Vitamin D and Immunomodulation in Bronchial Asthma

By

Tanupriya Agrawal

A (THESIS/DISSERTATION) Submitted to the Faculty of the Graduate School of Creighton University in

partial fulfillment of the requirements for the Degree of Doctor of Philosophy in the Department of Biomedical Sciences

Omaha NE, 2012

(June 11th 2012)

i

Dedicated to my Parents, Dr. Sushma and Dr. Subhash Agrawal

who sacrificed so much for me to be where I am today &

My Beloved Husband

Dr. Gaurav Gupta

1

Abstract

Asthma is one of the most common chronic inflammatory diseases of the airways.

The chronic inflammation is associated with airway hyperresponsiveness (AHR), airway

obstruction and airway remodeling. In asthmatics the airway epithelium produces

increased amounts of proinflammatory cytokines including TNF-α (Tumour Necrosis

Factor-α), IL-1(Interleukin-1), IL-6(Interleukin-6), IL-8(Interleukin-8),and

Transforming Growth Factor-β (TGF-β). Airway Smooth Muscle Cells (ASMCs) when

activated in an antigen sensitized state, behave in an autocrine and paracrine manner by

producing and responding to mediators such as TNF-α, IL-1β, IL-5, IL-6, IL-8, IL-17,

GM-CSF (Granulocyte macrophage colony-stimulating factor), TGF-β, RANTES

(Regulated upon Activation, Normal T-cell Expressed, and Secreted), Eotaxin, MCP-

1,2,3 (monocyte chemotactic protein). NF-κB (nuclear factor kappa-light-chain-

enhancer of activated B cells) is a key transcriptional factor that regulates and co-

ordinates the expression of various inflammatory genes and inflammatory mediators.

The p50 and RelA subunits of NF-κB are translocated to the nucleus by importin α3 and

importin α4.

The potent immunomodulatory role of Vitamin D in both innate and adaptive

immunity has recently been appreciated. Calcitriol exerts its action through the Vitamin

D receptor (VDR), which is a high affinity nuclear receptor. VDR functions as a

transcription factor that alters the transcription of target genes involved in a wide

spectrum of biological responses. The effects of vitamin D on the immune system make

it a potent immunoregulator and immunomodulator. Individuals with a vitamin D

deficiency are predisposed to asthma and allergies in the presence of other

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environmental stimuli. Lower serum vitamin D levels are associated with airway

hyperresponsiveness and increased asthma severity. The central hypothesis of these

studies is that vitamin D reduces allergic airway inflammation and airway

hyperresponsiveness by inhibiting the transcription and translation of importin α3 and

attenuating the migration of NF-κB to the nucleus of airway cells.

Cultured human bronchial smooth muscle cells (HBSMCs) were serum starved for

24 hrs and then stimulated with calcitriol in the presence and absence of cytokines,

TNF-α, IL-1β, and/or IL-10 for 24 hrs. The mRNA transcripts and protein expression of

importin α3 and α4 were analyzed using qPCR and immunoblotting respectively. The

nuclear proteins were extracted and RelA expression was also analyzed by

immunoblotting. In vivo studies included female BALB/c mice that were fed either a

vitamin D-deficient or 2,000 IU/kg or 10,000 IU/kg of vitamin D-supplemented diet.

Allergic airway inflammation in mice was induced by OVA- sensitization and challenge.

AHR to methacholine was measured using invasive and non- invasive methods .

Bronchoalveolar lavage fluid (BALF), blood, and lungs were collected at sacrifice.

In HBSMCs, calcitriol significantly decreased mRNA and protein expression of

importin α3 and protein expression of NF-кBp65 (RelA) in the nucleus. Calcitriol

attenuated the effects of TNF-α and IL-1β and upregulated mRNA and protein

expression of importin α3. IL-10 significantly decreased the TNF-α induced expression

of importin α3 and this effect was further potentiated by calcitriol.

Vitamin D-deficient OVA-sensitized and challenged mice exhibited exaggerated

response to methacholine, with increased airway inflammation compared to vitamin D

sufficient and supplemented OVA-sensitized and challenged mice. Interestingly, the

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OVA-sensitized and challenged supplemented group had reduced allergic airway

inflammation compared to vitamin D sufficient OVA-sensitized and challenged mice but

not to the level of PBS control animals. Similarly, the levels of cytokines IL-5, IL-6, IL-

13, IL-17 and TNF-α were higher in the vitamin D-deficient group of OVA-sensitized

mice. On the contrary, high serum levels of vitamin D in OVA-sensitized mice were

associated with lower levels of the cytokines, although cytokine levels were significantly

higher than those observed in control PBS mice of vitamin D-deficient group. Vitamin D

did not alter the levels of IL-4, RANTES or IP-10 in OVA-sensitized group. Serum

vitamin D levels positively correlated to the levels of T-regulatory cells in the PBS

control group; OVA-sensitization decreased the number of T-regulatory cells. This was

more marked in the vitamin D deficient group. Similar trend was observed with the IL-

10 levels. Vitamin D deficiency was associated with increased expression of TNF-α and

importin α3, and decreased expression of VDR. Supplementation with vitamin D

reduced the levels of TNF-α and importin α3, and increased expression of VDR. We

postulate that increased VDR activation due to high serum vitamin D levels (25(OH) D)

may be responsible for reducing allergic airway inflammation.

The findings in this study suggest a therapeutic role for vitamin D in allergic

asthma. These results show that vitamin D supplementation alleviates allergic airway

inflammation but does not completely reverse the inflammation. We support the

practice of vitamin D supplementation as an adjunct to the treatment of asthma.

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ACKNOWLEDGEMENT

First and foremost I would like to thank Almighty God for everything He has given

me in my life.

I offer my sincerest gratitude to my supervisor, Dr. Devendra K Agrawal, for his

excellent guidance, care, patience, and providing me with an excellent atmosphere for

doing research. His deep knowledge in medicine has enabled me to transform my basic

science research into evidence based medicine. The joy and enthusiasm he has for his

research was contagious and inspirational to me. He supported me throughout my Ph.D.

with his motivation, enthusiasm, and immense knowledge whilst allowing me the room

to work in my own way and encouraging me to generate my own ideas. As a result of his

dynamic training, he developed my confidence to carry out my own independent

research in the future.

My special thanks to Dr Againdra Bewtra for he as friend, teacher, and as a

guardian gave me constant encouragement, guidance, and moral support during

difficult times.

I would like to thank the members of my Graduate Advisory Committee Dr Kristen

Drescher, Dr Diane Cullen, and Dr. Eric Patterson for their continuous support,

thoughtfulness and insightful comments.

I also want to thank Creighton University Graduate School, as well as the

Department of Biomedical Sciences, for providing me with the opportunity to pursue my

graduate degree. Also, I am thankful to all my lab members for their help and

encouragement.

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More importantly, I thank my best friend and beloved husband, Dr. Gaurav

Gupta without his invaluable assistance, support and meticulous feedback this

dissertation could not be completed.

Lastly, and most importantly, I would like to present my deepest gratitude to my

parents Dr. Subhash Agrawal and Dr. Sushma Agrawal for their dedication and the

hardships they faced to educate me which provided the foundation of this work. From

the core of my heart I thank my sisters Dr. Swati Agrawal and Dr. Shikha Agrawal and

my brother Dr. Deepesh Agrawal for their unconditional love, constant support and

encouragement to believe in myself and pursue my dreams.

The study was funded by NIH grant R01HL085680 and R01AI75315 to Dr.

Devendra K. Agrawal.

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TABLE OF CONTENTS

ABSTRACT........................................................................................ 1 ACKNOWLEDGEMENTS................................................................... 4 TABLE OF CONTENTS.......................................................................6 LIST OF FIGURES……........................................................................13

LIST OF TABLES……..........................................................................15 ABBREVIATIONS..............................................................................16 CHAPTER 1 : Introduction……...........................................................20 1.1 Asthma ........................................................................................21 1.1.1 Definition of Asthma ..........................................................21 1.1.2 Epidemiology of Asthma.....................................................21

1.1.3 Economic Burden of Asthma .............................................22

1.1.4 Etiology of Asthma............................................................22

1.1.5: Classification of Asthma....................................................24

1.1.6 Pathogenesis of Asthma .....................................................25

1.1.6-1 Airway Inflammation…............................................26

1.1.6-2 Airway Obstruction ……………………………….….……… 27

1.1.6-3 Bronchial Hyperresponsiveness ......................... 29

1.1.7 Structural Changes in the Airways .....................................29

7

1.1.8 Airway Epithelium in Asthma ...........................................30

1.1.9 Airway Smooth Muscles in Asthma…………………………..……31

1.1.10 Cytokines in Asthma…………………………………………………..31

Interleukin-4…………………………………..……………………………………..…....32

Interleukin-5………………………………..………………………………….…………..33

Interleukin-9…………………………………..………………………………….………..33

Interleukin-13…………………………………….……………………...…………....…..34

Interleukin-17………………………………………….…………………………….…..34

Tumour Necrosis Factor -α ………………………………….……..……….………34

Interleukin -1β……………………………………………………………….………….35

Interleukin-10 ……………………………………………..……………………………36

Chemokines………………………………………………………………………………36

1.2 NF-κB……………………………………………………………………….……..…..38

1.2.1 Role of NF-κB in Asthma………………….……………………………39

1.2.2 Rel Protein Family ………………………………….……………..…….40

1.2.3 Nuclear Transport of NF-κB………………………………………….40

1.3 Nuclear-Cytoplasmic Transport…………….…………………….………….43

1.3.1 Importins and Exportins………………………………………………43

1.3.2 Import ……………………………………………….………………………46

1.3.3 Export ……………………………………………………….……………….46

1.3.4 Importin α …………………………………………..……………….……..49

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1.3.5 Importin β ……………………………...………………………………….52

1.3.6 The RanGTPase System……………….………………………………52

1.4. Immunomodulators in Asthma ………………………………………………54

1.4.1 Vitamin D …………………………………….……………………………54

1.4.2 Vitamin D Receptor………...…………………………..………………55

1.4.3 Vitamin D Metabolism…….………………………….…………….….56

1.4.5 VDR Signaling Pathway……….…………………….………………..59

1.4.6 Epidemiology of Vitamin D Deficiency…………………………..61

1.4.7 Vitamin D Optimal Status/Dietary Requirements …….….…62

1.4.8 Immunomodulating Effects of Vitamin D……..………………..63

1.4.8-1 Monocyte/Macrophages and Vitamin D …………..…..64

1.4.8-2 Dendritic Cells and Vitamin D……….………………………65

1.4.8-3 T Cells and Vitamin D ……….………………………………...65

Regulatory T-cells or Tregs………………………….………...66

Th17 cells……………………………………………………………..67

1.4.8-4 B Cells and Vitamin D ……………………………………….…67

1.4.9 Vitamin D and Asthma………………….……………………….…….68

1.5 : Central Hypothesis………………………………..………………………………72

Section 2: Materials and Methods …….………….……………………………….73

2.1 Cell Culture and Cell Stimulation……………………………………..74

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2.2 RNA Isolation: …………………………………………..………….……..75

2.3 Reverse Transcription……………..………………………..…………..76

2.4 Real-Time PCR…………………………………………………..…………76

2.5 Transfection of the Cells………………………….…..…………………78

2.6 Protein Isolation…………………………………….………..……………79

2.7 Nuclear Protein Extraction …………………………………..………..79

2.8 Immunoblotting ……………………………………………………………80

2.9 Annexin V Assay for Apoptosis……………..………………………...81

2.10.1 Animals and diets …………………………………….………………82

Vitamin D-Deficient mice model………………………….……..……82

Vitamin D-Sufficient mice model…………………………..…….…..82

Vitamin D-Supplemented mice model………….……………....……83

2.10.2 OVA-Sensitization and Challenge……………..….………83

2.10.3 Induction of Allergic Airway Inflammation ……...…84

2.10.4 Non-Invasive Measurement for Assessment of

Pulmonary Function…………………………………..….……85

Advantages of Non-Invasive WBP………………………...…..85

Disadvantages of Non-Invasive WBP……..…….………85

2.10.5 Invasive Measurement for Assessment of

Pulmonary Function………….…………………….……....…86

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2.11 Measurement of Serum 25(OH) D Levels and Serum

Calcium Levels…………………………………………..….……….……..87

2.12 Bronchoalveolar Lavage Fluid (BALF) and Multiplex

Cytokine Analysis………………………………….………….…………………..87

2.13 Histology and Lung Tissue Staining……………………….………..89

2.14 Trichrome Staining in Lung Sections Showing

Collagen Deposition………………………………….…………..………89

2.15 PAS Staining in Lung Sections Showing

Mucus Deposition.………………………….….……..……….………….90

2.16 Immunofluorescence ………………………………..……..…………..91

2.17 T-regulatory Cell Identification …………………..………..……….92

2.18 Data Processing and Statistical Analysis ………….………………92

2.19 Power Analysis ……………………………….…………………….………93

Section 3: Results……………………………………………….………………….……94

3.1: Effect of Calcitriol on Expression of VDR and CYP24A1

and CYP27B1…………………...............………………………………………….….95

3.2: Effect of Calcitriol on Expression of importin α3 and

importin α4…… ….……………………………………………………………..……..….97

3.3: Time dependent Effect of Calcitriol on VDR, CYP24A1, CYP27B1

and Importin α3 .…………………..…………………………..……………..…………99

3.4: Effect of Calcitriol on Apoptosis …………………………………………..…101

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3.5: Effect of Calcitriol on VDR Knockdown Cells on

Importin α3 Expression ……………………………………………………………….104

3.6: Nuclear Expression of RelA on Treatment with TNF-α……….………106

3.7: Effect of Calcitriol on RelA Expression on Importin α3

Knockdown Cells……………………………………….………………………….…….108

3.8: Effect of Calcitriol and TNF-α on Importin α3 ………………….………111

3.9: Effect of Calcitriol and IL-1β on Importin α3 …………………….……..113

3.10: Effect of Calcitriol, TNF-α and IL-10 on Importin α3 ….…………..115

3.11: Serum 25(OH)D Levels in Mice …….……………………………………….117

3.12: Pulmonary Function Measurement ……….………………………………118

3.13: Histology of Lungs ……………………………………………………………….122

3.14: Total and Differential Leukocyte Counts in

BALF ………………..……………………………..………………………………………..124

3.15: Th2 cytokines Levels in BALF ………………………..……………………..126

3.16: Levels of TNF-α, IL-6and IL-17 in BALF …………………………………128

3.17: Levels of RANTES and IP-10 in BALF ………………….…………………130

3.18: Percentage of CD4+CD25+ T-cells in Blood ……………………….……132

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3.19: Expression of TNF-α in Lungs……………………..…………………………135

3.20: Expression of VDR in Lungs………………………………………………….137

3.21: Expression of importin α3 in Lungs…………………………….………….140

Section 4: Discussion …………….………………..…………………………………..143

Conclusion…………………………………..…………………………………….153

Outstanding Questions and Future Directions……………………….154

Section 5: Bibliography…………..………...………………..……………………….156

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LIST OF FIGURES

Figure-1: Pathogenesis of Allergic Asthma……………………………………………………………25

Figure-2: Histological Features of Asthmatic Bronchiole ………………………………………28

Figure-3: NF-κB Signaling Pathway……………………………………………………………………..42

Figure-4: Nuclear Import and Export …………………………………………………………………45

Figure-5: Mechanism of Action of Importins and Exportins:…………………………………48

Figure-6: Metabolism of Vitamin D…………………………………………………………………….58

Figure-7: VDR signaling pathway ………………………………………………………………………60

Figure-8: Protocol for OVA-sensitization And Challenge ………………………………….…84

Figure-9: Effect of Calcitriol on Expression of VDR and CYP24A1

And CYP27B1……………………………………………………………………………………..….……..…96

Figure-10: Effect of Calcitriol on Expression of Importin α3 and

Importin α4 ……………………………………………………………………………………………...…..98

Figure-11: Time Dependent Effect of Calcitriol on VDR, CYP24A1, CYP27B1

and Importin α3 …………………………………………………………………….…..…………..……100

Figure-12: Effect of Calcitriol on Apoptosis …………………………………………….……..102

Figure-13: Effect of Calcitriol on VDR Knockdown Cells on Importin α3

Expression:………………………………………………………………………………………..…………105

Figure-14: Nuclear Expression of RelA on Treatment with TNF-α……………………..107

Figure-15: Effect of calcitriol on RelA Expression on Importin α3

Knockdown Cells…………………………………………………..………………………………………110

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Figure-16: Effect of Calcitriol and TNF-α on Importin α3 ……………………………………112

Figure-17: Effect of Calcitriol and IL-1β on Importin α3 …………………………………..…114

Figure-18: Effect of Calcitriol, TNF-α and IL-10 on Importin α3 ………………….………116

Figure-19: Pulmonary Function Measurement: …………………………………………………120

Figure-20: Histology of Lungs: ……………………………………………………………….………123

Figure-21: Total and Differential Leukocyte Counts in BALF ……………………….……125

Figure-22: Th2 Cytokines Levels in BALF ………………………………………….…..………127

Figure-23: Levels of TNF-α, IL-6and IL-17 in BALF ……………………………..……….…129

Figure-24: Levels of RANTES and IP-10 in BALF …………………………………..…………131

Figure-25: Percentage of CD4+CD25+ T-cells in Blood ………………….……….…..……133

Figure-26: Expression of TNF-α in the Lungs…………………………………………………..136

Figure-27: Expression of VDR in the Lungs…………………………………………….………138

Figure-28: Expression of importin α3 in the Lungs…………………………..………………141

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LIST OF TABLES

Table-1: Classification of Importin alpha………………………………………………49

Table-2: Isoforms of Importin α and β and Examples of Cargo Molecules ………………………………………………………………………………………….…………….50

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ABBREVIATIONS Ag Antigen AHR Airway Hyperresponsiveness ANOVA Analysis of Variance APC Antigen Presenting Cell ASMCs Airway Smooth Muscle Cells BAL Bronchoalveolar Lavage BALB/c Bagg Albino substrain C BALF Bronchoalveolar Lavage Fluid BSA Bovine Serum Albumin CD Cluster of Differentiation CYP27B1 25 hydroxyvitamin D3 1α hydroxylase CYP24A1 1α-25 dihydroxyvitamin D3 24 hydroxylase DAPI 4,’6-diamidino-2-phenylindole ELISA Enzyme-Linked Immunosorbent Assay FACS Fluorescence Activated Cell Sorter FBS Fetal Bovine Serum Fc Fragment Crystalizable FEV1 Forced Expiratory Volume in 1 second GAPDH Glyceraldehyde-3-phosphate dehydrogenase GFP Green Fluorescent Protein GM-CSF Granulocyte Macrophage Colony Stimulating Factor HBSMCs Human Bronchial Smooth Muscle Cells

17

i.p. Intraperitoneally

IACUC Institutional Animal Care and Use Committee

IL-1β Interleukin-1 Beta

IFN-γ Interferon-Gamma

Ig Immunoglobulin

IgE Immunoglobulin E

IL Interleukin

IκB Inhibitory Kappa Beta IKK Inhibitory κB kinase IL-4 Interleukin-4 IL-5 Interleukin-5 IL-6 Interleukin-6 IL-8 Interleukin-8 IL-10 Interleukin-10 IL-13 Interleukin-13

IL-17 Interleukin-17

IF Immunofluorescence

KPNA4 Importin α3

KPNA3 Importin α4

M-CSF Macrophage Colony Stimulating Factor

MHC Major Histocompatibility Complex

MIP Macrophage Inflammatory Protein

mRNA messenger RNA

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NE Nuclear Envelope

NES Nuclear Export Signal

NF-κB Nuclear Factor Kappa-B

NIK NF-κB-inducing kinase

NLS Nuclear Localization Signal

NPC Nuclear Pore Complex

OVA Ovalbumin

PAS Periodic acid-Schiff Stain

PBMC Peripheral Blood Mononuclear Cell

PBS Phosphate Buffered Saline

Penh Enhanced Pause

PI Propidium Iodide

RanGAP Ran GTPase activating protein

RanGEF Ran guanine nucleotide exchange factor

RanBP Ran-binding protein

RANTES Regulated upon Activation, Normal T Cell

Expressed and Presumably Secreted

RL Airway Resistance

RNA Ribonucleic Acid

RNase Ribonuclease

RT Reverse Transcription

RXR Retinoid X Receptor

SDS Sodium Dodecyl Sulphate

SEM Standard Error of the Mean

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STAT Signal Transducer and Activator of Transcription

TCR T Cell Receptor

TGF Transforming Growth Factor

TH T Helper Cell

TLR Toll Like Receptor

TNF Tumor Necrosis Factor

T regs Regulatory T Cell

VDR Vitamin D Receptor

VDRE Vitamin D Response Element

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Section 1:

INTRODUCTION

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1.1 Asthma:

1.1.1 Definition of Asthma

Asthma is a chronic inflammatory disorder of the airways [1]. The chronic

inflammation is associated with airway hyperresponsiveness (AHR), airway obstruction,

airway remodeling [2,3]. These pathological features lead to episodic and reversible

attacks of wheezing, chest tightness, shortness of breath, and coughing [4].

1.1.2 Epidemiology of asthma:

There is a worldwide increase in the prevalence of mortality and morbidity due to

asthma in the past 40 years. This upward trend is particularly apparent in children.

Nearly 300 million people worldwide are suffering from asthma, and every decade there

is increase in nearly 50% of the cases of asthma severity [5]. The prevalence of asthma

has increased in US. In 2005, an estimated 7.7% of persons living in the United States

(22.2 million) had asthma, 4.2% (12.2 million) had at least 1 asthma attack in the

previous year [6].

The annual mortality rate of asthma in US is approximately 5,000 [5]. The morbidity

rate in US is even more severe, there are 0.5 million hospitalizations and 1.5 million

emergency room visits annually [7].

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1.1.3 Economic burden of asthma

Asthma remains a significant burden in United States as well as globally, due to

patient morbidity and mortality, rising healthcare costs, and employee absenteeism.

Although asthma affects people of all ages, there is a higher prevalence among children

less than 18 years of age [4]. Uncontrolled asthma can limit daily life and is sometimes

fatal. The World Health Organization has estimated that 15 million disability-adjusted

life years are lost annually due to asthma, representing 1% of the total global disease

burden [1]. Asthma is a leading cause of activity limitation and costs our nation $56.0

billion in health care costs annually [4]. The estimated medical costs for asthma in 2010

were $5.5 billion for hospital care, $5.9 billion for prescription care, and $4.2 billion for

physicians’ services, totaling $15.6 billion in direct medical expenses[8]. Asthma

expenditure accounted loss in 11.8 million workdays and 14.7 million school days [6].

1.1.4 Etiology of asthma

Both host factors and environmental factors contribute to asthma. The host

factors include genetics, obesity and gender, all of these can influence asthma

development. Environmental factors can trigger asthma symptoms. These factors

include outdoor and indoor allergens, occupational sensitizers, infections, tobacco

smoke, and air pollution.

The indoor and outdoor allergens include house dust mites, cockroach, domestic pets,

rodents and molds [9]. Childhood infections with Respiratory Syncitial Virus or

parainfluenza lead to development of asthma later in life in an estimated 40% of the

children [10]. Over 300 substances are associated with occupational asthma

23

development [11]. Very high exposures of inhaled irritants may cause development of

“irritant induced asthma” even in non-atopic individuals [1]. Exposure to tobacco smoke

is associated with the risk of development of asthma-like symptoms in early childhood

[12]. Smoking in pregnancy has a great influence in lung development of child [13] and

infants born to smoking mothers are more prone to wheezing a hallmark of asthma in

their first year of life [13].

The hygiene hypothesis

The hygiene hypothesis states that a lack of early childhood exposure to infectious

agents and parasites increases susceptibility to allergic diseases by suppressing natural

development of the immune system [14]. Allergic diseases such as hay fever and eczema,

are observed to be less common in children from larger families, which were presumably

exposed to more infectious agents through their siblings, than in children from families

with only one child [15]. The allergic diseases are dominated by Th2 cells such as

asthma, allergic rhinitis and atopic dermatitis [16]. The mechanism of the hygiene

hypothesis is that there is insufficient stimulation of the Th1, stimulating the cell

defense of the immune system, leads to an overactive Th2, stimulating the antibody-

mediated immunity of the immune systems, leading to allergic disease [17].

24

1.1.5: Classification of Asthma

Asthma is classified into two major types depending on the stimuli-

1. Extrinsic asthma (allergic asthma or atopic asthma), is usually diagnosed on the basis

of the medical history, recurrent airway obstruction, bronchial hyperresponsiveness and

the presence of an underlying allergic diathesis as assessed by skin provocation, variably

increased eosinophil numbers in the blood and sputum, and the detection of allergen-

specific immunoglobulin E (IgE) [18-20].

2. Intrinsic asthma is found in individuals who fulfill most diagnostic criteria for

asthma, but do not have detectable skin test reactivity to common allergens or have no

increase in total or antigen-specific IgE, despite a raised eosinophil count in the blood

and/or sputum. Intrinsic asthma often triggered by non-allergic factors such as cold air,

occupational substance exposures, medication, or exercise [18-20].

1.1.6 Pathogenesis of Asthma

The pathophysiology of asthma involves the following components:

Airway inflammation

Intermittent airflow obstruction

Bronchial hyperresponsiveness

25

Figure-1: Pathogenesis of Allergic Asthma: Th2 cells release cytokines including IL-4, IL-

5, IL-9 and IL-13. These cytokines induce high levels of IgE, eosinophil recruitment, growth and

maturation of mast cells, histamine, leukotrienes and prostaglandin release.

26

1.1.6-1 Airway Inflammation

An asthmatic attack can be precipitated by inhalation of antigens as pollen, dust

mite, cat dander, and moulds. The inhaled antigen is taken up by antigen presenting

cells (APC) such as dendritic cells or macrophages which then process then present the

antigenic peptide that is bound to major histocompatibility complex (MHC) class II

molecules expressed on the APC to CD4+ T cells [21]. This causes the activation and

proliferation of CD4+T cells which differentiate into Th2 cells. The key transcription

factor regulating Th2 cell differentiation is GATA3. GATA3 is selectively expressed in

Th2 cells and its ectopic expression induces TH2 cell differentiation [22,23]. The

resultant Th2 cells release cytokines including IL-4, IL-5, IL-9 and IL-13 [22] (Figure

1). The IL-4 and IL-13 secreted by these activated Th2 cells bind to receptors on B cells

and cause isotype switching in B cells to IgE [24,25]. The B cells differentiate into

plasma cells producing high levels of soluble IgE, which in turn binds to FcεR1 receptors

on mast cells [26,27] (Figure-1). A second exposure of the immune system to the

allergen causes the allergen molecules to bind cell bound IgE resulting in the cross-

linking of IgE [28] and subsequent activation and degranulation of the mast cells

inducing transcription ,translation, and release of high levels of proinflammatory

cytokines and de novo synthesis of histamine, prostaglandins and leukotrienes [29].

These mediators and cytokines induce contraction of the airway smooth muscle

contraction, mucous secretion, and vasodilatation [29]. Mast cells, eosinophils,

epithelial cells, macrophages, and activated T lymphocytes are implicated in airway

inflammation. Inflammation is observed throughout the airways including the upper

respiratory tract and nose but the pathological effects are more pronounced in medium

sized bronchiole [1].

27

1.1.6-2 Airway Obstruction

The inflammatory mediators released to the circulation increase vascular

permeability and microvascular leakage with exudation of plasma into the airways [30].

Leakage of plasma proteins into the airways induces edema resulting in narrowing of

the airway lumen [31]. Plasma exudates in the lumen of the airways compromises

epithelial integrity and reduce mucus clearance [32] from the airway resulting in the

formation of a chronic mucous plug consisting of exudates mixed with mucus and

inflammatory and epithelial cells [29]. Together these processes contribute to the

airflow obstruction in asthmatics.

28

Figure-2: Schematic diagram showing histology of an asthmatic bronchiole- in

asthmatic bronchiole, there is thickening of the basement membrane because of sub-epithelial

fibrosis; inflammatory infiltrate in the bronchial walls, with a prominence of eosinophils and

mast cells; an increase in size of the mucosal glands and hypertrophy and hyperplasia of the

bronchial smooth muscle cells

29

1.1.6-3 Bronchial Hyperresponsiveness

Airway hyperresponsiveness is defined as an increased ability of the airways to

narrow after exposure to constrictor agonists [33] and is the characteristic functional

abnormality of asthma [1] (Figure-2). The airway narrowing results in limitated

airflow and intermittent symptoms, which causes increased resistance to airflow and

decreased expiratory flow rates [33]. Patients with airway hyperresponsiveness have a

decreased ability to expel air resulting in hyperinflation of bronchiole. The over

distention of the bronchiole helps maintain airway patency, and improves expiratory

flow. However, the over-distention of the bronchiole also modifies pulmonary

mechanics and increases the effort required to breathe [33]. Airway

hyperresponsiveness may be due to excessive contraction of airway smooth muscle

which is caused by increased volume of airway smooth muscle cells [34] or it may be due

to excessive airway narrowing by thickened airway wall by edema and structural

changes in the airways [35] .

1.1.7 Structural Changes in the Airways:

In response to cytokine milieu that is found in asthmatics, structural changes

occur in the airways of asthmatics, which are often collectively described as airway

remodeling [36]. These changes are often related to the severity of the disease and may

cause irreversible airway narrowing [37]. There is goblet cell hyperplasia/metaplasia in

the airway epithelial cell layer resulting in enormous amounts of mucus secretion [38].

Collagen fiber and proteoglycan deposition result in thickening of the subepithelial layer

30

of the bronchiole (Figure-2). The blood vessels in the airway walls undergo

proliferation (angiogenesis) due to the presence of growth factors such as vascular

endothelium growth factors (VEGF) [39]. Airway smooth muscle cells experience

hypertrophy and hyperplasia, thereby contributing to the increase in the thickness of the

airway wall [37] (Figure-2).

1.1.8 Airway Epithelium in Asthma

The epithelial cells lining the airways acts as a barrier by protecting the lungs from

injurious substance such as bacteria or cigarette smoke [40]. The bronchial epithelium

a stratified structure composed of ciliated columnar cells, goblet cells, submucosal

glands, serous cells, and basal cells [41]. Recent studies have revealed that the

respiratory epithelium is not only a physical barrier but is also highly active

metabolically. The respiratory epithelium produces a variety of inflammatory cytokines

and growth factors [42] and also plays a major role in eosinophils and neutrophils

recruitment [38] which penetrate the basement membrane resulting in hypertrophy

and sloughing of the epithelial cell layer [43]. In asthmatics the epithelium produces

higher levels of amount of pro-inflammatory cytokines and growth factors [44]. These

mediators include Tumor Necrosis Factor –α (TNF-α), IL-1, IL-6, IL-8, and TGF-β [42].

The resultant effect of the overproduction of these soluble mediators is myofibroblast

differentiation, fibroblast proliferation, and collagen production [45].

31

1.1.9 Airway Smooth muscles in Asthma

Traditionally, it was believed that airway smooth muscle cells (ASMCs) have a

passive role in airway inflammation and contract in response to pro-inflammatory

cytokines [46]. Recent reports have shown that ASMCs, in addition to their contractile

function, undergo structural changes including an increase in the ASMC mass resulting

in narrowing of the bronchial lumen [47]. The thickening of ASMCs occurs through

increased proliferation, decreased apoptosis or excessive growth and migration of

existing smooth muscle populations [47]. When ASMCs are activated in a sensitized

state produce and respond to proinflammatory and broncho-protective mediators [47].

Some of the main pro-inflammatory cytokines and chemokines secreted by the smooth

muscle cells in asthma are TNF-α, IL-1β, IL-5, IL-6, IL-8, IL-17, GM-CSF, TGF-β,

RANTES, Eotaxin, MCP-1,2,3 [48,49]. This secretory role of smooth muscles and the

secretion of these cytokines are regulated by numerous transcriptional factors such as

nuclear factor-κB (NF-κB) [50], activator protein-1 (AP-1) [51], nuclear factor of

activated T-cells (NF-AT) [52], cyclic AMP response element binding protein (CREB)

[52] and signal transduction-activated transcription factors (STAT) [53] play an

important role in the pathogenesis of asthma [54].

1.1.10 Cytokines and Chemokines in Asthma

The airway inflammation in asthma is the result of interacting network of cytokines

[55]. However, the precise functional role of each cytokine in asthma pathogenesis is not

fully understood [55]. Type 2 T-helper (Th2) cells are believed to play a central role in

airway inflammation [56]. There are a number of cytokines released from Th2 cells

32

during asthma attack [2,3]. The cytokine classification is not simply as they are

pleiotropic in nature [55]. With regard to the specific abnormalities of asthma and our

current understanding of the pathogenesis of asthma, they may be grouped as follows:

Interleukins released from Th2 cells -IL-4, IL-5, IL-9, IL-13.

Pro-inflammatory cytokines-TNF-α and IL-1β

Chemokines-RANTES, IP-10, MCP-1,Eotaxin

Growth Factors- TGF-β, EGF (Epidermal growth factor)

Anti-inflammatory cytokines- IL-10, IL-12, IL-18

Interleukin-4

IL-4 is a significant contributor in the pathogenesis of allergic inflammation [55].

IL-4 is produced by lymphocytes, eosinophils, basophils and mast cells [56]. IL-4

increases expression of class II MHC molecules and enhances expression of the low-

affinity IgE receptor (FcεRII; CD23) [57,58]. IL-4 promotes immunoglobulin synthesis

by B lymphocytes and plays an essential role in immunoglobulin class switching of

activated B lymphocytes from IgG to IgE [57,58]. IL-4 promotes the development of Th2

cells and inhibits the development of Th1 cells [57,58]. Asthmatics using IL-4 as an

inhaler had an increase in eosinophils in the sputum and increase in bronchial

responsiveness [59]. Transgenic mice overexpressing IL-4 selectively within the lung

were associated with the induction of the mucin MUC5AC gene and mucin

hypersecretion, suggesting a role for IL-4 in mucus hypersecretion [60].

33

Interleukin-5

IL-5 is required for eosinophil production, maturation, and activation and thus

contributes to bronchial hyperresponsiveness in asthma [55]. IL-5 monoclonal

antibodies can suppress allergen induced airway eosinophilia supporting a role for IL-5

in asthma [61]. IL-5 knockout mice did not demonstrate allergen induced eosinophilia

and airway hyperresponsiveness. On the contrary, mice overexpressing IL-5 in lung

epithelial cells showed increased levels of IL-5 in BAL fluid and serum and lung

histology suggestive of asthma [62].

Interleukin-9

IL-9 has a key role in the proliferation and differentiation of mast cells [63]. IL-9

stimulates proliferation of activated T cells and enhances immunoglobulins production

by B cells [64]. Mice overexpressing IL-9 have eosinophilia, mucus hyperplasia,

mastocytosis, AHR, and increased expression of other Th2 cytokines and IgE [65].

Blockage of IL-9 in allergen challenged sensitized mice results in inhibition of

pulmonary eosinophilia, mucus hypersecretion, and AHR [64]. Increased expression of

IL-9 and its receptor has been observed in asthmatic patients [66].

34

Interleukin-13

IL-13 expression is upregulated in the airway mucosa of patients with asthma [67]

Administration of IL-13 to mice increases airway eosinophilia and airway

hyperresponsiveness in animal models of asthma [67]. IL-13 is associated with

structural changes seen in chronic asthma, including goblet cell hyperplasia, airway

smooth muscle proliferation, and subepithelial fibrosis [68]. IL-13 induces AHR and

mucus hypersecretion by activating STAT6 in the airway epithelium [69].

Interleukin-17

The IL-17 cytokine family is a recently discovered group of cytokines [70]. The members

of the IL-17 family include IL-17A, IL-17B, IL-17C, IL-17D, IL-17E (also called IL-25),

and IL-17F [71]. In bronchial epithelial cells IL-17A induces the expression of mucin

genes, MUC5AC and MUC5B [72]. Increased expression of IL-17A or overexpression of

IL17 F is associated with enhanced mucous production by increased mucin gene

expression and goblet cell hyperplasia [73]. In addition this, IL-17A and IL-17F induces

several profibrotic cytokines. In fibroblasts isolated from bronchial biopsy of subjects

with asthma, IL-17A enhances the production of profibrotic cytokines, IL-6 and IL-11

[74].

TNF-α

Tumor necrosis alpha (TNF-α) is secreted by macrophages, mast cells, T cells, epithelial

cells, and airway smooth muscle cells [65]. Low levels of TNF-α helps in maintaining

homeostasis by regulating the body's circadian rhythm and promoting the remodeling

35

or replacement of injured and senescent tissue by stimulating fibroblast growth [75].

TNF-α has a important role in maintaining the immune response to bacterial, and

certain fungal, viral, and parasitic invasions as well as in the necrosis of specific tumors

[76]. TNF-α amplifies asthmatic inflammation in various cells mainly through activation

of NF-κB [77]. In normal subjects, inhalation of TNF-α induces AHR and airway

inflammation mediated by neutrophils [78]. TNF-α is a pro-inflammatory cytokine [77]

and has a major role in airway inflammation and airway remodeling in asthma [79].

Decreased TNF-α expression or TNF-α receptor or treatment with anti-TNF-α antibody

results in the attenuation of antigen-induced airway hyperresponsiveness and

eosinophilic airway inflammation [77,80]. Blocking TNF-α with etanercept or infliximab

reduced AHR in patients with refractory asthma and reduced exacerbations in patients

with moderate asthma respectively [79,80].

IL-1β

Interleukin-1 beta (IL-1β) is a member of the IL-1 family of cytokines which have

potent pro-inflammatory properties [81]. IL-1β is produced by moncytes/macrophages,

neutrophils, B cells, T cells, epithelial cells, and airway smooth muscle cells [56]. In

asthmatic subjects the levels of IL-1β in the BALF were increased as compared to non-

asthmatics subjects [82]. There is increased expression of IL-1β in the airway epithelium

and macrophages in patients with asthma that causes pulmonary inflammation and

enhanced mucin production [81,83].

36

IL-10

IL-10 is secreted by monocytes, macrophages, mast cells, T and B lymphocytes, and

dendritic cells (DCs) [84]. It is also produced by a subset of Tregs [65]. IL-10 is a

pleiotropic cytokine that can exert either immunosuppressive or immunostimulatory

effects on various cells [55]. In addition, IL-10 inhibits the transcription of various pro-

inflammatory cytokines including TNF-α, IL-1, IL-5 and IL-6 [85,86]. In response to

allergic challenge IL-10 production and release by monocytic cells is upregulated by

TNF-α, and by negative feedback regulation of itself [84]. IL-10 is a inhibitor of NF-κB

by dual mechanisms-1) inhibition of degradation of I-κB results in preventing NF-κB

translocation to the nucleus, (ii) inhibition of degradation of NF-κB already present in

the nucleus from binding to DNA [86]. Therefore, IL-10 can be considered an important

agent in resolving of inflammation. Steroid therapy and allergen specific

immunotherapy elevate endogenous IL-10 levels, suggesting that IL-10 could be

targeted to treat airway inflammatory diseases such as asthma and COPD [84].

Chemokines

Chemokines are family of small cytokines induce chemotaxis and activation of

leukocytes, and are involved in the pathogenesis of asthma [87].

RANTES (regulated on activation, normal T expressed and secreted) is a potent

chemokine for eosinophils, T cells and monocytes, and plays an important role in

recruiting leucocytes to inflammatory sites [88]. It is produced by T-cells

[89],peripheral blood mononuclear cells [90], and activated platelets [91].

37

Interferon gamma-induced protein 10 (IP-10 or (CXCL10)) is a potent inducible

chemokine and acts as a chemoattractant for monocytes/ macrophages, T cells, NK cells

and mast cells. Neutralizing IP-10 results in inhibition of mast cell migration towards

the asthmatic airway smooth muscle cells [88,92].

All the above mentioned cytokines, chemokines and growth factors contribute to

the induction of rapid-response genes and are regulated by various transcriptional

factors NF-κB stands out as a central coordinating regulator in the transcription of

many such genes [93].

38

1.2 NF-κB

Nuclear factor-kappa B (NF-κB) is a stimulus-responsive pleiotropic regulator of

gene control present in all cell types [94]. It is a structurally and evolutionary conserved

family of ubiquitously expressed transcription factors regulated a great variety of

normal cellular processes, such as immune and inflammatory responses, development,

proliferation, differentiation and apoptosis [95,96]. NF-κB plays a major role in the

pathogenesis of various diseases including cancer, chronic inflammation,

neurodegenerative diseases, and diabetes [97-101].

NF-κB is activated by more than 150 physiological and non-physiological stimuli,

physical and chemical stress, lipopolysaccharides (LPS), double-stranded RNA (dsRNA

), single-stranded RNA (ssRNA), T and B cell mitogens including bacterial, viral, and

fungal products, as well as inflammatory cytokines, oxidative stress, ultraviolet and

ionizing radiation [102,103]. On activation, NF-κB, participates in transcription control

of over 150 target genes [102]. Because this transcriptional factor regulates and co-

ordinates the expression of various inflammatory genes and inflammatory mediators,

including cytokines, chemokines, adhesion molecules, immunoreceptors and growth

factors NF-κB has often been termed a `central mediator of the human immune

response' [93,104].

39

1.2.1 Role of NF-κB in Asthma

There is enhanced NF-κB activation in asthmatic tissues [103]. Nuclear extracts

from peripheral blood mononuclear cells (PBMCs), bronchial biopsies, lung

parenchyma, sputum cells, cultured bronchial epithelial cells and cultured airway

smooth muscle cells from asthmatics have greater levels of NF-κB p65, p50 activation

and increased nuclear protein expression of NF-κB p65 compared to non-asthmatics

[103,105,106]. The cytokines secreted by smooth muscle cells in asthma including TNF-

α, IL-1β and IL-17 are responsible for activation of NF-κB [107]. Activated NF-κB

induces the rapid expression of multiple genes that play significant roles in airway

hyperresponsiveness and airway smooth muscle proliferation in asthma [107,108].

Some of these include proinflammatory cytokines (IL-lβ, TNF-α, GM-CSF (granulocyte

macrophage colony stimulating factor), IL-2, IL-4, IL-5, IL- 11, IL-13, IL- 17)

chemokines (IL-8, RANTES, eotaxin,IP-10), enzymes (inducible nitric oxide synthase,

inducible cyclooxygenase), adhesion molecules (ICAM- 1, VCAM-1, E-selectin) and

receptors (IL-2 receptor) [103,109,110].

IL-10 is a well known inhibitor of NF-κB activity in various cells [85,95]. IL-10

suppresses inflammation mediated damage to the lungs through its inhibitory effects on

NF-κB activation [110,111]. In monocytes/macrophages IL-10 suppresses the production

of inflammatory cytokines TNF-α, IL-1β, IL-6, and IL-8 [112]. IL-10 also inhibits the

stimulated release of RANTES and IL-8 from human airway smooth muscle cells in

culture [112]. The activation of NF-κB subunits p50-RelA is significant in the

pathogenesis of asthma and may lead to death [105,113]. For this reason, the NF-κB

signaling pathway is an attractive target for novel asthma therapies [105].

40

1.2.2 Rel Protein Family

NF-κB has an essential role in regulating the immune response to infection as

well as in cell differentiation and apoptosis [114]. The NF-κB family has 5 distinct

members; c-Rel, RelA( also called p65), RelB, p50(and its precursor p105), and p52 (and

its precursor p100), which are able to homodimerize or heterodimerize [114,115]. For

sequence specific DNA binding all these have an N-terminal Rel homology domain for

forming the homodimers and heterodimers of family members [114]. There is also a C-

terminal transcription activation domain (TAD) which is only present in RelA, RelB, and

c-Rel [114]. Neither p50 nor p52 have transcription activation domains (TAD) and they

rely on the other sub-units to regulate gene transcription [114]. RelB heterodimerizes

with p100 as well as the processed form of p52. RelA and c-Rel predominantly

heterodimerize with p50 [116,117].

1.2.3 Nuclear Transport of NF-κB

Under normal physiological conditions, NF-κB is sequestrated in the cytoplasm in

association with an inhibitory protein called IκB. The activation and regulation of NF-κB

is under the control of the IκB protein that masks the Nuclear Localizing Sequence

(NLS) of the NF-κB subunit [118]. IκB prevents the NF-κB subunit from moving into

the nucleus. Under basal conditions, due to dynamic shuttling of NF–κB from the

cytoplasm to the nucleus, a basal expression of RelA subunit is observed in the nucleus

[119]. The activation of cytoplasmic NF-κB can occur by various physiological and non-

physiological stimuli such as cytokines, growth factors, bacterial or viral infection and

UV irradiation [95,120]. Upon activation NF-κB is imported from the cytoplasm to the

nucleus [121,122]. The enzyme I-κB kinase phosphorylates I-κB on its N-terminal serine

41

residues resulting in degradation of IκB by a process called ubiquitination [116]. The

phosphorylation of I- κB is catalyzed by the IκB kinase(IKK) complex consists of two

catalytic subunits, IKKα and IKKβ, and the regulatory subunit IKKγ [116,123,124]. The

cytoplasmic NF –κB complexes, especially p50-RelA (p65), rapidly translocate to the

nucleus. In the nucleus, NF-κB binds to functional κB sites in the promoter regions of

target genes and modulates expression of several hundred target genes, including those

encoding cytokines, chemokines, growth factors, pro- and anti-apoptotic proteins, cell

surface receptors, cell adhesion molecules, and transcription factors [122,125,126]

(Figure 3). The p50 and RelA are large molecules, and cannot translocate into the

nucleus via passive diffusion. Nuclear import of p50 and RelA subunits is mediated by

receptor proteins called importins [127]. Importin α3 and Importin α4 are reported to

be the main importin α Isoforms utilized for the nuclear translocation of NF-κB p50-

RelA heterodimers upon stimulation of cells with TNF-α [113,127,128].

42

Figure-3: NF-κB Signaling Pathway: Schematic diagram showing activation and

translocation of NF-κB into the nucleus- Upon activation by various physiological and non-

physiological stimuli NF-κB is imported from the cytoplasm to the nucleus. The enzyme I-κB

kinase phosphorylates I-κB resulting in degradation of IκB by a process called ubiquitination.

The cytoplasmic NF –κB complexes, p50-RelA (p65), translocate to the nucleus. In the nucleus,

NF-κB binds to functional κB sites in the promoter regions of target genes where it induces

expression of various pro-inflammatory cytokines, chemokines, and adhesion molecules.

43

1.3 Nuclear-Cytoplasmic Transport

1.3.1 Importins and Exportins

In eukaryotic cells, the nucleus stores genetic information and the cytoplasm is

involved in protein synthesis [129]. Large amount of genetic information is transferred

between the nucleus and the cytoplasm [130]. A variety of proteins such as

transcriptional factors, ribonucleoproteins must be transported into the nucleus while

RNA and nuclear proteins are exported from the nucleus to the cytoplasm for protein

synthesis [131].

Nuclear transport is highly organized and efficient. The transport of molecules in

and out of the cell is monitored by Nuclear Pore Complexes (NPC). NPC’s are

approximately 125 MDa [131,132]. The NPC is composed of 30 different proteins, called

nucleoporins or Nups, many of which are present in multiple copies per NPC [133,134].

Molecules less than 20 KDa it can easily enter the NPC through passive diffusion, but

larger molecules require receptor-mediated nuclear transport [131,135]. The diameter of

the diffusion channel of NPC is ~9 nm, but during an active transport the channel opens

to a maximum size of approximately 40 nm [136] (Figure-4).

A protein destined for nuclear import or export contains a specific signal, a

nuclear localization signal (NLS) or nuclear export signal (NES), which are recognized

by importins and exportins, respectively [137]. The NLS is amino acid sequence

separated by a spacer arm which on a exposed surface of protein acts as a ‘tag’. The NLS

is found in cellular nuclear proteins and acts as a target sequence recognized by a

receptor (importin) molecule [138-140]. NLS are characterized as either basic

monopartite (one stretch of basic amino acids) or bipartite (two stretches of basic amino

44

acids sequences) [141,142]. A well characterized example of a monopartite and a

bipartite NLS are the single cluster in the SV40 large T antigen (126PKKKRKV132) [143]

and the two in nucleoplasmin (150KRPAAIKKAGQAKKKK170) respectively [141,142].

The NLS of NF-κB is characterized as monopartite EEKRKR [144-146]. The NLS has

receptors through which soluble proteins importin α and β attach and form a

heterodimer. The bidirectional trafficking of the macromolecules across the nuclear

envelope is mediated by a family of target receptors known as karyopherins which

comprise both importins and exportins [144-146].

Importins help molecules enter into the nucleus and exportins help molecules

back to the cytoplasm from the nucleus. The import of nuclear proteins is a two-step

process; in the first step the nuclear protein binds to the nuclear pore complex. This

process does not require energy. The second step is an energy dependent translocation

of the nuclear protein through the channel of nuclear pore complex [147]. Both

importins and exportins are regulated by the small Ras related GTPase, Ran. Ran binds

with karyopherins (importins and exportin) when it is bound to GTP. In the cytoplasm

Ran is bound to GDP and in the nucleus Ran is bounded to GTP [148,149].

45

Figure-4: Schematic diagram showing import and export of small and large

molecules in the nucleus: Molecule less than 20 KDa enter the NPC through passive

diffusion but large molecules greater than 40 KDa the transport is facilitated by importins and

exportins.

46

1.3.2 Import

The NLS has receptors for the soluble proteins importin α and β to bind and form

a heterodimer. Importin α acts as an adaptor protein which binds to importin β

through its amino-terminally located importin β binding (IBB) domain and to NLS

via its two NLS binding sites in the central area of the molecule [150,151] [152,153].

When importins (karyopherin) recognize the NLS receptor they are referred to as the

Karyopherin Cargo complex .This complex is can translocate to the nucleus through the

Nuclear Pore Complex. The Nuclear pore proteins (Nucleoprotein) are involved in this

process. Importin β is the transport receptor that moves the importin α and NLS

complex from the cytoplasm into the nuclear side of the NPC [131,154]. In the presence

of Ran GDP , the Karyopherin Cargo complex enters the nucleus and Ran GDP is

catalyzed to Ran GTP by the RanGDP/GTP exchange factor RCC1 known as Guanine

nucleotide exchange factor (GEF). Ran GTP causes a conformational change in the

importin molecule and the whole complex dissociates [148,149] (Figure 5).

1.3.3 Export

Importin β attaches to RanGTP and this complex translocates back to the

cytoplasm. RanGTP is converted to RanGDP and this reaction is catalysed by GTPase

activating protein (GAP) in the cytoplasm and causes importin β to dissociate from

RanGTP. Importin α requires an additional cargo or protein (exportin) to be transported

back to the cytoplasm. Importin α binds to RanGTP but this binding is not sufficient to

transport importin α back to the cytoplasm. A nuclear export factor known as Cellular

47

Apoptosis Susceptibility (CAS) protein attaches to importin α in presence of RanGTP

and translocates importin α to the cytoplasm. In the presence of RanGAP and its

cofactor RanBP1 this CAS protein-importin α-RanGTP complex dissociates

[129,132,149].

The protein or the transcriptional factor destined to be translocated to the nucleus

binds to the promoter region of DNA, and initiates transcription, resulting in mRNA

synthesis. This protein or the transcriptional factor also contains a specific signal known

as Nuclear Export Sequence (NES) is translocated back to the cytoplasm. The CRM1

protein or the chromosomal region maintenance (now also known as Exportin1) is a

nuclear export receptor and this recognizes NES and this complex in presence of

RanGTP is transported to the cytoplasm. In presence of RanGAP this CRM1-NES-

RanGTP complex dissociates in the cytoplasm.[86,132,148,149] (Figure 5)

48

Figure 5: Mechanism of Action of Importins and Exportins: A protein destined for

nuclear import or export contains a specific signal, a nuclear localization signal (NLS) or nuclear

export signal (NES), which are recognized by importins and exportins, respectively. Importin α

acts as an adaptor protein where on one side it binds to importin β on another side to NLS.

Both importin and exportin are regulated by small Ras related GTPase Ran. Ran binds with

karyopherins (importins and exportin) when it is bound to GTP. In the cytoplasm Ran is bound

to GDP and in the nucleus Ran is bounded to GTP. The NLS binds to the promoter region of the

DNA and starts the transcription process and mRNA is synthesized. This mRNA formed

49

possesses a NES. The Exportin1 is a nuclear export receptor and this recognizes NES and the

complex in presence of RanGTP is transported to the cytoplasm for translation.

1.3.4 Importin Alpha

Throughout evolution importin α molecules have remained structurally and functionally

conserved and can be found in eukaryotes from yeast to human. In humans, six different

importin α molecules have been identified: importin α1, importin α3, importin α4,

importin α5, importin α6, and importin α7 [131,155-158].

The importin α molecules can be classified into three distinct subfamilies, based on the

similarity of their primary structure. The first family consists of importin α1 and is the

only member of this subfamily. Importins α3 and α4 belong to the second subfamily.

The third subfamily consists of importins α5, α6, and α7 [131,159]. Members of the

different importin subfamilies have ~ 50% sequence identity, but within one importin

subfamily the sequence identity is approximately 80% [157,160].

Table-1: Classification of Importin alpha Molecules

50

Table-2 Isoforms of Importin α and β and examples of cargo molecules

transported. (Modified from- N.Okada, et al. Importins and exportins in cellular

differentiation 2008) [128]

51

Although all isoforms of importin α can mediate the import of NLS-containing

substrates into the nucleus, importin α molecules are functionally divergent. Importin α

isoforms display differences in their cell- and tissue-specific expression patterns.

Whereas some substrates can be transported into the nucleus by all importin α

isoforms, there is experimental evidence that several substrates are recognized and

transported specifically by a particular, or a subset of, importin α molecules

[127,131,159,161,162]

Importin α molecules have a N terminal domain that binds to the Importin Beta

Binding(IBB) Domain which cosist of ~ 40 highly conserved basic residues that are

necessary to bind importin-β and permit efficient nuclear entry [142]. Importin α also

has a large NLS-binding domain possessing a flexible linker, made up of ten tandem

armadillo (ARM) repeats [142,144,151,163]. The ARM repeats are arranged in a helical

manner and get twisted to assemble in a slug like structure, where the cNLS binds in its

belly in the cNLS binding groove [144,153]. The tenth ARM repeat is the binding site for

Exportin CAS. Its trans form IBB interacts with Importin β and in its cis form IBB

interacts with the NLS- binding groove [144,153]. In short, Importin β binds to the N

terminal domain , the cNLS-binding pocket is in the central ARM domain and binding

site of export receptor CAS is in tenth ARM repeat in C-terminal region of Importin α

[144,153].

52

1.3.5 Importin Beta

Importin β consists of helical protein composed of 19 tandem HEAT repeats (The HEAT

repeat domains are protein domains found in cytoplasmic proteins- HEAT-Huntingtin,

elongation factor 3 (EF3), protein phosphatase 2A (PP2A), and the yeast PI3-kinase

TOR1), arranged in a right-handed superhelix with both a pitch and a mean diameter of

approximately 50A° [142]. The individual HEAT motifs form a helical hairpin structure

suggestive of an armadillo repeat, consistent with the hypothesis that ARM and HEAT-

repeat-containing proteins are evolutionarily related [142]. Importin β binds to the IBB

domain of importin α. This complex takes a shape of a snail with IBB domain at the

centre and importin β wraps around it from outside. This result in N- and C-terminal

HEAT repeats nearly touching each other, giving the molecule a ‘closed’ appearance

[142].

1.3.6 The RanGTPase system

There is asymmetrical distribution of Ran GTP between the nucleus and the

cytoplasm that determines the direction of nuclear transport which is towards higher

Ran GTP. This lack of balance in the levels of Ran GTP is maintained by regulators

which control the state and cellular location of Ran GTP [164].

RanGAP: In mammalian cells, the Ran-specific GTPase activating protein (RanGAP) is

only present in the cytoplasm. RanGAP increases Ran’s rate of GTP hydrolysis by five-

fold thereby reducing the levels of Ran GTP in the cytoplasm. RanGAP is not present in

the nucleus and in its absence Ran hydrolyses GTP very slowly. The half life of the

53

RanGTP complex in the nucleus is very long, thus ensuring the steady state levels of

RanGTP [164].

RanBP1 and RanBP2: RanGTP-importin complexes are stable and have a half life of

several hours [164]. RanGTP-importin/exportin complexes inhibit RanGAP-induced

GTP hydrolysis by Ran [164]. The binding of importins and exportins to RanGTP but

not RanGAP would make it impossible for transport receptors to dissociate and shuttle

repeatedly across the NPC [164]. RanBP1 and RanBP2 are another family of Ran-

binding proteins [165] that promote the dissociation of RanGTP-importin/exportin

complexes. RanBP1/ RanBP2 binds to the transport receptor complex causing

dissociation of the complex and renders RanGTP in the complex accessible to RanGAP

[164].

54

1.4. Immunomodulators in Asthma

Immunomodulators are a group of drugs that either provide long-term control of

asthma or are considered steroid sparing. These medications modulate the immune

response to asthma triggers. There is a need for development of novel therapies that

inhibit key immunopathogenic mechanisms resulting in induction of immune tolerance

[166]. Novel therapeutic approaches under study include toll-like receptor (TLR)

agonists [167], oral and parenterally administered cytokine blockers, specific cytokine

receptor antagonists [168,169], and transcription factor modulators [166,170,171]. Over

the last several years, various studies have shown significant impact of vitamin D on

immune function have suggested vitamin D functions as an immunomodulator in

diseases such as asthma [172].

1.4.1 Vitamin D

Vitamin D is synthesized in skin in presence of adequate sunlight exposure [173].

The main function of vitamin D is to maintain extracellular fluid concentrations of

calcium and phosphorous within normal range [174]. This action of vitamin D is

achieved by increasing the absorption capacity of small intestine for calcium and

phosphorous vitamin D also stimulates calcium and phosphorous mobilization from the

bones [174]. Thus, vitamin D plays a significant role in normal adult bone remodeling by

regulating bone and mineral homeostasis [175]. 1, 25-Dihydroxyvitamin D3 [1, 25(OH)

2D3)], is an active metabolite of Vitamin D3, is synthesized in a highly regulated multi-

step process [176]. Calcitriol modulates the Vitamin D functioning and signaling in an

autocrine and paracrine manner [176]. Calcitriol is a pleiotropic hormone it not only

55

regulates calcium homeostasis in the body but also has role in the differentiation and

inhibition of proliferation of various normal and cancer cells [177].

Vitamin D may play a role in disorders including cancer [178,179], infection [180],

cardiovascular disease [181], schizophrenia [182], and immune-mediated diseases such

as multiple sclerosis [183], insulin-dependent diabetes [184], and asthma [185]. Vitamin

D and its active metabolite appear to have unique roles in a large number of tissues

[186]. However, the central feature of response to calcitriol in each of these tissues is the

presence of vitamin D receptor (VDR) [186].

1.4.2 Vitamin D Receptor

Calcitriol exerts its action through vitamin D receptor (VDR) which is a high

affinity nuclear receptor [187]. VDR is a member of the steroid nuclear receptors

superfamily [188]. VDR is also a transcription factor that interacts with its co-regulators

and alters the transcription of target gene which are involved in a wide spectrum of

biological responses [189].VDR was first discovered in the extracts prepared from

chicken intestine, as a result of this chicken was considered a model system in defining

the function of vitamin D in calcium and phosphorous homeostasis [190]. The receptor

was found in mammalian intestine as well [191]. Additional target tissues for vitamin D

including kidney, bone and parathyroid gland cells were defined in both avian as well as

mammalian system [192]. Further studies determined that the biological effects of

calcitriol are not limited to calcium and phosphorous homeostasis, but calcitriol also has

role in wide variety of functions including cellular proliferation and differentiation

[193,194]. VDR is also expressed in other tissues such as pancreas [192,195], placenta

56

[192], pituitary [192,196], ovary [197], testis [198], mammary gland [199,200] and heart

[201].

1.4.3 Vitamin D Metabolism

The process starts from the synthesis of Vitamin D3 in the skin by sunlight,

although it can be obtained from naturally from the diet and from foods that are

fortified with vitamin D, or from vitamin D supplements [174]. Exposure of skin to UV

light (270–300 nm) catalyzes the first step in vitamin D3 biosynthesis, converting 7-

dehydroxycholesterol into previtamin D3 which is then converted to vitamin D3

(cholecalciferol) [202]. Very minimal exposure (10-15 minutes) is needed to synthesize

cholecalciferol in the skin and provides more vitamin D3 than any known dietary source

[203].

Vitamin D3 is carried to the liver by vitamin D binding protein [204]. The hepatic

enzyme 25–hydroxylase, encoded by the gene CYP27A1, places a hydroxyl group in the

25 position of the vitamin D molecule, resulting in formation of 25-

hydroxycholecalciferol (calcidiol) that is the major circulating form of vitamin D and is

measured by routine laboratory testing [176,205]. The mitochondrial enzyme 1 α-

hydroxylase (encoded by gene CYP27B1) further hydroxylates the 25-

hydroxycholecalciferol to form the biological active form of vitamin D called is calcitriol

(1 α, 25(OH)2D3) [176] (Figure6).

The kidney is the primary site where the active form of vitamin D, 1, 25(OH) 2D, is

produced. The circulating levels of 1,25(OH)2D result from CYP27B1 (cytochrome p450

27B1)- mediated conversion of 25OHD in the proximal tubule of the kidney [206].

57

Renal CYP27B1 gene expression is activated by parathyroid hormone (PTH) and

suppressed by 1, 25(OH) 2D. The serum levels of these hormones are inversely related to

dietary calcium intake [207]. Once formed in the proximal tubule, 1,25(OH)2D is

released into the serum and acts as an endocrine hormone on the intestine, bone and

kidney to control calcium metabolism [207] (Figure 6). Studies have shown expression

of 1 α-hydroxylase in extra-renal sites such as brain, placenta, pancreas, lymph nodes

and skin, allowing local conversion of 25(OH)D3 to 1 α,25(OH)2D3 [208].

As 1 α, 25(OH)2 D3 is highly potent in elevating serum calcium and phosphate

levels it requires a mechanism to attenuate its activity. This is achieved by 1 α,

25(OH)2D3 inducible 25(OH)D3- 24-hydroxylase (encoded by the genes CYP24), which

catalyzes a series of oxidation reactions leading to formation of biliary excretory

product calcitroic acid [187].

58

Figure-6: Metabolism of Vitamin D Vitamin D is either produced in the skin by exposure to

UVB radiation or is ingested in the diet. Vitamin D3 is converted by the 25-hydroxylase (25-

OHase) in the liver to 25(OH)D. 25(OH)D is converted in the kidneys by enzyme 1-alpha-

hydroxylase (CYP27B1) to 1,25(OH)2D. Once formed, 1, 25(OH)2 D increases calcium and

phosphorus absorption in intestine and stimulates bone remodeling. In addition, 1, 25(OH)2 D

inhibits the renal CYP27B1 and stimulates the expression of the renal 25(OH)D-24-hydroxylase

(24-OHase). The induction of the 24-OHase results in the destruction of 1, 25(OH)2 D into a

water-soluble inactive metabolite calcitroic acid.

59

1.4.5 VDR Signaling Pathway

The key molecular action of calcitriol (1, 25(OH) 2D) is to initiate or suppress gene

transcription of multiple genes across diverse target tissues by binding to the VDR

[209]. VDR can be found in both the cytoplasm and nucleus of vitamin D target cells,

but in many cells it is predominantly a nuclear protein [210]. Binding of 1,25(OH)2D to

the VDR promotes association of VDR with the RXR (retinoid X receptor) an interaction

essential for VDR transcriptional activity [210]. Heterodimerization of the RXR–VDR–

ligand complex is required for migration of the complex from the cytoplasm to the

nucleus where the 1,25(OH)2D–VDR–RXR complex binds to VDREs (vitamin D-

response elements) in DNA to initiate gene transcription [210] (Figure 7).

60

Figure 7: VDR signaling pathway VDR is located in the cytoplasm of vitamin D target cells.

Binding to 1, 25(OH) 2 D changes the structure of VDR and allows formation of a heterodimer

with retinoid X receptor (RXR). In the nucleus, the VDR-RXR heterodimer binds to unique

DNA binding sites in the promoter of the vitamin D target genes called vitamin D response

elements (VDRE). Once bound to VDRE, the VDR-RXR complex recruits protein complexes to

promote events leading to transcription initiation. In some target cells such as kidney, bone and

intestine l, 25(OH) 2 D directly enters in the cell and binds to VDR. Some of the non-target cells

such as macrophages, monocytes, and dendritic cells express the membrane proteins megalin or

cubulin to concentrate the 25(OH)D3. These cells express the protein CYP27B1 and convert

25(OH)D to l, 25(OH) 2 D locally. It is believed that in this way, these target cells boost

61

1,25(OH)2D production by augmenting the circulating 1,25(OH)2D concentrations produced by

the kidney [204].

1.4.6 Epidemiology of Vitamin D Deficiency

In humans the primary determinant of vitamin D status is the amount of exposure

of an individual to sunlight, especially for people living in northern latitudes [211]. From

November to March, there are insufficient UV-B rays to produce vitamin D at the

northern latitude [212]. However, the vitamin D levels in populations living near the

Equator (e.g., in Saudi Arabia, Israel, India, and Costa Rica) and in the southeastern

United States have been found to be low, suggesting that lifestyle and diet can also have

major effects on vitamin D status [213].

Over the last decade, vitamin D deficiency or insufficiency has increased in United

States. From the 1988-1994 to the 2001-2004 data collections the mean serum 25(OH)

D level in the US population dropped by 6 ng/mL [212]. This drop in serum 25(OH) D

was associated with an overall increase in vitamin D insufficiency to nearly 3 of every 4

adolescent and adult Americans [212]. Nearly all non-Hispanic blacks (97%) and most

Mexican- Americans (90%) now have vitamin D insufficiency [212]. It may be due to

changes in behavior (eg less time outdoors) and diet [212]. In a recent study of 9,757

United States subjects from 1 to 21 years of age, approximately 9% had vitamin D

deficiency and approximately 61% of participants had vitamin D insufficiency [214].

Some of the causes of vitamin D deficiency included older age, female gender, African-

62

or Mexican-American ethnicity, obesity, the use of electronic devices, and reduced dairy

intake [214].

1.4.7 Vitamin D Optimal Status/Dietary Requirements

Vitamin D is both a nutrient and a hormone. The ability of the skin to metabolize

in skin that is influenced by many factors including the melanin content of the skin, age,

factors affecting sun exposure (latitude, season, time outdoors, and clothing), body fat,

and sunscreen use [215]. In contrast to other nutrients, vitamin D is found in limited

number of foods. Food sources of vitamin D include such as oily fish and fish liver oil,

egg yolk, and offal [216]. There is a wide variation in the content of vitamin D in these

natural sources due to the origin of the food (eg, farmed versus wild salmon). Cooking

methods (e.g. frying versus baking) can influence the amount of vitamin D in these

foods [217]. Therefore, most of the vitamin D that is ingested by humans comes from

secondary sources of vitamin D such as fortified foods (in the United States milk, some

milk products such as yogurt and margarine, and breakfast cereals are fortified with

vitamin D) and from supplements [218] .

25(OH) D3 is the major circulating form of the vitamin D in the serum, and has a

half life of 15 days. As calcitriol, the active form of vitamin D, has a short half life of ~15

hours, 25(OH) D3 is routinely used to measure the serum concentration of vitamin D

[204,219-221]. The current recommendation of vitamin D intake by Institute of

Medicine (IOM) is 600 IU/day from age 1 yrs through age 70 years (including pregnant

and lactating women), and 800 IU/d for those older than 70 years.

Each additional 100 IU of vitamin D per day raises serum 25(OH)D concentration

by approximately 1 ng/mL [222]. For infants the recommended dose of vitamin D is

63

400IU/day. However, current recommendations by IOM stating that these doses of

vitamin D are not sufficient for overall health [222].

Most experts define vitamin D deficiency as a circulating serum level of 25-

hydroxyvitamin D3 (25[OH]D) of <50 nmol/L (20 ng/mL); 25(OH)D levels between

50 and 75 nmol/L (20-30 ng/mL) are considered to be a relative insufficiency [174]. A

desirable (or sufficient) serum level of 25[OH]D is 75 to 100 nmol/L (30-40 ng/ mL)

[174]. The IOM committee considers the safe upper limit for vitamin D consumption to

be 10,000 IU/d [223,224]. The committee also recommended that hypercalcemia could

be used as an indicator of toxicity [223].

The most common symptoms of vitamin D intoxication are related to symptoms of

hypercalcemia due to primary renal failure. These symptoms include thirst, itchiness,

diarrhea, malaise, wasting, polyuria, and diminished appetite. Vitamin D intoxication

can also result in calcification of kidney, aorta, heart, lung and subcutaneous tissues

[225]. Hypercalcemia results only when 25(OH)D levels have consistently been above

375–500 nmol/L, which requires a daily intake of 40,000 IU of vitamin D [204,226].

1.4.8 Immunomodulating Effects of Vitamin D

Recent studies provide a strong evidence that vitamin D has a role in the innate

and adaptive immune systems [219,227]. The immunomodulating properties of vitamin

D were first described ~20 years ago [174,228,229]. Vitamin D has a significant role

both directly and indirectly in regulation of proliferation, differentiation and function of

immune cells [230]. The expression of VDR has been described in various immune cells

including circulating monocytes, macrophages, dendritic cells, dendritic cells and T cells

64

[231]. CYP27B1 (i.e.1-alpha-hydroxylase) is the enzyme responsible for the last step in

vitamin D-metabolism, i.e., hydrolyzing 25(OH) D3 to 1, 25(OH) 2D3. Interestingly,

CYP27B1 is present in T cells (and probably B cells), macrophages and dendritic cells

signifying that 1,25(OH)2D3 can be produced locally. This suggests that vitamin D plays

a role in the differentiation, biosynthetic activity, and function of leukocytes [232].

Antigen-presenting cells are not subject to negative regulation of 1-alpha-hydroxylase by

PTH or 1,25(OH)2D3 as happens in renal cells [233]. Thus, the local concentration of

calcitriol in antigen presenting cells is higher than the circulating levels.

1.4.8-1 Monocyte/Macrophages and Vitamin D

Calcitriol stimulates the differentiation of monocyte precursors into macrophage-

like cells, suggesting that there is an immunoregulatory effect of vitamin D [186]. The

expression of VDR in monocytes sensitizes the cells to calcitriol, and subsequent

maturation of the monocytes to macrophages occurs in an autocrine manner [186]. The

presence of CYP27B1 during differentiation of monocytes to macrophages suggests local

synthesis of calcitriol in these cells. When normal human macrophages were stimulated

with interferon gamma (IFNγ), the cells synthesized calcitriol [234]. The endogenous

presence of VDR and CYP27B1 in monocytes/macrophages is strongly suggestive of an

autocrine or intracrine system for vitamin D actions in normal

monocytes/macrophages.

65

1.4.8-2 Dendritic Cells and Vitamin D

Calcitriol reduces the proliferation and differentiation of dendritic cells and

inhibits the differentiation of monocytes to DCs and differentiation survival and

maturation of DCs [230]. Calcitriol indirectly inhibits Th1 responses by inhibiting IL-12

production by DCs [235]. This inhibition involves the direct interaction of calcitriol with

the VDR and NF-κB, which interferes with transcription of the IL-12 gene [235].

Calcitriol also decreases the expression of costimulatory molecules including CD40,

CD80 and CD86 on DCs thus decreasing the ability of cells to present antigens and

activate T cells [236,237]. Calcitriol also stimulates IL-10 production by DCs. Calcitriol

also diminishes the production of proinflammatory cytokines (e.g., IL-1 and TNF-α),

leading to decreased IL-2 and IFN-γ production and increased IL-10 and TGF-β

production [236,237].Calcitriol promotes the induction of CD4+CD25+ regulatory T

cells (Treg) by DCs [238], and increases Foxp3 and IL-10 expression, two crucial factors

for Treg induction [239].

1.4.8-3 T Cells and Vitamin D

VDR expression is almost undetectable in quiescent T cells but following antigenic

stimulation there is five-fold increase in VDR expression level on the T cells as they

proliferate [240]. There are both direct and indirect effects of vitamin D on T and B

cells. Calcitriol directly inhibits T cell proliferation. Approximately 102 genes have been

identified as targets for calcitriol [241]. Calcitriol decreases Th1 cell proliferation and

inhibits IL-2, IFN-γ, and TNF –α production by Th1 cells [242]. The role of vitamin D

in inhibiting Th1 immune responses have been well studied, but its effects on Th2

66

responses are more complex and not fully elucidated [243]. Studies suggest that

vitamin D induces a shift in the balance between Th1 and Th2 cytokines towards Th2

dominance [175,244] resulting in reduced secretion of Th1 cytokines IL-2 and IFN-γ

[244-246] and an increase in the Th2 cytokine, IL-4 [247,248]. Vitamin D

administration markedly enhances TGF-β1 and IL-4 transcript levels [242]. In human

cord blood T cells vitamin D not only inhibits IL-12–generated IFN-γ production but

also suppresses IL-4 and IL-4–induced expression of IL-13 [249] These studies seem

contradict the effects of vitamin D on Th1-Th2 dominance suggesting that there are

effects related to cell type, dose and timing of vitamin D administration [215,244].

Regulatory T-cells Vitamin D promotes the induction of Treg cells [250,251]. Vitamin D

in combination with glucocorticoids may stimulate T-regulatory cell generation to

produce IL-10 which suppresses the immune response [252]. In patients with steroid

resistant asthma, vitamin D administration reversed steroid resistance through

induction of IL-10–secreting Treg cells [253].

Preferential differentiation of Tregs is thought to be a pivotal mechanism linking

vitamin D to adaptive immunity, with potential beneficial effects for autoimmune

disease and host-graft rejection [251,254,255]. Tregs develop and function normally in

the presence of vitamin D and suppresses inappropriate Th1 and Th2 responses to

environmental exposure (i.e. allergens, lack of infections), leading to a more balanced

immune response. If vitamin D is lacking, Tregs do not develop and function normally,

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and in the presence of the appropriate environmental influence, Th1 or Th2 responses

are allowed to proceed unabated leading to disease [243].

Th17 cells- Th17 cells are so termed due to their capacity to synthesize interleukin-

17 (IL-17) [256]. Th17 cells are associated with tissue damage and inflammation and

play an essential role in combating certain pathogens [257]. The exact role of vitamin D

as a regulator of Th17 cells remains to be elucidated. Studies of animal models of the

gastrointestinal inflammatory disease colitis have shown that calcitriol treatment

reduces IL-17 expression [239]. Another study in mouse model of gastrointestinal

inflammation demonstrated that on ablation of a CYP27b1 resulted in loss of 1,25

(OH)2D3 leading to elevated levels of IL-17 [258].Calcitriol inhibits the synthesis and

release of IL-23 and IL-6 that are the cytokines important for the development of Th17

cells [239]. Th17 cells secrete a proinflammatory cytokine- IL-17 and develop in

response to IL-6 and transforming growth factor (TGF)–β [3]. Th17 express IL-23R,

which is stabilized and/or maintained by IL-23 signaling [3]. These studies illustrate

that vitamin D may suppress Th17 cells by decreasing IL-17, IL-6 and IL-23.

1.4.8-4 B Cells and Vitamin D

Similar to T-cells, low levels of VDR expression is found in quiescent B cells and

activation of B cells causes increased expression of VDR in these cells [259]. Calcitriol

inhibits B-cell proliferation, differentiation of plasma cells and immunoglobulin (Ig)

production [259,260].Interestingly, CYP27b1 expression is detected in B-cells,

68

indicating that B-cells may be capable of autocrine/intracrine responses to vitamin D

[202]. Thus, vitamin D might be involved in humoral immunity.

1.4.9 Vitamin D and Asthma

Vitamin D is a potent immunoregulator and immunomodulator, and vitamin D

deficiency can predispose an individual to asthma and allergies in the presence of other

environmental stimuli [243]. Vitamin D or UVB radiation reduces airway inflammation

and IL 4 levels in the BALF in mouse models of allergic airway inflammation [241,261].

Other study of mouse model in the allergic airway inflammation support a key role for

VDR lung expression in airway inflammation, demonstrating that VDR knock-out mice

have elevated serum levels of IL-13 and IgE but do not develop airway inflammation

[262].

Recent studies demonstrated a role for vitamin D in airway smooth muscle cells

(ASM). In TNF-α treated ASM cells, calcitriol inhibited chemokines, RANTES and IP-10

secretion in a concentration-dependent manner [188]. These chemokines and play a

critical role in the pathogenesis of asthma and facilitate the recruitment of inflammatory

cells in the airways [88]. Vitamin D inhibits growth of human ASM cell growth through

growth factor-induced phosphorylation of retinoblastoma protein and checkpoint kinase

1 [188]. In ASM, the expression of many genes is regulated after vitamin D stimulation,

including genes that have a role in asthma predisposition and pathogenesis [47].

According to recent studies, there appears to be a strong relationship between

Vitamin D deficiency and increased incidence of asthma [263]. Vitamin D deficiency has

been associated with obesity which is likely due to the decreased bioavailability of

69

vitamin D3 from cutaneous and dietary sources because of its deposition in body fat

compartments [264]. There is increased prevalence of vitamin D deficiency in African

American race even after consuming adequate amounts of vitamin D, which suggests

genetic association between vitamin D levels and African Americans race [265]

(particularly in urban, inner-city settings), and recent immigrants to Westernized

countries [266], thus reflecting the same epidemiologic patterns observed in the asthma

epidemic [243].

There is increasing evidence that vitamin D plays a role in the development of

allergic airway diseases. Vitamin D is required for the lung growth in utero and a

deficiency in vitamin D during pregnancy may lead to the development of asthma in the

offspring [267-269]. A cross-sectional case-control study was conducted to examine the

prevalence of vitamin D insufficiency and deficiency among urban African-American

(AA) youth with asthma compared with control subjects without asthma [270]. The

study found that 86% of urban AA youth with persistent asthma were either vitamin D

deficient or insufficient compared to 19% in controls. The investigators strongly

recommended routine vitamin D testing in urban AA youth particularly those with

asthma [270].

In a study of 616 children in Costa Rica, the relationship between vitamin D levels

and markers of asthma severity were evaluated. The study concluded that low levels of

vitamin D are associated with increased total IgE and eosinophil count in asthmatic and

allergic children [271]. Lower levels of vitamin D was associated with increased number

of hospitalizations, increased use of anti-inflammatory medications, and increased

airway hyperresponsiveness [271].

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Another study on nonsmoking adults with asthma assessed the relationship

between serum 25(OH)D (vitamin D)concentrations and lung function, airway

hyperresponsiveness (AHR), and glucocorticoid response, as measured by

dexamethasone-induced expression of mitogen-activated protein kinase phosphatase

(MKP)-1 by peripheral blood mononuclear cells [272]. This study showed similar results

as the previous study on pediatric population, as low levels of 25(OH) D are also

associated with impaired lung function, increased AHR, and reduced response to

glucocortricoid, suggesting that vitamin D supplementation of patients with asthma may

improve multiple parameters of asthma severity and treatment response [272]. The

study also found that low serum vitamin D levels are associated with increased levels of

TNF-α [272].

A recent study involving 2,112 adolescents ages 16-19 found that 35% of

adolescents consumed less amount of vitamin D then what is recommended for this

age group [243]. Teens ingesting low dietary levels of vitamin D had significantly

decreased lung function compared with teens who consumed more vitamin D. No

differences in the lung function were found between males and females [243]. The study

authors recommended that dietary vitamin D intake should be promoted at

recommended levels to ensure optimal lung function as well as to form and maintain

healthy bones [243].

Vitamin D deficiency leads to decreased lung volume, decreased lung function

and altered lung structure [273]. Children with severe, therapy-resistant asthma having

lower vitamin D levels had increased ASM mass and worse asthma control and lung

function compared to moderate asthmatic and non-asthmatic control subjects [274].

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Supplementing the pregnant women with high doses of vitamin D reduced asthma risk

of their children by 40% in children 3-5 years after their births [243]. These studies

emphasize a major role of vitamin D in modulating the immune response in allergic

airway diseases.

In addition to the clinical trials, there is a need to examine the underlying

mechanism and the signal pathway regulated by vitamin D. These studies should

include studies in animal models, genetic association studies, and gene expression

studies to refine the vitamin D effect on lung function, smooth muscle contraction and

relaxation, airway inflammation, airway remodeling, and immune cell function. Our

studies are focused on exploring the molecular mechanism to define the role of vitamin

D in preventing asthma using human bronchial smooth muscle cells (HBSMCs). In

these in vitro studies we have studied the effect of vitamin D on importin α3, a molecule

associated with the NF-κB signaling pathway. In vivo studies in a mouse model of

allergic airway inflammation to study the effects of a vitamin D- deficient, vitamin D-

sufficient or vitamin D-supplemented diet. The effect of vitamin D on lung histology,

airway hyperresponsiveness, T-regulatory cells and BALF cytokines were also

performed.

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1.5: Central Hypothesis

Vitamin D reduces allergic airway inflammation and airway hyperresponsiveness by

inhibiting migration of NF-κB to the nucleus via inhibition of transcription and

translation of importin α3 in airway cells.

Specific Aim 1: To determine the effect of calcitriol on the expression of importin

α3, importin α4, vitamin D receptor and the vitamin D metabolizing enzymes CYP24A1

and CYP27B1 in HBSMCs.

Specific Aim 2: To study the effect of calcitriol on TNF-α induced translocation of

NF-κB in the nucleus and to study the effect of cytokines and calcitriol on the expression

of importin α3 in HBSMCs.

Specific Aim 3: To examine the effect of vitamin D deficiency, sufficiency and

supplementation on the AHR and allergic airway inflammation in OVA-sensitized and

challenged mice.

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SECTION 2

MATERIALS AND METHODS

74

2.1 Cell Culture and Cell Stimulation

Primary human bronchial smooth muscle cells (HBMSC) were obtained from

ScienCell Research laboratories (Carlsbad, CA). Cells were cultured in 25 cm2 cell

culture flasks in smooth muscle cell medium (SMCM) (Sigma, St. Louis, MO) containing

10% Fetal Bovine Serum (FBS) (Sigma, St. Louis, MO) cells were maintained at 5% CO2

at 37⁰C. Cells in passage 3-7 maintained their SMC phenotype and were used in all

experiments.

Cells were characterized for smooth muscle cell markers including smooth muscle

α-actin and smooth muscle heavy chain by immunofluorescence by the following

method: HBSMCs were cultured in Lab-Tek chamber slides (Thermo Fischer Scientific,

Rochester, NY) in smooth muscle cell medium (SMCM) containing 10% FBS and were

maintained at 5% CO2 at 37⁰C. Cells were growth arrested by serum starvation for 24 h

by replacing the FBS containing SMCM with FBS free DMEM (Dulbecco’s modified

eagle’s medium). HBSMCs were fixed with ice-cold 4% (wt/vol) paraformaldehyde for

10 minutes, incubated in 0.1% Triton X-100 for 5 minutes followed by blocking with 1 %

bovine serum albumin (BSA) for 30 minutes. HBSMCs were then incubated with anti-

alpha actin antibody (santacruz Biotech, Santa Cruz, CA; sc-58669) and myosin heavy

chain antibody (santacruz Biotech, Santa Cruz, CA; sc-53092) for 1 hr followed cyanine3

conjugated secondary antibody (Jackson ImmunoResarch, Westgrove, PA) for 1 hour.

Slides were mounted with Vectashield mounting medium (Vector Laboratories

Burlingame, CA) and examined using fluorescence microscopy.

All experiments were performed on three biologically independent samples.

Cultured HBSMCs (70-80% confluent cells) were growth arrested by serum starvation

75

for 24 h by replacing the FBS containing SMCM with FBS free DMEM (Dulbecco’s

modified eagle’s medium) (Sigma, Saint Louis, MO). After 24 hr cells were stimulated

with calcitriol (D1530 Sigma-Aldrich, Saint Louis, MO) at various concentrations (0.1 –

100 nM) or ethanol (<0.05%) as vehicle in fresh DMEM for 24 hr. Recombinant TNF-α,

IL-1β and IL-10 ( (PeproTech, Inc., NJ) were used. After stimulation, cells were

harvested for RNA and protein. Each experiment was run in triplicate.

2.2 RNA Isolation

Total cellular RNA was extracted from adherent cells and mice lung tissue. For

HBSMCs cells were rinsed with SMCM, and then lysed in 1 ml of TRI Reagent (Sigma,

Saint Louis MO) by repetitive pipetting. The homogenate was incubated for 10 minutes

at room temperature. For lung tissue, 100mg of whole lung tissue was minced and 1ml

of trizol was added. Bromochloropropane (0.1 ml) or chloroform (0.2 ml ) per 1 ml of

TRI Reagent was added to the homogenate and vortexed for 15 sec. The resulting

homogenate was incubated at room temperature for 15 minutes followed by

centrifugation at 12,000 g for 15 min at 4 ⁰C. The colorless upper aqueous phase was

transferred to a new microcentrifuge tube. A total of 0.5 ml of isopropanol was added to

aqueous solution. The samples were incubated at room temperature for 10 minutes and

centrifuged at 12,000 g for 8 min at 4 ⁰C. The supernatant was removed and the RNA

pellet washed with 1ml of 80% ethanol, followed by a centrifugation at 12,000 g for 5

minutes at 4 ⁰C. The supernatant was removed and the RNA pellet was left and air-dried

for 10 min. RNAase free distilled water (50 μl) was added to each tube to solubilize the

76

RNA. The RNA concentration was measured using a Nano Drop (Thermo Scientific,

Rockford, IL).

2.3 Reverse Transcription

Total RNA (1 µg) was used for the synthesis of first strand of cDNA with oligo-dT

(1μg),(Integrated DNA Technologies, Coralville, IA), 4μl of 5x reaction buffer, 4.8 μl

MgCl2, 1 μl of dNTPmix and 0.5 μl of Improm II reverse transcriptase as per Improm II

reverse transcription kit (Promega, Madison, WI). Cycling conditions for reverse

transcription procedure were 25⁰C for 5 min, 42⁰C for 60 min and 70⁰C for 15 min.

2.4 Real-Time PCR

Quantitative PCR was performed using 8 μl cDNA, 10 μl SYBR green PCR master

mix (Bio-Rad Lab, Hercules, CA) and 1μl of forward and reverse primers (10

picomol/μl) (Integrated DNA Technologies, Coralville, IA) using a 7500 Real Time PCR

system (CFX96, Bio-Rad Labs, Hercules, CA). The specificity of the primers was

analyzed by running a melting curve. The PCR cycling conditions were 5 min at 95ºC ,

40 cycles of 30 sec at 95ºC, 30 sec at 52-58ºC (depending upon the primer annealing

temperatures) and 30 sec at 72ºC.

Each real-time PCR assay was performed using 10 individual samples in

duplicates and the threshold cycle values were averaged. Calculation of relative

normalized gene expression was done using the BioRad CFX manager software

VERSION 2.1 based on fold change in expression between samples calculated by the fold

77

change (compare to control) = 2 –ΔΔCt method [275]. The results were normalized

against housekeeping gene, glyceraldehyde-3-phosphate dehydrogenase (GAPDH).

The primer sequences used are as follows:

HUMAN:

Gene Sequence 5′-3′ Genbank®

accession no.

Size (bp)

GAPDH FP: GGGAAGGTGAAGGTCGGAGT RP: TTGAGGTCAATGAAGGGGTCA

NM_002046.4 119

VDR FP:CTTCAGGCGAAGCATGAAGC RP: CCTTCATCATGCCGATGTCC

NM_000376.2 134

CYP24A1 FP: CAAACCGTGGAAGGCCTATC RP: AGTCTTCCCCTTCCAGGATCA

NM_001128915.1 71

CYP27B1 FP: TGGCCCAGATCCTAACACATTT RP: GTCCGGGTCTTGGGTCTAACT

NM_000785.3 73

Importin

α3(KPNA4)

FP: TGTGAGCAAGCAGTGTGGGCA RP: TGGTGGTGGGTCTTTGTGGCG

NM_002268.4 186

Importin

α4(KPNA3)

FP: ATCCCCCGCCGCCTATGGAG RP: CTGGGTCTGCTCGTCGGTGC

NM_002267.3 266

78

MICE:

Gene Sequence 5′-3′ Genbank®

accession no.

Size (bp)

VDR FP:ATGGAGGCAATGGCAGCCAGCACCTC

RP:GAAACCCTTGCAGCCTTCACAGGTCA

NM_009504.4 141

Importin α3(KPNA4)

FP- TGATGTCCTGTCCCACTTCCCAAA

RP-CTTGCTGCTGATTGCCTGCTGTTA

NM_008466.5 107

GAPDH FP: TCAACAGCAACTCCCACTCTTCCA RP: ACCCTGTTGCTGTAGCCGTATTCA

NM_008084.2 115

2.5 Transfection of the Cells

To knockdown VDR and importin α3 genes HBSMCs were transfected with

human (h)VDR-small interfering (si)RNA oligonucleotides (sc-106692) (Santa Cruz

Biotechnology, Santa Cruz, CA), (h)KPNA4 siRNA (H00003840-R01) (Novus

Biologicals, Littleton, CO) or scrambled siRNA oligonucleotides (sc-37007) (Santa Cruz

Biotechnology, Santa Cruz, CA) serving as a negative control using FuGENE-HD

Transfection reagent (Roche Applied Science, Roche, Germany) according to

manufacturer’s instruction. The transfection efficiency was measured using Green

Fluorescent Protein (GFP) as a marker, which demonstrated that >85% of the cells

expressed GFP after 30hrs. The viability of the transfected cells was ~95% at this

timepoint. The knockdown efficiency was analyzed by western blot analysis.

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2.6 Protein Isolation

The cell monolayer was washed with serum free media and the cells were

detached by trypsinization and centrifuged at 300 g for 5 min. Cell lysis was performed

by addition of 50 μl of ice-cold RIPA (radioimmunoprecipitation) buffer (Sigma

Aldrich, Saint Louis, MO) with 1% protease inhibitor (P8340, Sigma, Saint Louis, MO).

The cell extract was incubated on ice and vortexed periodically for 30 min. The lysate

was sonicated and transferred to Eppendorf microcentrifuge tubes and centrifuged at

14,000 g for 10 min at 4°C. The pellet was discarded, and the protein lysates stored at

−70°C for protein expression analysis.

2.7 Nuclear Protein extraction

The nuclear protein was extracted according to the manufacture’s protocol

(Active Motif, Carlsbad, CA #40010). The cell monolayer was washed with cold

PBS/phosphatase inhibitors provided in the kit. Cells were gently scraped with cell

lifter, collected in a pre-chilled 15 ml conical tube and centrifuged at 500 rpm for 5 min

at 4⁰C. The pellet was gently resuspended in 150 μl 1x HBSS (hypotonic buffer) and

incubated for 15 min on ice. Detergent (5 μl) was added, then vortexed for 10 sec, and

centrifuged at 14000 g for 30 sec at 4⁰C. The supernatant was collected (cytoplasmic

fraction) in a pre-chilled microcentrifuge tube. The pellet was resuspended in 30 μl

complete lysis buffer and vortexed for 10 sec. The suspension was incubated for 30 min

on ice on a rocking platform set at 150 rpm then vortexed for 30 sec. After

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centrifugation at 14000 g for 10 min at 4⁰C the supernatant (nuclear fraction) was

collected. The nuclear protein obtained was aliquoted and stored at –80ºC.

2.8 Immunoblotting

The protein was quantified and 50 μg of each protein sample was resolved by

electrophoresis using 10-20% polyacryamide gels (BioRad, Hercules, CA), after mixing

with Laemmli loading buffer with 10% mercaptoethanol. The lysate was separated by gel

electrophoresis and transferred onto nitrocellulose membranes (Bio-Rad Laboratories,

Hercules, CA). Non-specific proteins were blocked by incubating the membranes in

blocking buffer (5% w/v nonfat dry milk, 0.05% Tween 20 in PBS). The membranes

were blocked then probed with a specific primary antibody and incubated overnight at

4⁰C with gentle rocking. The following primary antibodies (1:500 dilutions) were used:

Antibody Species/Source Catalogue No Company City/State

VDR Mouse D-6, sc-13133 Santa Cruz

Biotechnology

Santa Cruz ,CA

CYP24A1 Mouse H00001591 Abnova Walnut,CA

CYP27B1 Rabbit H-90, sc-67261 Santa Cruz

Biotechnology

Santa Cruz ,CA

KPNA4 Goat ab6039 Abcam

Boston,MA

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KPNA3 Goat ab6038 Abcam Boston,MA

NF-κBp65 Rabbit sc-372 Santa Cruz

Biotechnology

Santa Cruz ,CA

Lamin B goat sc-6216 Santa Cruz

Biotechnology

Santa Cruz ,CA

Protein expression in whole cell lysate was normalized against GAPDH (Novus

Biologicals, Littleton, CO). After 5-6 washes (10 minutes each) with washing buffer

(0.05% Tween20 in PBS), HRP-conjugated secondary antibodies (1:2000) (Novus

Biologicals, Littleton, CO) was incubated with the membrane for 1 hr at room

temperature with gentle rocking. The membrane was washed three times with washing

buffer and the immunoreactive bands were visualized using ECL chemiluminescence

detection reagents (Pierce, Rockford, IL). The image was captured with BioChemi CCD

camera (UVP, LLC Upland, CA)

2.9 Annexin V Assay for Apoptosis

Cultured HBSMCs were stimulated with calcitriol (0.1nM-100nM) for 24 hrs. As

per the following protocol, Annexin V Assay was performed using Annexin V FITC

Apoptosis Detection Kit (eBioscience, SanDiego, CA Cat# 88-8005). The cell monolayer

was washed with serum free medium and the cells detached by trypsinization and

centrifuged at 300 × g for 5 min. The resulting pellet was washed with PBS then

resuspended in 1ml 1X binding buffer. The cell suspension (100μl) and 5μl of

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fluorochrome conjugated Annexin V were mixed together and incubated for 12 minutes

at room temperature. Propidium Iodide Staining Solution (5μl) (eBioscience, SanDiego,

CA) was added to the cell suspension (cat. 006990). Samples were analyzed by flow

cytometry by using FACS Aria Flow Cytometry System (BD Biosciences, San Jose, CA

)using the PE channel for Propidium Iodide.

2.10.1 Animals and Diets

Female BALB/c mice were used for these studies as these mice are more prone

to develop antigen-specific IgE, Th2 cytokine production, eosinophilic inflammation,

early- and late-phase bronchoconstrictor responses, and AHR. They are a well accepted

animal model of airway hyperresponsiveness [276].

Female BALB/c mice were purchased from Harlan Laboratories (Indianapolis, IN) and

maintained in a specific pathogen-free environment at Creighton University. Food and

water were provided ad libitum. Mice were divided into the following three groups.

Vitamin D Deficient Diet: Breeding trios were fed on Vitamin D-deficient diet

containing 0 IU/kg of cholecalciferol (TD 110272) (Harlan Laboratories, Madison, WI).

The diet also contained 1.2% calcium to prevent the mice from developing hypocalcemic

tetany. The diet was also fortified with vitamin A, E and K. Female offsprings obtained

from the breeding trios were weaned on the same diet.

Vitamin D Sufficient diet: At 21 days of age female pups were weaned on vitamin D-

sufficient diet containing 2000IU/kg of Cholecalciferol and 0.6% calcium (TD110273)

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(Harlan Laboratories, Madison, WI). This dose of vitamin D represents the control

vitamin D diet.

Vitamin D Supplemented diet: At 21 days of age female pups were weaned on vitamin

D-supplemental diet containing 10,000IU/kg Cholecalciferol and 0.6% calcium

(TD110742) (Harlan Laboratories, Madison, WI) This dose of vitamin D represents the

supplemented vitamin D diet.

The research protocol of this study was approved by the Institutional Animal Care and

Use Committee of Creighton University.

2.10.2 OVA-Sensitization and Challenge

Naïve mouse is artificially sensitized and challenged to a foreign antigen to

develop allergic airway inflammation. OVA-albumin is the most potent and preferred

allergen [277]. The OVA allergen is mixed with adjuvant, aluminum hydroxide (AlOH3)

and this mixture is injected in a naïve mouse by intraperitoneal route which then

produces an innate immune response. A second dose is given at a fixed time interval

(usually 1-2 weeks) which induces an adaptive immune response [278]. The following

dose of allergen is given in the form of aerosol in the lungs induces an acute allergic

inflammation and eosinophila in the lungs [278]. Subsequent exposures to the same

allergen at regular intervals induces a chronic allergic asthma characterized by epithelial

cell hypertrophy, thickening of the smooth muscle cell layer, increased AHR, increased

neutrophilic infiltration [278,279].

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Figure-8: Protocol for OVA-sensitization and challenge

2.10.3 Induction of Allergic Airway Inflammation

All three groups of mice were OVA- sensitized and challenged starting at six weeks

of age. Allergic airway inflammation was induced by intraperitoneal injection of 20 µg

OVA (Sigma-Aldrich, St. Louis, MO) emulsified in 2.25 mg of Inject Alum (Pierce

Biotechnology, Rockford, IL) in a total volume of 100 μl on Days 0 and 14 (Figure 8).

Animals were aerosol challenged with 1% OVA/PBS using an Aeroneb Pro nebulizer

(Aerogen, Somerset, PA) for 20 minutes on three consecutive days from day 28 to day

30 and with 5% OVA on day 32. On day 33, airway hyperresponsiveness (AHR) and

enhanced pause in response to aerosolized acetyl β-methylcholine in PBS (3.125 mg/ml-

100mg/ml) (Sigma-Aldrich) were measured in the all mice using whole-body

plethysmography (Buxco Electronics, Troy, NY). Mice with established airway

hyperresponsiveness (AHR) were then challenged with 5% OVA/PBS by aerosol on day

44 and non-sensitized control mice were treated only with vehicle (sterile PBS). On day

45, AHR and enhanced pause in response to aerosolized acetyl β-methylcholine (Sigma-

Aldrich) were measured in all mice using whole body plethysmography (Buxco

85

Electronics, Troy, NY) and the data reported in Penh values (Figure 1). On day 46, AHR

to methacholine was measured with the invasive tracheotomy method to measure

specific airway resistance followed by collection of BALF, blood, and lungs.

2.10.4: Non- Invasive Measurement for Assessment of Pulmonary Function

Pulmonary function in mice was measured using non-invasive single-chamber

barometric whole body plethysmography (WBP) (Buxco Electronics Inc, Troy, NY)

[280]. This method evaluates the AHR to increasing doses of methacholine which

causes cholinergic stimulation in unrestrained conscious mice. The methacholine elicits

a very characteristic waveform in the WBP, with a large deflection early in expiration,

followed by duration of very low amplitude (pause) for the rest of expiration [281]. The

pause measures the changes in the box pressure occurring during expiration [282]. The

enhanced pause (Penh) is an indicator of AHR and is calculated using the box pressure

signal during inspiration and expiration and the timing of early and late expiration

[283].

Advantages of Non-Invasive WBP: WBP method is extensively used in asthma research

[283]. This method allows animals to freely move inside the chamber while the

pulmonary function is being measured eliminating the need for anesthesia [284]. The

non-invasive WBP also allows repeated testing of animals as the mice are not sacrificed

during the experiment.

Disadvantages of Non-Invasive WBP: Non-Invasive WBP may be affected by

temperature, humidity and the motion of the animal inside the chamber. There is also

86

controversy as to whether or not Penh value correlates well with airway resistance [285-

287].

2.10.5 Invasive Measurement for Assessment of Pulmonary Function

For better assessment of lower airway mechanics and dynamic compliance of

lungs an invasive method of pulmonary function measurement involving anesthesia,

tracheotomy, tracheal intubation, and mechanical ventilation may be used [288]. The

disadvantage of this technique is that it is time consuming and limits data production.

Secondly, the dose of anesthesia needs to be standardized and lastly, the investigator

must have a considerable expertise in animal surgery and respiratory physiology [289].

The non-invasive method of WBP is used for screening purposes while the pulmonary

function data can be confirmed by the invasive method. To confirm AHR to

methacholine, OVA-sensitized and non-sensitized mice were randomly selected and

were anesthetized, and cannulated via tracheostomy.

Anesthesia was administered by intraperitoneal injection of sodium pentobarbital at a

dose of (1.6mg/20g) of body weight. Mice were placed in the supine position and

surgically implanted with a cannula connected through the tracheal manifold to a

ventilation pump in the single chamber plethysmography for anesthetized animals

(Buxco-Elan TMSeries RC, ver 1.0 r3, Buxco Research Systems, Wilmington, North

Carolina). After mice were stabilized, they were challenged with aerosolized PBS

followed by increasing doses of methacholine (3.1, 6.25, 12.5, 25, 50, and 100 mg/ml).

Pressure and flow data was continuously recorded by pulmonary software (BioSystem

XA, Buxco Electronics, Inc.) to calculate lung specific airway resistance (RL).

87

2.11: Measurement of serum 25(OH) D levels and serum calcium levels

After euthanasia, 800μl of blood was collected from the left ventricle of mice.

100μl was saved for Tregs analysis and the remainder then kept at room temperature for

3 hours then centrifuged at 5000 rpm for 15 min. Serum was separated and samples

were sent to Creighton University Medical Center Pathology laboratory for

measurements of serum 25(OH) D and calcium.

2.12: Bronchoalveolar lavage Fluid (BALF) and Multiplex Cytokine Analysis

After sacrificing the mice, lungs were gently lavaged with 1 ml of warm saline

(37º C) via a tracheal cannula. Total cell counts were performed coulter counter

(Beckman and Coulter, Fullerton, CA). All samples were centrifuge at 400 rpm, for 10

minutes, and the supernatant stored in -80⁰C until multiplex assays were performed.

Recovered cells in the BALF were suspended in sterile PBS and cytospin slides were

stained (Hema 3 stain set, Biochemical Sciences, Inc., Swedesboro, NJ) for differential

counts. Differential analysis was performed using standard morphological criteria. 300

cells were examined per cytospin slide and absolute cell numbers were calculated per

milliliter of the BALF based on the percentage of individual cell in a slide.

The levels of IL-4, IL-5, IL-6, IL-10, IL-13, IL-17,RANTES, IP-10 and TNF-α in

the BAL fluid supernatants were measured by the Milliplex Mouse Cytokine/Chemokine

kit (Millipore, Billerica, MA) on a Luminex 200 analyzer (Luminex Corp, Austin, TX,

USA) following manufacturer’s recommendations.

Individual vials of beads were sonicated for 30 seconds and vortexed for 1

88

minute. They were then diluted with assay buffer and vortexed again. Deionized water

(250 μl ) was added to the Quality control 1 and Control 2, they were then thoroughly

mixed and vortexed to ensure proper mixing. To prepare the Standard the Mouse

Cytokine Standard was reconstituted with 250 μl of deionized water resulting in a

10,000 pg/ml concentration.Serial dilutions were prepared by adding 50 μl of vial

containing a total of 200 μl resulting in final concentrations of 2,000, 400, 80, 16 and

3.2 pg/ml. The 0 pg/ml standard (Background) consisted of only Assay Buffer. The

microtitre filter plate was washed with 200 μl Wash Buffer per sample. The plate was

shaken for 10min on a plate shaker and the wash buffer was removed by vacuum.

Standard or control or samples (25 μl) were added to appropriate wells. Premixed beads

(25 μl) were added to each well. The plate was sealed and incubated on a shaker

overnight at 4⁰C. The fluid was then removed by vacuum and washed 2 times with wash

buffer. Detection antibody (25 μl ) was added and incubated on a plate shaker for an

hour at room temperature. Streptavidin-Phycoerythrin (25 μl) was added to each well

and incubated on a shaker for 30 mins at room temperature. The fluid was removed by

vaccum and the plate was washed 2 times with 200 μl of wash buffer. The analysis was

done in duplicate and the cytokine concentrations were calculated against the standards

using Beadview® software (ver. 1.03, Upstate). The minimum detection concentration

was <2 pg/ml for IL-4, IL-5, IL-17,IP-10 and TNF-α, 8.0 pg/mL for IL-10, 10.8 pg/ml

for IL-13, 2.5 pg/ml for RANTES and 3.4 pg/mL for IL-6.

89

2.13: Histology and Lung Tissue Staining

Mice were euthanized with 100 μl of sodium pentobarbital (5mg/20g). Lung

lobes were fixed in 4% formalin and paraffin embedded. Thin sections of 5µm thickness

were cut from paraffin blocks using a Microtome (IMEB, San Marcos, CA). Sections were

submerged in a water bath at 39 – 40 °C and mounted on microscope charged slides.

Slides were deparaffinized by incubation in xylene for 5 minutes on a shaker followed by

ethanol xylene mix (50:50)for another 5 minutes. They were then rehydrated using

different concentrations of ethanol that is 100%, 90%, 80% and 70% for five minutes in

each. These lung sections were stained using Hematoxylin and Eosin (H&E) Kit

(Newcomer Supply, Middleton, WI). Slides were placed in distilled water for 2 minutes

then placed in Harris hematoxylin solution for 30 sec and rinsed in running tap water

for 5 min. Slides were then differentiated using a bluing solution for 10 sec and rinsed in

distilled water. The sections were counterstained with eosin for 22 sec. The tissue was

dehydrated using 95% and 100% ethanol followed by dipping in xylene ethanol mix

(50:50) for 5 times and in xylene for five minutes. Cytoseal 60 (Richard-Allan Scientific,

Waltham, MA) mounting medium was used for mounting slides.

2.14: Trichrome Staining of Lung Sections to Identify Collagen Deposition

Masson’s trichrome staining was used to identify collagen following the standard

protocol recommended by the manufacturer (IMEB Inc., San Marcos, CA). The lungs

sections were deparrafinized as described above. These deparrafinized sections were

incubated in Bouin solution at 56⁰C for 1 hour, and rinsed in running tap water for five

90

min. Sections were then placed in working weigert’s iron hematoxylin stain for 10

minutes and rinsed. The slides were then stained with Biebrich’s Scarlet acid fuchsin

solution for 5-10 min and rinsed in DI water for 30 sec. These rinsed slides were

submerged in phosphotungstic and phosphomolybdic acid solution for 5 min, aniline

blue stain for 5-10 min,Hematoxylin solution for 5 min and then 1% acetic acid solution

for one minute. The slides were then rinsed in distilled water for 30 sec. The sections

were dehydrated using 100% ethanol for 2 minutes and xylene-ethanol mix (50:50) for 1

minute and xylene for 1 minute. The sections were again cleared by using addition

xylene for 1 minute. Cytoseal 60 (Richard-Allan Scientific, Waltham, MA) mounting

medium was used for mounting slides.

2.15: PAS staining in Lung Sections showing Mucus Deposition

Mucus secretion was identified by periodic acid-Schiff (PAS) reaction using the

standard protocol recommended by the manufacturer (Sigma-Aldrich, St. Louis, MO).

Lung sections were deparaffinzed as described, previously and placed in distilled water

for five min. The slides were submerged in periodic acid solution for 5 min and rinsed

in several changes of distilled water. This was followed by immersing the slides in

Schiff’s reagent for 15 min at room temperature (18–26°C) and washed with running tap

water for 5 min. The sections were counterstained in Hematoxylin Solution, Gill No. 3,

for 90 sec. Cytoseal 60 (Richard-Allan Scientific, Waltham, MA) mounting medium was

used for mounting slides.

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2.16 Immunofluorescence

Paraffin embedded lung sections were mounted on slides and deparaffinized and

rehydrated as described previously. The slides were stained using specific antibodies to

VDR and importin α3. The sections were treated with DAKO target retrieval solution in

a steamer for 20 min, then cooled for 20 min and washed with PBS. The non-specific

binding was blocked for 2 hr at room temperature by using a blocking solution of

normal goat or normal donkey serum (1:200 dilutions), 0.25% Triton and PBS. The

sections were incubated with the primary antibody (1:200 dilutions) in blocking

solution overnight at 4ºC. The following antibodies were used.

Antibody Species/Source Catalogue No. Company City/State

KPNA4 goat ab6039 Abcam Boston,MA

VDR rabbit sc-1008 SantaCruz Biotechnology

SantaCruz, CA

TNF-α rabbit ab6671 Abcam Boston,MA

After washing with PBS, the tissue sections were incubated with Cy3-conjugated

secondary antibody for 2 hr at room temp. They were washed with PBS and mounted

using Vectashield mounting media with DAPI. The tissues were observed using an

Olympus inverted fluorescence microscope (Olympus BX51, Shinjuku, Tokyo, Japan).

Negative controls of sections were run without incubation with the primary antibody.

Computer-assisted quantification of immunostaining was performed using NIH Image J

software (http://rsb.info.nih.gov/ij/). Mean fluorescent intensity (MFI) of VDR

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immunostaining in lung tissue measured in arbitrary units (AU). Data are shown as

mean ± SEM of values of 10 measurements per mouse.

2.17 T-regulatory cell identification

Upon euthanizing 100μl of blood was collected into a sterile K3 EDTA

VACUTAINER blood collection tube (BD vacutainer Franklin Lakes, NJ).

Anticoagulated blood was kept at room temperature (20° to 25°C). T-regs cells were

identified as per following protocol. One hundred μL of whole blood was added to a

12x75mm tube followed by addition of 1 µl of fluorochrome-conjugated monoclonal

antibody to anti-mouse CD4+(FITC) and anti-mouse CD25+(PE)[ T-regulatory cell

markers]. The tube was gently vortexed and incubated for 30 min in the dark at room

temperature. FACSLyse formulation solution -1x (2 ml) was added, vortexed and

incubated at room temperature for 10 min. The tube was centrifuge at 500g for 5 min

and supernatant was removed. PBS4 (2ml) was added followed by centrifugation at

500g for 5 minutes. The supernatant was removed and the cells were resuspended in 0.5

mL of FACSFix formulation solution and mixed. Flow cytometric analysis was

performed by using FACS Aria Flow Cytometry System (BD Biosciences, San Jose, CA).

FITC, or PE-labeled IgG (BD Pharmingen, San Jose, CA) was used as the isotype

control.

2.18: Data processing and Statistical Analysis

Flow cytometric data analyzed using Flowjo software (Tree Star Inc. Ashland,

OR). The Graph Pad Prism 4.0 biochemical statistical package (Graph Pad Software,

93

Inc, San Diego, CA) software was used to analyze data and plot graphs. Statistical

analysis was performed using one-way ANOVA to analyze statistically significant

differences between groups. Post –hoc test included either Dunnett (all colums were

compared to control) or Bonferroni’s test (all colums were compared to each other).

Values of all measurements are reported as mean ± SEM. For in vivo studies, an

Unpaired Student’s t test was used to determine differences between two groups.

Multiple group comparison was made using one-way ANOVA with the Bonferroni

correction. Values of all measurements are reported as mean ± SEM. The P< 0.05 was

considered significant. (*P < 0.05, **P < 0.01, ***P < 0.001)

2.19 Power Analysis

To determine group size, power analysis was performed using changes in

pulmonary function in vivo as the endpoint. To detect a difference in the change of at

least 30% in pulmonary function between two groups with standard deviation of 15%

between the animals in the experimental group and when the data are analyzed the

Student’s t-test, the sample size necessary to have at least 80% power to detect this

change is 6 mice per group. In our previous studies, we experienced about 2-3%

mortality following antigen challenge and/or after bronchoconstrictory response to

methacholine. We rarely find the animal dead right after challenge to antigen or

methacholine. Thus, on an average 1-2 mice out of 60 mice were reported dead. To

account for this, 7 mice per experimental group were used in Specific Aim 3.

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SECTION 3

RESULT

95

3.1: Effect of calcitriol on mRNA transcripts and protein expression of VDR

and its metabolizing enzymes CYP24A1 and CYP27B1 in HBSMCs

Cultured HBSMCs were serum staved for 24 hours. Calcitriol was then added at

doses ranging from 0.1nM-100nM for 24 hours. RNA was isolated from these cells as

previously described qPCR was performed. Protein was also isolated from the cells and

the protein expression examined by immunoblotting. Unstimulated cells expressed VDR

and its metabolizing enzymes CYP24A1 and CYP27B1. Following calcitriol (0.1-100nM)

treatment, there was significant increase in both mRNA and protein expression of VDR

and CYP27A1 (Figure 9 A-D). A decrease in CYP27B1 mRNA and protein expression was

observed with increasing doses of calcitriol (Figure 9 E-F). The expression of VDR and

its metabolizing enzymes in HBSMCs suggests that HBSMCs possesses the machinery to

locally metabolize vitamin D and that HBSMCs are vitamin D responsive [290].

96

Figure 9: Effect of Calcitriol treatment on mRNA and protein expression of

VDR (A, B), CYP24A1 (C, D) and CYP27B1 (E, F) in HBSMCs: Cultured

HBSMCs were serum starved for 24 hours followed by treatment with different doses of

calcitriol (0.1- 100 nM) for 24 hours. The mRNA and protein were isolated from cell

lysates and subjected to qPCR and immunoblotting, performed respectively. Data are

shown as mean ± SEM from three individual samples per experiment and repeated

three times ***p<0.001, **p<0.01.

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3.2: Calcitriol decreases mRNA and protein expression of importin α3 but

did not alter the mRNA and protein expression of importin α4 in HBSMCs

Unstimulated HBSMCs express both protein and mRNA transcripts of importin

α3 and importin α4. Following treatment with increasing doses of calcitriol (0.1-

100nM), a significant decrease in both mRNA and protein expression of importin α3

was observed in HBSMCs (Figures 10 A, B). Calcitriol treatment at any dose did not

effect mRNA or protein expression of importin α4 (Figure 10 C, D). These results

demonstrate that treatment with calcitriol decreased of importin α3 expression and had

no effect on importin α4. Since the nuclear translocation of RelA is mediated by

importin α 3 and importin α4 [127], these data prompted us to analyze the effect of

calcitriol on the nuclear expression of RelA both in unstimulated and stimulated

conditions.

98

Figure 10: Effect of calcitriol treatment on mRNA and protein expression of

importin α3 (A, B) and importin α4 (C, D) in HBSMCs: Cultured HBSMCs were

serum starved for 24 hours followed by treatment with different doses of calcitriol (0.1-

100 nM) for 24 hours. mRNA and protein were isolated from cell lysates and qPCR and

immunoblotting performed respectively. Data are shown as mean ± SEM from three

individual samples in each experiment and repeated three times ***p<0.001, **p<0.01.

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3.3: Effect of calcitriol on mRNA transcripts of VDR, CYP24A1, CYP27B1

and importin α3 in HBSMCs (time dependent response)

HBSMCs were treated with calcitriol (100nM) at different time periods (0 -36hrs). The

mRNA expression of VDR and CYP24A1 increased in a time-dependent manner

following calcitriol treatment with the maximum expression of VDR at 18-36hrs post-

treatment and CYP24A1 at times 6-36 hrs post-treatment. Since there was no statistical

difference between these time periods the 24hr time-point was chosen for the following

experiments (Figure 11A, 11B).

In HBSMCs, mRNA expression of CYP27B1 and importin α3 following calcitriol

treatment decreased in a time-dependent manner with minimum expression of both

CYP27B1 and importin α3 at 24 hrs. Therefore, a time period of 24 hrs was chosen for

the following experiments (Figure 11C, 11D).

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Figure 11: Time-dependent effect of calcitriol on mRNA expression of VDR,

CYP24A1, CYP27B1 and importin α3 in HBSMCs: Cultured HBSMCs were serum

starved for 24 hr followed by treatment with calcitriol (100 nM) for 0-36hr. The mRNA

were isolated from cell lysates and qPCR was performed. Data are shown as mean ±

SEM from three individual samples and repeated three times *p <0.05, **p<0.01, ***p

<0.001.

101

3.4: Effect of calcitriol treatment on apoptosis in HBSMCs

To determine the effect of calcitriol on apoptosis in HBSMCs, cells were serum starved

for 24 hrs and then treated with different concentration of calcitriol (0.1nM -100 nM)

for 24 hrs and the Annexin V Apoptosis Assay was performed. We examined the

percentage of apoptotic cells by Annexin V and PI staining. Early apoptotic cells were

Annexin V positive and late apoptotic cells were both Annexin V and PI positive. Our

results demonstrate that 24 hrs of calcitriol treatment had no significant effect on cell

apoptosis in HBSMCs at any of the doses studied (0.1nM-100nM) (Figure 12).

102

Figure 12: Effect of calcitriol on cell apoptosis HBSMCs. HBSMCs were

cultured in T 25 flask and after the cells were 70-80% confluent, they were serum

starved for 24 hours and treated with varying concentrations of calcitriol (0.1 nM-100

nM) for 24 hours. The Annexin V Apotosis Assay was performed. The cells were stained

with annexin V and PI and analyzed by flow cytometry. The cells in the upper left

quadrant are annexin V positive and representative of early apoptotic cells. The cells in

the upper right quadrant are Annexin V and PI positive and representative of late

103

apoptotic cells. Data are shown as the mean ± SEM and representative of three

individual samples and repeated three times.

104

3.5: VDR knockdown abolishes the reduced expression of importin α3 by

calcitriol in HBSMCs

To further explore the role of VDR in the regulation of importin α3, we

evaluated the effect of reduced VDR expression in the HBSMCs by Transfection of cells

with siRNA. Transfection efficiency was determined using GFP as a marker, which

demonstrated viability of 95% and > 85% of the cells expressing GFP 30hrs after

transfection (Figure 13 A, B). The VDR level was reduced using hVDR specific siRNA,

with unrelated (scramble) siRNA as control. The knockdown efficiency was analyzed by

western blot analysis. As shown in Fig. 5C, 30 hr after siRNA transfection, VDR protein

expression was markedly reduced, with the knockdown efficiency of almost 90%.

VDR knockdown also decreased the expression of importin α3 by calcitriol.

Calcitriol treatment failed to reduce the expression of importin α3 in the VDR-siRNA-

transfected cells in contrast to what was observed in the scrambled siRNA-treated cells

(Fig. 5D). These data demonstrate that VDR knockdown abolishes calcitriol-induced

reduced expression of importin α3. These results are consistent with the data presented

in Fig. 10 (A, B) which shows that calcitriol significantly decreased the expression of

importin α3 in HBSMCs.

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Figure 13: Effect of VDR siRNA on the expression of VDR (C) and the effect

of calcitriol treatment on the protein expression of importin α3 (D) in VDR

knockdown HBSMCs:

Figure 13 (A) Viability of cells after 30 hrs of transfection (B) Transfection efficiency

with GFP after 30 hrs (magnification 100X), quantified by counting the GFP positive

cells divided by total number of live cells. Cultured HBSMCs were transfected with VDR

siRNA or scrambled siRNA and were stimulated with calcitriol (100 nM) for 24 hours.

Protein was isolated from cell lysates and subjected to immunoblotting. Data is shown

as mean ± SEM from three individual samples in each experiment ***p<0.001.

106

3.6: TNF-α increases activated RelA expression in the nuclear fraction of

HBSMCs in a dose-dependent manner

The proinflammatory cytokines, IL-1β and TNF-α, have been shown to play a

prominent role in the development of airway responsiveness and airway inflammation

in bronchial asthma [54]. HBSMCs were treated with different concentration of TNF-α

(1-100 ng/ml) for 20 mins. An abbreviated time course was performed as the half life of

activated RelA is less than 30 mins [291]. Under unstimulated conditions RelA was

detected in the nucleus. TNF-α (10-50ng/ml) treatment significantly increased the

nuclear protein expression of RelA (Figure 14A). Although TNF-α (100ng/ml) also

increased RelA expression, but the viability of the cells was significantly affected.

Therefore, a dose of 10ng/ml TNF-α was chosen for the following experiments.

HBSMCs were treated with TNF-α (10ng/ml) for varying time periods (0 -60

mins). The nuclear protein expression of RelA in HBSMCs increased in a time-

dependent manner following TNF-α treatment with the maximum expression at 15 min.

RelA levels returned to the baseline at 30 min (Figure 14B). Therefore, in the following

nuclear experiments HBSMCs were treated with a dose of 10 ng/ml of TNF-α for 15

mins.

107

Figure 14: Effect of TNF-α on the expression of activated-RelA expression in

the nuclear fraction of HBSMCs in a dose-and time-dependent manner (A,

B)

HBSMCs were treated with different doses of TNF-α (1-100ng/ml) for 20 min (A) and

TNF-α (10ng/ml) at 0-60mins (B). Nuclear proteins were extracted and analyzed for

RelA by immunoblotting. Data is shown as mean ± SEM from three individual samples

in each experiment and repeated three times **p<0.01, ***p<0.001.

108

3.7: Calcitriol decreases expression of activated RelA in the nuclear fraction

of TNF-α stimulated HBSMCs and Importin α3 knockdown decreases the

TNF-α induced activation of RelA and abolishes the action of calcitriol on

RelA translocation in the nuclear fraction of HBSMCs.

The importin α3 was knocked down using human importin α3-specific siRNA.

Scrambled siRNA was used as the control. As shown in Figure 15A, 30 hr after siRNA

transfection, the importin α3 protein expression was markedly reduced and knockdown

efficiency was nearly 85%.

Upon stimulation with TNF-α (10 ng/ml) for 15 min, there was significantly

increased nuclear protein expression of RelA. This effect was attenuated by calcitriol,

indicating that calcitriol decreases the translocation of RelA to the nucleus (Figure 15B).

To verify the role of importin α3 in RelA translocation from the cytoplasm to the

nucleus, we evaluated the effect of TNF-α on nuclear RelA expression in importin α3

siRNA-transfected HBSMCs. Importin α3 knockdown decreased of RelA translocation

to the nucleus in response to TNF-α. Since there was not complete absence of RelA

translocation to the nucleus, this suggests a potential role of either importin α4 or

another protein [127].

In the TNF-α-treated importin α3-siRNA-transfected cells, RelA expression was

significantly downregulated as compared to TNF-α-treated non-transfected cells. Our

results are in accordance with previous studies confirming that importin α3 plays a

major role in RelA translocation to the nucleus[127,292]. Concomitant treatment of

TNF-α with calcitriol of importin α3-siRNA-transfected cells had no effect on RelA

expression as compared to TNF-α treated importin α3-siRNA-transfected cells.

109

These data confirm that the action of calcitriol to reduce RelA translocation is

mediated by decrease in importin α3 expression.

110

Figure 15: Effect of importin α3 siRNA on the expression of importin α3 (A)

Effect of calcitriol on nuclear protein expression of in importin α3

knockdown HBSMCs on stimulation with TNF-α:

Cultured HBSMCs were transfected with importin α3 siRNA or scrambled siRNA and

were treated with calcitriol (100 nM) for 24 hrs and TNF-α (10ng/ml) for 15 min.

Nuclear proteins (B) were extracted for RelA immunoblotting. Lamin B was used as a

housekeeping protein for the nuclear extracts. Data are shown as mean ± SEM from

three individual samples in each experiment repeated three times *p<0.05, ***p<0.001.

These data confirm that the action of calcitriol to reduce RelA translocation is mediated

by decrease in importin α3 expression.

111

3.8: Calcitriol decreases TNF-α -induced mRNA and protein expression of

importin α3 in HBSMCs

The effect of the pro-inflammatory cytokine TNF-α on the expression of importin

α3 was examined. HBSMCs were treated with calcitriol (100 nM) ± TNF-α (10ng/ml) for

24hrs, followed by RNA and protein isolation for qPCR and immunoblotting,

respectively. Following TNF-α treatment there was ~3-fold increase in mRNA

expression (Figure 16A) and a significant increase in protein expression (Figure 16B) of

importin α3. Calcitriol treatment attenuated the TNF-α induced increase in the mRNA

and protein expression of importin α3 (Figure 16 A, B).

112

Figure 16: Effect of calcitriol ± TNF-α on mRNA and protein expression of

importin α3 in HBSMCs: Cultured HBSMCs were serum starved for 24 hours

followed by treatment with TNF-α (10ng/ml), (A, B) ± calcitriol (100 nM) for 24 hours.

The mRNA and protein was isolated from whole cell lysates and qPCR and

immunoblotting were performed respectively. Data is shown as mean ± SEM from three

individual samples in each experiment and repeated three times *p <0.05, **p<0.01,

***p <0.001.

113

3.9: Calcitriol decreases IL-1β-induced mRNA transcript and protein

expression of importin α3 in HBSMCs

The effect of IL-1β, a pro-inflammatory cytokine, on the expression of importin

α3 was examined. HBSMCs were treated with calcitriol (100 nM) ± IL-1β (10ng/ml) for

24hrs, followed by RNA and protein isolation for qPCR and immunoblotting. After IL-1β

treatment, there was ~ 2 -fold increase in mRNA expression and a significant increase in

protein expression of importin α3 (Figure 17 A, B). Calcitriol treatment attenuated the

IL-1β-induced increase in the importin α3 expression.

114

Figure 17: Effect of calcitriol ±IL-1β on mRNA and protein expression of

importin α3 in HBSMCs: Cultured HBSMCs were serum starved for 24 hours

followed by treatment with IL-1β (10ng/ml) ± calcitriol (100 nM) for 24 hours (A,B).

The mRNA and protein were isolated from whole cell lysates and qPCR (A) and

immunoblotting (B) were performed. Data are shown as mean ± SEM from three

individual samples in each experiment and repeated three times *p <0.05, **p<0.01,

***p <0.001.

115

3.10: Effect of Calcitriol, TNF-α, and IL-10 on the mRNA and protein

expression of importin α3 in HBSMCs

HBSMCs were treated with calcitriol (100 nM) ± IL-10 (10ng/ml) ± TNFα

(10ng/ml) for 24hrs, followed by RNA and protein isolation for qPCR and Western

blotting. Upon stimulation with IL-10, there was no significant change in the mRNA and

protein expression of importinα3 as compared to control. This led us to conclude that

IL-10 may not effect importin α3 expression. However, IL-10 significantly attenuated

the TNF-α induced increase in the expression of importin α3 and this decrease in

expression of importin α3 was further potentiated by addition of calcitriol (Figure 18 A,

B).

116

Figure 18: Effect of Calcitriol, TNF- α, and IL-10 on the mRNA and protein

expression of importin α3 in HBSMCs

Cultured HBSMCs were serum starved for 24 hr followed by treatment with IL-10

(10ng/ml) ± TNF-α (10 ng/ml) ± calcitriol (100 nM) for 24 hr. mRNA and protein was

isolated from whole cell lysates and qPCR (A) and immunoblotting (B), were performed.

Data is shown as mean ± SEM from three individual samples in each experiment

repeated three times *p <0.05, **p<0.01,***p <0.001. (Figure 18 A, B).

117

3.11: The dietary vitamin D3 content was revealed in serum:

The vitamin D3 metabolite 25(OH)D in serum was analyzed in mice at 6weeks and at 13

weeks after the introduction of diet. The results illustrated the content of vitamin D3 in

the diet as a reflection of 25 (OH)D levels in mice serum. Serum 25 (OH) D levels in

mice fed on vitamin D deficient diet (0 IU/Kg) were 5.00 ± 0.25 ng/ml (n=7), serum 25

(OH)D levels in mice supplemented with 2,000 IU of vitamin D3 were 31.00 ± 0.75

ng/ml (n=7) and serum 25 (OH)D levels in mice supplemented with 10,000 IU of

vitamin D3 were 67.12 ± 0.5 ng/ml (n=7) at 13 weeks of age. However, no change in

serum calcium levels were observed in all three groups of mice on different vitamin D

diets at 13 weeks after the diets were started.

118

3.12: Effect of vitamin D diets on AHR to methacholine in OVA-sensitized

and challenged mice

The AHR to methacholine was examined and established on days 33 and 45 as

measured by non-invasive whole-body plethysmography (Figure 19 A, B). OVA-

sensitized and challenged mice demonstrated elevated AHR to methacholine compared

to PBS control in all dietary groups. Vitamin D-deficient OVA-sensitized and challenged

mice exhibited significantly higher AHR to methacholine compared to vitamin D

sufficient OVA-sensitized mice. Interestingly, OVA-sensitized and challenged mice fed

on vitamin D supplemented diet exhibited significant reduction in AHR compared to

vitamin D sufficient OVA-sensitized mice. However, the AHR exhibited by vitamin D

supplemented OVA-sensitized and challenged was significantly higher than PBS control

mice of the same group. These results were confirmed with a more rigorous invasive

method involving tracheostomy in anesthetized mice to measure specific airway

resistance (Figure 19 C) and a similar pattern was observed.

There was no significant difference in AHR among PBS control vitamin D-deficient, PBS

control vitamin D Sufficient, PBS control vitamin D Supplemented. These data indicate

serum levels of vitamin D had no effect on the AHR of control PBS mice.

Administration of 100 mg/ml methacholine at day 45, exhibited Penh values of

7.38 ± 0.64 in vitamin D deficient OVA-sensitized and challenged mice; 3.73 ± 0.55 in

vitamin D sufficient OVA-sensitized and challenged mice; 2.12 ± 0.39 in vitamin D

supplemented OVA-sensitized and challenged mice (n = 7 in each group) (Figure 19 B).

Specific airway resistance induced by 100 mg/ml methacholine exhibited mean values of

11.11 ± 0.59 in vitamin D deficient OVA-sensitized and challenged mice; 7.18 ± 0.58 in

vitamin D sufficient OVA-sensitized and challenged mice; 4.18 ± 0.41 cm H2O.s/ml in

119

vitamin D supplemented OVA-sensitized and challenged mice (Figure 19C).

120

Figure 19: AHR and Specific Airway Resistance to Methacholine. A. Day 33,

AHR to methacholine (Mch) was established (N = 7 mice). B. Day 45, AHR to

methacholine (Mch) was measured again (N = 7 mice). C. F0ur randomly selected

animals from each group of PBS and OVA-sensitized and challenged groups were

subjected to invasive method of tracheostomy and airway resistance (RL) to Mch was

recorded. - Vitamin D deficient OVA-sensitized and challenged mice vs Vitamin D

sufficient OVA-sensitized and challenged mice - (*p <0.05; **p <0.01; ***p <0.001); #

Vitamin D deficient OVA-sensitized and challenged mice vs Vitamin D supplemented

OVA-sensitized and challenged mice; (#p <0.05; ##p <0.001); Vitamin D sufficient

OVA-sensitized and challenged mice vs Vitamin D supplemented OVA-sensitized and

121

challenged mice ( p <0.05; p <0.01); Vitamin D deficient PBS control mice vs

Vitamin D deficient OVA-sensitized and challenged mice;( p <0.001) ; ø Vitamin D

sufficient PBS control mice vs Vitamin D sufficient OVA-sensitized and challenged mice

(øp <0.05; øøp <0.01; øøøp <0.001); Vitamin D supplemented PBS control mice vs

Vitamin D supplemented OVA-sensitized and challenged mice - ( p <0.05; p <0.01;

p <0.001);

122

3.13: Differences in the degree of Airway Remodeling after OVA

sensitization and challenge in vitamin D diet groups

After tracheostomy, seven animals from each group were sacrificed. The lungs were

harvested and sectioned and then stained with H&E, Trichrome and PAS to

demonstrate histological hallmarks of asthmatic airways. Airways of the PBS control

mice in all vitamin D groups exhibited normal airway parenchyma, with no mucus

staining and little collagen staining around the respiratory epithelium.

Vitamin D deficient OVA-sensitized and challenged mice exhibited exaggerated

features of severe airway remodeling compared to vitamin D sufficient OVA-sensitized

and challenged mice. The airway of vitamin D deficient OVA-sensitized and challenged

mice had marked epithelial cell hypertrophy indicated by an increase in epithelial cell

height, airway occlusion, and increase mucus staining and collagen deposition. The

lungs of vitamin D supplemented OVA-sensitized and challenged mice exhibited the

signs of mild airway remodeling, showing some airway epithelial cell hypertrophy,

moderate mucus staining, and less collagen deposition (Figure 20 A–Q).

123

Figure 20: Lung histological examination – (I) Hematoxylin and eosin (H&E)

showing differences in the airway remodeling (A-F), (II) Trichrome staining showing

collagen deposition in blue colour (G-L) and, (III) periodic acid-Schiff (PAS) showing

mucous staining in pink colour (M-Q), in the airway of PBS-control and OVA-sensitized

and challenged vitamin D deficient , sufficient and supplemented mice. The histological

sections shown are representative of seven mice in each experimental group.

124

3.14: Effect of vitamin D on total and differential leukocyte counts in the

BALF

Sensitization and challenge of mice with OVA significantly increased the total

number of cells in the BALF with predominant increases in the numbers of eosinophils,

neutrophils, and lymphocytes in all OVA-sensitized and challenged groups of mice

compared to PBS control mice. Vitamin D deficient OVA-sensitized and challenged mice

had increased eosinophils and compared to vitamin D sufficient OVA-sensitized and

challenged mice and vitamin D supplemented OVA-sensitized and challenged mice. A

decrease in the number of eosinophils was observed in the BALF of vitamin D

supplemented OVA-sensitized and challenged mice compared to vitamin D sufficient

OVA-sensitized and challenged mice (Figure 21). However, no difference in the counts

of macrophages, neutrophils and lymphocytes were observed among OVA-sensitized

diet groups.

125

Figure 21: Effect of vitamin D on total and differential leukocyte numbers in

bronchoalveolar lavage fluid of PBS- control and OVA-sensitized mice. Total Leukocyte

count (TLC) in the BALF were counted using Coulter counter and differential analysis

was performed using standard morphological criteria. 300 cells were examined per

cytospin slide and absolute cell numbers were calculated per milliliter of the BALF

based on the percentage of individual cell in a slide. Data is shown as mean SEM for

seven animals in each group (*p < 0.05; **p < 0.01; ***p < 0.001)

126

3.15: Effect of vitamin D on Th2 cytokines in the BALF of OVA-sensitized

and challenged mice

The effect of vitamin D on Th2 cytokines was analyzed in the BALF using

Milliplex mouse cytokine/chemokine kit. A significant increase in the levels of IL-4 (A),

IL-5 (B), and IL-13 (D) in the BALF of all the OVA-sensitized and challenged groups of

mice compared to the PBS control. However, the levels of IL-5 and IL-13 were more

pronounced in vitamin D deficient OVA-sensitized and challenged mice. Vitamin D

supplemented OVA-sensitized and challenged mice had substantially reduced levels of

these cytokines compared to vitamin D sufficient OVA-sensitized and challenged mice.

There was no significant effect of vitamin D on the level of BALF IL-4 among the OVA-

sensitized groups. Conversely, there was a significant reduction in the levels of IL-10 in

vitamin D deficient OVA-sensitized and challenged mice and vitamin D sufficient OVA-

sensitized and challenged mice with no significant change in PBS control groups (C).

The vitamin D supplemented OVA-sensitized and challenged mice had significantly high

levels of IL-10 compared to vitamin D deficient OVA-sensitized and challenged mice and

vitamin D sufficient OVA-sensitized and challenged mice (Figure 22).

127

Figure 22: The levels of Th2 cytokine in the BALF of and OVA-sensitized and -

challenged mice compared to PBS control mice with different vitamin D groups. (A) IL-

4; (B) IL-5; (C ) IL-10 and (D) IL-13. Data are shown as means ±SEM for six animals in

per experimental group. *p, 0.05, **p, 0.01, ***p, 0.001.

128

3.16: Effect of vitamin D on TNF-α, IL-6and IL-17 in the BALF of OVA-

sensitized and challenged mice

OVA-sensitization and challenge significantly increased the levels of BALF TNF-

α, IL-6 and IL-17 compared to the levels of these cytokines in PBS control mice (Figure

23 A,B & C). The levels of these cytokines were higher in vitamin D deficient OVA-

sensitized and challenged mice compared to vitamin D sufficient OVA-sensitized and

challenged mice. In contrast, vitamin D supplemented OVA-sensitized and challenged

mice had significantly lowered BALF levels of TNF-α, IL-6 and IL-17, but, significantly

higher than the levels in PBS controls (Figure 23).

129

Figure 23: After euthanizing the mice, BALF was immediately collected and

centrifuged. The supernatant was collected and frozen. The samples were analyzed for

TNF-α (A); IL-6 (B); and IL-17(C) in OVA-sensitized and -challenged mice compared to

PBS control mice in all dietary groups. Data are shown as means ±SEM for six animals

in each experimental group. *P <0.05, **P < 0.01, ***P < 0.001.

130

3.17: Effect of vitamin D on RANTES and IP-10 in the BALF of OVA-

sensitized and challenged mice

The effect of vitamin D on the levels of chemokines RANTES and IP-10 were analyzed

using Milliplex mouse cytokine/chemokine kit. The BALF levels of RANTES and IP-10

were higher in all OVA-sensitization and challenged mice compared to PBS control

mice. No significant difference in the levels of these cytokines among the OVA-

sensitized and challenged mice was detected between the dietary groups. (Figure 24 A

& B).

131

Figure 24: Levels of RANTES and IP-10 in the BALF. Mice were sensitized and

challenged with OVA-antigen for 45 days before euthanasia. The BALF was collected,

centrifuged and frozen. The cytokines, RANTES (A) and IP-10 (B), in all samples were

measured in the stored samples. Data are shown as means (±SEM) for six animals in

each experimental group. *P <0.05, **P < 0.01, ***P < 0.001.

132

3.18: Effect of vitamin D on CD4+CD25+ T-cells Isolated from the blood of

OVA-sensitized and challenged, and PBS Control Mice

We isolated cells from the blood and performed flow cytometry to determine the

percentage of CD4+CD25+ T-cells in circulation, in each of the diet groups in both PBS

and OVA-sensitized and challenged mice (Figure 25). A substantial decrease in the

percentage of CD4+CD25+T-cells was observed in vitamin D-deficient and-sufficient

OVA sensitized and challenged mice compared to PBS control mice. Interestingly, the

vitamin D supplemented mice had significantly higher percentage of CD4+CD25+T-cells

in the peripheral blood suggesting a positive correlation between serum 25(OH)D levels

and T-regulatory cells.

133

Figure 25: Flow cytometry data showing % of CD4+ CD25 + lymphocytes in mice

peripheral blood. The CD4+ CD25 + cells (regulatory T cell markers) are in the quadrant

2. FITC conjugated monoclonal anti-mouse CD4 and PE-labeled anti-mouse CD25 was

used. Flow cytometric analysis was performed by using FACS Aria Flow Cytometry

134

System. FITC, PE-labeled IgG served as the isotype control. Data are shown as means

(±SEM) of six animals in per experimental group. *P <0.05, ***P < 0.001.

135

3.19: Effect of vitamin D on the expression of TNF-α in the lung of OVA –

sensitized and challenged mice: Low levels of TNF-α expression was observed in

the lung of all PBS control mice. There was a significant increase in TNF-α expression

in all OVA- sensitized and challenged mice, however a significantly higher expression of

TNF-α was found in vitamin D deficient OVA-sensitized and challenged mice had

compared to Vitamin D sufficient OVA-sensitized and challenged mice and vitamin D

supplemented OVA-sensitized and challenged mice. On the other hand, vitamin D

supplemented OVA-sensitized and challenged mice had a significantly lower expression

of TNF-α compared to vitamin D deficient OVA-sensitized and challenged mice or

vitamin D sufficient OVA-sensitized and challenged mice, but significantly higher than

PBS control mice. (Figure 26).

136

Figure 26: (A) TNF-α expression in lung tissue Immunofluorescence staining for TNF-

α in lung of PBS control vitamin D-deficient, vitamin D sufficient and vitamin D

supplemental mice compared to OVA sensitized and challenged vitamin D-deficient,

vitamin D-sufficient and vitamin D supplemented mice. All images are at a

magnification of 600 x. Sections were stained using rabbit anti- TNF-α antibody and

goat anti-rabbit Cy3 as secondary antibody. DAPI was used to stain the nuclei. The

histological slides are representative of seven mice in each experimental group.

(B) Mean fluorescent intensity (MFI) of TNF-α immunostaining in lung tissue

measured in arbitrary units (AU) (using NIH Image J software). Data are shown as

mean ± SEM of values of 10 measurements per mouse. Seven mice were analyzed per

group.

137

3.20: Effect of vitamin D on the expression of VDR in the lung of OVA –

sensitized and challenged mice:

The mRNA and protein expression in the lungs of all mice was analyzed by qRT-

PCR and immunoflourescence. There was significant mRNA and protein expression of

VDR in all vitamin D-PBS groups of mice, however as vitamin D-supplemented control

PBS mice and as vitamin D-sufficient control PBS mice had a higher expression of VDR

compared to moderate expression of VDR in as vitamin D-deficient control PBS mice.

Significant decrease in the expression of VDR was observed in OVA-sensitized and

challenged mice compared to PBS control mice in all groups. Interestingly, in vitamin D

deficient OVA-sensitized and challenged mice the expression of VDR was much lower

compared to vitamin D sufficient OVA-sensitized and challenged mice and vitamin D

supplemented OVA-sensitized and challenged mice. Vitamin D supplemented OVA-

sensitized and challenged mice showed a higher expression of VDR compared to

moderate expression in vitamin D sufficient OVA-sensitized and challenged mice, but

not as high as vitamin D-supplemented control PBS mice (Figure 27).

138

Figure 27: VDR expression in lung tissue- (A) Immunofluorescence showing

protein expression of VDR in lung of PBS control vitamin D-deficient, vitamin D

sufficient, and vitamin D supplemented mice compared to OVA sensitized and

challenged vitamin D-deficient, vitamin D sufficient and vitamin D supplemented mice

All images are at a magnification of 600 x. Sections were stained using rabbit anti- VDR

antibody and goat anti-rabbit cy3 as secondary antibody. DAPI was used to stain the

139

nuclei. (B) Mean fluorescent intensity (MFI) of VDR immunostaining in lung tissue

measured in arbitrary units (AU) (using NIH Image J software). Data are shown as

mean ± SEM of values of 10 measurements per mouse. (C) mRNA was isolated from

lung tissue and qPCR was performed to compare the expression of VDR among

different vitamin D groups. . Seven mice were analyzed per group *p <0.05, **p<0.01,

***p <0.001.

140

3.21: Effect of vitamin D on the expression of importin α3 in the lung of OVA

–sensitized and challenged mice

The mRNA and protein expression in the lung tissue of all groups of mice was

also analyzed by qRT-PCR and immunofluorescence. A moderate expression of importin

α3 was observed in the lung tissue of PBS control mice in all vitamin D groups. OVA-

sensitized and challenged mice exhibited a significantly increased mRNA and protein

expression of importin α3 in all vitamin D groups. However, the expression of importin

α3 was higher in vitamin D deficient OVA-sensitized and challenged mice compared to

vitamin D sufficient OVA-sensitized and challenged mice and vitamin D supplemented

OVA-sensitized and challenged mice. Additionally, vitamin D supplemented OVA-

sensitized and challenged mice demonstrated a lower expression than vitamin D

sufficient OVA-sensitized and challenged mice, but higher than control PBS mice

(Figure 28).

141

Figure 28: Importin α3 expression in lung tissue

(A) Immunofluorescence showing protein expression of importin α3 in lung of PBS

control vitamin D deficient, vitamin D sufficient and vitamin D supplemented mice

compared to OVA sensitized and challenged vitamin D-deficient, vitamin D-sufficient

and vitamin D-supplemented mice. All images are at a magnification of 600 x. Sections

were stained using goat anti- importin α3 antibody and Donkey anti-goat as secondary

antibody. DAPI was used to stain the nuclei. (B) Mean fluorescent intensity (MFI) of

142

importin α3 immunostaining in lung tissue measured in arbitrary units (AU) (using

NIH Image J software). Data are shown as mean ± SEM of values of 10 measurements

per mouse. (C) The mRNA was isolated from lung tissue and qPCR was performed to

compare the expression of importin α3 among different vitamin D groups. Data is

shown as mean ± SEM from seven individual mice in each experiment *p <0.05,

**p<0.01, ***p <0.001.

143

Section 4

Discussion

144

The immunomodulatory potential of vitamin D has been implicated in various

inflammatory diseases [293]. Vitamin D deficiency predisposes to many chronic

diseases, including type 1 diabetes, rheumatoid arthritis, crohn’s disease, infectious

diseases, and cancers [294]. Calcitriol, the biologically active form of vitamin D, exerts

its effects through the VDR [187]. Recent studies have established the role of VDR in cell

proliferation, differentiation, and immunomodulation [295].VDR is present in most cell

types of the immune system, particularly in APCs such as macrophages and dendritic

cells, as well as in both CD4+ and CD8+ T cells [293,296].

CYP24A1 and CYP27B1 are involved in the metabolism of Vitamin D and

hence these are genes of interest in the vitamin D pathway [297]. CYP27B1 encodes for

1-α-hydroxylase, which converts 25-hydroxy-vitamin D [25(OH) D] into the active

vitamin D metabolite 1, 25(OH) 2D. CYP24A1 encodes for 24-hydroxylase, the enzyme

that catalyzes the inactivation of 1,25(OH)2D [297]. VDR and its metabolizing enzymes

are expressed in HBSMCs suggesting that these cells can locally metabolize vitamin D

and functionally respond to vitamin D [297]. Treatment with calcitriol increased VDR

and CYP24A1 expression and decreased CYP27B1 expression in HBSMCs in a dose-

dependent manner. Our data is in agreement with previous studies on vitamin D in

airway smooth muscles, revealing an increased expression of VDR and CYP24A1 with

vitamin D analogues [47,88].

NF-κB is a transcriptional factor that has a central role in a variety of acute and

chronic inflammatory diseases. NF-κB is expressed in various cell types and is a master

regulator of many proinflammatory genes, leading to the synthesis of cytokines,

adhesion molecules, chemokines, growth factors and enzymes [103]. NF-κB signaling

145

has been a focus of extensive research because of its involvement in the regulation

chronic inflammatory diseases [103].

Recent studies demonstrated that the VDR is directly involved in regulating

activation of NF-κB. In dendritic cells, calcitriol targets the NF-κB pathway by inhibiting

IL-12 expression [298]. In human fibroblasts and keratinocytes, calcitriol decreases the

DNA binding capacity of NF-κB [299,300]. Vitamin D is reported to significantly

downregulate proinflammatory chemokine expression in pancreatic islet cells, resulting

in the up-regulation of IκBα transcription and arrest of NF-κB RelA nuclear

translocation [301]. In mouse embryonic fibroblasts VDR inhibits NF-κB activation

[302]. Together these studies suggest that vitamin D has an inhibitory action on NF-κB

activation.

The importin and exportin transport system present the mechanism involved

in the nucleo-cytoplasmic transport [128]. Any change in the expression of the

components involved in the nucleo-cytoplasmic transport machinery may impact gene

transcription and translation [128]. Little is currently known about the regulation of

importins and exportins in the airway cells. The nuclear translocation of NF-κB p50-

RelA is reported to be mediated by importin α3 and importin α4 [113,127,128].

In this study, we report the effect of calcitriol on the mRNA and protein

expression of importin α3 and importin α4 in HBSMCs. We determined that calcitriol

downregulates mRNA and protein expression of importin α3 with no significant effect

on importin α4. This decrease in importin α3 correlated with a calcitriol-induced

decrease in the migration of activated RelA (NF-κB p65) from the cytoplasm to the

nucleus. As NF-κB is a key regulator of inflammation and importin α3 translocates NF-

146

κB to the nucleus our findings on the effect of vitamin D to decrease both importin α3

and activation of NF-κB are significant.

Our in vitro experiments could be directly relevant to the in vivo process

occurring in asthmatic subject because in asthmatics there is an increase in the levels of

various proinflammatory cytokines, such as TNF-α, IL-1β that activates NF-κB. Studies

demonstrate that administration of TNF-α to normal subjects can lead to the

development of airway hyperresponsiveness and airway neutrophilia [47,88,188].

Based on our findings in this study calcitriol inhibits nuclear translocation of NF-

κB by downregulating importin α3. These results were confirmed by knocking down

importin α3 in HBSMCs. The decrease in NF-κB activation by calcitriol was

accompanied by a decrease in nuclear protein expression of RelA when HBSMCs were

stimulated with TNF-α. These studies demonstrate that calcitriol may be used as a

therapeutic agent in prevention of airway remodeling manifested as increases in ASM

mass as described in asthmatic subjects [188]. This further highlights the potential

significance of vitamin D in the modulation of ASM function in asthma.

A study by Theiss and colleagues [113] revealed that in colonic mucosal biopsies

from moderately-to-severely inflamed Crohn’s disease patients had increased

expression of importin α3. TNF-α is known to play a central role in the pathogenesis of

inflammatory bowel disease (IBD) and its concentration is increased in serum and stool

of IBD patients [113]. Our finding in HBSMCs are in accordance with this study

illustrating that potent inflammatory cytokines, such as TNF-α and IL-1β, increased

importin α3 expression to increase NF-κB levels in the nucleus. Further, the increased

importin α3 expression is attenuated by calcitriol. Calcitriol acts through the vitamin D

147

receptor since VDR knockdown by siRNA abolished the effect of calcitriol on importin

α3 downregulation. These data show that calcitriol prevents the TNF-α induced increase

in importin α3 and RelA through a process mediated by the VDR.

Importin α3 knockdown significantly reduced TNF-α induced nuclear

accumulation of RelA and abolished the effect of calcitriol on RelA expression. These

data confirm that importin α3 mediates the translocation of RelA to the nucleus and

that the decrease in nuclear translocation of RelA occurs via downregulation of importin

α3 by calcitriol.

IL-10, an anti-inflammatory cytokine, inhibits NF-κB thereby inhibiting the

transcription of various proinflammatory cytokines including TNF-α, IL-1 and IL-6

[85,303]. Vitamin D status is positively correlated to the level of IL-10 secreting T-

regulatory cells [304]. IL-10 in allergic inflammation is primarily released from T-

regulatory cells and these cells can decrease allergic airway inflammation and AHR

[303]. Calcitriol has been shown to promote the production of IL-10 in human B-cells

[305]. Our results demonstrate that calcitriol potentiated the anti-inflammatory effect of

IL-10 when treated concomitantly with TNF- α. The mechanisms by which IL-10 in

presence of TNF-α decreases the expression of importin α3 warrant further studies.

Our in vitro study on HBSMCs suggests that vitamin D may have a beneficial

effect on airway inflammation. But the potential limitation of these studies is that they

are in vitro in HBSMCs and requires confirmation in vivo. To verify our in vitro results,

we developed vitamin D-deficient, -sufficient, and -supplemented diets that were fed to

asthmatic mice to further examine the role of vitamin D status on airway

hyperresponsiveness and allergic airway inflammation.

148

We report data demonstrating that in OVA-sensitized and challenged mice,

vitamin D deficiency were associated with higher AHR and mice receiving vitamin D

supplemented were less responsive to methacholine. Vitamin D status had no effect on

AHR or histological changes in lungs from PBS control mice. In OVA- sensitized and

challenged mice vitamin D-deficiency was associated with an exaggerated response to

OVA-antigen and exhibited marked signs of airway remodeling compared to the mice

fed a vitamin D sufficient diet. Although higher serum vitamin D levels were associated

with lower airway inflammation and reduced hallmarks of asthma, higher serum

vitamin D levels were not associated with complete reversal of asthma. Our results are

in agreement with Sutherland and colleagues that reduced levels of vitamin D are

associated with impaired lung function and increased AHR [272].

Vitamin D deficiency was also associated with high BALF eosinophilia and mice

fed a vitamin D supplemented diet had fewer eosinophils in the BALF.

Our results are in agreement with the study by Brehm and colleagues in children that

vitamin D levels were significantly and inversely associated with total IgE and

eosinophil count [271].

One source of proinflammatory cytokines TNF-α and IL-6 is from activated

macrophages [306,307]. The primary producer of IL-17 in allergic inflammation are

macrophages rather than Th17 cells [308]. 1α,25-dihydroxyvitamin D3 is a potent

suppressor of interferon γ–mediated macrophage activation [309]. We did not observe

any difference in the macrophage number in the BALF between any of our treatment

groups however, there was a significant decrease in the levels of proinflammatory

cytokines released from activated macrophages namely TNF-α, IL-6 and IL-17 in the

sensitized mice fed a vitamin D supplemented diet. Our data supports recent studies

149

that vitamin D treatment inhibits IL-17 production [239,310]. Another study in a mouse

model of inflammatory bowel disease demonstrated decreased levels of vitamin D were

associated with elevated levels of IL-17 [258]. Vitamin D inhibits the synthesis and

release of IL-23 and IL-6, cytokines important for the development of Th17 cells [239].

These results suggest that vitamin D might macrophage activation thereby leading to

decreased cytokine release during macrophage activation. Vitamin D diminishes the

production of proinflammatory cytokines (e.g., IL-1 and TNF-α) by affecting the

interactions of T-cell/DC [236,237]. These decreased interactions results in decreased

production of IL-2 and IFN-γ and increased production of IL-10 and TGF-β [236,237].

Sensitization with OVA antigen in mice fed on a vitamin D diet had a

substantial increase in IL-5 and IL-13 in the BALF. Conversely, a vitamin D

supplementated diet resulted in decreased levels of these cytokines in the BALF, but not

to the levels found in PBS control mice. The reduction in the levels of TH2 cytokines

(IL-4, IL-5, and IL-13) has been shown to be an effective therapy to suppress airway

inflammation and hyper-responsiveness [311]. Vitamin D status of the mice was

independent of the levels of IL-4, RANTES and IP-10.

Studies show that calcitriol decreased Th1 cell proliferation and inhibits of IL-2,

IFN-γ, TNF –α production from Th1 cells [242]. The role of vitamin D in inhibiting Th1

immune responses has been already established however its effects on Th2 responses

are controversial [243]. Vitamin D induces a shift in the balance between Th1 and Th2-

type cytokines toward Th2 dominance or simply vitamin D promotes Th2 responses

[175,244]. However, study reveals reports of both the inhibition and enhancement of

Th2 responses [215]. In addition, the effect of vitamin D on IL-4 levels of IL-4 is under

debate, as some studies show an increase in the levels of IL-4 with vitamin D treatment

150

[247,248]. In human cord blood T cells, vitamin D not only decreased Th1 cytokine

production but also suppresses IL-4 and IL-4–induced expression of IL-13 [249]. In

murine model of asthma, the administration of 1,25-dihydroxyvitamin D to inhibited

airway inflammation, decreased levels of IL-4 in bronchoalveolar lavage fluid, and

decreased T-cell migration thereby inhibiting the inflammatory response [242]. We

observed no change in the levels of IL-4 following dietary supplementation with vitamin

D.

Vitamin D promotes the stimulation of CD4+CD25+ regulatory T cells by DCs

[238]. Calcitriol Foxp3 and IL-10 expression, two factors crucial for Treg induction

[239]. Our results show, OVA-sensitization and challenge were associated with reduced

IL-10 levels and a decreased number of T-regulatory cells. This effect was more

pronounced in vitamin D deficient mice. Conversely, vitamin D supplementation

significantly increased IL-10 levels and T-regulatory cell numbers.

Studies have shown that T-regulatory cells are potent immunomodulators of a

disordered immune response as seen in asthma [312] and cytokines such as IL-10 and

IL-15, are critical for stimulating their proliferation in vitro [313]. Vitamin D promotes

Treg cell induction [250,251]. Vitamin D in combination with glucocorticoids potentially

stimulates the production of IL-10 producing T-regulatory cells thereby suppressing the

immune response [252]. Vitamin D was reversed steroid resistance in patients with

asthma by inducing IL-10–secreting Treg cells [253].

In presence of vitamin D, Tregs develop and function normally, by suppressing

inappropriate Th1 and Th2 responses thereby leading to a more balanced immune

response. Low levels of vitamin D affects normal development and function of Tregs

and under certain environmental conditions, Th1 or Th2 responses may proceed

151

unabated, leading to disease [243].Our results support the above studies, demonstrating

that a decrease in Tregs was associated with increase in proinflammatory cytokines and

decreased IL-10 levels. Vitamin D deficiency resulted in a marked decrease of T-

regulatory cells and further increase in proinflammatory cytokines and decreased IL-10

levels.

NF-κB is a central transcription factor that regulates inflammatory responses in

various tissues. Activation of NF-κB induces the rapid expression of multiple genes that

play significant roles in the induction of airway hyperresponsiveness and airway

inflammation in asthma [107,108]. Potent inducers of NF-κB activation are TNF-α and

IL-1β and upon activation, NF-κB controls the expression of these cytokines which are

essential mediators of chronic inflammation [314]. NF-κB also induces the expression of

proinflammatory cytokines such as IL-12[102] and IFN-γ [315] released from Th1 cells;

IL-4 [103], IL-5 [103], IL-8 [102] and IL-13 [103,314] released from Th2 cells; TNF-α

and IL-6 [316] released from macrophages [316]; and IL- 17 released from macrophages

and Th17 cells [308].These data suggest that one potential treatment of chronic

inflammatory diseases such as asthma is to administer drugs that decrease NF-κB

activation or nuclear translocation. Upon stimulation, the nuclear translocation of NF-

κB p50-RelA subunits is mediated by importin α3 and α4 [113,127,128]. Our studies in

HBSMCs demonstrate a decreased expression of importin α3 upon treatment with

calcitriol, which leads to a decreased nuclear expression of NF-κB resulting in decreased

secretion of proinflammatory cytokines. Based on these results from our in vitro

studies, we analyzed of importin α3 expression in the lungs of OVA- sensitized and

challenged mice. Our results are in agreement with studies in colonic mucosal biopsies

from Crohn’s disease patients [113] that increased expression of importin α3 was

152

observed in OVA-sensitized and challenged mice. Our results are consistent with our in

vitro studies in HBSMCs, that treatment with TNF-α increased the expression of

importin α3 confirming that inflammation results in increased importin α3 expression.

However, vitamin D deficiency was associated with higher expression of importin α3

and the vitamin D supplemented mice had a significantly reduced expression of

importin α3. These data suggest that vitamin D deficiency is associated with marked

inflammatory changes. To confirm our results we evaluated the expression of TNF-α in

all groups of vitamin D mice.

TNF-α is a pro-inflammatory cytokine that activates many potential genes

responsible for inflammation, production of other inflammatory cytokines, and

increases the severity of asthma [77]. In our study, vitamin D deficiency increases the

TNF-α expression in lungs of mice model of allergic airway inflammation. Our results

support the findings of Sutherland and colleagues that low vitamin D levels are

associated with increased levels of TNF-α in asthmatic human subjects [272]. Asthmatic

mice with diets supplemented with vitamin D had low expression of TNF-α, suggesting

an anti-inflammatory role of vitamin D. Our data strongly suggests that vitamin D

deficiency is responsible for increased inflammation in asthma.

The expression of VDR is considerably decreased in the colonic mucosa of patients

with inflammatory bowel disease compared to normal patients [317]. Dysfunction of

VDR expression and a vitamin D3 deficiency can cause poor bone development and

overall health, including increasing the risk of many chronic inflammatory diseases and

cancers .We also analyzed the effect of vitamin D deficiency on the expression of VDR in

lungs of mouse model of allergic airway inflammation. In accordance with a previous

study [317], we demonstrated that under inflammatory conditions there is decreased

153

expression of the VDR in OVA-sensitized and challenged mice. Reduced serum levels of

vitamin D were associated with a further decrease in VDR expression possibility due to

increased expression of TNF-α.

In the present study, the effect of vitamin D on allergic airway inflammation might

be due to two underlying mechanisms: 1) decreased activation of NF-κB as a result of

decreased expression of importin α3, leading to a decrease in NF-κB inducible gene

expression; 2) Increased numbers of T-regulatory cells and production of IL-10

responsible for the immunosuppressive action of vitamin D. These two mechanisms

may be acting in concert to reduce the allergic airway response in asthma.

Conclusion:

Our data suggests that an inflammatory environment increases the TNF-α

expression and is associated with increased expression of importin α3 and decreased

expression of VDR. Vitamin D deficiency causes further increase in TNF-α, leading to

increased inflammation, and further increasing the expression of importin α3 while

decreasing the expression of VDR. Dietary vitamin D supplementation was associated

with reduced TNF-α and importin α3 expression thus increasing VDR expression.

Higher serum levels of vitamin D were associated with increased numbers of T-

regulatory cells which stimulate secretion of the anti-inflammatory cytokine IL-10,

resulting in a significant reduction in the levels of many pro-inflammatory cytokines,

which play a major role in the pathogenesis of asthma.

We report data demonstrating that in OVA-sensitized mice there is a significant

and deleterious association between reduced serum vitamin D levels, AHR, and airway

154

inflammation, which together constitute important biomarkers of asthma severity,

respiratory impairment, and disease prognosis. Our study suggests a therapeutic role for

vitamin D in allergic asthma. Our data show that vitamin D supplementation alleviates

allergic airway inflammation but does not completely reverse this effect. Our study

demonstrates that, vitamin D can be used as an adjunct to the treatment of asthma.

Thus, data from our study support the continued exploration of vitamin D

supplementation and new approaches in prevention and treatment of asthma. Thus,

vitamin D could be a novel immunomodulator in allergic airway inflammation in

asthma

Outstanding Questions and Future Directions

The role of endogenous factors other than vitamin D that regulate the action of importin

α3 needs to be elucidated. In vivo experiments using importin α3 knockout mice to

examine AHR and airway inflammation are needed to confirm in vitro data. Although

the current study reports the altered levels of IL-17 in the BALF, the role of vitamin D on

Th17 cells needs to be elucidated.

155

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PEER-REVIEWED PUBLICATIONS

1) Halvor S McGee, Arthur L Stallworth, Tanupriya Agrawal, Zhifei Shao, Lindsey

Lorence, and Devendra K. Agrawal Fms-like tyrosine kinase 3 ligand decreases t helper

type 17 cells and suppressors of cytokine signaling proteins in the lung of house dust

mite-sensitized and -challenged mice. Am J Respir Cell Mol Biol; 2010 ;43(5):520-

529.

2) Tanupriya Agrawal, Gaurav K Gupta, Devendra K Agrawal Calcitriol Decreases

NF-κB Nuclear Translocation via Decreasing Expression of importin α3 in Human

Bronchial Smooth Muscle Cells (Journal of Clinical Immunology April 2012)

(PMID:22526597)

3) Tanupriya Agrawal, Gaurav K Gupta, and Devendra K Agrawal: Vitamin D

deficiency decreases the expression of VDR and prohibitin in the lungs of OVA-

sensitized and challenged mice. Experimental and Molecular Pathology; 2012;

Apr 16;93(1):74-81

4) Gaurav K Gupta, Tanupriya Agrawal, Michael G. DelCore, Devendra K Agrawal

Vitamin D deficiency induces cardiac hypertrophy and inflammation in epicardial

adipose tissue in hypercholesterolemic swine. (Experimental and Molecular

Pathology; 2012 ; Apr 17;93(1):82-90

5) Gaurav K Gupta, Tanupriya Agrawal, Michael G. DelCore, Devendra K Agrawal.

Decreased Expression of Vitamin D Receptors in Smooth Muscle Cells of Porcine

Coronary Arteries following Angioplasty: Potential Implication in Coronary Intimal

Hyperplasia. PLoS One 2012;7:e42789

6) Tanupriya Agrawal, Gaurav K Gupta, Toluwalope O. Makinde and Devendra K

Agrawal : Vitamin D Supplementation Reduces Airway Hyperresponsiveness and Lung

Pathology in a Mouse Model of Allergic Asthma( Submitted)

ABSTRACTS PUBLISHED

1) Agrawal T, McGee HS, Stallworth AS, Shao Z, and Agrawal DK: Th17 Cells and

SOCS Proteins in House-Dust Mite-Induced Allergic Airway Inflammation in a Murine

Model Journal of Allergy and Clinical Immunology Vol. 125, Issue 2,

Supplement 1, Page AB41 February 2010.Annual meeting of American Academy of

Allergy, Asthma and Immunology, New Orleans, LA.

2) Agrawal T, and Agrawal DK Calcitriol Decreases the Expression of Importin Alpha3

in Human Bronchial Smooth Muscle Cells Journal of Bone and Mineral

Research 25 (suppl1) MO 175, Annual meeting of American Society of Bone and

Mineral Research, Toronto, Canada.2010.

3) Agrawal T, and Agrawal DK: Calcitriol increases the Expression of Importin7

(IPO7) and Importin13 (IPO13) in Human Bronchial Smooth Muscle Cells -Selected in

featured poster session. Journal of Allergy and Clinical Immunology Vol.

127, Issue 2, Supplement, Page AB144 February 2011 Annual meeting of American

Academy of Allergy, Asthma and Immunology, San Francisco, CA

4) Agrawal T, Gupta GK, and Agrawal DK. Effect Of Cytokines On Expression Of

Importin Alpha3 (kpna4) In Human Bronchial Smooth Muscle Cells. American

Journal of Respiratory and Critical Care Medicine. 2011 May; 183(1): A2580.

Selected in special poster discussion session.

5) Agrawal T, Gupta GK, and Agrawal DK. Effect Of TNF-Alpha On Vitamin D

Receptor And Its Metabolizing Enzymes In Human Bronchial Smooth Muscle Cells:

Potential Implication In Allergic Inflammation In Asthma. American Journal of

Respiratory and Critical Care Medicine. 2011 May; 183(1): A2578. . Selected in

special poster discussion session.

5) Agrawal T, Gupta GK, and Agrawal DK : A Cross talk between Prohibitin and

Importin α3 in Vitamin D signaling pathway- A novel implication in allergic asthma,

Accepted in Annual meeting of American Society of Bone and Mineral

Research 2011, San Diego, CA

6) Agrawal T, Gupta GK, Kim MJ, and Devendra K. Agrawal- Increased Expression of

Importin α3 (KPNA4) and Decreased VDR in the Lung of OVA-Sensitized and

Challenged Mice. Accepted in Annual Meeting of American Academy of Allergy

Asthma and Immunology (AAAAI) 2012 Orlando, FL