during the last 5 min of incubation. Fragments of

digested lungs were passed through a 100 micron cell strainer (BD Biosciences) and hypotonic

lysis was used to remove erythrocytes. Similarly, draining MLN were digested, followed by

suspension at appropriate concentrations.

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4.4 Results

Sustained gene expression is observed in immunodeficient mice following readministration

of HD-Ad vectors to the lung

In order to assess the role of adaptive immune response in mediating loss of transgene expression

observed with HD-Ad readministrations to the lung, we used Rag-deficient mice, which lack

mature T and B cells and are deficient in adaptive immune response(Mombaerts et al., 1992).

Vector delivery was carried out as illustrated in Figure 4-1A. Single intranasal delivery of HD-

Ad vectors encoding LacZ under the control of K18 promoter (HD-AdK18LacZ) resulted in

similar β-galactosidase activity in lungs of Rag-deficient and wild-type counterparts (Figure 4-

1B). However, upon both double and triple delivery, where mice first received empty HD-Ad

particles (C4HSU) followed by HD-Ad K18LacZ particles, β-galactosidase activity was

significantly reduced in the lungs of wild-type mice compared to Rag deficient mice (Figure 4-

1C,D). Double delivery led to a 50% loss of β-galactosidase activity in the lungs of wild-type

mice, which further increased to 82% loss in activity after triple delivery compared to mice

which received single delivery (Figure 4-1E). In contrast, only 20-30% reduction in β-

galactosidase activity was observed in the lungs of Rag deficient mice upon both double and

triple delivery. Taken together these findings indicate that upon multiple readministrations of

HD-Ad particles to the lung, there is a substantial loss in transgene expression and the loss is

largely mediated by an adaptive T and B cell response, which is targeted towards the HD-Ad

vector derived antigens.

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Figure 4-1: HD-Ad mediated transgene expression following readministrations is sustained

in Rag deficient mice, which are devoid of an adaptive immune response.

(A) Gene delivery was carried out as described. (B-D) Beta galactosidase activity in lungs of

mice following single, double or triple delivery. (E) Percent loss in beta galactosidase activity in

lungs of mice following double or triple delivery, normalized to beta galactosidase activity

observed upon first delivery. (n = 9 mice per group) *P<0.05.

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DCs generated in the presence of IL-10 are refractory to HD-Ad induced maturation

In the absence of HD-Ad stimulation, CD11c+ DCs generated in the presence of only GM-CSF

(normal DC) are in an immature state, indicated by very low levels of CD86 expression; however

upon incubation with HD-Ad particles; there is massive induction of CD86 expression, with

approximately 80% of DCs undergoing maturation (Figure 4-2A). In contrast, DCs generated by

culturing bone marrow cells in presence of high concentrations of IL-10 (IL-10 modified DCs)

were refractory to HD-Ad induced CD86 expression, for only 12-13% of DCs became CD86+

following stimulation with HD-Ad particles (Figure 4-2A). Furthermore, expression of other

maturation markers such as CD80, MHC II and CCR7, which are induced upon exposure of DCs

to HD-Ad vectors, was also reduced on IL-10 modified DCs compared to normal DCs following

HD-Ad stimulation (Figure 4-2B). Addition of IL-10 to normal DCs prior to HD-Ad stimulation

did not prevent CD86 induction, highlighting the requirement of IL-10 throughout DC

differentiation for generation of DCs refractory to HD-Ad induced maturation (Figure 4-3).

Intracellular FACS staining for adenoviral hexon protein indicated similar propensity of the two

DC populations to uptake HD-Ad particles (Figure 4-2C). However, following HD-Ad

stimulation, the secretion of inflammatory cytokines IL-1β and IL-12, was significantly reduced

from IL-10 modified DCs compared to normal DCs (Figure 4-2D, E). Taken together these

studies indicate that DCs derived from bone marrow cells in presence of IL-10 are refractory to

HD-Ad induced maturation and inflammatory cytokine production.

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141

Figure 4-2: DCs cultured in presence of IL-10 are refractory to HD-Ad induced maturation

(A) Representative FACS plots looking at CD86 expression on DCs cultured in absence of IL-10

(normal DCs) or in presence of IL-10 (IL-10 mod. DC) upon stimulation with HD-Ad vectors

(+HD-Ad). (B) Representative FACS histogram of CCR7, CD80 and MHC II expression. (C)

Representative FACS profile of hexon staining upon incubation of DCs with HD-Ad vectors.

Concentrations of (D) IL-1β and (E) IL-12 released upon stimulation of DCs by HD-Ad vectors.

Representative of six independent experiments. ) *P<0.05.

Figure 4-3: HD-Ad induced maturation of normal DC is not suppressed upon addition of

exogenous IL-10.

Shown are representative FACS plots looking at CD86 expression on normal DCs upon addition

of exogenous IL-10 prior to HD-Ad induced maturation. Representative of five independent

experiments.

CD86

CD

11c

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DCs derived in presence of IL-10 induce generation of IL-10 producing Tr1 regulatory T

cells which suppress HD-Ad induced T cell proliferation

Under basal conditions, without incubation with HD-Ad particles, normal DCs are poor

stimulators of T cell proliferation (Figure 4-4A). In contrast, following HD-Ad incubation,

normal DCs induce robust T cell proliferation, which is also observed upon addition of

exogenous IL-10 to the co-culture. In contrast, following HD-Ad incubation, IL-10 modified

DCs were significantly impaired in their ability to induce T cell proliferation (Figure 4-4A). In

order to further confirm the impairment of HD-Ad pulsed, IL-10 modified DCs to prime HD-Ad

specific T cell proliferation, we also measured frequencies of responding T cells. The frequency

of T cells responding to HD-Ad pulsed IL-10 modified DCs was significantly reduced compared

to the frequency of T cells responding to HD-Ad-pulsed normal DCs (Figure 4-4B). Taken

together these findings indicate that HD-Ad-pulsed IL-10 modified DCs are impaired in driving

HD-Ad-specific T cell proliferation.

We tested the ability of HD-Ad-pulsed IL-10-modified DCs to drive generation of Type 1

regulatory T cells (Tr1 cells), characterized by IL-10 production. HD-Ad pulsed normal DCs

induced IL-10 production in less than 1% of T cells, which increased to 5% of T cells upon

culture with HD-Ad-pulsed IL-10-modified DCs (Figure 4-4C). Furthermore, T cells cultured

with HD-Ad-pulsed IL-10-modified DCs secreted significantly elevated levels of IL-10

compared to T cells cultured with HD-Ad-pulsed normal DCs, further confirming enhanced

propensity of HD-Ad-pulsed IL-10-modified DCs to drive Tr1 differentiation (Figure 4-4D). Tr1

regulatory T cells are characterized by their ability to suppress T cell proliferation. Therefore, we

assessed the ability of Tr1 cells generated upon culture of naive T cells with HD-Ad pulsed IL-

10 modified DCs to suppress HD-Ad induced T cell proliferation. Since surface markers for

identification of Tr1 cells have not been identified, all the T cells cultured with HD-Ad pulsed

IL-10 modified DCs were used as suppressors at very high ratios. Only the T cells isolated from

co-culture with HD-Ad pulsed IL-10 modified DCs had the ability to suppress HD-Ad induced T

cell proliferation, indicating that culture of naive T cells with IL-10-modified DCs led to

induction of Tr1 regulatory T cells with an ability to suppress HD-Ad-induced T cell

proliferation in vitro (Figure 4-4E).

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Figure 4-4: DCs derived in presence of IL-10 suppress T cell proliferation upon exposure to

HD-Ad and instead induce generation of IL-10 secreting Tr1 regulatory T cells.

(A) T cell proliferation upon culture of naive T cells with HD-Ad pulsed DCs with/without

exogenous IL-10 addition. (B) Frequency of responding CD4+ T cells. (C) Representative FACS

histograms looking at IL-10 production by CD4+ T cells cultured in presence of normal or IL-10

modified DCs pulsed with HD-Ad vectors. (D) Histogram comparing levels of IL-10 secreted by

T cells cultured in presence of normal or IL-10 modified DCs pulsed with HD-Ad vectors. (E)

Suppression of T cell proliferation in a culture of naive T cells with HD-Ad pulsed DCs upon

addition of T cells derived from culture of normal or IL-10 modified DCs cultured with naive T

cells, with/without HD-Ad pulsing. Representative of five independent experiments * P<0.05.

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HD-Ad pulsed IL-10 modified DCs suppress pulmonary DC maturation and migration in

vivo upon intranasal challenge with HD-Ad vectors

In order to test the ability of IL-10 modified DCs to induce tolerance towards HD-Ad vectors in

vivo, mice were immunized with either HD-Ad pulsed normal DCs or HD-Ad pulsed IL-10

modified DCs and subsequently we assessed for pulmonary DC maturation and migration

following intranasal challenge with HD-Ad particles. 24 hours after immunization with DCs,

DCs could be observed in the mediastinal lymph nodes (MLN) along with the lungs (data not

shown). Assessment of pulmonary DC maturation was carried out by monitoring expression of

CD86 on pulmonary DCs (Figure 4-5A,B). CD86 expression levels were reduced on pulmonary

DCs from HD-Ad challenged mice which received HD-Ad pulsed IL-10 modified DCs

compared to HD-Ad challenged mice which received HD-Ad pulsed normal DCs, indicating that

delivery of HD-Ad pulsed IL-10 modified DCs induced tolerance which prevented pulmonary

DC maturation following HD-Ad delivery (Figure 4-5A). Delivery of HD-Ad vectors to saline

treated mice or to mice immunized with HD-Ad pulsed normal DCs, led to approximately 40%

increase in proportions of pulmonary CD86+ DCs compared to mice exposed only to saline

(Figure 4-5B). In contrast, only 20% increase in proportions of pulmonary CD86+ DCs was

observed in HD-Ad challenged mice which were immunized with HD-Ad pulsed IL-10 modified

DCs, further confirming the ability of HD-Ad pulsed IL-10 modified DCs to prime immune

tolerance towards HD-Ad vectors.

In order to assess pulmonary DC migration, prior to HD-Ad challenge, mice were given FITC-

Dextran to label pulmonary DCs and following HD-Ad challenge, proportions of FITC-xtran+

DCs were quantified in the mediastinal lymph nodes (MLN) (Figure 4-5C). Upon HD-Ad

challenge, approximately 20% of the DCs in the MLN migrated from the lungs in saline treated

mice and mice immunized with HD-Ad pulsed normal DCs. However, in the lymph nodes of

mice immunized with HD-Ad pulsed IL-10 modified DCs, following HD-Ad challenge, only

10% of the DCs were migratory, indicating an impairment of DCs to migrate to the draining

lymph nodes following HD-Ad challenge. These findings further confirm that delivery of HD-

Ad pulsed IL-10 modified DCs induced tolerance which prevented pulmonary DC migration to

the draining lymph nodes following challenge with HD-Ad vectors.

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147

Figure 4-5: DCs derived in the presence of IL-10 suppress pulmonary DC maturation and

migration in response to HD-Ad vectors.

(A) Representative FACS histogram looking at CD86 expression on pulmonary DCs of mice

exposed to saline, mice exposed to saline followed by HD-Ad vectors (saline then HD-Ad), mice

immunized with HD-Ad pulsed normal DC followed by HD-Ad vectors (Normal DC then HD-

Ad) or mice immunized with IL-10 modified DC followed by HD-Ad vectors (IL-10 mod. DC

then HD-Ad). (B) Proportions of mature CD86+ pulmonary DCs. (C) Proportions of FITC-

Dextran+ pulmonary DCs in the mediastinal lymph nodes of mice which received FITC-Dextran

prior to delivery of HD-Ad vectors. Representative of five independent experiments. * P<0.05.

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HD-Ad-pulsed IL-10-modified DCs suppress T cell infiltration following intranasal

challenge with HD-Ad vectors

In response to a single delivery of HD-Ad vectors to saline-treated mice, approximately 25% of

the cells in BALF were T cells, which were also observed in mice immunized with HD-Ad

pulsed normal DCs (Figure 4-6A). Furthermore, upon three rounds of HD-Ad delivery, the

proportions of BALF T cells increased to 50% (Figure 4-6B). In contrast, T cell proportions in

the BALF of HD-Ad challenged mice immunized with HD-Ad pulsed IL-10 modified DCs were

significantly reduced to only 5-7% upon single HD-Ad delivery and only 10-15% following

three rounds of HD-Ad delivery, highlighting the ability of HD-Ad pulsed IL-10 modified DCs

to mediate tolerance towards HD-Ad vectors (Figure 4-6A,B).

We also measured the ratios of CD4:CD8 T cells in the lungs of mice following challenge with

HD-Ad vectors, with a lower ratio indicative of an increased CD8+ T cell infiltration (Figure 4-

6C). Following a single delivery of HD-Ad vectors, a CD4:CD8 ratio of 1.5 to 2 was observed in

the lungs of saline treated mice or mice immunized with HD-Ad pulsed normal DCs. However,

this ratio was further reduced following three rounds of HD-Ad delivery, indicating an increase

in pulmonary CD8+ T cell infiltration. However, CD4:CD8 ratio of 4 was observed in the lungs

of mice immunized with HD-Ad pulsed IL-10 modified DCs following challenge with a single

dose or three doses of HD-Ad vectors, indicating a reduction in infiltration of CD8+ T cells into

the lung and thereby mediating tolerance induction (Figure 4-6C). Furthermore, we also

measured IFN-γ levels upon stimulation of T cells from MLN with HD-Ad pulsed normal DCs

(Figure 4-6D,E). Following HD-Ad challenge, T cells from saline treated or mice immunized

with HD-Ad pulsed normal DCs secreted elevated IFN-γ levels following HD-Ad stimulation,

which were further elevated upon challenge of mice with three doses of HD-Ad vectors (Figure

4-6D,E). In contrast, T cells from mice immunized with HD-Ad pulsed IL-10 modified DCs and

challenged with a single or three doses of HD-Ad vectors secreted significantly reduced levels of

IFN-γ following HD-Ad stimulation (Figure 4-6D,E). These results indicate that HD-Ad-pulsed

IL-10-modified DCs mediate tolerance induction towards HD-Ad vectors in vivo with a reduced

CD8+ T cell response against HD-Ad vectors.

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150

Figure 4-6: Reduced T cell infiltration is observed in lungs of HD-Ad challenged mice

immunized with HD-Ad pulsed IL-10 modified DCs.

Mice were immunized with HD-Ad-pulsed normal or IL-10-modified DCs and then were

challenged intranasally with a single dose of HD-Ad (5 X 1010 viral particles) vectors or three

doses of HD-Ad vectors separated by three weeks intervals. Proportion of T cells in the

bronchoalveolar lavage (BALF) of mice following (A) first delivery or (B) third delivery of HD-

Ad vectors. (C) Ratio of CD4:CD8 T cells in the lungs of mice. Levels of IFN-γ secreted by T

cells from MLN cells following stimulation with HD-Ad pulsed normal DCs after (D) first or (E)

third delivery of HD-Ad vectors. n=12 mice per group. *P<0.05.

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HD-Ad-pulsed IL-10-modified DCs suppress T cell proliferation following HD-Ad vector

challenge

Following a single delivery of HD-Ad particles, approximately 18-20% of the T cells were

proliferating in the MLN of saline treated mice or mice immunized with HD-Ad pulsed normal

DCs (Figure 4-7A) (Fig. 6a). In contrast, the proportion of proliferating T cells was reduced to

only 6% in the MLN of mice immunized with HD-Ad pulsed IL-10 modified DCs. Following

three rounds of HD-Ad delivery, the proportions of proliferating T cells increased to 35-40% in

the MLN of saline treated mice or mice immunized with HD-Ad pulsed normal DCs, which was

reduced to only 12% in the MLN of mice immunized with HD-Ad pulsed IL-10 modified DCs

(Figure 4-7A). Similarly, proliferation of T cells from MLN in response to HD-Ad vector

stimulation was significantly reduced from mice immunized with HD-Ad pulsed IL-10 modified

DCs compared to saline treated mice or mice immunized with HD-Ad pulsed normal DCs

following either single or three rounds of HD-Ad delivery (Figure 4-7B). Taken together, these

findings confirmed the ability of HD-Ad pulsed IL-10 modified DCs to induce immunological

tolerance towards HD-Ad vectors.

Our in vitro studies identified the ability of HD-Ad pulsed IL-10 modified DCs to drive

induction of IL-10-producing Tr1 cells. In order to confirm induction of Tr1 cells in vivo, we

measured IL-10 levels following stimulation of T cells from MLN of mice immunized mice with

HD-Ad vectors (Figure 4-7C). T cells from MLN of saline-treated mice or mice immunized with

HD-Ad pulsed normal DCs secreted very low levels of IL-10 following HD-Ad stimulation. In

contrast, T cells from MLN of mice immunized with HD-Ad pulsed IL-10 modified DCs

secreted significantly elevated levels of IL-10 following HD-Ad stimulation, indicating that

delivery of HD-Ad pulsed IL-10 modified DCs was able to drive Tr1 generation in vivo.

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Figure 4-7: Reduced T cell proliferation in response to pulmonary HD-Ad challenge is

observed in mice immunized with HD-Ad-pulsed IL-10-modified DCs.

Mice were immunized with HD-Ad-pulsed normal or IL-10-modified DCs and then were

challenged intranasally with a single dose of HD-Ad vectors (5 X 1010 viral particles) or three

doses of HD-Ad vectors separated by three weeks intervals. (A)Representative FACS histogram

looking at BrdU+ cells among CD3+ T cells in the mediastinal lymph nodes (MLN). (B)

Proliferation of T cells from MLN upon stimulation with HD-Ad pulsed normal DCs in vitro. (C)

Levels of IL-10 released upon stimulation of T cells from MLN upon stimulation with HD-Ad

pulsed normal DCs in vitro. n=12 mice per group. *P<0.05.

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HD-Ad-pulsed IL-10-modified DCs suppress the antibody response following HD-Ad

vector challenge

Following single delivery of HD-Ad vectors, low levels of anti-Ad antibodies were observed in

the serum and BALF of saline-treated mice or mice immunized with HD-Ad pulsed normal DCs,

which were significantly reduced both in the serum and BALF of mice immunized with HD-Ad

pulsed IL-10 modified DCs (Figure 4-8). Upon three rounds of HD-Ad vector delivery, several

folds higher titers of anti-Ad antibodies were observed in the serum and BALF of saline-treated

mice or mice immunized with HD-Ad-pulsed normal DCs compared to single round of HD-Ad

vector delivery. However, the levels of anti-Ad antibody titer were significantly reduced both in

the serum and BALF of mice immunized with HD-Ad-pulsed IL-10-modified DCs following

three rounds of HD-Ad challenge (Figure 4-8). Taken together, these findings indicate that

delivery of HD-Ad-pulsed IL-10-modified DCs induce tolerance characterized by suppression of

antibody response against HD-Ad vectors.

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Figure 4-8: Reduced anti-Ad antibodies in response to pulmonary HD-Ad challenge is

observed in mice immunized with HD-Ad-pulsed IL-10-modified DCs.

Mice were immunized with HD-Ad pulsed normal DCs or HD-Ad-pulsed IL-10-modified DCs

and then were challenged intranasally with a single dose of HD-Ad vectors (5 X 1010 viral

particles) or three doses of HD-Ad vectors separated by three weeks intervals. Histogram

comparing anti-Ad antibody levels in (A) serum and (B) bronchoalveolar lavage fluid (BALF)

following HD-Ad challenge. n =8 mice per group. *P<0.05.

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Sustained gene expression is observed in the lungs of mice which receive HD-Ad pulsed IL-

10 modified DCs following multiple rounds of HD-Ad mediated gene delivery

The aim of tolerance induction towards HD-Ad vectors is to prevent loss of gene expression

associated with multiple rounds of HD-Ad vector readministrations. Therefore, to assess

sustained gene expression following multiple rounds of HD-Ad gene delivery in mice tolerant

towards HD-Ad vectors, we devised a gene delivery protocol as shown in Figure 4-9A. Single

delivery of HDK18LacZ was associated with robust β-galactosidase activity in the lung along

with X-gal staining all throughout the airways (Figure 4-9B,C). However, in non-tolerant mice

which received multiple rounds of readministrations, β-galactosidase activity in the lung was

significantly reduced with very faint LacZ staining in the airways (Figure 4-9B,C). Similar

results were observed in tolerant (OVA) mice which received ovalbumin (OVA)-pulsed IL-10

modified DCs followed by multiple rounds of readministrations, indicating that delivery of

OVA-pulsed IL-10-modified DCs failed to induce tolerance towards HD-Ad vectors. In contrast

β-galactosidase activity was only slightly reduced in the lungs of mice which received HD-Ad

pulsed IL-10 modified DCs following multiple rounds of vector delivery and was significantly

higher than in the non-tolerant group or the group with tolerance towards OVA. Taken together

these findings indicate that delivery of IL-10-modified HD-Ad-pulsed DCs induces tolerance

towards HD-Ad vectors which prevents loss in gene expression associated with readministrations

of HD-Ad vectors to the lung.

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Figure 4-9: Gene expression following multiple rounds of pulmonary HD-Ad vectors is

sustained in mice immunized with HD-Ad pulsed IL-10 modified DCs.

(A) Outline of the readministration strategy. (B) β-galactosidase activity in the lungs of HD-Ad

challenged mice. Results are shown as mean β-galactosidase activity (RLU/µg of protein) ± SD.

(C) X-gal staining of the lungs following HD-Ad delivery. N=15 per group. *P<0.05.

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4.5 Discussion

The adaptive immune response against HD-Ad vectors is a major barrier in limiting transgene

expression following vector readministration to the lung. This was confirmed in Rag-deficient

mice, which have a defective adaptive immune response, for upon HD-Ad vector

readministration, transgene expression was sustained in the lungs of these mice compared to

wild-type counterparts. Moreover, upon multiple rounds of HD-Ad vector readministration to the

lung, there was an increase in CD8+ T cell infiltration to the lung along with several fold

increases in T cell proliferation and anti-Ad antibody titers, confirming an association of HD-Ad

vector readministration with magnification of adaptive immune response towards these vectors.

Therefore, it is important to devise strategies to induce tolerance towards HD-Ad vectors so that

vector readministration can be carried out and gene expression can be sustained. Our results

show that generation of DCs in the presence of high concentrations of IL-10 (IL-10-modified

DCs) leads to generation of tolerogenic DCs that are impaired in undergoing maturation in

response to HD-Ad vectors and instead of priming T cell response against HD-Ad vectors,

induce generation of IL-10-secreting Tr1 Tregs with an ability to suppress HD-Ad induced T cell

proliferation. Moreover, upon delivery of HD-Ad-pulsed IL-10-modified DCs to mice,

maturation of pulmonary DCs in response to intranasal HD-Ad delivery is impaired. Moreover,

upon several rounds of HD-Ad challenge, in tolerant mice, the T cell response towards HD-Ad

vectors is significantly diminishment along with a significant reduction in anti-Ad antibody

titers, confirming the ability of HD-Ad-pulsed IL-10-modified DCs in inducing long term

tolerance towards HD-Ad vectors.

The effects of IL-10 on DCs have been investigated in maintaining the immature status of DCs.

Initial studies indicated that IL-10 results in inhibition of DC-driven IFN-γ production by

purified CD4+ and CD8+ T cells (Macatonia et al., 1993). Addition of IL-10 to in vitro cultures

of dermal DCs has been to shown to suppress dermal DC maturation along with an impairment

of their ability to activate T cells (Mitra et al., 1995). Moreover, IL-10 treatment also results in

enhanced phagocytosis by DCs, which when coupled with downregulation of costimulatory

molecules, results in an immature DC phenotype. The effects of IL-10 on DCs correlate well

with inhibition of primary T cell responses, for addition of anti-IL-10 antibodies promotes DC

maturation (Corinti et al., 2001). Furthermore, retroviral transduction of EBV IL-10 into DCs has

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also been shown to result in immature phenotype (Takayama et al., 1998). Recent studies have

highlighted the ability of immature DCs to drive tolerance induction, for delivery of IL-10-

differentiated DCs can mediate tolerance induction in asthmatic mice through generation of

Tregs (Huang et al., 2011; Lu et al., 2011).

In our study, we were able to induce tolerance towards HD-Ad vectors using IL-10-modified

DCs pulsed with HD-Ad vectors. The Tregs generated in vitro were IL-10-secreting and not

Foxp3+, and were able to suppress HD-Ad induced T cell proliferation. This was likely due to

generation of Tr1 regulatory T cells, for in vitro generation of human tolerogenic DCs using IL-

10 have been shown to prime differentiation of antigen-specific Tr1 cells (Gregori et al., 2011).

T cells from the draining lymph of HD-Ad-challenged mice immunized with HD-Ad pulsed IL-

10 modified DCs secreted elevated levels of IL-10, indicating that HD-Ad-pulsed DCs primed

induction of Tr1 cells in vivo. Transfer of Tr1 regulatory T cells have been shown to abrogate

DC-mediated immune responses in an IL-10-dependent manner (Zhang et al., 2005). Delivery of

HD-Ad-pulsed IL-10-modified DCs to mice suppressed pulmonary DC maturation and migration

in response to HD-Ad vectors. Since DCs have a short life-span, it is likely that Tr1 cells induced

by HD-Ad-pulsed IL-10-modified DCs, secreted IL-10 in response to intranasal challenge with

HD-Ad vectors, which had an anti-inflammatory affect on pulmonary DCs, thereby suppressing

their maturation (Morelli and Thomson, 2007). Moreover, Tr1 cells could also directly suppress

DC maturation, for Tregs have been shown to directly suppress DC maturation via binding of

lymphocyte activation gene-3 (LAG) on the Treg surface to MHC II on the DC surface (Liang et

al., 2008).

IL-10-modified DCs were also able to significantly inhibit HD-Ad-induced T cell proliferation in

mice. Furthermore, in tolerant mice, CD8+ T cell infiltration into the lungs was significantly

reduced compared to non-tolerant mice, further highlighting the ability of IL-10 modified DCs to

induce tolerance towards HD-Ad vectors. This was likely due to impairment of pulmonary DC

migration upon HD-Ad delivery, which prevented induction of the T cell response against HD-

Ad vectors and also due to induction of HD-Ad Tregs which further suppressed T cell response

(Lambrecht, 2001). The T cell response is critical for induction of B cell mediated antibody

responses (Parker, 1993). Therefore, suppression of the T cell response against HD-Ad vectors

upon immunization with HD-Ad-pulsed IL-10-modified DCs also suppressed generation of anti-

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Ad antibody titers. In accord with induction of tolerance, transgene expression was enhanced

significantly in the lungs of mice immunized with HD-Ad pulsed IL-10 modified DCs.

Our findings also show that generation of tolerance was specific towards HD-Ad vectors, since

delivery of IL-10-modified DCs pulsed with OVA was not associated with sustained gene

expression following multiple rounds of HD-Ad readministration. This indicates that induction

of tolerance towards HD-Ad vectors will not compromise immunity against other pathogens.

Further studies are needed to identify the immunodominant HD-Ad derived epitopes against

which an immune response is potentiated following vector readministrations. Subsequently,

antigen specific tolerance can be induced towards the specific epitopes to enhance the antigen

specificity of tolerance induction. This strategy to induce antigen specific tolerance has

important implications in regenerative medicine because this strategy can be applied to other

gene therapy vectors, protein and cell therapy along with transplantation.

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Chapter 5

Induction of antigen specific tolerance by apoptotic dendritic cells

The contents of this chapter have been published in the following journals:

Rahul Kushwah and Jim Hu. Dendritic cell apoptosis: regulation of tolerance versus immunity.

Journal of Immunology. 2010 Jul; 185(2):795-802.

Rahul Kushwah, Jing Wu, Jordan Oliver, George Jiang, Jinyi Zhang, Katherine A Siminovitch

and Jim Hu. Uptake of apoptotic DC converts immature DC into tolerogenic DC that induce

differentiation of Foxp3+ Treg. European Journal of Immunology. 2010 Apr;40(4):1022-35.

Rahul Kushwah, Jordan Oliver, Jinyi Zhang, Katherine A Siminovitch and Jim Hu. Apoptotic

dendritic cells induce tolerance in mice through suppression of dendritic cell maturation and

induction of antigen-specific regulatory T cells. Journal of Immunology. 2009 Dec

1;183(11):7104-18.

Acknowledgment: I would like to acknowledge Jing Wu for performing real-time qPCR

analysis (Figures 5-5A-E, 5-7A-B) and Jordan Oliver (Figure 5-14C) for analyzing airway

histology. I would also like to acknowledge, Drs. Jinyi Zhang and Katherine Siminovitch for

reading the manuscript and providing us with OT-II transgenic mice. Also, thanks to George

Jiang for coordinating breeding of OT-II mice for our studies.

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5.1 Abstract

Dendritic cell (DC) apoptosis has been shown to play a role in maintaining a balance between

tolerance and immunity. Environments with significant DC apoptosis are immunosuppressive,

promote regulatory T cell (Treg) generation, and display functional impairment of remaining

DCs, indicating that DC apoptosis might contribute to tolerance. The importance of DC

apoptosis is further highlighted by studies identifying defects in DC apoptosis as triggers of

autoimmune diseases. Therefore, I explored the role of DC apoptosis in regulation of tolerance to

identify if DC apoptosis can be employed as a tool for generation of antigen-specific tolerance.

Objective of Study: I examined the influence of apoptotic DCs and necrotic DCs on viable DCs

in vitro and went on to assess the effects of apoptotic DCs in vivo.

Summary of Results: I show that immature viable DCs have the ability to take up apoptotic DC

as well as necrotic DC without it being recognized as an inflammatory event by immature viable

DC. However, the specific uptake of apoptotic DC converted immature viable DC into

tolerogenic DC, which were resistant to LPS-induced maturation. In contrast, these tolerogenic

DC secreted increased levels of TGF-β1, which induced differentiation of naïve T cells into

Foxp3+ regulatory T cells (Treg). Furthermore, induction of Treg differentiation only occurred

upon uptake of apoptotic DC and not upon uptake of apoptotic splenocytes by viable DC,

indicating that it is specifically the uptake of apoptotic DC that gives viable immature DC the

potential to induce Foxp3+ Treg.

In vivo delivery of apoptotic DCs to mice, results in their uptake by viable DCs, which

suppresses the ability of viable DCs to undergo maturation and subsequent migration to the

lymph nodes in response to LPS. Additionally, delivery of apoptotic DCs to LPS inflamed lungs

results in resolution of inflammation, which is mediated by the ability of apoptotic DCs to

suppress response of viable DCs to LPS. Additionally, apoptotic DCs also induce TGF-β1

secretion in the mediastinal lymph nodes, which results in expansion of Foxp3+ Tregs. Most

importantly, I show that delivery of apoptotic DCs followed by ovalbumin (OVA) in Complete

Freund’s adjuvant (CFA) to mice suppresses the T cell response to OVA and instead induces de

novo generation of OVA-specific Tregs. Furthermore, delivery of apoptotic DCs followed by

OVA in CFA results in expansion of Tregs in T-cell receptor transgenic (OT-II) mice. These

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findings demonstrate that apoptotic DCs are taken up by viable DCs in vivo, which promotes

tolerance through suppression of DC maturation and induction of Tregs.

Conclusion: Uptake of apoptotic DCs converts viable DCs into TGF-β1-secreting tolerogenic

DCs which drive differentiation of Foxp3+ Tregs. Delivery of apoptotic DCs followed by

antigen delivery results in Treg mediated antigen-specific tolerance generation in mice.

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5.2 Introduction Dendritic cells (DCs) are potent antigen-presenting cells with the ability to initiate T cell

responses. DCs are present in the peripheral tissues where they are constantly engulfing antigens

along with dying cells. However, the ability to initiate a T cell response depends on the ability of

DC to undergo maturation and subsequently migrate to draining lymph nodes, where in the

presence of appropriate cytokines, T cells can be differentiated into a particular lineage in an

antigen-specific manner. In addition to priming effector T cell responses, DCs also play a role in

induction of tolerance.

DCs are well positioned in peripheral tissues to capture foreign antigens. DC are phagocytic and

can ingest apoptotic cells, and hence are affected by the death of other cells in close proximity

(Huang et al., 2000; Sauter et al., 2000; Steinman et al., 2003). Clearance of apoptotic cells

results in their removal from tissues, and provides protection from release of pro-inflammatory

contents. According to the danger hypothesis, it has been suggested that injured cells, analogous

to necrotic cells provide danger signals which induce activation of DCs and subsequent induction

of T cells (Kono and Rock, 2008). In contrast to necrotic cells, apoptotic cells are thought to be

cleared rapidly without any immunological response (Peng et al., 2007; Sauter et al., 2000).

Studies have identified necrotic cells acting as adjuvants whereas apoptotic cells have been

reported as immunogenic (Feng et al., 2002; Johansson et al., 2007; Winau et al., 2006) or

immunosuppressive (Chen et al., 2001; Fadok et al., 1998).

DC apoptosis in itself is an important event for maintenance of tolerance. Defects in DC

apoptosis have been linked to development of autoimmunity with systemic autoimmune diseases

modeled in transgenic mice harboring defects in DC apoptosis (Chen et al., 2006) but not in mice

with apoptosis defects in T and B cells (Doerfler et al., 2000; Newton et al., 1998; Walsh et al.,

1998). However, it is unclear how defects in DC apoptosis can trigger autoimmune responses.

Furthermore, spontaneous DC apoptosis has been reported in sepsis as well as breast cancer

patients with its significance being unclear (Ito et al., 2006; Pinzon-Charry et al., 2007; Pinzon-

Charry et al., 2006). Most patient deaths associated with sepsis occur at later time points and are

associated with prolonged immunosuppression (Zeni et al., 1997). In this later stage there is

marked apoptosis of DC, with no effects on macrophage and neutrophil apoptosis. In addition,

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immunostimulants such as CpG DNA inhibit DC apoptosis (Park et al., 2002), while the

deficiency of pro-apoptotic Bim protein in DC results in autoimmunity(Chen et al., 2007).

Immature DCs are highly phagocytic and have the ability to ingest entire cells. Immature DC

have the ability to acquire protein complexes or soluble antigen using many different pathways

such as macropinocytosis, endocytosis and even through ingestion of entire cells. Previous

studies have reported that ingestion of necrotic cells is recognized as immunostimulatory,

whereas ingestion of apoptotic cells appears to be an immunologically null event (Gallucci et al.,

1999; Sauter et al., 2000). Despite the importance of DC apoptosis in the immune response,

studies have not investigated the effects of ingestion of apoptotic or necrotic DCs by viable DCs.

Therefore, we investigated the effects of DC apoptosis on viable DCs by first performing in vitro

studies where apoptotic/necrotic DCs were cultured with viable DCs and second by performing

in vivo studies in mice through direct injection of apoptotic DCs.

Our in vitro studies show that viable immature DCs have the ability to uptake apoptotic DCs.

The uptake of apoptotic DC or necrotic DC is recognized as an immunologically null event.

However, it is the uptake of apoptotic DC that suppresses subsequent maturation of viable DC in

response to LPS and results in upregulation of TGF-β2 and preferential secretion of TGF-β1,

dependent on mTOR pathway, which mediates induction of naïve T cells into Foxp3+ Treg. In

contrast, the uptake of apoptotic splenocytes by viable immature DC does not result in TGF-β1

secretion, nor does it result in induction of Foxp3+ Treg. Altogether, the findings suggest that the

uptake of apoptotic DC by viable DC provides a potential to induce Foxp3+ Treg.

Our data further shows that delivery of apoptotic DCs in mice results in their rapid uptake by

viable DCs both in the spleen as well as the draining lymph nodes. Although the uptake of either

necrotic or apoptotic DCs by viable DCs in vivo does not affect their phenotype, it has an effect

on subsequent responses to LPS. Apoptotic DCs are able to suppress subsequent LPS induced

DC maturation and DC migration from the periphery to the lymph nodes. Furthermore, apoptotic

DCs are able to suppress LPS-induced airway inflammation when they are given together with

LPS or after LPS delivery. This has to do with the ability of apoptotic DCs to suppress IL-12

production by viable DCs in the lymph nodes and their ability to instead promote secretion of

TGF-β1 from the draining lymph nodes, which contributes to immunosuppression and induces

expansion of Foxp3+ Tregs. In addition, we also show that delivery of apoptotic DCs followed by

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OVA-CFA in mice results in induction of tolerance, via generation of OVA-specific Tregs. Taken

together, these findings clearly demonstrate that apoptotic DCs can be potentially used for

induction of tolerance via generation of antigen-specific Tregs and for suppression of pre-existing

inflammation.

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5.3 Materials and Methods

Mice, antibodies and other reagents

C57BL/6 mice were purchased from Charles River Laboratories (St. Constant, QC) and B6.SJL

mice were purchased from Taconic (Hudson, NY) and maintained as per guidelines of SickKids

animal facilities. All the animal studies were reviewed and approved by the SickKids

Institutional Committee for humane use of laboratory animals. OT-II mice were purchased from

Jackson Laboratories (Bar Harbor, ME). OT-II transgenic mice express the mouse α-chain and β-

chain T-cell receptor that pairs with the CD4 coreceptor and is specific for chicken ovalbumin

323-339 in the context of I-Ab. All the antibodies used were directed against mouse antigens.

The following antibodies were purchased from eBioscience (San Diego, CA): CD11c PE, CD11c

PE-Cy7, CD86 PE, CD80 PE, MHC II PE, IL-10 Alexa647, IL-12 APC, CD25 PE-Cy7, BrdU

FITC, IL-17 PE, Foxp3 PE along with neutralizing IL-4 and IFN-γ antibody and the following

from BD Biosciences (Mississauga, ON): CD11c-FITC, CD4-FITC, CD25-PE, CD3-PE. Anti-

TGF-β neutralizing antibody (MAB1835) was obtained from R&D Systems (Minneapolis, MN).

Isotype control IgGs were obtained from eBioscience and/or Serotec (Raleigh, NC). CFSE was

obtained from Molecular Probes (Burlington, ON); BrdU, Ovalbumin, Cytochalasin D,

Rapamycin and PI were obtained from Sigma-Aldrich (Oakville, ON). GM-CSF was obtained

from R&D Systems (Minneapolis, MN). IL-6 and TGF-β were obtained from Peprotech (Rocky

Hill, NJ). Cell proliferation ELISA based on BrdU incorporation and chemiluminescent

detection was obtained from Roche (Laval, QC). Aerrane was obtained from Baxter

(Mississauga, ON). Triton-X buffer was used for permeabilization for intracellular Foxp3

staining.

Generation of bone marrow derived dendritic cells

Bone marrow cells were isolated from tibia and femurs of adult mice and cultured in the

presence of GM-CSF as described (Kushwah et al., 2008). On day 3, the non-adherent

granulocytes, and T and B cells were gently removed and the respective fresh media were added,

and two days later the loosely-adherent proliferating DC aggregates were dislodged and re-

plated. On day 7, the released, weakly-adherent cells were harvested. 85-90% of the cells were

confirmed to be CD11c+ DCs via FACS analysis.

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CFSE labelling of DCs

DCs were harvested and suspended in balanced salt solution (BSS) containing 1 μM CFSE at a

concentration of 106 cells/ml. Subsequently, the cells were incubated at 37 C for 10 min. Fetal

calf serum was added to a concentration of 5% and the cells were washed twice.

Isolation of naïve CD4+CD25- T cells

Naïve CD4+CD25-CD62L+ T cells were isolated from spleens of wild-type mice or OT-II mice

using CD4+ CD62L+ naïve T-cell isolation kit in conjunction with MACS columns from Miltenyi

Biotec Inc. (Auburn, CA), according to the manufacturer’s instructions.

Induction of dendritic cell apoptosis and necrosis

Dendritic cells were cultured on a 6-well dish and irradiated for 2 min with a UV

transilluminator, with a peak intensity of 9000 mW/cm2 at the filter surface and a peak emission

of 313 nm. Induction of apoptosis was confirmed using apoptosis, necrosis and healthy cell

quantification kit (Biotium, Hayward, CA), following the manufacturer’s instructions. Necrosis

was induced by pelleting cells followed by 3 cycles of freeze and thaw. A similar protocol was

used for induction of splenocyte apoptosis, which were isolated from spleens of C57BL/6 mice

as described (Kushwah et al., 2008).

Live DC and apoptotic DC/splenocytes or necrotic DC co-culture experiments

Bone marrow-derived immature live DC (100,000 cells/well) were co-cultured with

apoptotic/necrotic DC or apoptotic splenocytes (1,000,000 cells/well). In some experiments,

Cytochalasin D (0.8 μg/ml) was added to inhibit phagocytosis. In order to inhibit mTOR

signalling pathway, rapamycin (100nM) was added to the co-culture of apoptotic DC with viable

DC. 24 hours later, cells were exposed to 1μg/ml LPS and FACS analysis was performed.

In vitro generation of Treg

Live DC (100,000 /well) were incubated with apoptotic / necrotic DC or apoptotic splenocytes

(1,000,000 cells/well) at a ratio of 1:10 and then pulsed with OVA, followed by co-culture with

naïve CD4+ T cells (250,000/well) from OT-II mice. Five days later, CD4+ T cells were analyzed

for foxp3 expression via FACS. In some experiments, neutralizing TGF-β antibody was added

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(50μg/ml). In transwell experiments, DC were added to the top chamber and naïve CD4+ T cells

from C57BL/6 mice were placed in the lower chamber and stimulated with plate bound CD3 and

soluble CD28 antibodies.

In vitro suppression assay

OVA-pulsed (0.5 mg/ml) DC were used as stimulators and naïve OT-II CD4+ T cells were used

as responders. The stimulators (2.5 X 105 cells/well) and responder cells (2.5 X 104 cells/well)

were cultured in 96-well round-bottom plates at a ratio of 10:1 and suppressors (CD25+) isolated

from co-culture of OT-II naïve T cells and OVA-pulsed viable DC that had taken up apoptotic

DC or from draining lymph nodes of mice, were added at a ratio of 1:2, 1:10 or 1:30.

Proliferation was assessed at day 4 of co-culture using BrdU cell proliferation assay following

manufacturer’s instructions (Roche, QC).

To isolate CD4+CD25+ T cells, lymph nodes from immunized mize were isolated and a single

cell preparation was prepared. Cell suspension was first enriched for CD4+ T cells by depletion

of non-CD4+ T cells using CD4+ T cell isolation kit (Miltenyi Biotec) in conjuction with LS

columns. The flow-through consisted of enriched CD4+ T cells. Cells were stained for CD25 and

cell-sorting was carried out to isolate CD4+CD25hi T cells. FACS analysis confirmed that greater

than 94% of the T cells were CD4+CD25+.

In vitro generation of Th17 cells

Naïve CD4+CD25- T cells were cultured for 4 days in the presence of LPS-treated live DC, LPS-

treated live DC incubated with necrotic DC or LPS-treated live DC incubated with apoptotic DC,

and were activated with plate-bound anti-CD3 and soluble anti-CD28 antibodies in the presence

of 5 ng/ml IL-6, 2.5 ng/ml TGF-β, 10 μg/ml anti-IL-4 and 10 μg/ml anti-IFN-γ.

Cytokine assays.

We quantified levels of TGF- β1 in BALF and culture supernatants by ELISA using commercial

kits following the manufacturer’s instructions (TGF-β1 kit, R&D Systems, Minneapolis, MN). 5

X 105 cells/well were cultured in X-VIVO 20 serum-free medium (Cambrex) and 48 hours later,

culture supernatants were used for ELISA.

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Real-time PCR

TaqMan real-time RT-PCR was carried out as described previously using primer sequences

listed in table 5-1(Wu et al., 2008).

Table 5-1: Primer sequences for assessment of cytokine induction in viable DCs upon

apoptotic DC uptake

NAME SEQUENCE REFERENCE

TGF-beta1

forward: 5'-CAACAATTCCTGGCGTTACCTTGG-3'

Am J Physiol Renal Physiol 295: F118-F127, 2008

reverse: 5'-GAAAGCCCTGTATTCCGTCTCCTT-3'

TGF-beta2

forward: 5'-CTTAACATCTCCCACCCAGC-3'

J. Immunol., 179: 6325-6335, 2007

reverse: 5'-TCACCACTGGCATATGTAGA-3'

IL-6

forward: 5'-CCCAACAGACCTGTCTATACC-3'

This study

reverse: 5'-CTGCAAGTGCATCATCGTTGTTC-3'

IL-1beta

forward: 5'-GACAGTGATGAGAATGACCTG-3'

This study

reverse: 5'-CCACAGCCACAATGAGTGATA-3'

IL-12p35

forward: 5'-CACCCTTGCCCTCCTAAACC-3'

JEM, vol201, No. 9, 1435-1446, 2005

reverse: 5'-CACCTGGCAGGTCCAGAGA-3'

IL-12p40

forward: 5'-ACAGCACCAGCTTCTTCATCAG-3'

JEM, vol201, No. 9, 1435-1446, 2005

reverse: 5'-TCTTCAAAGGCTTCATCTGCAA-3'

TNF-alpha

forward: 5'-CATCTTCTCAAAATTCGAGTGACAA-3'

JEM, vol204, No. 6, 1487-1501

reverse: 5'-TGGGAGTAGACAAGGTACAACCC-3'

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In vivo assays to assess suppression by apoptotic DCs

Wild-type C57BL/6 mice were injected with 100 μl saline alone or 4 X 106 apoptotic/necrotic

DCs in 100 μl volume saline in footpads and 6 hours later, 1 μg LPS was injected in a volume of

50 μl. FITC-Dextran was injected prior to LPS delivery, to label DCs in the periphery. 24 hours

later, popliteal lymph nodes were isolated and assessed for migration of FITC-Dextran+CD11c+

DCs along with CD86 expression on CD11c+ DCs via FACS analysis. Furthermore, experiments

were also repeated by delivering CFSE-labelled apoptotic or necrotic DCs, followed by

assessment of CD86, CD80 and MHC II expression on CFSE+CD11c+ DCs (indicative of DCs

that had taken up apoptotic/necrotic DCs). Additionally, experiments were also performed to

look at IL-12+ DCs in the popliteal lymph nodes. Single cell suspensions from popliteal lymph

nodes were prepared and cells were cultured in media containing Brefeldin A (1μg/ml) for 3

hours. Subsequently, cells were stained for CD11c, fixed and permeabilized using buffer

containing saponin and stained with IL-12 antibody or isotype control, followed by subsequent

analysis on a FACSCalibur.

In vivo lung assays

Mice that were 8-11 wks of age, were lightly anesthetized by Aerrane inhalation, and 100 μl of

saline alone or 20 μg of LPS in a volume of 100 μl saline with or without apoptotic DCs was

delivered intranasally. 10 X 106 apoptotic DCs were delivered together with LPS or 1 day after

LPS delivery. 6 hrs or 24 hrs later, mice were sacrificed and bronchoalveolar lavage fluid

(BALF) along with the mediastinal lymph nodes were isolated. In order to isolate BALF, mouse

tracheas were briefly exposed and were cannulated with a catheter and a syringe, and using ice-

cold PBS with 5mM EDTA, the lower respiratory tract was rinsed 6 times using 1 ml volume to

collect inflammatory cells from the airspaces. Total cell counts in BALF were determined by

light microscopy and supernatants were used for determination of TGF- β1 levels by ELISA.

Single cell suspensions were prepared from mediastinal lymph nodes which were stained with

CD4 and Foxp3 to assess percentage of Tregs via FACS analysis as described (Kushwah et al.,

2008). Additionally, in another set of mice, 48 hours after delivery, mice were sacrificed and

lungs were isolated for histological analysis. Formalin-fixed, paraffin-embedded mouse lung

tissue samples were sectioned at 4 μm and stained with haematoxylin and eosin (H&E) for

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histological examination of LPS-induced inflammation under a light microscope as described

previously in chapter 3.

Immunization with OVA and assessment of T cell proliferation in vivo

Mice were immunized with 100 µg OVA in a volume of 100 µl saline, emulsified in an equal

volume of CFA. BrdU labeling solution was prepared in PBS with a final concentration of 5

mg/ml. After treatment with OVA-CFA, mice were given daily intraperitoneal injections of

BrdU (1 mg/mouse) for 7 days. On 7th day, mice were sacrificed and draining lymph nodes were

isolated. Single cell suspension was prepared and cells were staining were stained for CD3 and

BrdU, as described previously in chapter 2.

Statistical analysis

Statistical analyses were performed using Student t-test to compare two groups and ANOVA to

compare multiple groups (SPSS 16.0). Significance was set at P<0.05.

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5.4 Results

Induction of DC apoptosis by UV radiation in vitro

Bone-marrow-derived DC were treated with UV light and apoptosis induction was assessed at 1

hour and 6 hours after UV treatment. Prior to UV treatment, cells were mostly positive for

Hoechst 3342 (a cell permeant DNA binding stain, blue) with very few cells being positive for

annexin V (a phosphatidylserine binding protein, green), indicative of live DC. One hour after

UV treatment, majority of cells were positive for both annexin V as well as Hoechst 3342, with

very few cells positive for Ethidium homodimer (EH) (a nuclei probe, impermeant to live or

apoptotic cells, red) (Figure 5-1A). In these cells, there was translocation of phosphatidylserine

on the membrane as indicated by positive annexin V staining, but the membrane integrity was

still maintained, since they were mostly negative for EH stain; hence they can be classified as

apoptotic cells. In contrast, 6 hours after UV treatment, there was a pronounced increase in EH

positive cells, indicating that the membrane integrity was compromised. However, these cells

were also positive for annexin V (Figure 5-1A). Therefore, these cells can be classified as late

apoptotic cells. In order to further confirm apoptosis in a quantitative manner, 1 or 6 hours after

UV treatment, DC were stained with annexin V and propidium iodide (PI), and apoptosis was

assessed via FACS analysis. Prior to UV treatment, approximately 10% of DC were annexin

V+PI-, whereas, 1 hour after UV treatment approximately 45% of DC were annexin V+PI-,

indicative of apoptotic cells and confirming our above findings (Figure 5-1B). At 6 hours post

UV treatment, approximately 80% of cells were annexin V+PI+, indicating that these cells were

in late apoptosis (Figure 5-1B). This proportion of annexin V+PI+ cells further increased to close

to 92.4%, 8 hours after UV treatment (Figure 5-1B). Therefore, we chose to use cells

immediately after UV treatment as apoptotic DC for further experiments. Similarly, apoptosis

was induced in splenocytes via UV radiation and 1 hour after UV treatment, approximately 40%

of splenocytes were annexin V+PI-, indicative of apoptotic splenocytes (Figure 5-1C).

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Annexin V - FITC

Pro

pid

ium

Iod

ide

Annexin V - FITC

Prop

idiu

m Io

dide

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Figure 5-1: UV radiation induces apoptosis in DC and splenocytes.

(A) Detection of apoptotic DC, 1 hr, and 6 hr after UV exposure, by staining with annexin V-

FITC, EH and Hoechst 33342. (B) FACS analysis for assessment of late and apoptotic DC by

staining with annexin V-FITC and PI. (C) Splenocytes were UV-irradiated and FACS analysis

was conducted for assessment of apoptotic splenocytes by staining with annexin V-FITC and PI.

Data are representative of 4-5 independent experiments.

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Uptake of apoptotic DC by live DC in vitro

In order to assess uptake of apoptotic DC by viable DC, apoptotic DC were labelled with CFSE

and incubated with immature viable DC. Eight hours later, FACS analysis was performed to

assess uptake of CFSE-labelled apoptotic DC by live DC (PI-CD11c+) (Figure 5-2A). Results

indicate that approximately 50% of viable DC had taken up apoptotic DC (Figure 5-2). In order

to confirm that there were no contaminating CFSE+ PI- apoptotic DC, a parallel experiment was

performed where apoptotic DC were labelled with CFSE, cultured for 8 hours, and subsequently

stained with PI; approximately 98% of the DC were PI+, indicating that gating for PI- cells,

would gate out any CFSE+ apoptotic DC. Furthermore, in order to distinguish binding of

apoptotic DC to live DC from uptake of apoptotic DC by live DC, the co-culture experiments

were carried out in the presence of cytochalasin D, a known inhibitor of phagocytosis (Figure 5-

2). In the presence of cytochalasin D, only 12% of the cells were CFSE+, which is probably

indicative of apoptotic DC that bound to live DC. Collectively, the results indicate that immature

viable DCs have the ability to phagocytose apoptotic DC.

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A

Pro

pid

ium

Iod

ide

CFSEForward scatter

Freq

uenc

y

No cytochalasin D

CD11c MHC II

Freq

uenc

y

B

Freq

uenc

y

CFSE

Pro

pid

ium

Iod

ide

Forward scatter

Cytochalasin D

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Figure 5-2. Viable DC uptake apoptotic DC in vitro and this uptake is inhibited by

cytochalasin D.

CFSE-labelled apoptotic DC were incubated with viable immature DC in (A) absence or (B)

presence of cytochalasin D at a ratio of 10:1 and 8 hours later, FACS analyses were conducted to

assess uptake of CFSE+ apoptotic DC by viable DC. Viable DC were gated based on PI-

exclusion and the proportion of CFSE+ cells was assessed among PI- viable DC. DC phenotype is

confirmed by staining with CD11c and MHC class II (A). Data are representative of 3

independent experiments.

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Viable DC do not undergo maturation upon uptake of apoptotic or necrotic DC in vitro

In order to assess the effects of apoptotic or necrotic DC on viable DC, viable immature DCs

were incubated with mature apoptotic, immature apoptotic and necrotic DC respectively. In order

to generate mature apoptotic DC, bone-marrow-derived DCs were treated with LPS for 24 hours

to induce maturation followed by exposure to UV radiation.

Viable immature DCs were characterized as CD11c+ DC with low levels of CD86, CD80 and

MHC II expression. LPS treatment of viable immature DC resulted in upregulation of CD86,

CD80 and MHC II (Figure 5-3A). Furthermore viable immature DC do not produce any IL-12;

however, in response to LPS, approximately 30% of DC were IL-12+, as expected (Figure 5-3B).

However, treatment with immature or mature apoptotic DC did not result in upregulation of

CD86, CD80 or MHC II; nor was there any induction of IL-12 production. Similar results were

also observed upon treatment of immature viable DC with necrotic DC. Taken together, these

findings indicate that immature/mature apoptotic or necrotic DC do not induce maturation of

viable immature DC.

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Figure 5-3. Immature/mature apoptotic DC or necrotic DC do not induce maturation of

viable DC.

Viable immature DC were incubated with immature apoptotic DC, mature apoptotic DC, LPS or

necrotic DC and 24 hours later FACS analysis was performed to assess expression of CD86,

CD80, MHC II on PI- CD11c+ viable DC (A) along with the proportion of IL-12+ cells among

CD11c+ DC (B). Data are representative of 3 independent experiments, with n=3-4 for every

experiment.

Immature apoptoticDCs

Mature apoptotic DCs LPS Necrotic DCs

MHC II

IL-12

CD 86

CD 80

Freq

uen

cyFL

1

A

B

Isotype control

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Viable DC upon uptake of apoptotic DC but not necrotic DC are resistant to LPS induced

maturation in vitro

We next assessed the effects of uptake of necrotic/apoptotic DC by viable immature DC on

subsequent treatment with LPS (Figure 5-4A). In the absence of inflammatory stimuli, viable

immature DC express very low levels of CD86, with approximately only 20% cells being

CD86+. This proportion increases to 50-60% upon treatment with LPS with a concomitant

increase in the intensity of CD86 expression (Figure 5-4B). Similar results are also observed

upon incubation of viable immature DC with necrotic DC followed by treatment with LPS.

However, upon incubation of viable immature DC with apoptotic DC followed by LPS

treatment, only 20-25% of viable immature DC become CD86+, which is in fact similar to the

levels seen in viable immature DC without any LPS treatment (Figure 5-4B,C). Furthermore,

incubation of viable immature DC with apoptotic splenocytes also resulted in suppression of LPS

induced subsequent DC maturation. However, the extent of immunosuppression induced by

apoptotic splenocytes was not as potent as apoptotic DC (Figure 5-4B, C). These results indicate

that uptake of apoptotic DC by viable immature DC prevents subsequent upregulation of CD86

in response to LPS.

In the absence of inflammatory stimuli, viable immature DCs do not produce any IL-12.

However, in response to LPS, approximately 22% of cells become IL-12+ (Figure 5-4D, E).

Similarly, viable immature DC incubated with necrotic DC followed by treatment with LPS

show similar proportion of IL-12+ DC. In contrast, viable DC incubated with apoptotic

splenocytes followed by LPS treatment, showed a slight reduction in IL-12 production, as only

8-11% of the cells became IL-12+. However, viable immature DC incubated with apoptotic DC

followed by treatment with LPS failed to induce IL-12, as only 1-2% of DC become IL-12+

(Figure 5-4D, E). The uptake of apoptotic DC by viable immature DC is critically important for

suppression of CD86 upregulation and IL-12 induction in response to LPS for no suppression is

observed in response to LPS if apoptotic DC and viable DC are separated in culture via

transwell.

In addition to IL-12, DC maturation is also characterized by upregulation of other inflammatory

cytokines. In order to assess the effects of apoptotic or necrotic DC uptake by viable immature

DC on induction of inflammatory cytokines in response to LPS, we looked at the mRNA

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expression levels of inflammatory cytokines, including IL-1β (Figure 5-5A), IL-6 (Figure 5-5B),

TNF-α (Figure 5-5C), IL-12p35 (Figure 5-5D) and IL-12p40 (Figure 5-5E). These inflammatory

cytokines are expressed at very low levels in viable immature DC at basal levels. However, in

response to LPS, there is massive and rapid induction of these cytokines at mRNA levels (Figure

5-5A-E). However, incubation of viable immature DC with apoptotic DC but not necrotic DC

suppressed induction of the aforementioned inflammatory cytokines in response to LPS. These

findings collectively indicate that the specific uptake of apoptotic DC converts viable immature

DC into tolerogenic DC.

Next, we looked at the ability of viable DC to prime ovalbumin (OVA)-specific T cell

proliferation upon apoptotic DC uptake (Figure 5-5F). Viable immature DC were incubated with

apoptotic or necrotic DC and then pulsed with OVA in the presence of LPS. Next, these were

cultured with naïve T cells to assess their ability to induce OVA-specific T cell proliferation.

Extensive T cell proliferation was observed upon culture of naïve T cells with viable immature

DC that were pulsed with OVA in the presence of LPS or with viable immature DC that were

first incubated with necrotic DC and then pulsed with OVA. However, OVA-pulsed viable DC

that had taken up apopotic DC failed to induce OVA-specific T cell proliferation (Figure 5-5F).

These results indicate that upon uptake of apoptotic DC but not necrotic DC, viable DC are

refractory to LPS-induced maturation.

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184

Figure 5-4. Viable DCs fail to upregulate CD86 expression and IL-12 production in

response to LPS upon uptake of apoptotic DC.

Viable immature DC were cultured as follows: without LPS (DC only), with LPS (DC+LPS),

incubated with apoptotic DC and then subsequently cultured with LPS (DC+ApoDC+LPS),

incubated with necrotic DC and subsequently cultured with LPS (DC+NecDC+LPS), incubated

with apoptotic splenocytes and subsequently cultured with LPS (DC+ApoSplen+LPS). (A)

Representative dot plots depicting gating strategy for viable CD11c+ DC and exclude

apoptotic/necrotic DC. (B) Comparison of proportion of CD86+CD11c+ DC 24 hours after

culture. (C) Representative histograms of CD86 expression on viable CD11c+ DC in response to

indicated treatments. (D) Representative histograms of IL-12 production by CD11c+ DC in

response to indicated treatments. (E) Comparison of proportion of IL-12+ DC 24 hours after

culture. Data show mean ± SD, and are representative of 4-5 independent experiments, with n=3-

4 in every experiment.*p<0.05, DC+ApoDC+LPS vs. all other groups except DC only. #p<0.05,

DC+ApoSplen+LPS vs. all other groups.

185

186

Figure 5-5. Viable DC become tolerogenic DC upon uptake of apoptotic DC.

Viable DC were cultured as follows: without LPS (DC only), with LPS (DC+LPS), incubated

with apoptotic DC and then subsequently with LPS (DC+ApoDC+LPS), or incubated with

necrotic DC and subsequently cultured with LPS (DC+NecDC+LPS). Real-time RT-PCR

analysis to look at expression levels of IL-1β (A), IL-6 (B), TNF-α (C), IL-12p35 (D), and IL-

12p40 (E). Results show relative expression of different cytokines normalized to expression

levels in viable immature DC without any treatment (DC only). (F) Naïve CD4+CD25- T cells

isolated from OT-II mice were cultured with viable DC (DC- No OVA), viable DC pulsed with

OVA (DC - OVA), viable DC incubated with apoptotic DC and then pulsed with OVA

(DC+ApoDC – OVA) or viable DC incubated with necrotic DC and then pulsed with OVA (DC

+ NecDC –OVA). 3 days later, proliferation was assessed via BrdU incorporation assay. Data

show mean ± SD, obtained from 4-5 independent experiments, with n=2-3 in every

experiment.*p<0.05, DC+ApoDC+LPS vs all other groups except DC only for D-E, DC+Apo-

OVA vs. all other groups except DC-No OVA for F. #p<0.05, DC+ApoDC+LPS vs. all other

groups for A-C.

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Viable DC upon uptake of apoptotic DC induce differentiation of naïve T cells into Treg

Since viable DC acquired a tolerogenic phenotype upon apoptotic DC uptake; we next went on

to assess the ability of viable DC to induce Treg differentiation upon apoptotic DC uptake.

Culture of naïve CD4+CD25- OT-II T cells with OVA-pulsed viable DC resulted in

approximately 4-5% of naïve T cells differentiating into Foxp3+ Treg, which increased to

approximately 23-24% upon culture with OVA-pulsed viable DC that had taken up apoptotic

DC. In contrast, culture of naïve CD4+CD25- T cells with OVA-pulsed viable DC that had taken

up necrotic DC only resulted in approximately 5-6% Foxp3+ Treg (Figure 5-6A, B). In order to

test if the induction of Foxp3+ Treg was induced specifically upon uptake of apoptotic DC by

viable immature DC and not by uptake of other types of apoptotic cells, we looked at the effects

of apoptotic splenocyte uptake on the ability of viable DC to induce Foxp3+ Treg. Results

indicate that the uptake of apoptotic splenocytes did not enhance the ability of viable DC to

induce Treg, as only 7-8% of naïve T cells differentiated into Foxp3+ Treg, which was similar to

the control group. Furthermore, we also assessed the ability of in vitro generated Foxp3+ Treg to

suppress T cell proliferation. Our findings identify that the CD4+CD25+ T cell subset only from

the co-culture of naïve T cells and OVA-pulsed viable DC that had taken up apoptotic DC, was

in fact enriched for suppressor T cells, since they were able to inhibit T cell proliferation in a

dose-dependent manner (Figure 5-6C). Overall, these results indicate that it was specifically the

uptake of apoptotic DC which was primarily responsible for induction of Foxp3+ Treg by viable

DC.

Next, we wanted to assess whether the ability to induce Foxp3+ Treg by viable DC upon

apoptotic DC uptake was dependent on interaction with naïve T cells or soluble factors. This was

tested by separating T cells from DC using a transwell plate followed by an assessment of

Foxp3+ Treg induction. Naïve CD4+CD25- T cells from wild-type mice were placed in the lower

chamber, stimulated with plate-bound anti-CD3 as well as soluble CD28 antibody, and with

viable DC and apoptotic DC in the upper chamber, Foxp3 induction was observed in

approximately 20% of T cells, indicating that soluble factors secreted by viable DC that have

taken up apoptotic DC may be involved in Foxp3 induction (Figure 5-6D, E). In contrast,

addition of only viable DC, necrotic DC, viable DC and necrotic DC, or apoptotic DC alone or

viable DC and apoptotic splenocytes, even with a very high ratio of apoptotic splenocytes to the

upper well of the transwell only resulted in approximately 5-6% of naïve CD4+CD25- T cells

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differentiating into Foxp3+ Treg. Overall, these findings indicate that only upon uptake of

apoptotic DC, viable DC acquire the ability to induce Foxp3+ Treg, which is mediated by soluble

factors released by viable DC upon apoptotic DC uptake.

Additionally, since tolerance is a balance between effector and suppressor T cells, we looked at

the effect of apoptotic/necrotic DC on the ability of viable DC to induce Th17. Our findings

demonstrated that it is only upon apoptotic DC uptake, that viable DC had a diminished ability to

induce Th17 (Figure 5-6F).

189

190

Figure 5-6. Viable DC take up apoptotic DC and induce differentiation of naïve T cells into

Foxp3+ Treg in vitro.

Naïve OT-II CD4+CD25- T cells were cultured with the following: live DC pulsed with OVA

(Live DC), necrotic DC (NecDC), live DC incubated with necrotic DC and then pulsed with

OVA (Live + NecDC), apoptotic splenocytes (ApoSplen), live DC incubated with apoptotic

splenocytes and then pulsed with OVA (Live + ApoSplen), apoptotic DC (ApoDC), or live DC

incubated with apoptotic DC and then pulsed with OVA (Live + Apo DC). Five days later, T

cells were analyzed for Foxp3 expression. (A) Representative dot plots of CD4+Foxp3+ Treg in

OT-II T cells cultured with DC under various conditions. (B) Histogram comparing percentages

of Treg. Percentages are normalized to total CD4+ T cells in the culture. (C) 5 days after culture,

CD4+CD25hi T cells were isolated from the co-culture and were added to a co-culture of naïve

OT-II CD4+ T cells and OVA-pulsed DC at different ratios. 4 days later, cell proliferation was

assessed by BrdU incorporation assay and data are presented as % suppression of T cell

proliferation compared to that of OT-II CD4+ T cells cultured in the presence of OVA-pulsed DC

without addition of any CD4+CD25hi T cells. (D-E) Naïve wild-type CD4+CD25- T cells were

cultured for 5 days with plate-bound anti-CD3 Ab and soluble anti-CD28 Ab, under a transwell

containing treatments as described above without pulsing live DC with OVA. 5 days later, FACS

analysis was performed to assess percentages of CD4+ Foxp3+ Treg. (D) Representative

histogram of Foxp3 on CD4+ T cells cultured under a transwell containing Live DC or

Live+ApoDC. (E) Comparison of % Treg induced as a proportion of CD4+ T cells. (F) Naïve

CD4+CD25- T cells were cultured under Th17 inducing conditions in presence of indicated

treatments. Representative histogram of the proportion of IL-17+ cells after 4 days of culture.

Data show mean ±SD and are representative of 4 independent experiments, with n=3 for each

experiment.*p<0.05 for Live+ApoDC vs. all the other groups.

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Viable DC secrete TGF-β1 upon apoptotic DC uptake in vitro

Since TGF-β is a known inducer of Foxp3, we looked at the induction of TGF-β1 and TGF-β2 at

the mRNA level in viable DC that had taken up apoptotic DC in the presence/absence of LPS.

Our findings indicate that at basal levels without any stimulation there is some expression of

TGF-β1 in DC wh ich is suppressed in response to LPS. This suppression is also observed in

viable DC incubated with necrotic DC followed by LPS exposure (Figure 5-7A). However, no

suppression of TGF-β1 expression is observed in viable DC incubated with apoptotic DC prior to

LPS exposure. At the same time, no induction of TGF-β1 is observed in this group. In contrast to

TGF-β1, TGF-β2 levels were upregulated approximately 12-13 fold in viable DC incubated with

apoptotic DC followed by LPS exposure compared to viable immature DC without any treatment

(Figure 5-7B).

Since cytokines are also regulated at translational level; we also looked at the protein levels of

total as well as active TGF-β1 by ELISA. Results show that upon uptake of apoptotic DC, there

was a significant increase in secretion of total as well active TGF-β1 by viable DC (Figure 5-7C,

D). However, this was not observed upon uptake of necrotic DC or apoptotic splenocytes by

viable DC. In addition, viable immature DC upon incubation with apoptotic DC followed by LPS

exposure, also secreted significantly higher levels of both total as well as active TGF-β1

compared to viable immature DC treated with LPS or viable immature DC incubated with

necrotic DC and then treated with LPS. Collectively, these findings clearly show that only upon

uptake of apoptotic DC, viable DC secrete increased levels of TGF-β1, which is regulated at the

protein level. In order to confirm that it was specifically the release of TGF-β upon uptake of

apoptotic DC by live DC which was mediating induction of Foxp3+ Treg, we repeated Treg

differentiation experiments in the presence of TGF-β neutralizing antibody (Figure 5-7E). Our

results indicate that addition of TGF-β neutralizing antibody to the culture was able to reduce the

proportion of Foxp3+ Treg from 23% to 3%, indicating that it is in fact the release of TGF-β

upon uptake of apoptotic DC by live DC, which mediates differentiation of naïve T cells into

Foxp3+ Treg.

The release of TGF-β1 by live DC upon apoptotic DC uptake was regulated at the translational

level, since no upregulation of TGF-β1 mRNA was observed. In order to investigate the

underlying mechanism, we looked at the role of the mammalian target of rapamycin (mTOR).

192

mTOR, a serine/threonine protein kinase, is a regulator of translation and its major substrates

include p70S60K serine/threonine kinase and 4E Binding protein (4EBP-1). Live DC were co-

cultured with apoptotic DC in the presence of rapamycin, a known inhibitor of the mTOR

pathway. Next, we looked at the levels of total and active TGF-β1 released in the media (Figure

5-8A). Our findings indicate that pre-treatment with rapamycin resulted in significant reduction

of both total as well as active TGF-β1 released in the media, indicating a role of mTOR in the

observed TGF-β1 release upon uptake of apoptotic DC by viable DC. Furthermore, TGF-β1

secretion in response to LPS stimulation of viable DC that had taken apoptotic DC, was also

suppressed in presence of rapamycin (Figure 5-8B).

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194

Figure 5-7. In response to LPS, viable DC that have taken up apoptotic DC, induce

secretion of TGF-β1 and upregulate TGF-β2 gene expression, which mediates generation of

Foxp3+ Treg.

(A-B) Real-time RT-PCR analysis was conducted to detect expression levels of TGF-β1 (A) and

TGF-β2 (B) from cultured DC with the following treatments: no LPS (DC only), incubation with

apoptotic DC with no LPS (DC+ApoDC), LPS (DC+LPS), incubation with apoptotic DC

followed by culturing in the presence of LPS (DC+ApoDC+LPS) or incubation with necrotic DC

followed by culturing in the presence of LPS (DC +NecDC +LPS). (C) Concentration of total

TGF-β1 and active TGF-β1 released into medium by viable DC (DC only), necrotic DC

(NecDC), viable DC incubated with necrotic DC (DC + NecDC), apoptotic splenocytes

(ApoSplen), viable DC incubated with apoptotic splenocytes (DC + ApoSplen), apoptotic DC

(ApoDC), or viable DC incubated with apoptotic DC (DC + ApoDC). (D) Concentration of total

TGF-β1 and active TGF-β1 released into the media by: viable DC cultured in the presence of

LPS (DC + LPS), viable DC incubated with necrotic DC and then cultured in the presence of

LPS (DC + NecDC + LPS) or viable DC incubated with apoptotic DC and then cultured in the

presence of LPS (DC + ApoDC + LPS). (E) Naïve CD4+CD25- T cells, isolated from spleens of

OT-II mice, were cultured with the following: live DC pulsed with OVA (DC), live DC

incubated with necrotic DC and then pulsed with OVA (DC + NecDC), or live DC incubated

with apoptotic DC and then pulsed with OVA (DC + ApoDC). Furthermore, TGF-β neutralizing

antibody or a control antibody was added to the culture. Five days later, T cells were analyzed

for Foxp3 expression. Data show means ± SD, obtained and pooled from 4 independent

experiments, n=2-3 for each experiment. *p<0.05, DC+ApoDC+LPS vs. all other groups for B,

DC+ApoDC vs. all other groups for total TGF-β1 o r activ e TGF-β1 levels for C,

DC+ApoDC+LPS vs. all other groups for total TGF-β1 or activ e TGF-β1 levels for D, DC +

ApoDC with control antibody versus DC + ApoDC with neutralizing TGF-β antibody for E.

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Figure 5-8. mTOR pathway is involved in induction of TGF-β1secretion upon uptake of

apoptotic DC by viable DC.

(A) Total and active TGF-β1 levels released in the media upon uptake of apoptotic DC by live

DC in the absence/presence of rapamycin. (B) Total and active TGF-β1 levels released in the

media in response to LPS stimulation upon uptake of apoptotic DC by live DC in the

presence/absence of rapamycin. Data show means ± SD, representative of 3 independent

experiments. *p<0.05, rapamycin treated groups vs. untreated groups.

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Uptake of apoptotic DCs by viable DCs in vivo

In order to assess whether viable DCs can take up apoptotic DCs in vivo, CFSE labelled

apoptotic DCs were injected intravenously in mice through the tail vein and 24 hours later,

spleen and draining lymph nodes were isolated to assess for uptake of CFSE+ apoptotic DCs. We

gated on PI- CD11c+ DCs for viable DCs and among them we assessed the proportion of CFSE+

DCs, which were indicative of viable CD11c+ DCs that had taken up apoptotic DCs. Our data

show that approximately 14-15% of CD11c+ DCs in spleen were able to take up apoptotic DCs,

whereas 11 % of CD11c+ DCs in draining lymph nodes had taken up apoptotic DCs (Figure 5-

9A, B). In contrast, only 3% of CD11c- cells (i.e. non-DCs) had taken up CFSE-labelled

apoptotic DCs, indicating that the uptake of apoptotic DCs was largely restricted to CD11c+ DCs

(Figure 5-9B).

To further eliminate the possibility of CFSE+ apoptotic DCs being PI- upon in vivo delivery, we

performed delivery of CFSE+ CD45.1 apoptotic DCs to CD45.2 mice (Figure 5-10A) and

delivery of CFSE+ CD45.2 apoptotic DCs to CD45.1 mice (Figure 5-10B). Our results show that

upon delivery of CD45.1+ DCs to CD45.2 mice, 99% of PI-CD11c+ DCs were CD45.2+, among

which 11-12% were CFSE+, indicating that it was in fact the host CD45.2+ DCs which took up

donor CFSE+ apoptotic DCs (Figure 5-10A). Similarly, upon delivery of CFSE+ CD45.2

apoptotic DCs to CD45.1 mice, 98-99% of PI-CD11c+ DCs were CD45.1+, among which 13%

were CFSE+, indicating that it was the host CD45.1+ DCs which took up donor CFSE+ apoptotic

DCs (Figure 5-10B).

CFSE-labelled apoptotic DCs were also injected into footpads of wild-type mice. 24 hours later

popliteal lymph nodes (PLN) were isolated, and the uptake of CFSE+ apoptotic DCs by CD11c+

viable DCs was assessed via FACS analysis. We gated on PI- CD11c+ DCs for viable DCs,

among which approximately 15% DCs were CFSE+ (Figure 5-9B). In contrast, when we gated

on PI- CD11c+ DCs from mesenteric lymph nodes, only 1.6% of the cells were CFSE+, which

was expected, since most of the lymphatic drainage from the footpads is into the PLN (Figure 5-

9B). Taken together, our findings demonstrate that delivery of apoptotic DCs in mice results in

their uptake by viable CD11c+ DCs in both the spleen as well as in the draining lymph nodes.

Similarly, CFSE-labelled necrotic DCs were delivered via footpad injections to mice and 24

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hours later, approximately 11-12% of PI- CD11c+ DCs were CFSE+, indicating uptake of

necrotic DCs by viable DCs in vivo (Figure 5-9C).

Next we went on to assess the phenotype of DCs that take up apoptotic DCs (Figure 5-9D).

CFSE labelled apoptotic DCs were injected intravenously in mice and 24 hours later FACS

analysis was performed to assess the uptake of CFSE labelled apoptotic DCs by different DC

subsets. Gating was performed based on CD11c expression to classify CD11c+ DCs as CD11clo

or CD11chi. Among the two populations, further gating was performed based on CD11b and

CD8 expression. Our findings identify, that it is primarily the CD11chi DCs that take up

apoptotic DCs (48-50%), with much lower proportions of CD11clo DC taking up apoptotic DCs

(10-11%). When we further classified DCs based on CD11b expression, no differences were

observed, with similar proportions of both CD11chiCD11b- and CD11chiCD11b+ DCs taking up

apoptotic DCs. Further gating based on CD8 expression revealed that higher proportions of

CD11chiCD8+ (30.4%) and CD11cloCD8+ (7.4%) took up apoptotic DCs compared to

CD11chiCD8- (20.6%) and CD11cloCD8- (4.2%), respectively. Overall, these findings indicate

that CD11chi DCs are primarily responsible for taking up apoptotic DCs, with preferential uptake

by CD11chiCD8+ DCs.

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Freq

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Popliteal lymph n odesFootpad inje ction

A

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Gated on CD11c+ cells

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CFSE

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CFSE

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Figure 5-9: Apoptotic or necrotic DCs are taken up by viable DCs in vivo.

CFSE-labelled apoptotic DCs were injected intravenously or into footpads of mice and 24 hours

later, spleen and draining lymph nodes or popliteal lymph nodes (PLN) were isolated. FACS

analysis was performed to look at the uptake of apoptotic CFSE+ DCs by CD11c+ viable DCs

from the spleen and draining lymph nodes (mediastinal, mesenteric, inguinal, brachial and

superficial cervical nodes) (A, B). Similarly FACS analysis was also conducted to look at

uptake of necrotic CFSE+ DCs by CD11c+ viable DCs in PLN (C). Propidium iodide (PI)

exclusion was performed to gate on viable CD11c+ DCs. Among the PI- viable cells, gating was

performed on CD11c+ cells to assess for presence of CFSE+CD11c+ DCs. Controls included

injection of unlabelled apoptotic or unlabelled necrotic DCs along with assessment of CFSE+

cells among CD11c+ viable DCs in the mesenteric lymph nodes upon footpad injections (B). (D)

FACS analysis was performed to assess uptake of CFSE+ DCs by different subsets of DCs based

on different levels of CD11c, CD11b and CD8 expression. Data shown is representative of 3-4

independent experiments.

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Figure 5-10: Uptake of apoptotic DCs by viable DCs in vivo.

(A)CD45.1+ apoptotic DCs from B6.SJL mice were injected into C57BL/6 mice (CD45.2+). 24

hours later, gating was performed to look at viable PI-CD11c+CD45.2+ host DCs among which

the proportion of CFSE+ cells was assessed to identify DCs that had taken up apoptotic DCs. (B)

CD45.2+ apoptotic DCs from C57BL/6 mice were injected into B6.SJL mice (CD45.1+).24

hours later, gating was performed to look at viable PI-CD11c+CD45.2+ host DCs among which

the proportion of CFSE+ cells was assessed to identify DCs that had taken up apoptotic DCs.

Pro

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11c

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Delivery of apoptotic or necrotic DCs does not induce viable DC maturation in vivo

In order to test whether apoptotic or necrotic DCs have any inflammatory effect on viable DCs,

footpad injections of either saline, apoptotic DCs, LPS or necrotic DCs were performed in mice

and 24 hours later, PLN were isolated to assess for maturation of viable DCs (Figure 5-11).

Gating was performed on PI- CD11c+ DCs, indicative of viable DCs. In response to saline,

approximately 56% of DCs in the PLN were CD86+, which indicates the basal levels of CD86

expression on DCs in the lymph nodes. Similarly, delivery of apoptotic DCs did not alter the

proportion of CD86+ DCs. In contrast, delivery of LPS resulted in an increase in the levels of

CD86+ DCs, which increased from 56% at basal levels to 85% in response to LPS. However,

similar to delivery of apoptotic DCs, injection of necrotic DCs did not result in upregulation of

CD86 expression on DCs in the PLN (Figure 5-11A). Furthermore, we also looked at the

proportion of IL-12+ DCs in the PLN (Figure 5-11B-D). At basal levels and upon injection of

apoptotic or necrotic DCs, only 5% of the DCs were IL-12+. In contrast, delivery of LPS resulted

in IL-12 production by approximately 30-35% of DCs in the PLN. Overall; these results

indicated that delivery of apoptotic or necrotic DCs does not result in maturation of viable DCs

in vivo. We also isolated the PLN and looked at TGF-β1 secretion from PLN derived cells in

vitro (Figure 5-11E). Our findings indicate that PLN cells from mice that received apoptotic DCs

secreted significantly elevated levels of TGF-β1.

We also looked at the phenotype of viable CD11c+ DCs in lymph nodes that had taken up

apoptotic or necrotic DCs. In order to test this, CFSE+ apoptotic or necrotic DCs were delivered

by footpad injections and gating was performed first on PI- CD11c+ DCs from the lymph nodes,

indicative of viable DCs and among them further gating was performed on CFSE+ DCs to look at

CD86, CD80 and MHC II expression levels on viable DCs that had taken up apoptotic or

necrotic DCs (Figure 5-11F). 59% of viable DCs that had taken up apoptotic DCs were CD86+,

which was similar to the proportion of CD86+ DCs normally observed in lymph nodes upon

saline injections (Figure 5-11G). Similar proportions of CD86+ DCs were observed among viable

DCs that had taken up necrotic DCs (57%). Similarly the proportions of CD80+ and MHC II+

viable CD11c+ DCs that had taken up apoptotic or necrotic DCs was similar to the proportions of

lymph node DCs observed upon saline injections (Figure 5-11G) Additionally, we also assessed

for mean fluorescent intensity (MFI) of CD86, CD80 and MHC II on viable DCs that had taken

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up apoptotic or necrotic DCs and our results show that uptake of apoptotic or necrotic DCs did

not change the MFI of CD86, CD80 or MHC class II compared to MFI of these markers on

lymph node DCs of mice which received saline only (Figure 5-11G). Taken together these

findings indicate that uptake of apoptotic or necrotic DCs by viable DCs in vivo, is not

recognized as inflammatory.

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Figure 5-11: Delivery of apoptotic DCs to mice results in their uptake by viable DCs, which

do not undergo maturation but produce TGF-β1.

Mice were injected in footpads with saline, apoptotic DCs, LPS or necrotic DCs and 24 hours

later, PLN were isolated. (A) Representative FACS histograms depicting CD86 expression on

viable PI-CD11c+ DCs in PLN upon delivery as described above. (B) Representative FACS plots

depicting gating strategy, used for gating on CD11c+ DCs from the PLN. (C) Representative

FACS plots depicting IL-12+ cells among gated CD11c+ cells from the PLN. (D) Comparison of

proportions of IL-12+ CD11c+ DCs in PLN upon delivery as described above. (E) Release of

TGF-β1 from single cell suspensions of PLN cells isolated from mice treated as above upon

culture in vitro. (F-G) Mice were injected as above, but with CFSE-labelled apoptotic or necrotic

cells and 24 hours later assessed for CD86 levels on viable DCs in PLN that had taken up

apoptotic DCs. (F) Gating strategy to gate on viable CD11c+ DCs that had taken up CFSE

labelled apoptotic or necrotic DCs. Gating was first performed on PI-CD11c+ DCs to gate on

viable DCs (i), followed by gating on CFSE+ population among PI-CD11c+ DC population (ii),

then expression of CD86 (iii), CD80 (iv) and MHC II (v) was assessed on the gated PI-

CD11c+CFSE+ population (iii). (G) Proportions of CD86+, CD80+ and MHCII+ DCs among PI-

CD11c+CFSE+ population along with respective mean fluorescence intensity. All data are mean

+/- SD, and representative of n=4-5 and representative of 4 independent experiments. *P<0.05,

statistically significant compared to all the other groups.

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Apoptotic DCs can suppress LPS-induced viable DC migration in vivo

In order to confirm the immunosuppressive properties of apoptotic DCs in vivo, we looked at the

effects of apoptotic or necrotic DCs on LPS-induced DC migration from the periphery to PLN

upon footpad injections by gating on CD11c+PI- cells (viable cells) (Figure 5-12). FITC-Dextran

injection was used to label DCs in the periphery to monitor their migration to the lymph nodes as

described (Kushwah et al., 2008). Upon delivery of saline, approximately 14-15% of CD11c+

DCs in the lymph nodes were FITC-Dextran+, which indicates the basal levels of DC migration

from the periphery to the lymph nodes (Figure 5-12A, B). In contrast, in response to LPS

delivery, approximately 40% of CD11c+ DCs in PLN were FITC-Dextran+, indicating increased

migration of DCs from the periphery to PLN. This was expected, as LPS in a known inducer of

DC maturation and migration. Similarly, delivery of necrotic DCs prior to LPS delivery

produced similar results, indicating that necrotic DC delivery was not able to dampen LPS

induced DC migration from the periphery to the PLN. However, upon injection of apoptotic DCs

prior to LPS administration, only 18% of the DCs in PLN were FITC-Dextran+ i.e. were from the

periphery, indicating that apoptotic DCs suppressed LPS-induced DC migration (Figure 5-12A,

B). This was further confirmed by measuring absolute count of total DCs in the PLN along with

the absolute count of FITC-Dextran+ DCs (indicative of DCs that had migrated from the

periphery to PLN); both were significantly reduced upon injection of apoptotic DCs prior to LPS

delivery compared to mice that had received LPS alone (Figure 5-12C). In fact, the levels of DC

migration observed upon delivery of apoptotic DCs prior to LPS delivery was similar to the basal

levels, seen upon injection of saline only. Overall, the findings indicate that injection of

apoptotic DCs prior to LPS delivery was able to suppress LPS induced DC migration from the

periphery to the lymph nodes. In contrast, delivery of necrotic DCs prior to LPS delivery had no

effect on LPS induced DC migration from the periphery to PLN.

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FITC-Dextran

14.740.98

18.13

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Figure 5-12: Apoptotic DCs suppress LPS-induced DC migration from periphery to

draining lymph nodes in vivo.

Mice were injected in footpads with FITC-Dextran along with saline (saline), FITC-Dextran

prior to LPS delivery (LPS), FITC-Dextran along with apoptotic DCs prior to LPS delivery

(ApoDC + LPS), or FITC-Dextran along with necrotic DCs (NecDC +LPS) prior to LPS

delivery. 24 hours later, percent of FITC-Dextran+ cells among viable CD11c+ DCs were

analyzed in the PLN using FACS analysis. PI exclusion was used to exclude injected

apoptotic/necrotic DCs from viable DCs. (A) Representative FACS histograms looking at FITC-

Dextran+ DCs among CD11c+ DCs from the PLN of mice treated, as described above. (B)

Comparison of the proportion of FITC-Dextran+ DCs in the PLN of mice treated as described

above. (C) Absolute DC count in PLN was measured using FACS analysis with total DC count

corresponding to total CD11c+ count and FITC-Dextran+ CD11c+ DC count corresponding to

DCs that had migrated from the periphery to PLN. All data are means +/- SD obtained from n =

4-5 for each group. *P<0.05, statistically significant compared to NecDC+LPS and LPS.

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Apoptotic DCs suppress LPS induced DC maturation in vivo

In order to assess the effects of apoptotic DCs on DC maturation in vivo, we looked at the

expression of CD86 and CCR7 on DCs in PLN after delivery of apoptotic or necrotic DCs prior

to LPS delivery (Figure 5-13). DCs normally express very low levels of CD86, which is

upregulated in response to inflammatory stimuli, such as LPS. Upon saline delivery,

approximately 69% of CD11c+DCs in PLN were CD86+, which increased to 80-83% upon

injection of LPS or necrotic DCs prior to LPS delivery (Figure 5-13A). In contrast, injection of

apoptotic DCs prior to LPS delivery reduced this to 68%, which was similar to the proportion of

CD86+ DCs observed in PLN of mice which received saline only. In addition, a higher

proportion of DCs were CD86- upon injection of apoptotic DCs prior to LPS delivery (30%)

compared to LPS alone (21%) or LPS after necrotic DC injection (20%) (Figure 5-13C).

Moreover, though the proportion of CD86lo DCs was similar among all the groups, the

proportion of mature DCs (i.e. CD86hi DCs) was significantly reduced upon injection of

apoptotic DCs prior to LPS delivery compared to the rest of the groups, and was similar to the

levels seen with saline injections (Figure 5-13D, E). We also looked at the expression levels of

CCR7, which is a chemokine receptor, upregulated on DCs as they undergo maturation (Figure

5-13F). Our results indicate that though there was not a significant difference in the proportion of

DCs that were CCR7+, there was indeed a difference in the levels of CCR7 expression. Upon

saline delivery, the MFI of CCR7 on CD11c+ DCs in PLN was 52, which increased to 103 upon

LPS delivery and increased to 115 upon delivery of necrotic DCs prior to LPS delivery. In

contrast, injection of apoptotic DCs prior to LPS delivery reduced the MFI to 48, which was

indeed similar to the MFI of CCR7 expression seen only with saline delivery. These results

clearly show that apoptotic DCs but not necrotic DCs can suppress maturation of viable DCs in

response to an inflammatory stimulus in vivo.

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Figure 5-13: Apoptotic DCs suppress LPS-induced DC maturation in vivo.

Mice were injected in footpads with saline (saline), LPS (LPS), apoptotic DCs prior to LPS

delivery (ApoDC + LPS), or necrotic DCs (NecDC +LPS) prior to LPS delivery. 24 hours later

FACS staining was performed to assess expression of CD86 by CD11c+ viable DCs in the PLN.

PI exclusion was used to gate out dead DCs. (A) Representative FACS expression profile of

CD86 on PI-CD11c+ DCs from treated animals as described above. (B) Shown here is the gating

strategy used to classify DCs as CD86-, CD86lo and CD86hi. (C-E) Proportions of CD86- (C),

CD86lo (D) and CD86hi (E) CD11c+ DCs in PLN of groups of mice injected as described. (F)

Representative FACS expression profile of CCR7 on PI-CD11c+ DCs along with corresponding

MFI from treated animals as described above. All data are means +/- SD obtained from n = 4-5

for each group.*P<0.05 for ApoDC + LPS vs LPS and NecDC+ LPS for C and E.

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Intranasal delivery of apoptotic DCs can suppress LPS-induced inflammation and induce

expansion of Tregs

In order to further confirm the immunosuppressive and tolerogenic properties of apoptotic DCs,

we assessed the effects of apoptotic DCs in suppression of airway inflammation (Figure 5-14).

Delivery of CFSE+ apoptotic DCs resulted in uptake by viable CD11c+ DCs that could be

detected in the mediastinal lymph nodes (MLN) (Figure 5-14A). Using wild-type mice, we

performed intranasal delivery of apoptotic DCs both with LPS or 1 day after LPS delivery. In

response to delivery of LPS, there was approximately a 3-fold increase in absolute cell count in

bronchoalveolar lavage fluid (BALF) compared to mice that received saline only (Figure 5-14B).

In contrast, delivery of LPS with apoptotic DCs or delivery of apoptotic DCs 1 day after LPS

delivery resulted in significant reductions in absolute cell count in BALF compared to LPS

delivery alone. However, there was a slightly greater reduction in absolute cell count in BALF

upon delivery of apoptotic DCs at 1 day after LPS delivery compared to delivery of LPS together

with apoptotic DCs. In addition, we also performed histopathological assessment of

inflammation in the airways by staining lung sections with hematoxylin and eosin, followed by

observation under a light microscope (Figure 5-14C). Intranasal delivery of saline or apoptotic

DCs alone did not result in any infiltration within the airways. However, upon delivery of LPS,

perivascular inflammation as well as both parenchymal and intra-alveolar infiltrates of

neutrophils, macrophages and lymphocytes could be observed throughout the lungs, which was

also the case upon delivery of LPS followed by saline. In contrast, delivery of LPS along with

apoptotic DCs resulted in reduced levels of cellular infiltrates compared to LPS delivery alone.

This reduction in the extent of cellular infiltrate was even more profound upon delivery of

apoptotic DCs 1 day after LPS delivery.

In order to further study the extent of immunosuppression induced by apoptotic DCs, we also

looked at the proportion of IL-12+ DCs in MLN of treated mice (Figure 5-15A). Approximately

5% of CD11c+ DCs in the MLN were detected as IL-12+ DCs upon saline delivery, which

increased to 35-40% upon intranasal delivery of LPS. Similarly, 35-40% of DCs were IL-12+ in

MLN of mice that received LPS followed by saline delivery. In contrast, delivery of apoptotic

DCs with LPS or LPS followed by apoptotic DCs resulted in suppression of IL-12 production by

DCs in MLN (Figure 5-15A). Approximately, 15-17% of the DCs were IL-12+ in MLN of mice

which received LPS with apoptotic DCs and 8-10% were IL-12+ in MLN of mice which received

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LPS followed by apoptotic DCs. This was overall a significant reduction compared to the 35-

40% IL-12+ DCs observed in MLN of mice which received LPS only. Next, we assessed the

capacity of CD11c+ DCs from MLN of treated mice to respond to LPS restimulation in vitro.

MLN from treated mice were isolated and single cell suspension was prepared, which was

cultured in presence of LPS and 24 hours later, proportion of IL-12+CD11c+ DCs was assessed

by FACS analysis (Figure 5-15B-D). Exposure of cells from MLN of mice which received saline

had a robust response to LPS, which approximately 35-40% of DCs as IL-12+. Similarly, LPS

exposure of cells from MLN of mice which received LPS only or LPS followed by saline,

resulted in approximately 40-50% of DCs producing IL-12. In contrast, cells from mice treated

with LPS in combination with apoptotic DCs or LPS followed by apoptotic DCs had a

significantly diminished ability to respond to LPS (Figure 5-15B-D). Only 20% of DCs from

mice that received LPS with apoptotic DCs were IL-12+ upon restimulation with LPS. Similarly,

only 10-12% of DCs from mice which received LPS followed by apoptotic DCs were IL-12+

upon LPS restimulation. Taken together, these findings indicate that delivery of apoptotic DCs

with LPS or after LPS exposure, results in suppression of inflammation. This may partly be due

to the suppressive effects of apoptotic DCs on IL-12 production by CD11c+ DCs in the lymph

nodes in response to LPS.

Since our experiments showed that delivery of apoptotic DCs resulted in TGF-β1 secretion from

the lymph nodes, we looked at the secretion of TGF-β1 in BALF fluid after intranasal delivery of

apoptotic DCs (Figure 5-16A). Delivery of LPS or LPS followed by saline did not result in any

significant change in total TGF-β1 levels in BALF compared to the level in BALF of mice that

received saline alone. In contrast, delivery of apoptotic DCs (both in combination with LPS or 24

hours after LPS delivery) resulted in significant increase in TGF-β1 in BALF (Figure 5-16A).

However, the increase was transient, since it could only be observed at 6 hours after delivery and

returned to normal levels after 24 hours. We also looked at TGF-β1 secretion from the MLN of

mice that received apoptotic DCs (Figure 5-16B). MLN were isolated 24 hours after delivery of

apoptotic DCs in combination with LPS or apoptotic DCs 1 day after LPS delivery; a single cell

suspension was made, which was cultured in serum free media for 48 hours, after which TGF-β1

levels in the medium were assessed by ELISA. Our results indicate that there was a significant

increase in TGF-β1 secretion from MLN cells isolated from mice that received LPS in

combination with apoptotic DCs or apoptotic DCs alone 24 hours after LPS delivery as

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compared to mice that received saline alone, LPS alone or LPS followed by saline delivery

(Figure 5-16B). Since there was enhanced TGF-β1 secretion from lymph nodes, we assessed

whether there was Treg expansion occurring in MLN upon apoptotic DC delivery (Figure 5-16C,

D). Approximately 5-8% of CD4+ T cells in MLN of mice that received saline alone, LPS alone

or LPS followed by saline delivery were CD4+Foxp3+ Tregs. In contrast, in mice that received

LPS in combination with apoptotic DCs, the proportion of CD4+Foxp3+ Tregs increased to 15%.

However, in mice that had inflammation prior to apoptotic DC delivery (i.e. received LPS

followed by apoptotic DCs), there was a significant increase in the proportion of CD4+Foxp3+

Tregs, which increased to approximately 18-22 % of CD4+ T cells in the MLN (Figure 5-16C, D).

Similarly, increase in the absolute count of CD4+Foxp3+ Tregs was also observed in the MLN

(Figure 5-16E). Therefore, our findings indicate that intranasal delivery of apoptotic DCs can

mediate resolution of inflammation, firstly through induction of TGF-β1 secretion, which itself is

known to be immunosuppressive and secondly through their ability to suppress IL-12

production by DCs in lymph nodes. Additionally, our data also indicates that upon intranasal

delivery of apoptotic DCs, there is secretion of TGF-β1 in MLN, which promotes expansion of

Tregs.

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Figure 5-14: Apoptotic DCs can suppress LPS-induced airway inflammation.

The following treatments were delivered to mice by intranasal administration: (i) saline alone,

(ii) LPS alone, (iii) LPS then saline, (iv) apoptotic DC alone (ApoDC only), (v) LPS with

apoptotic DCs (LPS + ApoDC) or (vi) LPS followed by delivery of apoptotic DCs after 1 day

(LPS then ApoDC). (A) Representative FACS histogram looking at uptake of CFSE labelled

apoptotic DCs by viable PI-CD11c+ DCs in the mediastinal lymph node upon intranasal delivery.

(B) Total cell count in BALF of mice, 24 hours post delivery. (C) Histopathological analysis of

haematoxylin and eosin stained lung sections to assess inflammation in the airways 48 hours

after delivery conditions as described above (i-vi). Upon intranasal delivery of saline or apoptotic

DCs alone, no inflammation was present within the airways. Perivascular inflammation as well

as both parenchymal and intra-alveolar infiltrates of neutrophils, macrophages and lymphocytes

were detected throughout the lungs upon delivery of LPS, which was also the case upon delivery

of LPS followed by saline. In contrast, delivery of LPS along with apoptotic DCs resulted in a

reduction of inflammatory cell infiltrate compared to LPS delivery alone. This reduction was

even more obvious for delivery of LPS followed by apoptotic DCs. Images were taken at 50x

magnification and insets refer to a 200x-magnified region of the perivascular inflammation

indicated by a box within the same image. Images are representative of n = 3-4 mice for each

group. Data is presented as mean +/- SD, obtained from n=4 mice per group, representative of 3

independent experiments. *P<0.05, LPS + ApoDC or LPS then Apo DC vs LPS and LPS then

Saline.

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Figure 5-15: Intranasal delivery of apoptotic DCs suppresses LPS-induced IL-12

production in DCs in mediastinal lymph nodes (MLN).

Mice were administered the following by intranasal delivery: saline, LPS, LPS with apoptotic

DCs (LPS + ApoDC), LPS followed by saline after 1 day (LPS then saline) or LPS followed by

delivery of apoptotic DCs after 1 day (LPS then ApoDC). (A) Proportion of IL-12+ CD11c+ DCs

in MLN, 24 hours after delivery. (B-D) After 24 hours, MLN were isolated and a single cell

suspension was prepared which was cultured in presence of LPS for 24 hours. (B) Representative

FACS plots depicting gating strategy, used for gating on CD11c+ DCs from the MLN single cell

suspension. (C) Representative FACS plots looking at IL-12+ CD11c+ DCs in culture after LPS

exposure. (D) The graph compares the proportion of IL-12+CD11c+ DCs in culture after LPS

exposure. Data are presented as means +/- SD, obtained from n=4 mice for each group,

representative of 4 independent experiments. *P<0.05, LPS then ApoDC vs all the other groups

except LPS + ApoDC, #P<0.05, LPS+ApoDC vs all other groups except LPS then ApoDC.

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Figure 5-16: Intranasal delivery of apoptotic DCs results in the secretion of TGF-β1 and

Treg expansion.

Mice were administered the following by intranasal delivery: saline, LPS, LPS with apoptotic

DCs (LPS + ApoDC), LPS followed by saline after 1 day (LPS then saline) or LPS followed by

delivery of apoptotic DCs after 1 day (LPS then ApoDC). (A) Concentration of total TGF-β1

levels in BALF, 6 hours and 24 hours after delivery. (B) Concentration of total TGF- β1 that is

released into media 48 hours after culture of mediastinal lymph node (MLN) cells isolated from

the mice treated as above. (C) Representative FACS plots looking at Foxp3+ cells among CD4+

cells from the MLN of mice, 1 day after above-mentioned treatments. (D) Histogram compares

the percentage of CD4+Foxp3+ Tregs normalized to total CD4+ T cells in the MLN of mice, 1 day

after above-mentioned treatments, assessed by FACS analyses. (E) Comparison of absolute

count of CD4+Foxp3+ Tregs from MLN of mice, 1 day after above-mentioned treatments. All data

are presented as mean +/- SD, obtained from n=4-5 mice per group, representative of 4

independent experiments. * P<0.05, LPS + ApoDC or LPS then ApoDC vs all other groups for A

and B; LPS then ApoDC vs all other groups except LPS+ApoDC for D. #P=0.08, LPS+ApoDC

vs LPS then saline.

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Apoptotic DCs can induce immune tolerance in vivo

Since our previous findings suggested that in vitro viable DCs prime differentiation of naïve T

cells into Tregs and do not induce T cell proliferation upon apoptotic DC uptake, we assessed

whether apoptotic DCs can suppress T cell proliferation in vivo (Figure 5-17A). C57BL/6 mice

were injected with OVA in saline or OVA in Complete Freund’s adjuvant (CFA), which would

prime a robust T cell response directed against OVA. In order to test the efficacy of apoptotic

DCs in inducing tolerance, 6 hours prior to injection of OVA in CFA, mice were injected with

apoptotic DCs or necrotic DCs intravenously. Mice were given intraperitoneal injections of

BrdU to label proliferating cells and 7 days after delivery of OVA in CFA, mice were sacrificed

and T cell proliferation was assessed in draining lymph nodes by looking at the proportion of

BrdU+ CD3+ T cells. Results indicate that upon delivery of OVA in saline, approximately 3-4%

of the total CD3+ T cells in draining lymph nodes were proliferating (i.e. BrdU+) (Figure 5-17B,

C). However, upon injection of OVA in CFA, the number increased to approximately 21-22%,

indicating that CFA primed a robust immune response against OVA, as expected. Similarly, 22-

23% of the T cells in draining lymph nodes of mice that received necrotic DCs prior to delivery

of OVA in CFA were BrdU+, indicating that necrotic DCs had no effect on the ability of viable

DCs to prime a T cell response in vivo. In contrast, upon delivery of apoptotic DCs prior to

delivery of OVA in CFA, approximately 10% of the T cells were BrdU+ (i.e. proliferating),

which was a marked reduction compared to 23% BrdU+ T cells seen with delivery of OVA in

CFA (Figure 5-17B, C). Therefore, our results demonstrate that delivery of apoptotic DCs prior

to delivery of OVA in CFA was able to suppress the priming of immune response directed

towards OVA. Furthermore, 7 days after delivery, draining lymph nodes were isolated and total

cells were cultured in the presence of OVA for 4 days to assess for OVA-specific T cell

proliferation (Figure 5-17D). Cells isolated from lymph nodes of mice that received apoptotic

DCs prior to delivery of OVA in CFA had significantly lower levels of proliferation compared to

the cells isolated from mice which received OVA in CFA alone, or necrotic DCs prior to

delivery of OVA in CFA (Figure 5-17D).

We also wanted to confirm the ability of apoptotic DCs to induce antigen-specific Tregs in vivo

(Figure 5-17E,F). Mice were injected with OVA in saline, OVA in CFA, apoptotic DCs prior to

delivery of OVA in CFA or necrotic DCs prior to delivery of OVA in CFA. After 7 days,

draining lymph nodes were isolated, and magnetic separation was performed to enrich for CD4+

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T cells, followed by cell sorting for CD4+CD25hi cells, which would include CD4+Foxp3+ Tregs

(Figure 5-17E). Since wild-type mice are not expected to have tolerance towards OVA, they

should not have any Tregs specific to OVA. In contrast, if apoptotic DCs are able to induce OVA-

specific Tregs, then Tregs from mice treated with apoptotic DCs prior to OVA-CFA delivery

should be able to suppress OVA-specific T cell proliferation in vitro. In order to test this, the

CD4+CD25+ T cells isolated from the different groups of mice were added as “suppressors” to a

co-culture of naïve CD4+CD25- T cells isolated from OT-II mice which would have specificity

for OVA, acting as “responders” along with dendritic cells pulsed with OVA to stimulate OT-II

cells, acting as “stimulators”. CD4+CD25+ were added at ratios of 1:2, 1:10 and 1:30 to naïve

OT-II CD4+CD25- T cells in the co-culture. CD4+CD25+ T cells isolated from mice immunized

with OVA-CFA failed to suppress proliferation, indicating that there were no Tregs specific for

OVA in the immunized mice (Figure 5-17F). This was expected because immunization with

OVA in CFA led to a robust immune response. Similarly, CD4+CD25+ T cells isolated from

mice which only received apoptotic DCs, failed to suppress T cell proliferation. This was again

expected, since the animals, which received apoptotic DCs, were not exposed to OVA.

Therefore, there was no differentiation of naïve T cells into Tregs with specificity for OVA.

CD4+CD25+ T cells isolated from mice, which received necrotic DCs prior to OVA-CFA

delivery, were again not able to suppress T cell proliferation, indicating that there were no OVA-

specific Tregs generated in mice upon delivery of necrotic DCs prior to OVA-CFA. In contrast,

CD4+CD25+ T cells isolated from animals, which received apoptotic DCs prior to OVA-CFA

delivery, were able to suppress proliferation of OT-II CD4+CD25- T cells in a dose-dependent

manner, with approximately 70% suppression at 1:2 and 30% suppression at 1:10 ratio of

suppressors to responders (Figure 5-17F). This indicates that delivery of apoptotic DCs, prior to

delivery of OVA-CFA, was able to induce de novo generation of OVA- specific Tregs in mice.

In addition to the ability of apoptotic DCs to induce de novo generation of Tregs, we also wanted

to assess their ability to induce expansion of Tregs. OT-II mice were given intravenous injections

of saline, OVA-CFA or apoptotic DCs followed by OVA-CFA. 24 hours later, the proportions of

Tregs in spleen as well as the draining lymph nodes were assessed via FACS analysis. Upon

injection of saline, the proportions of Tregs relative to total CD4+ T cells were approximately 8%

and 5% in spleen and draining lymph nodes, respectively (Figure 5-18). After injection of OVA-

CFA, there was a slight increase in the proportion of Tregs, which increased to approximately

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10% in the spleen and 7% in the draining lymph nodes, albeit the difference compared to saline

delivery was not statistically significant. In contrast, 24 hours after delivery of apoptotic DCs

prior to OVA-CFA delivery, the proportions of Tregs increased to approximately 20% in the

spleen and 15% in the draining lymph nodes, which was around 2-3 fold expansion compared to

the proportions observed in mice that received saline only. The expansion of Tregs upon delivery

of apoptotic DCs prior to OVA-CFA was not observed in brachial lymph nodes, which was

expected since OVA-CFA was delivered via the intraperitoneal route and it is primarily the

lymphatic drainage from the thoracic region that reaches the brachial lymph nodes (Fig.ure 5-

18B). Therefore, these results clearly indicate that delivery of apoptotic DCs prior to antigen

stimulation can induce expansion of antigen-specific Tregs. Collectively, our results indicate that

delivery of apoptotic DCs prior to antigen delivery in vivo can induce de novo generation of

antigen-specific Tregs along with expansion of existing Tregs, thereby preventing induction of a T

cell response against the delivered antigen.

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Figure 5-17: Apoptotic DCs induce antigen-specific Tregs in vivo.

(A) Protocol for induction of OVA-specific tolerance in mice. C57BL/6 mice were injected

intravenously with ovalbumin (OVA) in saline (OVA-Saline), OVA in Complete Freund’s

Adjuvant (OVA-CFA), apoptotic DCs followed by OVA-CFA after 6 hours (ApoDC then OVA-

CFA) or necrotic DCs followed by OVA-CFA after 6 hours (NecDC then OVA-CFA). For the

next 7 days, one set of mice were given intraperitoneal injections of BrdU as described in

materials and methods section and the other set of mice was used after 7 days for isolation of

CD4+CD25+ cells. One week later, T cell proliferation in draining lymph nodes (inguinal and

mesenteric) was assessed via BrdU incorporation. (B) Representative histograms depicting BrdU

incorporation by CD3+ T cells in the draining lymph nodes of immunized mice. (C) Comparison

of % BrdU+ T cells, expressed as percent of total CD3+ T cells, from draining lymph nodes of

immunized mice. (D) After 7 days, draining lymph nodes were isolated and total cells were

cultured with OVA protein for 4 days and antigen specific T cell proliferation was determined by

BrdU incorporation assay. (E) After 7 days, CD4+CD25+ T cells were isolated from draining

lymph nodes of mice injected with OVA-CFA (CD25+ OVA-CFA mice), apoptotic DCs (CD25+

ApoDC mice), necrotic DC followed by OVA-CFA (CD25+ NecDC then OVA-CFA mice) or

apoptotic DCs followed by OVA-CFA (CD25+ ApoDC then OVA-CFA). Magnetic selection

was used to enrich for CD4+ cells (91%) purity, which was subjected to cell sorting for isolation

of CD25hi population (94% purity). (F) The isolated CD4+CD25+ T cells were then added to a

co-culture of naïve OT-II CD4+ T cells and OVA-pulsed BMDCs at 3 different ratios of 1:2,

1:10 and 1:30. 4 days later, cell proliferation was assessed by BrdU incorporation assay and data

is presented as % suppression of T cell proliferation compared to that of OT-II CD4+ T cells

cultured in presence of OVA pulsed BMDCs without addition of any CD4+CD25+ T cells. All

data presented as mean+/-SD, representative of n=4 mice per group and representative of 3

independent experiments. *P<0.05, ApoDC then OVA-CFA vs all other groups except OVA-

Saline.

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Figure 5-18: Apoptotic DCs induce expansion of Tregs in vivo.

(A) Representative FACS profile of Foxp3 expression on CD4+ cells from spleens and draining

lymph nodes of OT-II mice immunized with necrotic DCs (NecDC only), apoptotic DCs

(ApoDC only), CFA (CFA only), Saline (Saline only), OVA-CFA or with apoptotic DCs

followed by OVA-CFA (ApoDC then OVA-CFA). Immunization was performed as shown in

Figure 5-17A. One day after immunization, proportions of Tregs were assessed in spleen along

with draining lymph nodes (inguinal and mesenteric) via FACS analysis. (B) Representative

FACS profile of Foxp3 expression on CD4+ cells from brachial lymph nodes of mice immunized

intravenously with apoptotic DCs followed by OVA-CFA. (C) Comparison of percent

CD4+Foxp3+ Tregs in spleen and draining lymph nodes (iguinal and mesenteric) from OT-II mice

immunized as described above. Data presented as mean +/- SD, representative of n=3 mice per

group. *P<0.05, ApoDC then OVA-CFA vs all the groups.

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5.4 Discussion

Taken together, our results show that the impact of dying DC on the immune system is

dependent on the manner in which DC die. If DC undergo apoptosis and viable DC take them up,

then viable DC transform into tolerogenic DC. These tolerogenic DC are resistant to stimuli-

induced maturation, secrete TGF-β1, which is dependent on mTOR pathway and induce

generation of Foxp3+ Tregs. Surprisingly, our findings show that necrotic DC, irrespective of

their maturation status, are not immuno-stimulatory, which may be due to the paucity of presence

of certain immunosuppressive factors in primary DC, which renders them non-immunogenic

even after the cellular contents are released into the extracellular milieu. However, such factors

still need to be identified.

Furthermore, our results clearly indicate that apoptotic DCs have a tolerogenic effect on the

immune response in vivo. Our findings indicate that delivery of apoptotic DCs in mice results in

their rapid engulfment by viable DCs in both the spleen as well as draining lymph nodes.

However, the delivery of apoptotic or necrotic DCs has no effect on the surface phenotype of

viable DCs which take them up nor is their uptake recognized as inflammatory by viable DCs. In

contrast to the delivery of necrotic DCs, delivery of apoptotic DCs results in TGF-β1 production

in the draining lymph nodes. If an inflammatory stimulus such as LPS is delivered after delivery

of apoptotic DCs, then apoptotic DCs suppress migration of DCs from the periphery to the

draining lymph nodes. In addition, the maturation of DCs in responses to the inflammatory

stimuli is also inhibited. Furthermore, this results in suppression of T cell response to the

inflammatory agent and instead results in de novo generation of antigen-specific Tregs along with

expansion of existing Tregs. Overall, our findings identify that priming of the immune system in

vivo with apoptotic DCs results in tolerance induction to a subsequent immunogen. Our findings

establish a model whereby selective uptake of apoptotic DC induces immunologic tolerance via

suppression of DC function and induction of Tregs (Figure 5-19).

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Figure 5-19: Proposed model for tolerance induction by apoptotic DCs.

(A – B) Exposure to UV radiation induces apoptosis of both (A) lymphocytes and (B) DCs,

leading to exposure of phosphatidylserine on the cell surface of DCs and lymphocytes along with

another protein specifically on apoptotic DCs that could mediate preferential tolerance induction

by apoptotic DCs. In addition, there could also be an exposure of a trigger for αvβ8 integrins. (C)

Scavenger receptors expressed on the DC surface play a role in the uptake of apoptotic cells, and

interaction of phosphatidylserine on apoptotic cells with its receptor on DCs likely leads to

prevention of subsequent maturation in response to inflammatory stimuli, such as LPS, with no

evidence for secretion of TGF-β1. The suppression of NFκB pathway could be mediated by

signaling through the phosphatidylserine receptor, the role of which is controversial in DCs. (D)

In contrast to apoptotic lymphocyte uptake, apoptotic DC uptake by viable DCs results in

secretion of TGF-β1, suppression of maturation, and induction of Ag-specific Foxp3+ Tregs.

Secretion of TGF-β1 is dependent on the mTOR pathway, although the signal that results in

mTOR activation upon apoptotic cell uptake is not known. This could be mediated by a protein

specifically present on apoptotic DCs, which could also result in suppression of DC maturation.

Furthermore, activation of inactive TGF-β1 to the active form could be mediated through avb8

signaling, although the mechanism of how apoptotic DCs could trigger signaling is not known.

Suppression of DC maturation could be mediated by signals from an unidentified receptor on the

apoptotic DC surface via suppression of NFκB activation in addition to signaling through the

phosphatidylserine receptor.

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In addition to induction of tolerance, our findings also identify that apoptotic DCs can be used

for suppression of existing inflammation. Our findings show that apoptotic DCs are quite potent

at suppression of LPS-induced inflammation in the airways. The suppression is actually

mediated in three different steps. Firstly, apoptotic DCs suppress the induction of IL-12 by viable

DCs in the draining MLN and make them non-responsive to a further inflammatory insult.

Secondly, apoptotic DCs also induce transient TGF- β1 secretion in BALF, which is known to be

immunosuppressive. Thirdly, apoptotic DCs induce TGF-β1 secretion in the MLN, which results

in Treg expansion. Taken together, these findings indicate that apoptotic DCs are very potent in

suppression of existing inflammation. The tolerogenic effects of DCs reported in this study could

be relevant for generation of immunosuppression and antigen-specific tolerance for many

applications in transplantation, allergy and autoimmunity. These findings could have

implications in suppression of airway immune response as is the case in asthma, lung

transplantation and also upon delivery of gene therapy vectors. In human lung transplantation, it

has been shown that grafts with high levels of inflammatory cytokines develop severe graft

dysfunction following reperfusion (De Perrot et al., 2001; De Perrot et al., 2002; Fisher et al.,

2001). Perhaps, delivery of apoptotic DCs prior to harvesting the organ may prevent induction of

reperfusion injury by suppressing the inflammatory response, which could potentially enhance

graft survival and limit lung inflammation. In addition, airway immune responses are a barrier to

the success of airway gene therapy (Kushwah et al., 2007a). Our previous work has shown that

even so called non-immunogenic vectors such as helper-dependent adenoviral vectors are able to

potentiate cytotoxic T cell response upon airway delivery at low doses(Kushwah et al., 2008).

This poses a serious barrier towards readministration of gene therapy vectors. Therefore, it is

feasible that using apoptotic DCs, perhaps transient immunosuppression and tolerance can be

induced, which would prevent subsequent immune response against gene therapy vectors.

Studies have shown that DC can take up antigen from dying cells and cross-present the antigenic

material onto both MHC I as well as MHC II (Albert et al., 1998b; Inaba et al., 1998). However,

these studies relied on the use of mature DC to phagocytose apoptotic cells. We can speculate

that perhaps in a physiological setting, if the causative agent of DC apoptosis is an infection, then

it is usually the semi-mature or mature viable DC in close proximity that take up apoptotic DC.

Thereby, these viable DC can cross-present the antigen and then prime a T cell response rather

than induction of tolerance, as seen in our study.

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Previous studies have indicated that phosphatidylserine, an anionic aminophospholipid, which is

exposed on cell surface as cells undergo apoptosis, plays an important role in the recognition and

clearance of apoptotic cells by macrophages. Studies have also shown that this interaction with

phosphatidylserine results in suppression of macrophage activation and induction of TGF-β1

gene expression (De et al., 2002; Fadok et al., 1998; Huynh et al., 2002). It is tempting to argue

that conversion of viable immature DC to tolerogenic DC with a potential to induce Treg via

secretion of TGF-β1, is largely phosphatidylserine dependent. However, in our study, when

viable immature DC were exposed to apoptotic splenocytes, no increase in TGF-β1 secretion was

observed and previous studies have also indicated that exposure of murine DC to apoptotic cells

or phosphatidylserine does not induce TGF-β1 secretion (Chen et al., 2004; Morelli et al., 2003;

Takahashi and Kobayashi, 2003). Therefore, it is likely that, the ability to secrete TGF-β1 and to

induce Foxp3+ Treg may be dependent on the uptake of apoptotic DC by viable DC, which has

not been described previously, and could be independent of phosphatidylserine. It is feasible that

as DC undergo apoptosis, there is exposure of phosphatidylserine, which may play a passive role

in suppression of DC by suppressing the ability of DC to undergo maturation without any

induction of Foxp3+ Treg.. We propose that uptake of apoptotic DC in particular, triggers

signalling through a previously unidentified receptor in viable DC that induces TGF-β1

secretion.

Our findings identify that the release of TGF-β1 upon uptake of apoptotic DC by viable DC is

regulated at translational level via mTOR pathway.Mammalian target of rapamycin (mTOR), a

serine/threonine protein kinase is a regulator of translation and its major substrates include

p70S60K serine/threonine kinase and 4E Binding protein (4EBP-1). mTOR phosphorylates

4EBP-1 which results in release of protein translation initiation factor eIF4E. eIF4E plays a role

in enhancing rates of translation of capped mRNA which also includes TGF-β1. mTOR is likely

regulated upstream by PI3/Akt pathway and Rho A has previously been shown to induce PI3

pathway to prevent myoblast death (Reuveny et al., 2004). Therefore it is likely that RhoA

induces PI3K which phosphorylates mTOR resulting in release of eIF4E, which further results in

increased translation of TGF-β1 mRNA. Some studies have indicated that another mechanism

whereby DC can acquire tolerogenic potential is through induction of indoleamine-2,3-

deoxygenase (IDO) (Orabona et al., 2008; Williams et al., 2008). Our results show no

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upregulation of IDO upon uptake of apoptotic DC by viable DC , indicating that induction of

IDO is likely not the underlying mechanism for tolerance induction (data not shown).

The hallmarks of sepsis include impaired immune function along with immunosuppression

(Hotchkiss and Karl, 2003). Concominantly, there is substantial depletion of DC along with

increased levels of circulating Treg (Hotchkiss et al., 2002; Monneret et al., 2003; Tinsley et al.,

2003). However, the mechanism of how DC apoptosis can contribute to immunosuppression in

sepsis is unclear. Our findings suggest that perhaps enhanced DC apoptosis in sepsis may result

in their uptake by viable DC, resulting in immunosuppression and Treg induction/expansion. We

need to be cautious in interpreting our findings because our data indicate that several fold higher

amounts of apoptotic DC are required than live DC for tolerance induction. However, in certain

pathologies such as sepsis, there could be local environments within lymphoid organs where

apoptotic DC could potentially outnumber live DC.

Systemic autoimmune diseases can be modeled in transgenic mice harboring defects in DC

apoptosis (Chen et al., 2006) but not in mice with apoptosis defects in T cells and B cells

(Doerfler et al., 2000; Newton et al., 1998; Walsh et al., 1998). Our study shows that in addition

to the dogma of DC apoptosis as a mechanism to eliminate activated DC to prevent hyper-

activation of the immune response, DC apoptosis also plays an active role in induction and

maintenance of tolerance through induction of Treg, whereby defects in DC apoptosis may

trigger autoimmunity. Studies have documented a substantial depletion of DCs, both in sepsis

patients and septic mice linking this event to immunosuppression, which is one of the main

reasons for the fatality observed in sepsis patients (Hotchkiss et al., 2002; Tinsley et al., 2003).

In addition, increased levels of circulating Tregs have also been observed in sepsis patients

(Monneret et al., 2003). Mice over expressing anti-apoptotic proteins (selectively in DCs) are

resistant to sepsis-induced immunosuppression (Hotchkiss et al., 1999). However, the

mechanism of how DC apoptosis can contribute to immunosuppression is unclear. Our study

sheds light on immunosuppression induced by apoptotic DCs and demonstrates that DC

apoptosis in sepsis may impact the ability of viable DCs to undergo maturation and instead

promote induction and expansion of Tregs, thereby offering a potential explanation for the

observation of enhanced circulating levels of Tregs in sepsis patients. In addition to sepsis, high

levels of spontaneous DC apoptosis have also been observed in breast cancer patients (Pinzon-

Charry et al., 2007; Pinzon-Charry et al., 2006). Our study indicates that DC apoptosis in cancer

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patients may play a role in suppressing immune responses against the tumor by inducing

immunosuppression and tolerance. Therefore, prevention of DC apoptosis may enhance the

therapeutic effects of chemotherapy in tumor eradication (Pinzon-Charry et al., 2007; Pinzon-

Charry et al., 2006).

Taken together, our findings clearly show that apoptotic DCs have a potent ability to induce

immunological tolerance in vivo. Furthermore, even in cases of pre-existing inflammation,

apoptotic DCs are quite potent in suppressing inflammation and inducing expansion of Tregs. Our

findings may represent a therapeutic strategy in prevention of unwanted immune responses in

autoimmune diseases and transplantation along with inhibition of DC apoptosis to assist in tumor

eradication. Moreover, this strategy can likely be used for induction of antigen specific tolerance

towards gene therapy vectors such as HD-Ad which will allow for long-term sustained gene

expression.

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Chapter 6

Discussion and future directions

.

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6.1 Discussion:

Several vector systems, both viral and non-viral have been investigated for cystic fibrosis gene

therapy as discussed in chapter 1. However, viral based gene delivery systems have an upper

hand over non-viral methods particularly due to several folds higher efficiency of gene delivery

in vivo. Among viral vectors, the use of several vector systems is limited due to high

immunogenicity, integration into host genome, low packaging capacity and low efficiency of

gene transfer to the airway epithelial cells, which is the cellular target for cystic fibrosis gene

therapy. Adenoviral vectors in particular, are promising for CF gene therapy and several studies

have confirmed correction of CF defect using Ad mediated gene transfer to mice. Several

developments have been made in improving Ad gene vectors, specially the development of third

generation or so called helper-dependent or gutless adenoviral vectors. The HD-Ad gutless

vectors have demonstrated significant improvement over first generation vectors in terms of

toxicity, efficiency and using cell specific promoters such as cytokeratin 18, vectors can be

targeted specifically to the airway epithelium, which further reduces inflammatory response. At

the same time, due to reduced toxicity, HD-Ad systems can also be used for delivery of anti-

inflammatory siRNAs, which would have been a problem with first generation vectors.

HD-Ad vectors are associated with lowered adaptive immune response due to complete lack of

viral gene expression. However, it is important to note that although there is lack of viral gene

expression, delivery of HD-Ad particles is in fact delivery of antigen which could itself

potentiate an immune response. Upon readministration of HD-Ad vector particles, there is a

decrease in transgene expression which correlates with an increasing antibody titer against the

virus, indicating that there may be an adaptive response being mounted against HD-Ad particles

(Koehler et al., 2006). Nevertheless, none of the studies to date have assessed the ability of HD-

Ad particles to induce T cell immune response upon airway delivery (Koehler et al., 2006;

Morral et al., 1999). Therefore, it is important to study the immune response to empty HD-Ad

vectors, which lack a transgene so that the observed immune response is a direct consequence of

HD-Ad vectors. In chapter 2, we studied the immune response to HD-Ad vectors upon intranasal

administration.

Our findings show that HD-Ad vectors are able to induce maturation of DCs in vitro, which

subsequently potentiate a T cell response with approximately 6% of naive T cells undergoing

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proliferation in response to HD-Ad vectors. This indicates that even upon delivery of low dose of

HD-Ad particles, pulmonary DCs likely undergo maturation and potentiate a T cell response.

Upon delivery of a high vector dose (1.5 X 1010 vp), both CD4+ and CD8+ T cells could be

observed in the bronchoalveolar lavage fluid with approximately half of the infiltrating cells

being T cells around day 7 post vector delivery. In contrast, upon delivery of a low dose (3 fold

lower than the high dose), no T cell infiltration in bronchoalveolar lavage fluid was observed,

indicating that perhaps at low dose HD-Ad particles can evade the immune response. However,

upon looking at T cell proliferation, proliferating T cells could be observed both in the lungs and

draining lymph nodes, indicating that even at low dose HD-Ad vector can induce a T cell

response, comprised of both CD4+ and CD8+ T cells. Induction of T cell response was largely

mediated by conventional pulmonary DCs which underwent maturation and migrated to the

pulmonary draining lymph nodes where T cell response was potentiated. In addition to

conventional DCs, plasmacytoid DCs also underwent maturation but did not migrate to the

draining lymph nodes. Their role in initiating pulmonary immune response to HD-Ad vectors

warrants further investigation. It was suprising that though there was no viral gene expression,

cytotoxic CD8+ T cell was observed. This was likely primed by CD8α lymphoid DCs,

maturation of which was observed in the draining lymph nodes following delivery of HD-Ad

vectors. However, the mechanisms of how conventional DCs can transfer their HD-Ad antigen

load to CD8α lymphoid DCs in the pulmonary draining lymph nodes require further

investigation to better understand how cytotoxic immune response can be primed against HD-Ad

vectors in absence of any transgene expression.

In two recent gene therapy clinical trials using AAV serotype 2 vectors, which are thought to be

the least immunogenic; there were reports of apparent immune responses with appearance of

CD8+ effector T cells (Manno et al., 2006). This gave rise to the so-called capsid T-cell

hypothesis, whereby exogenous vector proteins may be shuttled into the MHC class I pathway

for T-cell priming (Wilson, 2007). Our results demonstrate similar phenomena occurring upon

pulmonary delivery of HD-Ad vectors, whereby we are seeing proliferation of CD8+ T cells

along with CD4+ T cells. This observation indicates that HD-Ad derived peptides are being

shuttled to MHC I pathway and cross-presented in context with MHC class I to induce CD8+ T

cell response along with presentation with MHC class II to induce CD4+ T cell response. Five

dominant CD8+ T cell epitopes on the capsid hexon protein of adenovirus have been identified

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and since HD-Ad vector does indeed has the capsid proteins, it is possible that perhaps cross-

presentation of these epitopes may play a role in inducing CD8+ T cell responses (Leen et al.,

2004). Our results indicate that perhaps CD8α DCs may play a role in cross-presenting HD-Ad

derived epitopes, since this subset of DC undergoes maturation within the draining MLN upon

HD-Ad delivery. We did not observe recruitment of CD8α DCs in the lungs as expected, since

these DCs are thought to be non-migratory. However, maturation of CD8α DCs indicates that

perhaps cDCs may be migrating to the draining MLN and transferring the antigens to CD8α

DCs, which thereby results in maturation of CD8α DCs. Mature CD8α DCs may then go on to

induce CD8+ T cell response, as has been observed with lung and sub-cutaneous infection using

HSV (Allan et al., 2006; Belz et al., 2004). Moreover, CD103+ DCs with the ability to cross-

present inhaled antigens and migrate to draining MLN have also been identified in the mouse

lung (del Rio et al., 2007). Therefore, there may be a specialized population of DCs that may be

initiating CD8+ T cell responses upon HD-Ad delivery via cross-presentation, which may

include CD103+ DCs along with CD8α DCs, which we have shown to mature in draining MLN

in response to HD-Ad. MHC:peptide complexes of immunodominant epitopes can have a half-

life over 7 days (Lazarski et al., 2005; Sant et al., 2005). Therefore, HD-Ad transduced epithelial

cells may present vector derived epitopes for substantial amount of time to allow for responding

T cells to migrate to the airways and gradually target/eliminate some transduced cells.

Furthermore, upon readministration, memory T cells are probably activated which are recruited

to the site within the first 3 days and mediate gradual clearing of transduced cells and hence may

account for observed loss in transgene expression upon vector readministration (Hikono et al.,

2006; Koehler et al., 2006). Moreover, the extent of both CD4+ as well as CD8+ proliferation in

the lung as well as draining MLN, peaked around day 6-7, which is similar to the timeline of

pulmonary T cell responses associated with other pulmonary infections such as influenza

(Lawrence and Braciale, 2004; Lawrence et al., 2005).

The pulmonary immune response to HD-Ad vector is primarily orchestrated by cDCs which

mature and migrate to draining MLN within the first 48 hours after HD-Ad delivery. This raises

the possibility that perhaps, pulmonary delivery of known DC maturation inhibitors such as

Prostaglandin E2, Bortezomib, FK778, a derivative of the active leflunomide-metabolite, within

the first 24 hours after HD-Ad vector delivery, may inhibit priming of pulmonary adaptive

immune responses by preventing maturation and thereby migration of cDCs (Sa-Nunes et al.,

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2007; Straube et al., 2007; Zeyda et al., 2005). Moreover, since T cell proliferation peaked

around a week after HD-Ad delivery, transient immunosuppression within a week after HD-Ad

vector delivery using cyclosporine A or CTLA4Ig to inhibit CD28/B7 pathway or inhibition of

CD40/CD40L pathway via CD40LIg administration, will prevent T-cell proliferation and hence

suppress adaptive immune response to HD-Ad vectors (Jooss et al., 1998; Nishiyama et al.,

2005; Yang et al., 1996b). Strategies to inhibit adaptive immune responses will inhibit induction

of memory T cells. This may go on to prevent the observed loss of transgene expression and anti-

HD-Ad antibody titer, observed in mice upon subsequent re-administrations. Our findings

indicate that with a lower dose of HD-Ad vectors, the pulmonary adaptive immune response

against HD-Ad vectors can be reduced.

Therefore, it is likely that by further increasing the efficacy of HD-Ad mediated gene transfer,

the vector dose can be reduced even further, which will likely limit the immune response against

HD-Ad vectors. Adenoviral entry into the cells requires an initial binding of viral fiber to the

CAR (Coxsackie and adenoviral receptor) receptor followed by internalization. Unfortunately,

the localization of CAR receptors to the basolateral side of epithelial cells creates a new obstacle

by reducing the efficiency of gene transfer (Grubb et al., 1994). Additionally, the airways of CF

patients are covered with a thick mucus layer due to chloride channel defects, which can further

reduce the efficiency of gene delivery to the epithelial cells (Sanders et al., 2000). In order to

increase the efficiency of Ad mediated gene delivery to the airways to reduce vector dose, in

chapter 3, we explored the use of nacystelyn (NAL), a mucolytic agent used clinically in CF

patients to reduce viscosity in the airways, in increasing the efficiency of HD-Ad mediated gene

transfer. NAL has been shown to possess antioxidant(Gillissen et al., 1997), mucolytic and anti-

inflammatory properties, thus indicating that it may be beneficial for CF patients.(Tomkiewicz et

al., 1994; Tomkiewicz et al., 1995).

Pretreatment of airways using intranasal delivery of NAL prior to gene delivery did result in a

significant enhancement of gene transfer as indicated by increased expression of viral encoded

transgene. However, the overall levels of gene expression were low, indicating that NAL by

itself may not open tight junctions to allow vector particles to reach the basolateral surface, and

therefore, some other formulation was also needed in addition to NAL, to partially fulfill this

requirement. Therefore, we explored the use of DEAE-Dextran in conjunction with NAL to

increase Ad mediated gene delivery of the airways. Pre-treatment with NAL followed by

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adenoviral vector delivery in 20 μg/ml DEAE-Dextran resulted in approximately 64 fold

enhancement in gene transfer compared to viral delivery in saline without NAL pre-treatment.

The effects of NAL and DEAE-Dextran in enhancing gene transfer were observed with both first

generation as well as HD-Ad vectors and the enhancement effects were not localized for gene

expression was enhanced all throughout the lungs. Additionally, NAL was also able to suppress

acute inflammation for mice which received NAL prior to vector delivery in DEAE-Dextran had

lower grade of pulmonary inflammation. In principle, our findings establish the efficiency of

NAL in increasing Ad mediated gene transfer to the airways. Furthermore, human airways both

in normal and particularly in CF patients secrete a lot of mucus (Fehrenbach, 2002); therefore,

we can speculate that pre-treatment with NAL may result in even higher levels of enhancement

of adenoviral vector-mediated gene delivery than seen in mice, mostly due to its mucolytic

activity in combination with anti-inflammatory effects. Further work is needed to better identify

the molecular mechanisms used by NAL and DEAE-Dextran in enhancing gene transfer to the

airways.

These findings made as conclude that although the vector dose needed to achieve substantial

levels of gene expression can be reduced using a combination of NAL and DEAE-Dextran,

immune response will eventually ensue and reduce long term gene expression due to a

requirement for vector readministration. Therefore, it is highly important to develop strategies

that can specifically suppress immune response towards Ad vectors. Moreover, such

interventions need to be devised in a manner where immune response only towards certain HD-

Ad antigens is suppressed but immune response to other pathogens is maintained. Therefore,

there is a need to develop antigen specific tolerance strategies which specifically target HD-Ad

vectors.

Immature DCs are potent inducers of tolerance for they prevent induction of T cell response and

promote generation of antigen specific Tregs as discussed in chapter 1. Recent studies have

highlighted the ability of immature DCs to drive tolerance tolerance induction, for delivery of

IL-10 differentiated DCs can mediate tolerance induction in asthmatic mice through generation

of Tregs (Huang et al., 2011; Lu et al., 2011). Hence, to induce tolerance towards HD-Ad

vectors, we explored the use of IL-10 to generate immature DCs, which were refractory to HD-

Ad vectors and can subsequently be used to induce HD-Ad specific tolerance as discussed in

chapter 4. DCs generated from bone marrow cells under high concentration of IL-10 are

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refractory to HD-Ad induced maturation and instead prime generation of IL-10 secreting Tr1

Tregs which can suppress HD-Ad induced T cell proliferation. Additionally, delivery of HD-Ad

pulsed IL-10 modified DCs to mice suppressed maturation and migration of pulmonary DCs in

response to intranasal delivery of HD-Ad vectors. Since, DCs have a short life-span, it is likely

that Tr1 cells induced by HD-Ad pulsed IL-10 modified DCs, secreted IL-10 in response to

intranasal challenge with HD-Ad vectors, which had an anti-inflammatory affect on pulmonary

DCs, thereby suppression their maturation (Morelli and Thomson, 2007). Moreover, Tr1 cells

could also directly suppress DC maturation, for Tregs have been shown to directly suppress DC

maturation via binding of lymphocyte activation gene-3 (LAG) on Treg surface to MHC II on

DC surface (Liang et al., 2008).

Furthermore, there was suppression of T cell as well as antibody response against HD-Ad

vectors. This was likely due to impairment of pulmonary DC migration upon HD-Ad delivery,

which prevented induction of T cell response against HD-Ad vectors and also due to induction of

HD-Ad Tregs which further suppressed T cell response (Lambrecht, 2001). T cell response is

critical for induction of B cell mediated antibody response (Parker, 1993). Therefore,

suppression of T cell response against HD-Ad vectors upon immunization with HD-Ad pulsed

IL-10 modified DCs also suppressed generation of anti-Ad antibody titers. The suppression of

HD-Ad induced T cell response could be observed even after three rounds of HD-Ad delivery.

Additionally, T cells from the pulmonary draining lymph nodes of tolerance mice secreted

elevated levels of IL-10 following in vitro stimulation with HD-Ad pulsed DCs, indicating

generation of HD-Ad specific Tr1 cells in vivo. Furthermore, following three rounds of HD-Ad

vector readministrations, gene expression was sustained in the lungs of tolerant mice compared

to non-tolerant mice. In contrast, sustained gene expression was not observed in the lungs of

mice which received IL-10 modified DCs pulsed with ovalbumin, indicating that in tolerant

mice, tolerance induction was specific towards HD-Ad vectors.

Further studies are needed to identify the immunodominant HD-Ad derived epitopes against

which an immune response is potentiated following vector readministrations. Subsequently,

antigen specific tolerance can be induced towards the specific epitopes to enhance the antigen

specificity of tolerance induction.

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Environments with significant DC apoptosis are associated with increased levels of circulating

Tregs along with an impairment of DC function as discussed in chapter 1. Furthermore, mature

DCs play a very important role in induction of immune response. Therefore, impairment of DC

function could also promote tolerance by preventing induction of a T cell response. In chapter 5,

we explored the interactions between apoptotic and viable DCs and developed a novel apoptotic

DC based strategy to generate antigen specific tolerance in vivo. Our findings show that

apoptotic DCs are rapidly taken up by viable DCs, which prevents their subsequent maturation

and instead promotes TGF-β1 secretion via mTOR signalling pathway, which promotes

generation of Tregs. Induction of Treg was specific to uptake of apoptotic DCs by viable DCs for

uptake of necrotic DCs or apoptotic splenocytes failed to induce Treg generation. In vivo

delivery of apoptotic DCs results in selective uptake by viable DCs in the spleen and draining

lymph nodes. Furthermore, if an antigen is delivered after delivery of apoptotic DCs in vivo,

there is suppression of DC maturation and migration to the draining lymph nodes, which

suppresses immune response to the antigen. Moreover, T cell response against the antigen is also

suppressed due to generation of antigen specific Foxp3+ Tregs. These findings illustrate that

delivery of apoptotic DCs followed by delivery of an antigen can be used to generate antigen

specific tolerance. It is likely that this particular strategy can be employed for generation of HD-

Ad specific tolerance, but this requires further investigation.

Studies have shown that DC can take up antigen from dying cells and cross-present the antigenic

material onto both MHC I as well as MHC II (Albert et al., 1998b; Inaba et al., 1998). However,

these studies relied on the use of mature DC to phagocytose apoptotic cells. We can speculate

that perhaps in a physiological setting, if the causative agent of DC apoptosis is an infection, then

it is usually the semi-mature or mature viable DC in close proximity that take up apoptotic DC.

Thereby, these viable DC can cross-present the antigen and then prime a T cell response rather

than induction of tolerance, as seen in our study.

Studies have also shown that this interaction with phosphatidylserine results in suppression of

macrophage activation and induction of TGF-β1 gene expression (De et al., 2002; Fadok et al.,

1998; Huynh et al., 2002). It is tempting to argue that conversion of viable immature DC to

tolerogenic DC with a potential to induce Treg via secretion of TGF-β1, is largely

phosphatidylserine dependent. However, in our study, when viable immature DC were exposed

to apoptotic splenocytes, no increase in TGF-β1 secretion was observed and previous studies

242

have also indicated that exposure of murine DC to apoptotic cells or phosphatidylserine does not

induce TGF-β1 secretion (Chen et al., 2004; Morelli et al., 2003; Takahashi and Kobayashi,

2003). Therefore, it is likely that the ability to secrete TGF-β1 and to induce Foxp3+ Treg, may

be dependent on the uptake of apoptotic DC by viable DC, which has not been described

previously, and could be independent of phosphatidylserine. It is feasible that as DC undergo

apoptosis, there is exposure of phosphatidylserine, which may play a passive role in suppression

of DC by suppressing the ability of DC to undergo maturation without any induction of Foxp3+

Treg.. We propose that uptake of apoptotic DC in particular, triggers signalling through a

previously unidentified receptor in viable DC that induces TGF-β1 secretion.

Our findings identify that the release of TGF-β1 upon uptake of apoptotic DC by viable DC is

regulated at translational level via mTOR pathway.Mammalian target of rapamycin (mTOR), a

serine/threonine protein kinase is a regulator of translation and its major substrates include

p70S60K serine/threonine kinase and 4E Binding protein (4EBP-1). mTOR phosphorylates

4EBP-1 which results in release of protein translation initiation factor eIF4E. eIF4E plays a role

in enhancing rates of translation of capped mRNA which also includes TGF-β1. mTOR is likely

regulated upstream by PI3/Akt pathway and Rho A has previously been shown to induce PI3

pathway to prevent myoblast death(Reuveny et al., 2004). Therefore it is likely that RhoA

induces PI3K which phosphorylates mTOR resulting in release of eIF4E, which further results in

increased translation of TGF-β1 mRNA.

Taken together, our findings clearly show that apoptotic DCs have a potent ability to induce

immunological tolerance in vivo. Furthermore, even in cases of pre-existing inflammation,

apoptotic DCs are quite potent in suppressing inflammation and inducing expansion of Tregs.

Moreover, this strategy can likely be used for induction of antigen specific tolerance towards

gene therapy vectors such as HD-Ad which will allow for long term sustained gene expression.

Altogether, in this thesis we have tried to overcome the barriers associated with Ad mediated

gene delivery to the airways. Although with the use of NAL and DEAE-Dextran, viral dose can

be substantially reduced, immune response to HD-Ad vectors upon intranasal delivery remains a

challenge. To overcome this challenge, we describe two new strategies: first one is based on

using HD-Ad pulsed DCs generated in presence of IL-10 to generate tolerance towards HD-Ad

vectors and the second strategy is based on delivery of apoptotic DCs followed by delivery of

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antigen i.e. HD-Ad to generate antigen specific tolerance in vivo. Although our findings provide

the proof of principle in inducing antigen specific tolerance using the two approaches, further

work is needed as discussed under future directions to further develop these strategies for clinical

use.

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6.2 Future Directions

6.2.1 Dissecting the role of different DC subsets in initiating immune response against HD-Ad

vectors

In our study, described in chapter 2, we identified a major role of conventional pulmonary

CD11c+ DCs in initiating immune response against HD-Ad vectors. However, it needs to be

noted that the lung environment consists of 3 DC sub-populations which include

CD103+CD11chigh DCs in the intraepithelial network, CD103-CD11chigh DCs in the lamina

propria of conducting airways along with plasmacytoid DCs(Lambrecht and Hammad, 2009).

Although it is likely that CD103+CD11chigh DCs in the intraepithelial network are primarily

involved in HD-Ad induced T cell response due to their physical location, studies need to be

carried out to better identify the role of various pulmonary DC subsets in orchestrating the

immune response against HD-Ad vectors. In our study, although we observed maturation of IFN

producing plasmacytoid DCs, their migration to the draining lymph nodes was not observed.

TLR recognition of virus leads to IFN production, which positively feedbacks via IFNR to drive

further IFN production by pDCs (Kumagai et al., 2009). In addition to serving as a source of

IFN, pDCs have also been shown to be critical for differentiation of activated B cells to plasma

cells via secretion of Type I interferons and IL-6 (Jego et al., 2003). The role of pDCs in

inducing immune response against HD-Ad vectors can be assessed by performing selective

depletion of pDCs in the lungs. 120G8 antibody has been shown to be highly effective in

depleting pDC populations in vivo (de Heer et al., 2004; Smit et al., 2006). pDC depletion can be

carried out prior to HD-Ad delivery and adaptive immune response can be subsequently assessed

after intranasal HD-Ad and compared to mice not depletion of pDC population, to identify

whether pDCs play a role in initiating immune response against HD-Ad vectors. If mice depleted

of pDCs mount an impaired immune response against HD-Ad and the role of pDC is mainly to

act as local producers of IFN, then systemic delivery of IFN-α can be carried out in mice

depleted of pDC to assess if adaptive immune response to airway HD-Ad delivery can be

restored. In case the immune response is restored upon delivery of IFN-α, then that would

indicate that the major role of pDCs is to act as local producers of IFN during an immune

response against HD-Ad vectors.

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In our studies we observed maturation of CD8α+ DCs in the draining lymph nodes, which we

think could mediate induction of CD8+ T cell response against HD-Ad vectors. However, it is

extremely important to understand how HD-Ad peptide: MHC complexes are transferred from

conventional pulmonary DCs to CD8α+ DCs in the draining lymph nodes. In order to confirm

whether lymphoid CD8α+ DCs are the true initiators of CD8+ T cell response against HD-Ad

vectors, Langerin-DTREGFP transgenic mice can be used. Langerin-DTREGFP mice express

human diphtheria toxin receptor under control of langerin promoter. Therefore, treatment of

these mice with diphtheria toxin selectively depletes langerin+ DCs, which includes CD8α+ DCs

(Kissenpfennig et al., 2005; Petersen et al., 2011). If the induction of CD8+ T cell response upon

HD-Ad delivery is impaired in langerin-DTREGFP mice following CD8α+ DC depletion, then

that would indicate that CD8α+ DCs are primarily responsible for inducing CD8+ T cell response

against HD-Ad vectors. Subsequent studies can be carried out to identify the mechanisms of

HD-Ad antigen transfer by conventional DCs to CD8α+ DCs. To assess if upon HD-Ad delivery,

conventional DCs undergo apoptosis in the draining lymph nodes and antigen is subsequently

transferred to CD8α+ DCs, apoptosis of conventional pulmonary DCs labelled by intranasal

delivery of FITC-Dextran can be monitored in the draining lymph nodes upon HD-Ad delivery

by Flow cytometry. In case apoptosis is observed, the next step will be to identify whether

exosomes released by dying conventional DCs can be taken up by CD8α+ DCs which

subsequently prime a CD8+ T cell response. This can be assessed by isolating exosomes derived

from HD-Ad treated conventional DCs followed by the culture of CD8α+ DCs with the

exosomes. Next, the ability of CD8α+ DCs cultured with exosomes to prime CD8+ T cell

response can be assessed in vitro.

DCs are the key initiators of immune response and tolerance. Therefore it is important to

understand the mechanisms of HD-Ad recognition by DCs. Culture of DCs with HD-Ad vectors

drives DC maturation. Macrophages have recently been shown to recognize adenoviral DNA in

the cytosol by the inflammasome, which is independent of TLR recognition (Muruve et al.,

2008). The inflammasome is comprised of innate cytosolic molecular complex among which

ASC and NALP3 appear to be important for recognition of adenoviral DNA by macrophages

(Muruve et al., 2008). In order to assess if similar mechanism is employed by DC mediated

recognition of HD-Ad particles, ASC-/- and NALP3-/- DCs can be generated from ASC and

NALP3 deficient mice respectively and can subsequently be treated with HD-Ad particles. In

246

case, the maturation of ASC-/- and NALP3-/- DCs is impaired then that would indicate that

inflammasome mediated recognition plays an important role recognition of HD-Ad particles by

DCs. Furthermore, DCs can also be treated with HD-Ad capsids deprived of any packaging DNA

to identify presence of nucleic acid independent mechanisms in recognition of HD-Ad particles

by DCs.

6.2.2 Identification of mechanisms involved in NAL and DEAE-Dextran mediated

enhancement of HD-Ad gene transfer to the airways

DEAE-Dextran is a cationic molecule which forms complexes with Ad particles and enhances

gene transfer to the airway epithelial cells. The enhancement is largely mediated by the cationic

nature of DEAE-Dextran which allows for Ad entry into cells which are relatively resistant to Ad

infection (Fasbender et al., 1997; Kaplan et al., 1998). However, it is still important to

investigate whether complex of Ad particles with DEAE-Dextran have any effect on tight

junctions for compounds such as sodium caprate increase the efficiency of gene transfer by

opening tight junctions (Gregory et al., 2003). The complex of Ad particles and DEAE-Dextran

is needed to be used shortly after preparation for optimum gene transfer. Therefore, studies need

to be carried out to identify the time span after complex preparation within which the ability of

complexes to infect airway epithelial cells remains intact. Additionally, in our study, described in

chapter 2, we looked at 10ug/ml and 20 ug/ml DEAE-Dextran concentrations to generate Ad

complexes. However, thorough studies need to be carried out using different ranges of DEAE-

Dextran concentration to identify the optimum concentration for maximizing Ad mediated gene

delivery to the airways.

In our study, described in chapter 3, pretreatment with NAL led to further increase in the

efficiency of gene transfer. NAL has previously been shown to reduce IL-8 production by

macrophage cell lines and has also been shown to reduce LPS induced neutrophil recruitment to

the mouse lungs (Antonicelli et al., 2004). Additionally, NAL has also been shown to suppress

DC maturation (Vosters et al., 2003). Furthermore, studies have confirmed mucolytic effects of

NAL (Ferrari et al., 2001). In our study, we identified anti-inflammatory effects of NAL in

suppressing pulmonary inflammation in response to Ad particles. It is likely that since mice do

not have lots of mucus in their airways, beneficial effects of NAL were most likely due to its

anti-inflammatory effects. Studies need to be carried out to look at pulmonary DC maturation in

247

mice exposed to Ad vectors with or without NAL pretreatment to identify whether NAL

suppressed Ad vector induced pulmonary DC maturation. Moreover, cytokine analysis of BALF

from mice exposed to Ad vectors with or without NAL pretreatment shall be carried out to

identify pro inflammatory cytokines suppressed by NAL. Once these target cytokines have been

identified, primary airway epithelial cells can be isolated and pretreated with NAL followed by

infection with Ad particles and molecular analysis can be carried out to better dissect out the

mechanism of NAL induced immunosuppression.

6.2.3 Identification of the mechanisms of tolerance induction towards HD-Ad particles by

immature DCs

Our findings in chapter 4 provide the proof of principle of tolerance induction towards HD-Ad

vectors in mice upon delivery of HD-Ad pulsed DCs cultured in presence of IL-10. However, it

is important to identify the mechanisms of tolerance induction. Our in vitro findings indicate that

HD-Ad pulsed DCs cultured in presence of IL-10 likely induce generation of IL-10 producing

Tr1 regulatory T cells. Furthermore, in vivo findings indicate that delivery of HD-Ad pulsed IL-

10 cultured DCs led to induction of IL-10 producing T cells, with specificity towards HD-Ad

vectors. To identify if tolerance in vivo was achieved through generation of Tr1 cells, lymph

nodes can be isolated from mice which receive HD-Ad pulsed IL-10 cultured DCs and T cells

can be isolated, which will include the Treg population. The isolated T cells can be adoptively

transferred to naive mice and then the animals can be exposed to HD-Ad particles. If mice which

received T cells are impaired in induction of immune response against HD-Ad vectors, then that

would indicate that induction of HD-Ad specific Tr1 was the mechanism employed by HD-Ad

pulsed IL-10 cultured DCs in inducing tolerance towards HD-Ad particles.

Although we confirmed suppression of HD-Ad specific T cell proliferation in tolerance mice, the

ultimate goal of tolerance induction is to achieve sustained gene expression in tolerant mice. In

our studies we have confirmed sustained gene expression in mice which recieived HD-Ad pulsed

IL-10 modified DCs following three rounds of HD-Ad vector readministrations. However,

studies are needed to perform more readministrations to identify for how long tolerance can be

maintained in mice following delivery of HD-Ad pulsed IL-10 modified DCs. Furthermore, once

a break of tolerance is observed, studies need to be conducted to identify if further rounds of DC

248

administrations can be carried out to increase the effectiveness of tolerance induction towards

HD-Ad vectors.

Additionally, in the study, LacZ was used as a transgene. Studies need to be carried out with a

therapeutic gene such as CFTR to identify if CFTR expression can be sustained in mice

following delivery of HD-Ad pulsed IL-10 modified DCs following multiple rounds of vector

readminstrations.

6.2.4 Tolerance induction towards HD-Ad vectors using apoptotic DCs

Our findings in chapter 5 establish the use of apoptotic DCs in inducing antigen specific

tolerance and also shed light on the underlying mechanisms. However, in our studies we used

ovalbumin as a prototypic antigen, delivery of which after delivery of apoptotic DCs to mice

induced ovalbumin specific tolerance induction. Experiments need to be carried out, whereby

mice are given apoptotic DCs followed by delivery of HD-Ad vectors to assess if apoptotic DCs

can induce HD-Ad vector specific tolerance.

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6.3 Conclusion:

The success of adenoviral mediated gene therapy is hindered by reduced efficiency of gene

transfer to the airway epithelial along with an induction of immune response to vectors.

Although, HD-Ad vectors have lowered immunogenicity than earlier generation of adenoviral

vectors due to lack of viral gene expression, immune response still remains a problem due to a

need to perform vector readministrations. I have determined that by formulating adenoviral

vectors in DEAE-Dextran, and performing delivery to the airways after pretreatment with NAL,

the efficiency of gene transfer can be increased by several folds and hence the quantity of vector

particles can be effectively reduced. However, my findings also indicate that intranasal delivery

of HD-Ad particles even at a low dose can lead to an induction of pulmonary adaptive immune

response, which includes both helper and cytotoxic T cells. Therefore, to circumvent the immune

response, I have developed two novel strategies for induction of antigen specific tolerance. I

have shown that delivery of HD-Ad pulsed DCs derived in presence of IL-10 to mice can

suppress HD-Ad induced T cell proliferation for upto three rounds of gene delivery and lead to

stable gene expression following vector readministrations. Additionally, I have shown that

delivery of antigen prior to delivery of apoptotic DCs can also induce antigen specific tolerance

through generation of Tregs. Further studies are needed to confirm long lasting gene expression

upon use of these two strategies to induce HD-Ad specific tolerance. Overall my findings

identify that delivery of HD-Ad vectors can potentiate an adaptive immune response and at the

same time offer three strategies which can be used to increase efficacy of gene transfer and to

reduce immune response against HD-Ad vectors, which can allow for multiple vector

readministrations to the lung along with sustained gene expression.

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