MOLECULAR PATHOGENESIS OF NON- EOSINOPHILIC ASTHMA
Katherine Joanne Baines
B.BiomedSci(Hons)
A Thesis Submitted for the Degree of Doctor of Philosophy October 2007 Faculty of Health School of Biomedical Sciences University of Newcastle
STATEMENT OF ORIGINALITY
This work contains no material which has been accepted for the award of any other degree or diploma in any university or other tertiary institution and, to the best of my knowledge and belief, contains no material previously published or written by another person, except where due reference has been made in the text. I give consent to this copy of my thesis, when deposited in the University Library, being made available for loan and photocopying subject to the provisions of the Copyright Act 1968.
ACKNOWLEDGEMENT OF AUTHORSHIP/COLLABORATION
I hereby certify that the work embodied in this Thesis is the result of original research, the greater part of which was completed subsequent to admission to candidature for the degree (except in cases where the Committee has granted approval for credit to be granted from previous candidature at another institution).
Signature:……………………………….. Date:…………………………………..
ii Acknowledgements
When I first began thinking about writing this section of my thesis it dawned on me how lucky I am to have so many people I need to thank that have played an important role in both my PhD studies and my life. Doing a PhD has been a great challenge in which I have learnt so much and grown so much as a person and a researcher.
First and foremost I would like to acknowledge the The Asthma Foundation of NSW and the Asthma CRC for providing funding for my PhD scholarship. I would like to thank my supervisors Prof Peter Gibson and Prof Rodney Scott for your ongoing support and encouragement throughout this process. I have a lot of respect for you both and thankyou for your sound advice and ideas, and for being patient with me, but also for pushing me when I needed it. Thanks Rodney for your guidance, confidence in me, and recommending me to the Respiratory group in the beginning, that change was the best thing I could have done. Peter, thankyou for all the opportunities you have offered and continue to offer me, I really admire your wealth of knowledge and ability to see ‘the big picture’.
A big thankyou to all my colleagues and friends in the Respiratory Medicine and Childhood Cancer research groups at the Hunter Medical Research Institute. Thanks to Naomi Fibbens, Rebecca Oldham, Terry Grissell and Joanna Mimica for your lab work and skill, for teaching me methods, answering my many questions and also for a bit of a chat and laugh now and then. It’s loads of fun working with you guys, and you have become great friends. Thanks to Noreen Bell for your tireless work seeing the patients and collecting the samples for this study. Thanks to Deborah Hall for always getting me organised with everything I can think of, and for editing this thesis. Thanks to Glenda Walker for your help with the zymography. Thanks to Dr Nikola Bowden for your help with the microarrays. Thanks to Dr Vanessa Murphy for your help with formatting this thesis, and to Dr Lisa Wood for reviewing sections of this thesis.
To Dr Jodie Simpson (Country Star), it’s great working with you, thanks for all your guidance and encouragement. Thanks for bringing me out of my ‘shell’ and for being such a great friend and workmate. To my office buddies Nicole Ryan and Heather Powell, thanks for always being ready for a coffee and for being so kind and supportive.
iii
To my dear friends Allison Thomas and Carolyn Brooks, thanks for always being there for me over the years; you are the best friends a girl could ask for. To the Uni girls (you know who you are), thanks for your friendship, for all the trips to Gloria’s, and for understanding the science talk and the PhD pressures. Thanks to my housemates Andrew and Bronwyn Rundle and Katie Brooker for putting up with me being stressed and having my papers everywhere, and for being so fun to live with.
To Shane Nolland, thankyou for being my ‘rock’, and my best friend, I am very lucky to have you in my life. I don’t know if I would have got this far if it wasn’t for your love, patience and support. Thanks to my sister Penny Baines and to Ryan Chan, we really enjoy spending time with you guys. Pen, you are a cool little sister, thanks for never being afraid to have a go at anything, you always inspire me. To my Grandma, Audrey Johnson, thanks for always giving me perspective, you may not realise it, but you always remind me what is important in life. Finally I want to thank my parents, Lesley and Lee Baines, for your unconditional love and support in so many ways. Thankyou for always having faith in me and for encouraging me to go after whatever I want in life. Dad you can relax now, I have finally finished!
iv Publications
Simpson, J.L., Baines, K.J., Gibson, P.G. Chapter 17: The Biology of Neutrophils. Middleton’s Allergy 7th Edition. In Press August 2007.
Abstracts
Baines, K.J., Bell, N.V. Simpson, J.L., Scott, R.J., Boyle, M.J. Gibson, P.G. 2005. Enhanced IL-8 Release from Neutrophils in Non-Eosinophilic Asthma. Inflammation Research. 54: S137. (Poster)
Baines, K.J., Simpson, J.L., Scott, R.J., Bell, N.V., Boyle, M.J. Gibson, P.G. 2006. Enhanced IL-8 Release from Neutrophils in Non-Eosinophilic Asthma. Respirology. 11(2): A29. (Poster)
Baines, K.J., Bowden, N.A., Scott, R.J., Simpson, J.L., Gibson, P.G. 2007. Molecular Analysis of Neutrophils in Asthma Subtypes. Respirology. 12(1): A20. (Oral)
Baines, K.J., Bowden, N.A., Scott, R.J., Simpson, J.L., Gibson, P.G. 2007. Molecular Analysis of Neutrophils in Asthma Subtypes. American Journal of Respiratory and Critical Care Medicine. 175: A683. (Poster)
Gibson, P.G., Baines, K.J., Simpson, J.L., Scott R.J. 2007. Gene Expression Profiling Identifies Neutrophilic and Eosinophilic Asthma as Distinct Subtypes. European Respiratory Journal. 30(51): A3169. (Oral)
v TABLE OF CONTENTS
Table of Tables………………………………………………………….…………….xi Table of Figures……………………………………………………………………....xv Abbreviations……………………………………………………………………...... xvii Abstract……...………………………………………………………………..………..1
CHAPTER 1: GENERAL INTRODUCTION ...... 3 1.1 ASTHMA...... 3 1.1.1 Allergy...... 4 1.2 INFLAMMATORY SUBTYPES OF ASTHMA...... 4 1.2.1 Eosinophilic Asthma ...... 5 1.2.1.1 Adaptive Immune Response ...... 5 1.2.1.2 Eosinophils...... 7 1.2.2 Non-Eosinophilic Asthma ...... 8 1.3 NEUTROPHILS IN ASTHMA...... 12 1.4 NEUTROPHILS IN CHRONIC OBSTRUCTIVE PULMONARY DISEASE (COPD)...... 13 1.5 BIOLOGY OF NEUTROPHILS ...... 14 1.5.1 Neutrophil Migration ...... 15 1.5.1.1 Myeloid Development...... 16 1.5.1.2 Neutrophil Trafficking and Margination...... 18 1.5.1.3 Cellular Adhesion Molecules...... 18 1.5.1.4 Integrins ...... 19 1.5.1.5 Inflammatory Stimulus ...... 20 1.5.1.6 Endothelial Cell Interactions...... 20 1.5.1.7 Epithelial Cell Interactions...... 20 1.5.1.8 Chemotactic Mediators...... 22 1.5.1.8.1 Interleukin-8...... 23 1.5.2 Neutrophil Phagocytosis ...... 24 1.5.3 Innate Immune Activation ...... 24 1.5.3.1 Toll Like Receptors (TLRs)...... 25 1.5.3.2 Nucleotide-Binding Oligomerisation Domain (NOD) Molecules ...... 27 1.5.4 Neutrophil Granules ...... 27 1.5.4.1 Mechanisms of Degranulation ...... 29 1.5.4.2 Proteolytic Enzymes ...... 29 1.5.5 Respiratory Burst ...... 31 1.5.5.1 Reactive Oxygen Species...... 32 1.5.6 Neutrophil Clearance and Death...... 33 1.5.7 Cytokine Synthesis...... 34 1.5.8 Neutrophil Priming and Activation...... 36 1.6 NEUTROPHIL GENE EXPRESSION ...... 37 1.7 CONTRIBUTION OF NEUTROPHILS TO THE ADAPTIVE IMMUNE RESPONSE ...... 39 1.8 NEUTROPHIL FUNCTION IN AGEING ...... 39 1.9 NEUTROPHILIC ASTHMA ...... 41 1.9.1 Mechanisms of Neutrophilic Asthma ...... 41 1.9.2 Triggers of Neutrophilic Airway Inflammation ...... 44 1.9.2.1 Endotoxin ...... 44
vi 1.9.2.2 Respiratory Viruses...... 45 1.9.2.3 Air Pollution...... 45 1.10 NEUTROPHILS AND CORTICOSTEROIDS ...... 46 1.11 AIMS & HYPOTHESES...... 47 1.12 SPECIFIC HYPOTHESES ...... 47 CHAPTER 2: MATERIALS AND METHODS ...... 48 2.1 CLINICAL INFORMATION ...... 48 2.1.1 Collection of Clinical Information...... 48 2.1.2 Spirometry...... 48 2.1.3 Saline Challenge and Sputum Induction...... 49 2.1.4 Allergy Skin Prick Testing...... 50 2.1.5 Ethics...... 51 2.2 CELL ISOLATION ...... 51 2.2.1 Isolation of Sputum Neutrophils ...... 51 2.2.1.1 Induced Sputum Processing ...... 51 2.2.1.2 Sputum Differential Cell Count ...... 51 2.2.2 Sputum Cell Isolation via Magnetic Cell Separation...... 52 2.2.3 Isolation of Peripheral Blood Granulocytes...... 52 2.2.3.1 Percoll Density Gradient...... 52 2.2.3.2 Magnetic Cell Separation...... 53 2.2.3.3 Cell Culture and Stimulation...... 53 2.3 ASSAYS ...... 54 2.3.1 Interleukin-8 (IL-8) ...... 54 2.3.2 Interleukin-1β (IL-1β) ...... 54 2.3.3 Tumor Necrosis Factor-α (TNF-α) ...... 54 2.3.4 Oncostatin M (OSM)...... 54 2.3.5 Neutrophil Elastase...... 55 2.3.6 Matrix Metalloproteinase-9 (MMP-9) ...... 55 2.3.7 Zymography of MMP-9 Activity...... 56 2.4 MOLECULAR METHODS ...... 56 2.4.1 RNA Extraction ...... 56 2.4.2 RNA Quantitation for Real-Time PCR...... 57 2.4.3 Reverse Transcription for Real-Time PCR ...... 57 2.4.3.1 Primers and Probe Sequences ...... 58 2.4.4 Real-Time PCR ...... 59 2.4.4.1 Analysis of Relative Real-Time PCR results...... 59 2.4.4.2 PCR Controls...... 59 2.4.4.2.1 Endogenous Control...... 59 2.4.4.2.2 Specific Target Positive Controls (Calibrators) ...... 60 2.4.4.2.2.1 IL-8 mRNA ...... 60 2.4.4.2.2.2 IL-1β and TNF-α mRNA ...... 60 2.4.4.2.2.3 TLR2, TLR4, and OSM mRNA...... 60 2.4.5 Gene Expression Studies with Illumina BeadArrays ...... 60 2.4.5.1 RNA Quantitation for BeadArrays ...... 60 2.4.5.2 RNA Amplification for BeadArrays...... 61 2.4.5.2.1 Reverse Transcription to Synthesise First Strand cDNA...... 61 2.4.5.2.2 Second Strand cDNA Synthesis...... 61 2.4.5.2.3 cDNA Purification...... 62 2.4.5.2.4 In Vitro Transcription (IVT) ...... 62
vii 2.4.5.2.5 cRNA Purification...... 62 2.4.5.3 Illumina BeadChip Protocol...... 62 2.4.5.3.1 Hybridisation...... 63 2.4.5.3.2 Washing ...... 63 2.4.5.3.3 Scanning...... 63 2.5 DATA ANALYSIS ...... 63 2.5.1 Microarray Data Analysis...... 64 2.5.1.1 Resting versus Stimulated Neutrophils ...... 64 2.5.1.2 Resting Neutrophils in Asthma Subtypes ...... 64 2.5.1.3 LPS Stimulated Neutrophils Asthma Subtypes ...... 65 CHAPTER 3: ACTIVATION OF BLOOD GRANULOCYTES ...... 66 3.1 INTRODUCTION...... 66 3.2 METHODS...... 68 3.3 RESULTS ...... 68 3.3.2 Protease Release ...... 69 3.3.3 Chemokine and Cytokine Release ...... 71 3.4 DISCUSSION ...... 74 CHAPTER 4: DIFFERENCES BETWEEN AIRWAY AND SYSTEMIC INNATE IMMUNE FUNCTION...... 79 4.1 INTRODUCTION...... 79 4.2 METHODS...... 81 4.3 RESULTS ...... 82 4.3.1 Magnetic Cell Separation of Neutrophils ...... 82 4.3.2 Effects of LPS Stimulation on Peripheral Blood Neutrophils...... 86 4.3.3 Effects of LPS Stimulation on Sputum Neutrophils...... 90 4.3.4 Comparison of Sputum to Blood Neutrophils ...... 91 4.3.5 Effects of DTT Treatment of Peripheral Blood Neutrophils ...... 94 4.4 DISCUSSION ...... 95 CHAPTER 5: INNATE IMMUNE RESPONSES OF NEUTROPHILS IN AGEING ...... 100 5.1 INTRODUCTION...... 100 5.2 METHODS...... 101 5.3 RESULTS ...... 102 5.3.1 Clinical Features...... 102 5.3.2 Inflammatory Cells...... 103 5.3.3 Sputum Supernatant IL-8 ...... 103 5.3.4 Chemokine and Cytokine Production from Sputum Neutrophils ...... 104 5.3.5 Chemokine and Cytokine Production from Peripheral Blood Neutrophils ...... 105 5.3.6 Total MMP-9 Release from Peripheral Blood Neutrophils ...... 107 5.3.7 Neutrophil TLR Expression...... 108 5.4 DISCUSSION ...... 110 CHAPTER 6: INNATE IMMUNE RESPONSES OF NEUTROPHILS IN AIRWAY DISEASE ...... 114 6.1 INTRODUCTION...... 114 6.2 METHODS...... 115 6.3 RESULTS ...... 116
viii 6.3.1 Comparison of Asthma and Healthy Controls ...... 116 6.3.1.1 Clinical Features...... 116 6.3.1.2 Inflammatory Cells...... 117 6.3.1.3 Sputum Supernatant IL-8 ...... 118 6.3.1.4 Chemokine and Cytokine Production from Sputum Neutrophils .....119 6.3.1.5 Chemokine and Cytokine Production from Peripheral Blood Neutrophils...... 120 6.3.1.6 Total MMP-9 Release from Peripheral Blood Neutrophils ...... 122 6.3.1.7 Neutrophil TLR Expression...... 122 6.3.2 Comparison of COPD and Healthy Controls ...... 125 6.3.2.1 Clinical Features...... 125 6.3.2.2 Inflammatory Cells...... 125 6.3.2.3 Sputum Supernatant IL-8 ...... 126 6.3.2.4 Chemokine and Cytokine Production from Sputum Neutrophils .....127 6.3.2.5 Chemokine and Cytokine Production from Peripheral Blood Neutrophils...... 127 6.3.2.6 Total MMP-9 Release from Peripheral Blood Neutrophils ...... 130 6.3.2.7 Neutrophil TLR Expression...... 130 6.3.3 Comparison of Asthma and COPD...... 133 6.3.3.1 Clinical Features...... 133 6.3.3.2 Inflammatory Cells...... 133 6.3.3.3 Sputum supernatant IL-8...... 134 6.3.3.4 Chemokine and Cytokine Production from Sputum Neutrophils .....134 6.3.3.5 Chemokine and Cytokine Production from Peripheral Blood Neutrophils...... 135 6.3.3.6 Total MMP-9 Release from Peripheral Blood Neutrophils ...... 136 6.3.3.7 Neutrophil TLR Expression...... 137 6.4 DISCUSSION ...... 139 CHAPTER 7: INNATE IMMUNE RESPONSES OF NEUTROPHILS IN ASTHMA SUBTYPES ...... 143 7.1 INTRODUCTION...... 143 7.2 METHODS...... 144 7.2.1 Asthma Subtype Classification...... 145 7.3 RESULTS ...... 145 7.3.1 Comparison of Eosinophilic and Non-Eosinophilic Asthma ...... 145 7.3.1.1 Clinical Features...... 145 7.3.1.2 Inflammatory Cells...... 146 7.3.1.3 Sputum Supernatant IL-8 ...... 149 7.3.1.4 Chemokine and Cytokine Production from Sputum Neutrophils .....149 7.3.1.5 Chemokine and Cytokine Production from Blood Neutrophils...... 151 7.3.1.6 MMP-9 Release from Peripheral Blood Neutrophils...... 153 7.3.1.7 Neutrophil TLR Expression...... 154 7.3.2 Comparison of Eosinophilic, Neutrophilic and Paucigranulocytic Asthma...... 157 7.3.2.1 Clinical Features...... 157 7.3.2.2 Inflammatory Cells...... 157 7.3.2.3 Sputum Supernatant IL-8 ...... 160 7.3.2.4 Chemokine and Cytokine Production from Sputum Neutrophils .....160 7.3.2.5 MMP-9 Release from Sputum Neutrophils...... 163
ix 7.3.2.6 Chemokine and Cytokine Production from Blood Neutrophils...... 163 7.3.2.7 MMP-9 release from blood neutrophils ...... 168 7.3.2.8 Neutrophil TLR Expression...... 169 7.3.3 Associations ...... 171 7.4 DISCUSSION ...... 172 CHAPTER 8: MOLECULAR ANALYSIS OF NEUTROPHILS IN ASTHMA SUBTYPES ...... 179 8.1 INTRODUCTION...... 179 8.2 METHODS...... 180 8.2.1 Data Analysis ...... 180 8.3 RESULTS ...... 181 8.3.1 Clinical Features...... 181 8.3.2 Induced Sputum Inflammatory Cells...... 181 8.3.3 Altered Neutrophil Gene Expression with LPS Stimulation ...... 182 8.3.4 Altered Gene Expression Profiles in Asthma Subtypes...... 185 8.3.4.1 Resting Blood Neutrophils...... 185 8.3.4.2 LPS Stimulated Blood Neutrophils...... 193 8.4 DISCUSSION ...... 196 CHAPTER 9: GENERAL DISCUSSION...... 201 9.2 RELATIVE CYTOKINE PRODUCTION FROM ISOLATED NEUTROPHILS ...... 203 9.3 DISTINCT DIFFERENCES BETWEEN AIRWAY AND BLOOD NEUTROPHILS ...... 204 9.4 ENDOTOXIN TOLERANCE ...... 205 9.5 IMPACT OF AGEING ...... 206 9.6 INNATE IMMUNE DYSFUNCTION IN AIRWAY DISEASE...... 207 9.7 ACCUMULATION OF NEUTROPHILS IN THE AIRWAYS IN NEUTROPHILIC ASTHMA ...... 208 9.7.1 Increased Migration...... 208 9.7.2 Delayed Apoptosis...... 210 9.7.3 Impaired Efferocytosis ...... 211 9.8 COMPLICATIONS OF REPORTING NEUTROPHIL DATA: VOLUME OR CELL NUMBER?...... 213 9.9 CONCLUSIONS ...... 214 CHAPTER 10: REFERENCES...... 215 APPENDIX A…………………………………………………………...……………257
x TABLE OF TABLES
Table 2.1 Trade Names of Medication Withheld for 4.5% Saline Challenge...... 50 Table 3.1 Cell Viability of Granulocytes at Specified Time Points...... 69 Table 3.2 Levels of TLR4 and IL-1β mRNA at 2 hours...... 74 Table 4.1 Proportions of neutrophils, eosinophils and other (lymphocytes or monocytes) of the CD16- and CD16+ blood cell fractions created using MACS...... 83 Table 4.2 Proportions of neutrophils, macrophages and eosinophils in the CD16- and CD16+ sputum cell fractions created using MACS...... 85 Table 4.3 Cell viability before and after MACS...... 86 Table 4.4 Levels of chemokine and cytokine mRNA expression in isolated blood neutrophils (1 x 106 cells/mL) at rest and with LPS stimulation...... 87 Table 4.5 Total and active MMP-9 (pg/mL) release from isolated peripheral blood neutrophils...... 88 Table 4.6 Levels of chemokine and cytokine mRNA expression in sputum neutrophils (1 x 105 cells/mL) at rest and with LPS stimulation...... 90 Table 4.7 Cell viability of blood compared to sputum samples before and after MACS ...... 91 Table 4.8 Levels of chemokine and cytokine mRNA expression in resting sputum and blood neutrophils (1 x 105 cell/mL) ...... 92 Table 4.9 Levels of chemokine and cytokine mRNA expression in LPS stimulated sputum and blood neutrophils (1 x 105 cell/mL)...... 93 Table 5.1 Clinical characteristics of healthy control participants under and over 55 years of age...... 102 Table 5.2 Induced sputum inflammatory cell counts healthy control participants under and over 55 years of age...... 103 Table 5.3 Levels of chemokine and cytokine mRNA expression in resting sputum neutrophils from healthy control participants under (n=5) and over (n=4) 55 years of age...... 105 Table 5.4 Levels of chemokine and cytokine mRNA expression in resting blood neutrophils isolated from participants under (n=5) and over (n=8) years of age..106 Table 5.5 Levels of chemokine and cytokine mRNA expression in LPS stimulated blood neutrophils isolated from participants under (n=5) and over (n=8) years of age..107 Table 5.6 Total MMP-9 release from blood neutrophils (106 cells/mL) isolated from participants under (n=5) and over (n=8) years of age...... 108 Table 5.7 Relative mRNA levels of TLR4 in sputum and blood neutrophils isolated from participants under (n=5) and over (n=8) years of age ...... 108 Table 5.8 Relative mRNA levels of TLR2 in sputum and blood neutrophils isolated from participants under (n=5) and over (n=8) years of age ...... 109 Table 5.9 Summary of results for the comparison between older versus younger healthy control participants...... 109 Table 6.1 Clinical characteristics of participants with asthma and healthy control subjects...... 117 Table 6.2 Induced sputum inflammatory cell counts from participants with asthma compared to healthy controls ...... 118 Table 6.3 Levels of chemokine and cytokine mRNA expression in resting sputum neutrophils from participants with asthma compared to healthy controls ...... 120 Table 6.4 Levels of chemokine and cytokine mRNA expression in resting neutrophils from subjects with asthma and healthy controls ...... 120
xi Table 6.5 Levels of chemokine and cytokine mRNA expression in LPS stimulated neutrophils from participants with asthma compared with healthy controls ...... 121 Table 6.6 Total MMP-9 release from blood neutrophils (106 cells/mL) isolated from participants with asthma compared with healthy controls ...... 122 Table 6.7 Active MMP-9 release from blood neutrophils (106 cells/mL) isolated from participants with asthma compared with healthy controls ...... 122 Table 6.8 Relative mRNA levels of TLR4 in sputum and blood neutrophils from participants with asthma compared with healthy controls ...... 123 Table 6.9 Relative mRNA levels of TLR2 in sputum and blood neutrophils from participants with asthma compared with healthy controls ...... 123 Table 6.10 Summary of results for the comparison between participants with asthma and age matched healthy controls ...... 124 Table 6.11 Clinical characteristics of participants with COPD compared to healthy controls...... 125 Table 6.12 Induced sputum inflammatory cell counts for participants with COPD compared with healthy controls ...... 126 Table 6.13 Levels of chemokine and cytokine protein release from sputum neutrophils isolated from participants with COPD compared to healthy controls...... 127 Table 6.14 Levels of chemokine and cytokine mRNA expression in sputum neutrophils isolated from participants with COPD compared to healthy controls...... 127 Table 6.15 Levels of chemokine and cytokine mRNA in resting blood neutrophils isolated from participants with COPD and healthy controls...... 128 Table 6.16 Levels of chemokine and cytokine mRNA expression in LPS stimulated neutrophils from participants with COPD compared with healthy controls...... 129 Table 6.17 Active MMP-9 release from blood neutrophils (106 cells/mL) isolated from participants with COPD compared with healthy controls...... 130 Table 6.18 Relative mRNA levels of TLR4 in sputum and blood neutrophils from participants with COPD compared with healthy controls...... 131 Table 6.19 Relative mRNA levels of TLR2 in sputum and blood neutrophils from participants with COPD compared with healthy controls...... 131 Table 6.20 Summary of results for the comparison between participants with COPD and age matched healthy controls...... 132 Table 6.21 Clinical characteristics of participants with asthma compared to those with COPD ...... 133 Table 6.22 Induced sputum inflammatory cell counts for participants with asthma compared with COPD ...... 134 Table 6.23 Chemokine and cytokine production of sputum neutrophils isolated from participants with asthma compared with COPD ...... 135 Table 6.24 Levels of chemokine and cytokine mRNA expression in resting blood neutrophils isolated from participants with asthma compared with COPD...... 135 Table 6.25 Levels of chemokine and cytokine mRNA expression in LPS stimulated neutrophils isolated from participants with asthma or COPD ...... 136 Table 6.26 Total MMP-9 release from blood neutrophils (106cells/mL) isolated from participants with asthma compared with COPD ...... 137 Table 6.27 Relative mRNA levels of TLR4 in sputum and blood neutrophils from participants with asthma and COPD ...... 137 Table 6.28 Relative mRNA levels of TLR2 in sputum and blood neutrophils from participants with asthma and COPD ...... 137 Table 6.29 Summary of results for the comparison between participants with asthma and COPD ...... 138
xii Table 7.1 Clinical characteristics of eosinophilic asthma, non-eosinophilic asthma and healthy controls ...... 147 Table 7.2 Inflammatory cell counts for subjects with eosinophilic asthma, non- eosinophilic asthma and healthy controls ...... 148 Table 7.3 Levels of chemokine and cytokine mRNA expression in resting sputum neutrophils from participants with eosinophilic asthma, and non-eosinophilic asthma compared to healthy controls...... 150 Table 7.4 Levels of chemokine and cytokine mRNA expression in resting blood neutrophils from participants with eosinophilic asthma and non-eosinophilic asthma compared to healthy controls...... 152 Table 7.5 Relative messenger RNA levels of IL-8, IL-1β and TNF-a of LPS stimulated blood neutrophils from participants with asthma compared to healthy controls ..153 Table 7.6 Total MMP-9 release from blood neutrophils (106 cells/mL) isolated from participants with non-eosinophilic asthma, eosinophilic asthma and healthy controls...... 153 Table 7.7 Active MMP-9 release from blood neutrophils (106 cells/mL) isolated from participants with non-eosinophilic asthma, eosinophilic asthma and healthy controls...... 154 Table 7.8 Relative mRNA levels of TLR4 in sputum and blood neutrophils from subjects with non-eosinophilic asthma, eosinophilic asthma and healthy controls ...... 155 Table 7.9 Relative mRNA levels of TLR2 in sputum and blood neutrophils from subjects with non-eosinophilic asthma, eosinophilic asthma and healthy controls ...... 155 Table 7.10 Summary of results for the comparison between participants with non- eosinophilic asthma, eosinophilic asthma and healthy controls...... 156 Table 7.11 Clinical characteristics of neutrophilic asthma, eosinophilic asthma, paucigranulocytic asthma and healthy controls ...... 158 Table 7.12 Table 12: Inflammatory cell counts for subjects with eosinophilic asthma, non-eosinophilic asthma and healthy controls. ‡ p<0.004 versus healthy controls159 Table 7.13 Relative messenger RNA levels of IL-8, IL-1β and TNF-α of resting sputum neutrophils in asthma subtypes and healthy controls...... 162 Table 7.14 Levels of chemokine and cytokine mRNA expression in resting blood neutrophils in asthma subtypes and healthy controls...... 165 Table 7.15 Levels of chemokine and cytokine mRNA expression in LPS stimulated blood neutrophils in asthma subtypes and healthy controls...... 167 Table 7.16 Total MMP-9 released from blood neutrophils (106 cells/mL) in asthma subtypes and healthy controls ...... 168 Table 7.17 Active MMP-9 released from blood neutrophils (106 cells/mL) in asthma subtypes and healthy controls ...... 168 Table 7.18 Relative mRNA levels of TLR4 in sputum and blood neutrophils in asthma subtypes and healthy controls ...... 170 Table 7.19 Relative mRNA levels of TLR2 in sputum and blood neutrophils in asthma subtypes and healthy controls ...... 170 Table 7.20 Summary of results for the comparison between EA, NA, PGA versus healthy controls ...... 171 Table 7.21 Spearman correlations of innate immune mediators measured in resting sputum neutrophils and clinical parameters (*p<0.05)...... 172 Table 8.1 Clinical characteristics of neutrophilic and eosinophilic asthma subjects....181
xiii Table 8.2 Induced sputum inflammatory cell counts for subjects with eosinophilic asthma, non-eosinophilic asthma and healthy controls...... 182 Table 8.3 OSM, TLR2 and IL-8 fold change increase with LPS stimulation from baseline for microarrays and real-time PCR ...... 185 Table 8.4 LPS regulated genes that are also upregulated in resting neutrophils from subjects with neutrophilic asthma versus eosinophilic asthma...... 189 Table 8.5 Upregulated genes in resting neutrophils from subjects with neutrophilic asthma versus eosinophilic asthma involved in signal transduction...... 190 Table 8.6 Differentially regulated genes in resting neutrophils from subjects with neutrophilic asthma versus eosinophilic asthma related to cell motility...... 190 Table 8.7 Genes that were altered in resting neutrophils from subjects with neutrophilic asthma versus eosinophilic asthma involved in apoptosis...... 191 Table 8.8 Genes that were upregulated in resting neutrophils from subjects with neutrophilic asthma versus eosinophilic asthma involved in the NF-κB cascade 191 Table 8.9 Selected genes with immune related function that were altered in resting neutrophils from subjects with neutrophilic asthma compared to eosinophilic asthma ...... 192 Table 8.10 Selected genes with immune related function that were altered in resting neutrophils from subjects with neutrophilic asthma compared to eosinophilic asthma ...... 195 Table 9.1 Summary of Results...... 202 Table 9.2 Level of IL-8 measured in sputum supernatant corrected for the number of neutrophils present in the sample...... 213
xiv TABLE OF FIGURES
Figure 1.1 Pathways of Inflammation in Asthma ...... 10 Figure 1.2 Induced sputum cytospins of the four inflammatory subtypes of asthma, including neutrophilic (a), eosinophilic (b), mixed granulocytic (c), and paucigranulocytic (d) [9]...... 11 Figure 1.3 Important neutrophil functions ...... 15 Figure 1.4 Migration of neutrophils from the blood to the airways...... 22 Figure 1.5 Contents of Neutrophil Granules...... 28 Figure 1.6 Innate Immune Activation Pathway in Neutrophilic Asthma...... 43 Figure 2.1 Neutrophil Elastase Standard Curve...... 55 Figure 3.1 Total MMP-9 release from stimulated blood granulocytes at 24 hours ...... 70 Figure 3.2 Total MMP-9 release from LPS stimulated granulocytes (A) and NE release from control granulocytes (B) over 24 hours (n=10)...... 71 Figure 3.3 IL-8 release from granulocytes at 24 hours ...... 72 Figure 3.4 IL-8 release from LPS stimulated granulocytes over 24 hours ...... 72 Figure 3.5 Kinetics of TNF-α mRNA and protein levels over time ...... 73 Figure 3.6 TNF-α mRNA levels at 2 hours...... 74 Figure 4.1 Magnetic cell separation of peripheral blood eosinophils (A) and neutrophils (B) using CD16 microbeads, magnification 400x...... 84 Figure 4.2 Isolated Sputum Neutrophils, Magnification 1000x...... 85 Figure 4.3 Chemokine (A: IL-8) and cytokine (B: IL-1β, C: TNF-α, D: OSM) release from blood neutrophils (1 x 106 cell/mL) ...... 87 Figure 4.4 mRNA expression positively correlates with protein release...... 88 Figure 4.5 Zymography gel confirms the presence of MMP-9...... 89 Figure 4.6 TLR4 and TLR2 mRNA expression in isolated blood neutrophils (1 x 106 cells/mL) ...... 89 Figure 4.7 Chemokine (A: IL-8) and cytokine (B: IL-1β and C: TNF-α) release from sputum neutrophils (1 x 105 cell/mL)...... 90 Figure 4.8 TLR4 and TLR2 mRNA expression in sputum neutrophils (1 x 105 cells/mL) ...... 91 Figure 4.9 Chemokine (A: IL-8) and cytokine (B: IL-1β and C: TNF-α) release from resting sputum neutrophils compared to blood neutrophils (1 x 105 cell/mL)...... 92 Figure 4.10 Levels of TLR4 (A) and TLR2 (B) mRNA expression in resting sputum and blood neutrophils (1 x 105 cell/mL) ...... 93 Figure 4.11 Chemokine (A: IL-8) and Cytokine (B: IL-1β and C: TNF-α) release from LPS stimulated sputum neutrophils compared to LPS stimulated blood neutrophils (1 x 105 cell/mL) ...... 93 Figure 4.12 Levels of TLR4 (A) and TLR2 (B) mRNA expression in LPS stimulated sputum and blood neutrophils (1 x 105 cell/mL)...... 94 Figure 4.13 DTT treatment of neutrophils had no effect on IL-8 production...... 94 Figure 5.1 Level of IL-8 detected in sputum supernatant from participants under 55 (n=5) and over 55 (n=6)...... 104 Figure 5.2 Chemokine (A: IL-8) and Cytokine (B: IL-1β, C: TNF-α) release from resting sputum neutrophils (105 cells/mL) isolated from participants under (n=5) and over 55 (n=4)...... 105 Figure 5.3 Chemokine (A: IL-8) and Cytokine (B: IL-1β, C: TNF-α D: OSM) release from LPS stimulated blood neutrophils (106 cells/mL) isolated from participants under (n=5) and over (n=8) years of age...... 107 Figure 6.1 Levels of IL-8 detected in sputum supernatant...... 118
xv Figure 6.2 Chemokine (A: IL-8) and Cytokine (B: IL-1β, C: TNF-α) release from resting sputum neutrophils (105 cells/mL) ...... 119 Figure 6.3 Chemokine (A: IL-8) and Cytokine (B: IL-1β, C: TNF-α D: OSM) release from LPS stimulated blood neutrophils (106 cells/mL) ...... 121 Figure 6.4 Level of IL-8 in sputum supernatant from subjects with COPD compared to healthy controls ...... 126 Figure 6.5 Chemokine (A: IL-8) and Cytokine (B: IL-1β, C: TNF-α D: OSM) release from LPS stimulated blood neutrophils (106 cells/mL) isolated from participants with COPD compared with healthy controls...... 129 Figure 6.6 Total MMP-9 release from isolated blood neutrophils at rest (A) and stimulated with 100ng/mL LPS (B) in COPD compared with healthy controls...130 Figure 6.7 Chemokine (A:IL-8) and Cytokine (B: IL-1β, C: TNF-α, D: OSM) release from LPS stimulated blood neutrophils (106cells/mL) isolated from participants with asthma compared with COPD...... 136 Figure 7.1 Level of IL-8 detected in sputum supernatant ...... 149 Figure 7.2 Chemokine (A: IL-8) and Cytokine (B: IL-1β, C: TNF-α) release from resting sputum neutrophils isolated from subjects with non-eosinophilic asthma, eosinophilic asthma and healthy controls ...... 150 Figure 7.3 Enhanced IL-8 release from neutrophils in non-eosinophilic asthma ...... 151 Figure 7.4 Chemokine (A: IL-8) and Cytokine (B: IL-1β, C: TNF-α) release from LPS stimulated blood neutrophils (106 cells/mL) from subjects with non-eosinophilic asthma, eosinophilic asthma and healthy controls ...... 152 Figure 7.5 Subjects with neutrophilic asthma have significantly higher levels of sputum supernatant IL-8 ...... 160 Figure 7.6 Chemokine (A: IL-8) and cytokine (B: IL-1β, C: TNF-α) release from resting sputum neutrophils (105 cells/mL) asthma subtypes and healthy controls ...... 161 Figure 7.7 Zymography to assess levels of MMP-9 in culture supernatants of sputum neutrophils (105 cells/mL) at 24 hours...... 163 Figure 7.8 IL-8 release from resting blood neutrophils in asthma subtypes...... 164 Figure 7.9 Chemokine (A: IL-8) and Cytokine (B: IL-1β, C: TNF-α) release from LPS stimulated blood neutrophils (106 cells/mL) in asthma subtypes and healthy controls...... 166 Figure 7.10 Zymography to assess levels of MMP-9 in culture supernatants of isolated blood neutrophils (106 cells/mL) at 24 hours...... 168 Figure 8.1 Gene expression profiles of resting versus LPS stimulated neutrophils...... 184 Figure 8.2 Gene expression profiles of resting neutrophils from subjects with eosinophilic asthma versus those with neutrophilic asthma ...... 187 Figure 8.3 Schematic representation of genes altered both by LPS and in neutrophilic asthma ...... 188 Figure 8.4 Gene expression profiles of LPS stimulated neutrophils from subjects with neutrophilic asthma versus those with eosinophilic asthma ...... 194 Figure 9.2 Relative cytokine production (IL-8 protein expressed as average fold change from resting blood neutrophils, 1 x 105 cells/mL) ...... 203 Figure 9.3 Cycle of neutrophilic airway inflammation in neutrophilic asthma ...... 209
xvi ABBREVIATIONS
AHR Airway Hyperresponsiveness APCs Antigen Presenting Cells BAL Bronchoalveolar Lavage CAM Cellular Adhesion Molecule cDNA Complimentary Deoxyribonucleic Acid CF Cystic Fibrosis cRNA Complimentary Ribonucleic Acid COPD Chronic Obstructive Pulmonary Disease DNA Deoxyribonucleic Acid DTT Dithiothreitol EA Eosinophilic Asthma ECM Extracellular Matrix ELISA Enzyme Linked Immuno Sorbent Assay
FEV1 Forced Expiratory Volume in 1 Second FVC Forced Vital Capacity GINA Global Initiative for Asthma G-CSF Granulocyte – Colony Stimulating Factor GM-CSF Granulocyte Macrophage - Colony Stimulating Factor HBSS Hanks Balanced Salt Solution ICAM Intercellular Adhesion Molecule ICS Inhaled Corticosteroids IFN Interferon IKK IκB kinase IL Interleukin IRAK IL-1R associated kinase IVT In Vitro Transcription JAM Junction Adhesion Molecule LBP LPS Binding Protein LPS Lipopolysaccharide LTA Lipoteichoic Acids
LTB4 Leukotriene B4 LTRAs Leukotriene Receptor Antagonists
xvii MACS Magnetic Cell Separation MD-2 Myeloid Differentiation-2 MIP Macrophage Inflammatory Protein mg Milligram MGG May Grunwald Giemsa mL Millilitre mRNA messenger RNA MMP Matrix Metalloproteinase MPO Myeloperoxidase NA Neutrophilic Asthma NE Neutrophil Elastase NEA Non-eosinophilic Asthma NF-κB Nuclear Factor κB NOD Nucleotide Binding Oligomerisation Domain OCS Oral Corticosteroids OD Optical Density PAF Platelet Activating Factor PBS Phosphate Buffered Saline PAMPs Pathogen Associated Molecular Patterns PBMCs Peripheral Blood Mononuclear Cells PEF Peak Expiratory Flow PCR Polymerase Chain Reaction PGA Paucigranulocytic Asthma PMA Phorbol Myristate Acetate PRR Pattern Recognition Receptor RANTES Regulated on Activation Normal T cell Expressed and Secreted RNA Ribonucleic Acid ROS Reactive Oxygen Species RSV Respiratory Syncytial Virus SP-A Surfactant Protein-A TH T Helper TIMP-1 Tissue Inhibitor of Metalloproteinases-1 TLR Toll-like Receptor TNF-α Tumor Necrosis Factor-alpha
xviii ABSTRACT
Asthma involves chronic inflammation of the airways that is heterogeneous in nature. Eosinophilic airway responses are well described in asthma, however non-eosinophilic subtypes of asthma have been recently reported, and can involve the influx of neutrophils into the airways (neutrophilic asthma). Neutrophils are important effector cells of the innate immune system. These cells are the first to migrate to inflammatory sites, where they contain and eliminate pathogenic microorganisms. Neutrophils also release cytokines and chemokines that initiate and amplify inflammatory responses.
The mechanisms of neutrophilic asthma remain largely unknown; however activation of the innate immune response is implicated, particularly increased levels of proinflammatory cytokines Interleukin (IL)-8 and IL-1β and gene expression of Toll Like Receptor (TLR)-4 and TLR2 have been demonstrated in induced sputum samples. This thesis examines innate immune responses of airway and circulating neutrophils, with a focus on neutrophilic asthma. Innate immune neutrophil activation occurs in response to exposure to Lipopolysaccharide (LPS), which activates TLR4. The activation response consists of the release of preformed granule associated mediators such as Matrix Metalloproteinase (MMP)-9 and Oncostatin M (OSM), new gene transcription and release of inflammatory cytokines such as IL-8, IL-1β and Tumor Necrosis Factor (TNF)-α, and new gene transcription of TLR2 & TLR4 which serve to amplify neutrophil responses. In addition, this thesis examines whole genome gene expression profiles of circulating neutrophils in neutrophilic and eosinophilic asthma. The aims of this thesis are based on the hypothesis that dysregulation of innate immune neutrophil responses occurs with ageing and airway disease, particularly neutrophilic asthma and chronic obstructive pulmonary disease (COPD).
With advancing age, there were alterations in the innate immune responses of neutrophils, which were characterised by enhanced spontaneous activation of both airway and circulating neutrophils, and a decreased response of circulating neutrophils to LPS. There was a decreased activation of airway neutrophils in airway disease that was most pronounced in neutrophilic asthma and COPD, with decreased production and release of proinflammatory cytokines most likely due to a downregulation of TLR4. TLR2 was downregulated in resting and LPS stimulated circulating neutrophils in
1 asthma, particularly neutrophilic asthma. Circulating neutrophils had a decreased spontaneous release of total MMP-9, and downregulation of OSM, TLR2 and TLR4 at rest in COPD. However when stimulated with LPS, subjects with COPD had an enhanced proinflammatory cytokine release, with increases in IL-8 and TNF-α compared to subjects with asthma or healthy controls. Analysis of whole genome gene expression of circulating neutrophils in asthma revealed distinct gene profiles relating to asthma subtype. There was upregulation of genes relating to cell motility, inhibition of apoptosis and the NF-κB in neutrophilic asthma, which would contribute to their accumulation in the airways.
The innate immune response is critical in controlling infections by bacteria and viruses. The reduced innate immune response of airway neutrophils in airway disease could contribute to impaired local defense, which may lead to an increased susceptibility to infection by invading pathogens. Systemically, the molecular mechanisms of neutrophilic asthma are distinct from eosinophilic asthma and may involve the enhancement of neutrophil chemotaxis and survival, contributing to their accumulation in the airways.
2 Chapter 1: General Introduction
1.1 Asthma
Asthma is a chronic inflammatory disease of the airways. Airway inflammation in asthma is related to airways hyperresponsiveness (AHR), airflow obstruction and respiratory symptoms. The acute symptoms of asthma are usually reversible spontaneously or with treatment. The current definition of asthma has been adopted since 1997: “Asthma is a chronic inflammatory disorder of the airways in which many cells and cellular elements may play a role. The chronic inflammation is associated with airway hyperresponsiveness that leads to recurrent episodes of wheezing, breathlessness, chest tightness and coughing, particularly at night or in the early morning. These episodes are usually associated with widespread but variable airflow obstruction within the lung that is often reversible either spontaneously or with treatment.” [1]. This definition acknowledges the heterogeneity of the inflammatory response in asthma.
Studies of post mortem tissue show important pathological features of asthma. Macroscopic changes in asthma include lung hyperinflation, and clogging of the airways with mucus, cells and cell secretions. Microscopically there is an influx of inflammatory cells including eosinophils, lymphocytes and neutrophils into the airway lumen, as well as epithelial cell disruption, vasodilation and vascular leakage. Other features include an increased amount of smooth muscle, formation of new blood vessels, an increase in the number of epithelial goblet cells, and thickening of the subepithelial basement membrane due to fibrous tissue deposition [1, 2].
AHR and airflow limitation are two physiological manifestations of asthma, both of which are related to airway inflammation. AHR is defined as an excessive airway narrowing to not only allergens but to non-specific stimuli as well, for example chemicals such as methacholine, cold air, or smoke. The degree of hyperresponsiveness of the airways is related to the severity of asthma [3]. Airflow limitation in asthma is due to a number of reasons, including acute bronchoconstriction, airway wall swelling, intraluminal mucus, and airway remodeling [1].
3 Asthma is a significant problem worldwide, which causes substantial social burden and costs to the public and private health systems [1]. The prevalence of asthma is increasing worldwide, however the reasons for this increase remain unknown [4]. In Australia, a significant proportion of the population has asthma. It affects approximately 14-16% of children and 10-12% of adults, and these rates are relatively high compared with other countries around the world [5]. Over the past 5 to 10 years, asthma related deaths continue to decline. This is due to improvements in asthma management, in particular drug treatments for asthma, and increase in expenditure on asthma, particularly for pharmaceuticals. In 2003, 314 people died from asthma, which represents 0.3% of all deaths in Australia [5]. This rate has decreased since the 1980’s, but continues to be high by international standards.
1.1.1 Allergy
Allergy is frequently associated with asthma in both children and adults. Atopic (allergic) individuals are distinguished from nonatopic individuals via increased production of IgE to common environmental allergens such as house dust mite, animal proteins, pollens and fungi. Atopy can be assessed by skin prick tests observing a wheel and flare reaction for a positive result, or specific serum IgE measures of sensitisation against a panel of common allergens [6].
Atopy is acknowledged as a major risk factor for asthma, and it has been associated independently with AHR, asthma prevalence, persistence and severity [7]. However, many people who are atopic by skin prick tests do not have symptoms of asthma, indicating that other factors influence development of the disease [6]. The contribution of atopy to asthma is estimated at 40% in both children and adults [8]. A systematic review conducted by Pearce et al [8] determined that the proportion of asthma caused by atopy is usually less than 50%.
1.2 Inflammatory Subtypes of Asthma
Airway inflammation is a characteristic feature of asthma, which can contribute to airway hyperresponsiveness, airway obstruction, respiratory symptoms and disease severity. Many factors can play a role in inducing airway inflammation; however
4 studies so far have focused on cells recruited to the airways, specifically their activation and release of inflammatory mediators.
The pathogenesis of asthma is complex and depends on many genetic and environmental factors. Much of the current literature describes asthma as an allergic disease in which eosinophils are primarily responsible. However, inflammation in asthma now appears to be far more complicated than eosinophilic inflammation alone. It is now recognised that asthma can be divided into subtypes of Eosinophilic and Non- Eosinophilic Asthma, based on induced sputum cell count, and classified by the presence or absence of eosinophils, respectively.
1.2.1 Eosinophilic Asthma
Eosinophilic asthma is defined as asthma symptoms and AHR in the presence of sputum eosinophils of >1% [9]. Mechanisms underlying eosinophilic asthma include exposure to allergens, leading to activation of Th2 cells, release of IL-5 and eosinophil influx (Figure 1.1). The adaptive immune response is heavily involved in the pathogenesis of eosinophilic asthma.
1.2.1.1 Adaptive Immune Response
There are two phases to the adaptive immune response to allergen, the early and the late phase reaction. The most important component of the adaptive immune reaction in asthma is the activation of T cells to produce helper type 2 (Th2) cytokines including IL-4, IL-5 and IL-13, which play important roles in the production of IgE and recruitment of eosinophils [10].
Following exposure to allergen, individuals may become sensitised. This process involves dendritic cells and subsequent T cell responses leading to allergen specific IgE production and Th2 responses. The antigen presenting dendritic cells in the airway mucosa intercept allergen that manages to avoid mucociliary clearance and penetrate the epithelial layer. The dendritic cells then migrate to the regional lymph nodes where they present the antigen to the B and T cells and IgE production is initiated by the B cell in the presence of IL-4 and IL-13 [7]. IL-4 and IL-13 are important for the production of
5 IgE from B cells, and lead to a switch from IgG to IgE production in these cells. In particular, IL-13 is an important regulator of and may play a central role in asthma and allergic diseases, demonstrated by studies of IL-13 deficient mice, which have impaired Th2 responses along with decreased IgE production [11]. Antigen specific IgE is released into the blood stream, where it can bind to the high affinity IgE receptor (FcεR1) on the surface of mast cells, basophils and dendritic cells, or the low affinity receptors (FcεR2, CD23) on monocytes/macrophages and lymphocytes [12].
When a person next encounters an inhaled allergen, then the early and late phase allergic reactions may be elicited. The early phase response, lasting 30-60 minutes, begins with the inhalation of a particular allergen, and is characterised by mediator release from mast cells leading to constriction of airway smooth muscle, vascular leakage and mucus production. The inflammatory cascade is initiated when IgE bound to effector cells is cross-linked by allergen. Mast cell activation via IgE cross linking results in cellular degranulation, and release of a comprehensive array of preformed and newly generated proinflammatory mediators including histamine, chymase, tryptase, lipid mediators, reactive oxygen species and cytokines [10]. These substances are toxic and have been associated with acute asthma symptoms. They can induce contraction of smooth muscle cells, mucus secretion and vasodilation.
IgE plays a critical role in allergic inflammation. Genetic analysis in families has proven an association between bronchial hyperresponsiveness and IgE levels [13]. Anti-IgE (omalizumab) attenuates both early and late phase responses in mild asthma patients after the inhalation of allergen [14]. Omalizumab also improves asthma symptoms in patients with moderate to severe allergic asthma who are taking corticosteroids [15].
Allergen induced responses that are clinically significant result in a biphasic response where intense inflammation follows the immediate IgE hypersensitivity (early) reaction. This is termed the late phase response, and occurs 3 to 4 hours after the early phase response. The late phase response is characterised by excessive inflammation of the airways resulting in airway narrowing [16]. Central to the late phase response is the activation of Th2 cells to release proinflammatory cytokines including IL-5, resulting in influx and activation of eosinophils [7]. Eosinophil influx into the airways is a common pathological feature of allergen-induced asthma.
6
1.2.1.2 Eosinophils
Eosinophils are multifunctional leukocytes that are known to be involved in the pathogenesis of parasitic helminth infections and allergic disease [17]. Activated eosinophils degranulate, and release granule derived proteins, such as major basic protein (MBP) and eosinophil peroxidase (EPO), as well as producing lipid derived mediators for example LTC4 and platelet activating factor (PAF), and reactive oxygen species [18]. The eosinophil can regulate immune function via the synthesis of at least 28 cytokines such as IL-4, IL-5 GM-CSF, chemokines such as eotaxin, IL-8 and RANTES and growth factors such as TGF-α and β [18]. In the airways of asthmatic subjects, eosinophils have the ability to exacerbate and prolong inflammatory responses through the release of inflammatory mediators. Major basic protein (MBP) can cause epithelial damage, airway constriction, and AHR. MBP instillation into mouse airways in vivo can induce AHR through effects on the respiratory epithelium [19].
Processes essential to the development and recruitment of eosinophils include hematopoietic development release from the bone marrow, endothelial adhesion, chemotaxis, and survival.
Eosinophils are derived from pluripotent stem cells in the bone marrow. IL-5 is responsible for eosinophil growth, differentiation and mobilisation from the bone marrow, as well as prolonging eosinophil survival [20]. Airway eosinophilia can be reproduced in both mice and human models of allergic asthma. IL-5 deficiency in mice results in an absence of eosinophilia after allergen sensitisation and challenge with ovalbumin. These mice also did not develop AHR to methacholine [21]. Inhalation of IL-5 in human patients with allergic asthma induces blood eosinophilia and increases in serum ECP, a marker of eosinophil activation [22]. IL-3 and GM-CSF have also been found to play a role in the priming and activation of eosinophils [23]. Cytokine stimulation of eosinophils results in their rapid migration and further production of inflammatory mediators [12].
The recruitment of eosinophils to the airways involves adhesion between the cell and the endothelium at the sites of inflammation and expression of adhesion molecules is
7 critical for this process. Binding initially occurs as low affinity rolling, mediated by E and P selectins, of the cell on the vascular endothelium [24]. Further activation is required for firm adhesion to the endothelium and subsequent extravasation, which involves upregulation of adhesion molecules including very late antigen (VLA)-4 and
β2-integrins on eosinophils and intercellular adhesion molecule (ICAM)-1 or vascular cell adhesion molecule (VCAM)-1 on the endothelium [25].
Numerous chemokines are important to the recruitment of eosinophils, and are expressed at the sites of allergic inflammation after challenge with allergen. Eotaxin is a lineage specific chemokine for eosinophils, and has been associated with early recruitment of eosinophils after allergen challenge. At later time points, chemokines such as RANTES (regulated on activation, normal T cell expressed and presumably secreted), monocyte chemotactic proteins (MCP)-2, 3, 4, 5, and macrophage inflammatory protein (MIP)-1α are also important in tissue recruitment of eosinophils [26]. Eosinophil responses to eotaxin, RANTES, MCP-3 and MCP-4 can be blocked by a monoclonal antibody directed against the chemokine receptor CCR3 [27]. Production of RANTES by eosinophils has been shown to affect the chemotactic activity of eosinophils in vitro [18]. This indicates that eosinophils may also have the potential to promote its own migration.
Enhanced survival of eosinophils as a result of reduced apoptosis may be a factor in the persistence of airways inflammation in asthma [28]. Increased production of IL-5 and GM-CSF has been shown to delay apoptosis of monocytes and eosinophils [29], whereas corticosteroid treatment increases the rate of eosinophil apoptosis [30]. Eosinophil apoptosis is delayed in steroid naive asthma, and this delay can be explained partially by the presence of GM-CSF [31].
1.2.2 Non-Eosinophilic Asthma
A well-characterised pathway of inflammation implicated in asthma pathogenesis involves the inhalation of allergens, which induce Th2 lymphocyte, and IL-5 mediated eosinophil influx [1] (Figure 1.1). Despite effective treatments targeting this inflammatory process, asthma symptoms are known to persist in the absence of eosinophilia [32-36]. This inflammatory subtype is termed non-eosinophilic asthma
8 [37]. Subjects with non-eosinophilic asthma have clinical symptoms of asthma with AHR and without elevated sputum eosinophils [9, 32, 37].
Non-eosinophilic asthma (NEA) was first demonstrated by Turner et al [35] in mild exacerbations of asthma, and now has been shown to occur in all grades of severity of asthma, including moderate asthma [38], severe refractory asthma [36], persistent asthma [32], and steroid resistant asthma [34]. NEA is proven to be stable and reproducible over the short (4 weeks) and long (5 years) term [9]. NEA is associated with corticosteroid unresponsiveness [34], and poor short term responses to inhaled corticosteroid treatment [39]. Corticosteroid treatments have little impact on non- eosinophilic inflammation, and they potentially promote neutrophilic inflammation by reducing neutrophil apoptosis [40]. NEA is a significant subtype of asthma occurring in both children and adults, and accounting for up to 50% of cases [37].
Pavord et al [34] associated NEA with smoking and atopy, as the participants in their study were more likely to be non-atopic and current smokers. Gibson et al [32] demonstrated an association between age and disease duration and non-eosinophilic inflammation. Basyigit et al [38] observed that subjects with NEA had a longer disease duration and tended to be non-atopic. These data suggest that older, non-atopic patients with more severe airway obstruction and longer disease duration tend to develop non- eosinophilic airway inflammation.
9 Allergens e.g. house dust mite, Endotoxin, Viruses, Air cat dander, pollen Pollution
Activated Th2 cells Activated Epithelial Cells & Macrophages IL-5 IL-8
Eosinophils Neutrophils
Eosinophilic Non-Eosinophilic
Asthma Asthma Figure 1.1 Pathways of Inflammation in Asthma
Recently, the use of induced sputum inflammatory cell counts has suggested the presence of four distinct subtypes of asthma based on sputum eosinophil and neutrophil proportions [9]. This study identified four inflammatory subtypes of asthma, including those with increased sputum eosinophils (eosinophilic asthma), increased sputum neutrophils (neutrophilic asthma), increased sputum eosinophils and neutrophils (mixed granulocytic asthma) and normal levels of sputum eosinophils and neutrophils (paucigranulocytic asthma). Sputum cytospins of the four subtypes of asthma can be viewed in Figure 1.2.
10
Figure 1.2 Induced sputum cytospins of the four inflammatory subtypes of asthma, including neutrophilic (a), eosinophilic (b), mixed granulocytic (c), and paucigranulocytic (d) [9].
The mechanisms underlying non-eosinophilic subtypes of asthma remain largely unknown. Most studies of NEA have associated it with elevated IL-8 levels and neutrophil numbers [32, 36, 41-43], suggesting that non-allergic neutrophil mediated inflammation is important in driving this type of airway inflammation (Figure 1.1). Gibson et al [32] identified a role for neutrophilic inflammation in non-eosinophilic persistant asthma. Participants with NEA in this study had increased numbers of neutrophils in induced sputum samples, as well as increased levels of myeloperoxidase and the potent neutrophil chemoattractant IL-8.
Airway inflammation in non-eosinophilic asthma shows similarities to that seen in occupational asthma [44-46]. The mechanism underlying non-allergic occupational asthma is the release of innate immune cytokines, including IL-1, IL-6, IL-8 and TNF-α,
11 leading to massive infiltration and activation of neutrophils [44]. This supports the hypothesis that innate immune mechanisms involving neutrophil activation mediate the inflammatory process in NEA [37].
1.3 Neutrophils in Asthma
Significant evidence exists to implicate neutrophils in the pathogenesis of asthma. Neutrophils are prominent in nocturnal asthmatics, and the presence of neutrophils is correlated with severity of nocturnal asthma [47]. In addition, neutrophils are found in large numbers in the airways of subjects with acute exacerbations of asthma, and this was mostly associated with respiratory tract infections [48]. High neutrophil counts in bronchoalveolar lavage (BAL) have also been demonstrated in patients having an acute asthma exacerbation that was not associated with infection [49].
Neutrophils are increased in more severe forms of asthma including in subjects with severe asthma requiring intubation [41], sudden onset fatal asthma and life threatening asthma [49-51]. Large numbers of neutrophils, with minimal numbers of eosinophils, have been demonstrated in the airway mucosa of patients with sudden onset fatal asthma [51, 52]. Moreover, studies in severe asthmatics suggest a subgroup of patients that had no eosinophil infiltration, but rather predominantly neutrophil infiltration [36, 43, 53]. This group of patients had poor airway function despite very high doses of inhaled and oral corticosteroids.
Peripheral blood neutrophils from subjects with allergic asthma release increased amounts of myeloperoxidase upon stimulation with N-formyl-methionyl-leucyl- phenylalanine (fMLP) [54], which is thought to be mediated by activation of IgE receptors [55]. There is also increased production of reactive oxygen species from peripheral blood neutrophils obtained from subjects with asthma both spontaneously [56] and with stimulation [57, 58].
AHR is considered a hallmark feature of asthma. Inhalation of a variety of triggers of neutrophilic airway inflammation, including ozone [59], infection [60], endotoxin [61], smoking [62] and organic dust [63], are associated with AHR. Evidence suggests that neutrophil depletion is associated with improvements in AHR [59, 64]. Inflammatory
12 mediators involved in neutrophil accumulation such as LTB4, IL-17 and IL-8 have also been associated with AHR [65, 66].
Neutrophils may play a role in airway remodeling through the release of growth factors such as transforming growth factor (TGF)-β, which may lead to activation of fibroblasts and alterations in the turnover of the extracellular matrix. Increased TGF-β production from neutrophils has been demonstrated in patients with asthma compared to healthy controls [67]. Formation of new blood vessels (angiogenesis) is also an important feature of airway remodeling. Neutrophils are an important source of angiogenic factors and release VEGF upon stimulation with bacterial products, the released VEGF can then activate endothelial cells to induce angiogenesis [68].
1.4 Neutrophils in Chronic Obstructive Pulmonary Disease (COPD)
Although the main focus of this thesis was to investigate the innate immune responses of neutrophils in asthma subtypes, neutrophils remain a hallmark feature of chronic obstructive pulmonary disorder (COPD) [69, 70]. Therefore a small group of participants with COPD were included in this study to investigate both the similarities and the differences of COPD and neutrophilic asthma. The presence of neutrophils in the airways of COPD patients is associated with increased levels of neutrophilic inflammatory mediators including IL-8, TNF-α and MMP-9 [71-73]. Markers of airway neutrophilic inflammation are correlated with COPD disease progression [74], and the degree of neutrophilic inflammation is also correlated with clinical severity [75]. Neutrophilic airway inflammatory markers are further increased with COPD exacerbations [76].
Peripheral blood neutrophils from subjects with COPD show altered activity in both stable disease and exacerbations, including increased presence of cell surface adhesion molecule expression [77-79], upregulation of genes relating to inflammation [80] and enhanced respiratory burst [78].
13 1.5 Biology of Neutrophils
Neutrophils are highly specialised short-lived cells that arrive first at the site of inflammation or infection. Neutrophils account for 50-75% of circulating leukocytes in humans, and their numbers are further increased in acute and chronic inflammatory diseases [81]. The bulk of their life span is spent proliferating and differentiating in the bone marrow where the cells are stored for a few days and then released into the circulation. The cells circulate in the blood for a short time before they migrate into the tissues where they function as mobile phagocytes.
Neutrophils are an important part of the innate immune defense against injury and infection due to their ability to engulf and kill pathogenic microorganisms [82]. Any defect in the functionality of phagocytic cells can result in fatal diseases due to the lack of protection from invading pathogens.
The inflammatory response mediated by neutrophils is a multi-step process that involves adhesion of circulating neutrophils to the activated vascular endothelium, and the transmigration of the neutrophil to the inflammatory site, where they become activated and execute a number of defence mechanisms. Neutrophils recognize and ingest foreign organisms through phagocytosis, where they kill and degrade microbes by the production of reactive oxygen species and antimicrobial and proteolytic granule proteins. In addition to this, neutrophils synthesise cytokines and chemokines, which recruit and regulate the inflammatory response of other effector cells, such as macrophages, T lymphocytes and neutrophils themselves. The neutrophil then undergoes apoptosis, which facilitates resolution of inflammation (Figure 1.3). The studies in this thesis focus on the neutrophil proinflammatory response as part of innate immune activation.
14 Rolling IL-8 Adhesion IL-8
Local injury or infection Diapedesis
Phagocytosis & killing of Apoptosis & microorganisms phagocytosis by macrophages
Degranulation Pro-inflammatory response
Figure 1.3 Important neutrophil functions In response to local infection or injury, neutrophils attach to the activated endothelium via a series of interactions between adhesion molecules and their receptors. Neutrophils engulf and kill microorganisms by the production of reactive oxygen species, and antimicrobial and proteolytic granule proteins. Neutrophils synthesize cytokines and chemokines, which recruit and regulate the inflammatory response of other effector cells. Neutrophils then undergo apoptosis, which facilitates resolution of inflammation.
1.5.2 Neutrophil Migration
During airway infection or inflammation, neutrophils migrate from the blood to the airways where they play a crucial role in innate immune defense. This is a complex process that first requires the maturation of neutrophils in the bone marrow, their release into the circulation and migration across the endothelial and epithelial interface, which occurs under the influence of chemotactic factors and adhesion molecules [83]. This process gradually changes the functional state of the neutrophil from a passive circulating cell into a highly activated effector cell of innate immunity.
Mature neutrophils do not undergo cell division. They are generated continuously from the bone marrow (approximately 1011 cells per day). The numbers of circulating neutrophils are tightly regulated, but can be greatly amplified in times of stress, e.g., infection. The maturation of neutrophils in the bone marrow involves the highly
15 controlled process of myelopoiesis, where pluripotent stem cells divide and differentiate into myeloid precursors. During maturation, neutrophil granules are formed, which contribute to the inflammatory response in the fight against microorganisms.
A variety of pre-formed and newly synthesized compounds are important involving neutrophils including serine and metalloproteinases, reactive oxygen species, lipid mediators and defensins. These toxic molecules are released from activated neutrophils, and have the ability to cause significant tissue damage to the lung and airways in asthma. This damage occurs when neutrophils accumulate in large numbers, and their activation is inappropriate or uncontrolled.
1.5.2.1 Myeloid Development
Neutrophils are produced continuously from hematopoietic stem cells of the bone marrow, and this process is called myelopoiesis. Developing neutrophils can be divided into 6 subtypes, including myeloblasts, promyelocytes, myelocytes, metamyelocytes, band cells and mature neutrophils. Myeloblasts have a large oval nucleus, large nucleoli and no granules in the cytoplasm. This stage of the cell development is followed by two secretory stages; the promyelocyte and the myelocyte, where neutrophil granules are formed. The metamyelocyte and band forms are nonproliferating, nonsecretory stages, which develop into the mature neutrophil. Mature neutrophils are characterised by their multilobed nucleus and cytoplasm containing granules [84].
Neutrophil granules are formed during the differentiation process. Immature transport vesicles in early promyelocytes bud off the Golgi and fuse, creating granules with a high content of myeloperoxidase (MPO), termed ‘azurophil granules’ or more simply ‘primary granules’[85]. Azurophil granules also contain serine proteases and antibiotic proteins, and therefore these granules are thought to be the true microbicidal compartment that is mobilised upon phagocytosis [86]. At the promyelocyte/myelocyte transition, the production of MPO stops, and granules formed at later stages are MPO negative. Peroxidase negative granules are divided into 2 groups, the specific (secondary) granules and the gelatinase (tertiary) granules. Specific granules have a high content of lactoferrin and low levels of gelatinase and are formed in myelocytes and metamyelocytes, and gelatinase granules form in band cells and segmented
16 neutrophils and are high in gelatinases but low in lactoferrin [87]. Secretory vesicles are formed by endocytosis as they contain plasma proteins such as albumin, and they appear in segmented neutrophils [88, 89].
Two very rare genetic diseases Chediak-Higashi syndrome and specific granule deficiency exhibit the importance of functioning neutrophil granules. Chediak-Higashi syndrome results from the fusion of specific and azurophilic granules [90]. Whereas specific granule deficiency results from the absence of secondary granule proteins [91]. These diseases are characterised by recurrent infections and shortened life expectancy, illustrating the fundamental importance of the granule contents in host defense.
Many factors influence the development of neutrophils in the bone marrow. These include stromal cells such as fibroblastoid cells, endothelial cells, adipocytes, reticular cells and macrophages; components of the extracellular matrix such as collagens, glycoproteins and proteoglycans; as well as adhesion molecules such as CD11b/CD18 and growth factors such as G-CSF and GM-CSF [92, 93]. Important also in this process are specific changes in gene expression patterns controlled by transcription factors such as C/EBPs and PU.1 [94, 95]. During maturation neutrophils increase their mobility, deformability and responsiveness to chemokines.
Neutrophil maturation in the bone marrow takes approximately 10-15 days, and depends on the detachment of the cells from the marrow microenvironment, and the mechanical ‘pumping’ of the cells into the bone marrow sinuses. Immature neutrophils can be released prematurely into the circulation in times of infection or inflammation, and these cells preferentially sequester into the lung microvessels [96]. Exposure to inhalants, such as cigarette smoke, can decrease the transit time of neutrophils through the bone marrow, and cause the release of immature neutrophils into the blood stream [97]. Contact with cytokines (e.g., G-CSF, GM-CSF, IL-1), and chemokines (e.g. IL-8) can influence this process through the release of proteases (e.g., MMP-9) and the shedding of L-selectin [98-100]. Once released into the blood stream neutrophils have a half-life of 4 to 10 hours, and can migrate into the tissues.
17 1.5.2.2 Neutrophil Trafficking and Margination
Peripheral blood neutrophils are divided between a circulating pool, present in large and small blood vessels, and a marginating pool that are arrested in capillaries (mainly pulmonary) [101]. The pulmonary capillary bed is the main site containing marginating neutrophils and measuring 20-60 times that of the concentration of large systemic blood vessels [102]. Most neutrophils have to deform and elongate to travel through the pulmonary capillaries due to the vast network of the capillary bed, and the vessels being of a smaller diameter in comparison to spheric neutrophils [103]. The requirement of neutrophils to deform to travel through the pulmonary capillaries increases their transit time, resulting in a higher concentration of neutrophils in this space [104].
Normal margination should not be confused with neutrophil sequestration, which is defined as amplified intravascular neutrophil numbers induced by inflammatory mediators and complement factors. Initial stages of sequestration are thought to involve cytoskeletal rearrangements such as increases in F actin at the periphery of the cell to reduce neutrophil deformability. Prolonged sequestration of neutrophils requires CD11b/CD18 [102]. The migration of neutrophils into tissues involves neutrophil rolling, activation and firm adhesion to endothelial cells, followed by migration through the endothelial cell layer, the basement membrane and the epithelial interface and accumulation in the airway lumen (Figure 1.4). These events involve complex interactions between neutrophils and the endothelium, extracellular matrix and epithelium, and are largely mediated by cellular adhesion molecules (CAMs) such as the β2 integrins. This process gradually changes the functional state of the neutrophil from a passive circulating cell into a highly activated effector cell of innate immunity.
1.5.2.3 Cellular Adhesion Molecules
Inflammatory mediators released by damaged tissues, induces adhesion molecule expression on the endothelium. This, along with the slow flow rate in the capillary and vessel dilation at sites of inflammation, allows neutrophils to loosely adhere, roll and tether along the endothelial surface. The recruitment of neutrophils can be described as a sequential process having 3 distinct adhesive events [105]. The first adhesive event is
18 the primary adhesion, followed by neutrophil activation and finally activation dependent firm adhesion.
Neutrophil adherence to the endothelium involves cellular adhesion molecules on the neutrophil and endothelial cell, whose expression is tightly regulated. Rolling adhesion of neutrophils to the endothelium is mediated by L-selectin on the neutrophil and P- and E-selectin on the endothelium [86, 106]. Rolling allows interaction between CXC chemokines such as IL-8 presented on the surface of endothelial cells, which activates
β2 integrin expression. Interaction between the integrins CD11a/CD18 and CD11b/CD18 and the endothelial immunoglobulin (Ig) superfamily members, intercellular adhesion molecule (ICAM)-1 and ICAM-2, are required for effective neutrophil transmigration and firm adhesion to the endothelium [107, 108].
1.5.2.4 Integrins
The integrins are a family of heterodimeric transmembrane glycoproteins that mediate direct cell-cell, cell-extracellular matrix and cell-pathogen interactions. They contain 2 functional units: α and β chains. The β2 integrins are expressed on neutrophils and consist of 4 different heterodimers: CD11a/CD18 or leukocyte function associated antigen-1 (LFA-1), CD11b/CD18 or Mac-1, CD11c/CD18 or p150, 95 and CD11d/CD18. Leukocyte Adhesion Deficiency (LAD) results from a mutation in the gene for CD18 and is associated with recurrent bacterial infections due to an inability to recruit these cells to a site of infection [109].
The functional state and presence of integrins on neutrophils is regulated by lipid, cytokine and chemokine signalling molecules as well as ‘cross talk’ from other adhesion molecules. Integrins exist in predominately inactive states on circulating immune cells. Multiple mechanisms, including conformational change (affinity regulation) and clustering associated with the cytoskeleton (avidity regulation) are responsible for integrin activation, arising from or caused by ligand binding. Interestingly, the ability of the extracellular domains of integrins to bind ligands can be activated in less than 1 second via signals from within the cell (inside out signalling) [110].
19 1.5.2.5 Inflammatory Stimulus
In the pulmonary circulation, neutrophil migration occurs through at least two pathways: CD11b/CD18-dependent and –independent, and is dependent on the inflammatory stimulus. Inflammatory stimuli that invoke CD18 dependent neutrophil migration include LPS, Pseudomonas aeruginosa, immunoglobulin G (IgG), IL-1, immune complexes and phorbol myristate acetate (PMA). Stimuli that induce CD11b/CD18 independent neutrophil migration include Streptococcus pneumoniae; group B Streptococcus, Staphylococcus aureus, hydrochloric acid, hypoxia and C5a [102]. In vitro models have demonstrated that the bacterial derived chemoattractant fMLP stimulates CD18-dependent neutrophil migration, whereas the host derived chemoattractants IL-8 and LTB4 stimulate CD18-independent neutrophil migration [111]. It is unclear if one pathway is preferentially used over another in relation to neutrophil influx into the lung. However, a recent study adding cystic fibrosis sputum, which is a cocktail of both bacterial and host derived chemoattractants, to neutrophils in vitro preferentially induced CD18-independent migration [112].
1.5.2.6 Endothelial Cell Interactions
Although a significant amount of information is known about the first three steps of neutrophil migration (rolling, activation and adhesion), the mechanisms that underlie transendothelial migration remain unclear. Generally, leukocytes traverse the endothelial barrier through the cleft between two to three adjacent cells. Trans- endothelial migration, but also acquisition of cell polarity of the neutrophil, is thought to be mediated by platelet/endothelial cell adhesion molecule (PECAM)-1 [113] and junction adhesion molecules (JAMs) expressed at intercellular tight junctions of endothelial and epithelial cells [114]. Recently, the binding of JAM-C to Mac-1 was found to be of importance in neutrophil transendothelial migration [115].
1.5.2.7 Epithelial Cell Interactions
Mechanisms underlying neutrophil migration through the epithelium are only beginning to emerge. This process involves 3 stages, which include epithelial adhesion, migration and post-migration. Initially neutrophil firm adherence to the basolateral epithelial
20 membrane is mediated exclusively by Mac-1 [116]. Transepithelial migration of neutrophils involves both cell-cell interactions that include adhesion molecules and signalling events to open the epithelial tight junctions, allowing the passage of cells without disturbance of the epithelial barrier. The interaction between CD47 and signal regulatory protein-α (SIRPα) enhances the migration rate of neutrophils through the epithelium [117]. Further to this, JAMs are likely to be important in the migratory process, as well as the formation of a seal around migrating cells to preserve barrier function. After migration through the epithelium, neutrophils can adhere to ICAM-1 present on the apical surface of the epithelial cells.
21
Post-migration AIRWAY LUMEN ICAM-1
EPITHELIUM Adhesion Migration CD11b/CD18 CD47 SIRPα JAMs
Chemotactic gradient e.g. IL-8 EXTRA-VASCULAR TISSUE
ENDOTHELIUM IL-8 Migration Cell polarity Firm Adhesion PECAM-1 Β2 Integrins JAMs Rolling ICAM-1 & 2 Selectins
BLOOD
Figure 1.4 Migration of neutrophils from the blood to the airways
1.5.2.8 Chemotactic Mediators
Once through the endothelial basement membrane, neutrophils migrate along a chemotactic gradient. Neutrophil chemotactic proteins include chemokines (e.g. IL-8),
bacterial products (e.g. N-formyl methionyl peptides), lipid mediators (e.g. LTB4) and complement split products (e.g. C5a).
22 The chemokine family constitutes about 50 low molecular weight proteins that exert their effects through activation of one of 19 G-protein coupled chemokine receptors [118]. There are two main subfamilies of chemokines, CXC and CC, which are classified according to the position of the first two cysteines in their amino acid sequence (separated by one amino acid – CXC, or adjacent CC). Many chemokines can bind to more than one receptor and most chemokine receptors can bind more than one chemokine. Chemokines are produced by inflamed tissues and activate signal cascades in the neutrophil that lead to increase in cell motility, adhesion and survival.
1.5.2.8.1 Interleukin-8
IL-8 is a potent chemotactic factor for neutrophils and is proinflammatory in its effects [119, 120]. It is the main chemoattractant in the lung, since blocking of IL-8 with a neutralising antibody resulted in a 75-98% inhibition of its chemotactic activity [121]. High concentrations of IL-8 have been detected at inflammation sites in vivo [122]. IL- 8 is a member of the CXC subfamily. Other members of this family include epithelial cell-derived neutrophil activator-78 (ENA-78), growth regulatory gene (Gro)-α, Gro-β and neutrophil-activating peptide-2 (NAP-2); and granulocyte chemotactic protein-2 (GCP-2) [123]. IL-8 is produced by many cell types, including epithelial cells, alveolar macrophages, lymphocytes, fibroblasts, endothelial cells and neutrophils and is released upon proinflammatory stimulation [124]. IL-8 is involved in the whole process of neutrophil transmigration, including the shedding of L-selectin, upregulation of β2 integrins and adhesion to the endothelium. Furthermore, IL-8 can activate various functions of the neutrophils, including degranulation and respiratory burst [125, 126]. IL-8 also causes the secretion of neutrophil elastase and matrix metalloproteinase –9 (MMP-9), which are capable of causing tissue damage [127].
Peripheral blood neutrophils express 2 main chemokine receptors for IL-8 on the cell surface, CXCR1 and CXCR2. Engagement of CXCR2 is thought to promote neutrophil recruitment in response to many ligands, including IL-8 [128]. Engagement of CXCR1 which binds only IL-8 and IL-6 at high affinity, is thought to promote neutrophil activation [129]. CXCR1 and CXCR2 expression is downregulated by exposure to bacterial products such as LPS (acting through TLR4), and lipopeptides (acting through TLR2) [130].
23 1.5.3 Neutrophil Phagocytosis
Throughout the phagocytic process, neutrophils release a range of inflammatory mediators that contribute to the local inflammatory process, including lipid mediators
(e.g. leukotriene B4), proteolytic enzymes (e.g. neutrophil elastase), reactive oxygen species (e.g. superoxide), cytokines (e.g. TNF-α) and chemokines (e.g. IL-8) [131, 132]. This results in the pathogen being destroyed and resolution of the inflammatory response.
The mechanism underlying the uptake of pathogens begins with the binding of receptors on the neutrophil membrane with specific molecules on the membrane of the particle to be engulfed. The ligand/receptor complex then induces rearrangements in the cytoskeleton, which leads to internalisation of the complex, forming a phagosome. Fusion of the phagosome with intracellular granules allows the formation of the phagolysosome. Bacteria in the phagolysosome are killed by exposure to enzymes, reactive oxygen species and antimicrobial peptides, and this can be characterised into oxygen dependent (respiratory burst) or oxygen independent mechanisms (degranulation) [133]. The efficiency of neutrophils depends on these mechanisms, which rely on the activation of NADPH-oxidase generating reactive oxygen intermediates, and the release of granule contents via degranulation.
1.5.4 Innate Immune Activation
The lung, particularly the respiratory epithelium, has a large area that is exposed to the external environment and the introduction of potentially pathogenic microorganisms through inspiration. The airways are protected via nonspecific mechanisms such as ciliary beat, cough reflex, and mucus clearance, as well as defense against invading pathogens through recruitment of phagocytic cells of the innate immune system. Dysregulation of the innate immune system may contribute to the pathogenesis of inflammatory diseases including asthma.
The innate immune system was previously thought to be non-specific, however recent evidence proves its extraordinary complexity and specificity. This specificity allows the body to distinguish pathogenic and nonpathogenic particles that enter the respiratory
24 tract. The main action of the innate immune system is the recognition of conserved sequences on microbes, known as pathogen-associated molecular patterns (PAMPs). PAMPs are recognized by pattern recognition receptors (PRRs) that are present on epithelial cells and leukocytes [134]. Examples of well-recognised PAMPs are bacterial lipopolysaccharide (LPS), peptidoglycan, lipoteichoic acids, mannans, bacterial DNA, double stranded RNA and glucan. PAMPs are only found on microbial pathogens, allowing for the innate immune system to distinguish between structures that are ‘self’ and ‘non-self’, and they are usually essential for the survival or pathogenicity of the organism. The same PAMP is usually shared by the entire class of pathogen, for example, LPS is present on all gram negative bacteria [135, 136].
The receptors of the innate immune system are encoded in the germ line, and are expressed on many effector cells including macrophages, dendritic cells and B cells. Stucturally, PRRs can be divided into 3 classes, which are secreted, endocytic, and signalling [135, 136]. Secreted PRRs such as the collectins (mannose binding lectin (MBL) or the surfactant proteins), bind to the cell wall of the pathogen, which flags them for recognition by phagocytes and the complement system. Endocytic PRRs are expressed on the cell surface of phagocytic cells. These molecules assist in the uptake and delivery of the pathogen to the lysosomes of the cell, where the pathogen is destroyed. An example of an endocytic PRR is the macrophage mannose receptor, which specifically identifies carbohydrates with large numbers of mannoses that are characteristic of microorganisms, and activation of this receptor leads to the phagocytosis of the foreign organism by the macrophage. Signalling PRRs recognise PAMPs, this resulting in the activation of signalling pathways that induce the expression of immune response genes, which include proinflammatory cytokines. The Toll Receptor family are the main signalling receptors of innate immunity [135, 136].
1.5.4.1 Toll Like Receptors (TLRs)
Toll Like Receptors (TLRs) play a crucial role in the detection of invading microorganisms and the initiation of host defences. TLRs are highly conserved type 1 transmembrane receptors. They are characterised by a NH2-terminal extracellular leucine rich repeat domain and a COOH-terminal intracellular tail homologous to the IL-1 receptor (Toll/IL-1 receptor (TIR) domain) [137]. Upon ligand binding to the
25 extracellular portion of these receptors, the TIR domain recruits the adapter molecule MyD88, which leads to a signalling cascade involving IL-1 receptor associated kinase (IRAK), TNF-associated factor 6 (TRAF6), the end result being activation of the transcription factor NF-κB and transcription of inflammatory cytokines [135, 136].
Currently the TLR family contains 10 members, and at the mRNA level, neutrophils appear to express all of these receptors except for TLR3 [138]. TLR4 was the first toll to be characterised. TLR4 is the receptor for LPS, present on gram-negative bacteria. A number of molecules are important in the detection of LPS. LPS first binds to a serum protein called LPS-binding protein, which transports the LPS to the CD14 receptor present on macrophages and B cells. There is also a soluble form of CD14 (sCD14) that binds endotoxin with high affinity. The CD14-LPS complex binds with TLR4 and also with another molecule MD2, and once this has occurred then the signalling cascade is initiated [139]. TLR4 and MD-2 are associated with each other on the cell surface, and CD14 is recruited to the complex after binding to LPS [136].
Subsequent signal pathways activated by TLR4 can either be dependent on myeloid differentiation factor (MyD88), which occur early or independent of MyD88, which occur later and involve the adapters TIR-containing adapter molecule (TRIF) and TRIF related adapter molecule (TRAM) [140]. Early LPS signalling leads to the activation of NF-κB, IRF3 and MAPK pathways, which are mediated by MyD88 and the MyD88 adapter like protein (Mal). Following the phosphorylation of IL-1 associated kinase (IRAK), TNF-receptor associated factor (TRAF)-6 is activated which leads to the production of proinflammatory genes. The later response to LPS is mediated by TRIF and TRAM and results in the activation of TRAF-6 and TANK-binding protein (TBK)- 1 and finally NF-κB and IRF3 and the induction of cytokines, chemokines and other transcription factors [140]. Genes which are induced by activation of NF-κB include GM-CSF, TNF-α, IL-1β, IL-6 and IL-8 [141].
TLR2 is a receptor for a variety of microbial ligands, including lipoteichoic acids (LTAs) and peptidoglycans associated with gram-positive bacteria [142]. TLR2 combines TLR1 or TLR6 to initiate a signalling cascade. Activation of both TLR2 and TLR4 regulates several important proinflammatory neutrophil functions through the activation of the NF-κB pathway, and these include neutrophil activation, migration and
26 survival [130]. Exposure to purified LPS increases neutrophil expression of TLR2 and CD14 but does not change expression of TLR4. Upon stimulation with PAMPs including peptidoglycan, zymosan and araLAM (a component of Mycobacterium tuberculosis) neutrophils produce IL-8 and superoxide and also have increased phagocytosis [138].
Genetic polymorphisms of genes encoding TLR4 or TLR2 may modulate the inflammatory response to microbial products. Genetic variation in TLR2 has been reported to determine the susceptibility of a population of farmers to develop asthma and allergies [143]. The TLR4 (Asp299Gly) polymorphism is linked with a reduced response of PBMCs to LPS stimulation and a 4-fold higher prevalence of asthma [144]. There is also evidence that CD14 polymorphisms are associated with the development of asthma and atopy [145].
1.5.4.2 Nucleotide-Binding Oligomerisation Domain (NOD) Molecules
More recently, another class of PRRs has been identified, called nucleotide-binding oligomerisation domain (NOD) molecules, including NOD1 and NOD2. NODs are cytoplasmic surveillance proteins and may be activated following internalisation of bacteria by phagocytic cells [146]. NOD1 is important in the host response to gram- negative peptidoglycan. NOD1 activation has been shown to enhance the inflammatory response of TLRs, particularly proinflammatory cytokine release [147]. Genetic variation in NOD1 is associated with increased susceptibility to asthma [148].
1.5.5 Neutrophil Granules
Neutrophil granules differ in their contents and primary functions (Figure 1.5). Azurophilic granules contain proteins and peptides that are crucial for the killing and digestion of microbes. Azurophilic granules contain the MPO and three important serine proteinases: cathepsin G, neutrophil elastase and proteinase 3. These proteases degrade proteins including bacterial virulence factors [133]. Important antimicrobial proteins found in these granules include bactericidal/permeability increasing protein (BPI), which can permeabilise and kill gram negative bacteria, and defensins, which also kill gram negative and gram positive bacteria by causing permeabilisation of the bacterial
27 membrane. Defensins can also regulate the inflammatory response by binding to protease inhibitors such as α1-antitrypsin [86].
Specific granules participate in antimicrobial activities by mobilisation of their contents to either the phagosome or the extra cellular space. They contain many substances that include lactoferrin, lysosyme, hCAP-18, and neutrophil gelatinase associated lipocalin (NGAL) [149]. Gelatinase granules are primarily important as a source of matrix degrading enzymes and membrane receptors needed during neutrophil migration. They contain 3 matrix metalloproteinases including gelatinase (MMP-9), collagenase (MMP- 8) and leukolysin (MMP-25). Both specific and gelatinase granules also contain cytochrome b558, which is transferred to the phagosomal membrane upon neutrophil activation, where it participates in the respiratory burst [150].
NEWLY SYNTHESISED PREFORMED
Cytokines Azurophilic Granules e.g. IL-1β, TNF-α e.g. Elastase, Defensins, Myeloperoxidase, Cathepsin G Chemokines e.g. IL-8 Gelatinase Granules e.g. Gelatinases (including Growth Factors MMP-9) e.g. G-CSF, GM-CSF Specific Granules Reactive Oxygen Species e.g. Flavocytochrome b558 e.g. Superoxide
Figure 1.5 Contents of Neutrophil Granules
Secretory vesicles are crucial for the replenishment of membrane components, and these vesicles are needed at early stages of neutrophil activation. They are mobilised in response to a wide range of inflammatory stimuli, and contain central membrane receptors including CD11b/CD18 [151], and CD14 [152]. The mobilisation of secretory vesicles is associated with shedding of L-selectin and associated with firm adhesion of the neutrophil to the vascular endothelium [153].
28 1.5.5.1 Mechanisms of Degranulation
Degranulation into the phagolysosome or in the extracellular space involves fusion of the neutrophil granules to the plasma membrane, involving protein – protein interactions that dock a vesicle to its final destination [86]. Each granule differs in its ability for exocytosis. Secretory vesicles have the highest potential for extracellular release, followed by gelatinase granules, specific granules and azurophilic granules [154]. Stimulation of neutrophils with small amounts of fMLP in vitro results in rapid release of secretory vesicles, but not other granules [151], whereas stimulation of neutrophils with PMA results in complete release of gelatinase granules, moderate release of specific granules and low release of azurophilic granules [155].
The underlying signalling pathways leading to degranulation of neutrophils are not fully understood. However, the mobilisation of neutrophil granules has been linked to elevations in intracellular Ca2+ levels, the end result of the activation of a number of signalling cascades [150]. High concentrations of Ca2+ can activate annexin proteins that mediate vesicle aggregation and membrane fusion [156], as well as promoting interactions between SNAP receptor (SNARE) proteins [157]. SNAREs, consisting of vesicle and target-membrane-specific types (v- and t-SNAREs) are components of protein machinery that participate in vesicle budding and fusion. Several SNAREs have been identified in neutrophils, including vesicle associated membrane protein-2 (VAMP-2) and secretory carrier membrane protein (SCAMP), found in gelatinase and specific granules, syntaxin-4 and 6 which are located in the plasma membrane [158, 159], and SNAP-23 and 25 which are found in peroxidase-negative granules [158, 160].
1.5.5.2 Proteolytic Enzymes
Proteolytic enzymes play an essential role in remodelling and maintenance of the extracellular matrix (ECM). Proteolytic enzymes can remain active for long periods of time meaning that they can repetitively participate in enzymatic reactions. Proteases also influence essential physiologic functions such as angiogenesis, vasculogenesis, apoptosis and cell migration [161]. Neutrophil granules contain over 20 enzymes, however the serine protease neutrophil elastase (NE) and the gelatinase matrix
29 metalloproteinase (MMP)-9 are recognised to have the greatest potential to cause tissue destruction.
Both active NE and MMP-9 are increased in asthma [162, 163], and more recently it has been demonstrated that proteolytic activity in asthma is dependent on the inflammatory phenotype [164]. More specifically, Simpson et al demonstrated that subjects with eosinophilic asthma had higher levels of active MMP-9 compared to subjects with neutrophilic asthma and healthy controls. Neutrophilic asthma subjects had a higher level of total MMP-9, however this was bound to tissue inhibitor of metalloproteinase-1 (TIMP-1) and therefore inactive. NE activity was demonstrated in neutrophilic asthma, but not other asthma phenotypes or healthy controls [164].
Matrix Metalloproteinases (MMPs) are a family of zinc containing enzymes that are capable of degrading proteins of the extracellular matrix (ECM), and modulating chemokine and cytokine function. MMPs are divided into several subgroups including collagenases (MMP-1, -8, -13), gelatinases (MMP-2, -9), stromelysins (MMP-3, -10, - 11), matrilysin (MMP-7), elastase (MMP-12) and MT-MMPs (MMP-14, -15, -16, -17) [165]. MMPs and their inhibitors tissue inhibitors of metalloproteinases (TIMPs) play an important role in the pathogenesis of a number of inflammatory diseases including asthma. MMPs have the ability to influence the function and migration of inflammatory cells as well as matrix deposition and degradation. Upsets in the balance of MMPs/TIMPs may cause tissue injury (excess of MMPs) or increased tissue fibrosis (excess of TIMPs) [166]. MMP-9 has been shown to be enhanced in asthma patients, especially during exacerbations [167]. LPS and IL-8 have been shown to cause the release of MMP-9 in whole blood [168, 169].
MMP-9 is thought to play a key role in tissue remodeling and repair through the degradation of type IV collagen, the major component of basal membranes. MMP-9 is expressed from a large array of cells, including neutrophils [168] and eosinophils [170]. In neutrophils, this enzyme is thought to be synthesised during the cells maturation in the bone marrow and stored in specific granules until needed [87]. MMP-9 is released as an inactive pro-form, containing a pro-peptide that interacts with the zinc ion in the active site, keeping the enzyme inactive. MMP-9 is activated by proteolytic cleavage of
30 the pro-peptide domain by serine proteases and other MMPs such as MMP-2 or -3 [165]. Recent evidence shows that MMP-9 is also capable of being activated by free radicals [171] and bacterial proteinases [172]. MMP-9 is inhibited by TIMP-1, which binds to both active and pro-forms of MMP-9 to keep them inactive [173]. Interestingly, 10% of the MMP-9 is complexed with TIMP-1 when secreted from human neutrophils [173].
Neutrophil elastase (NE) is a serine protease that is found in the azurophilic granules of neutrophils. NE has a wide range of functions including the breakdown of elastin and collagen types I-IV, proteolyses surfactant proteins, and activation of MMPs [131]. The main function of NE is its antimicrobial effect against gram negative bacteria, however it is also important for effective neutrophil migration due to the activation of MMPs [174, 175]. NE can aggravate inflammation by increasing the release of IL-8 through stimulation of TLR4 [176].
Mucus hypersecretion is a frequent characteristic of asthma, with goblet cell hyperplasia and degranulation along with submucosal glands being responsible for the mucus production [177, 178]. Studies have shown that the release of active NE from neutrophils can stimulate goblet cell degranulation, and therefore mucus hypersecretion and possible airflow obstruction [177-179]. Activation of neutrophils causes translocation of NE from the azurophilic granules within the cells cytoplasm where it normally resides, to the cell surface where it is enzymatically active and can interact with the goblet cells [178].
1.5.6 Respiratory Burst
The respiratory burst involves the activation of NADPH-oxidase, which is an enzymatic phox phox phox complex composed of cytosolic (p40 , p47 , and p67 ) and flavocytochrome b558 phox phox which is made up of membrane proteins (p22 and gp91 ). Flavocytochrome b558 is located between the plasma membrane and the membrane of the specific granules, and it is incorporated into the phagocytic vacuole, where it pumps electrons from NADPH in the cytosol to oxygen in the vacuole [149]. When neutrophils are activated, p47phox is phosphorylated, which causes cytosolic components to migrate to the plasma membrane, where they are able to associate with flavocytochrome b558, assembling the
31 active oxidase [86]. Reactive oxygen species (ROS) are generated as a result of NADPH - oxidase activity, which produces superoxide (O2 ). Superoxide can be rapidly converted into hydrogen peroxide (H2O2) by the enzyme superoxide dismutase. Superoxide and hydrogen peroxide can also react to generate the highly reactive hydroxyl radical (HO•). In addition, myeloperoxidase (MPO), a constituent of the azurophilic granules, generates hyperchlorous acid (HOCl) from hydrogen peroxide [86, 133, 149].
NADPH-oxidase, through the formation of ROS, plays a fundamental role in the killing of microbes. Dysfunction of NADPH-oxidase occurs in patients with chronic granulomatous disease (CGD), who have profound immunodeficiency that makes these patients susceptible to bacterial and fungal infections [180]. This dysfunction of NADPH oxidase in CGD results from a genetic defect in any of the genes encoding the four phox subunits of the enzyme complex. Phagocytes from CGD patients fail to mount a respiratory burst, illustrating the importance of the respiratory burst in combating infections [181].
1.5.6.1 Reactive Oxygen Species
It is well known that exposure of the lung to reactive oxygen species (ROS) results in pulmonary injury. Recent findings demonstrate an expanded role for ROS, in that they can activate granule proteins, interact with various signalling cascades, and modulate neutrophil functions. Superoxide can activate granule proteins through the recruitment of K+ to the phagosome, allowing cationic proteases of the azurophilic granules such as neutrophil elastase (NE) and cathepsin G (CG) to go from highly organized intra- granule structures into solution where they can kill ingested microbes [182]. ROS can inhibit a variety of protein tyrosine phosphatases through oxidation of key residues, allowing the phosphorylation of other molecules to proceed [183]. ROS can disrupt intercellular tight junctions, and increase the permeability of the endothelial barrier via the phosphorylation of focal adhesion kinase in endothelial cells [184]. ROS also modulate neutrophil function by inducing apoptosis through a caspase-8 dependent manner [185].
32
1.5.7 Neutrophil Clearance and Death
Apoptosis is a genetically programmed mechanism of cell death, which plays an important role in embryonic development, cell turnover, and control of the immune response. Human neutrophils are produced at a rate of 1-2 x 1011 cells a day, and they survive in the circulation for about 24-36 hours [186]. After killing and digesting invading microbes, the neutrophils fate at the inflammatory site is to undergo apoptosis and be phagocytosed by macrophages (efferocytosis). The correct regulation of neutrophil apoptosis is crucial to maintain neutrophil numbers in the blood, as well as for the effective removal of invading pathogens and resolution of inflammation. Apoptosis allows inflammation to resolve, preventing neutrophils from a necrotic death where they would release their toxic contents into the surrounding tissue causing damage. Dysregulation of apoptosis could in fact lead to the persistence of neutrophils at the inflammatory site and the development of chronic inflammation.
Neutrophil apoptosis appears to occur randomly in vitro, and the rate of apoptosis is dependent on the time of culture, the cell density, the presence or absence of serum and any contact with artificial surfaces [187]. In vivo, this process may limit neutrophils destructive capability. Within a few minutes, neutrophil apoptosis results in irreversible chromatin condensation, nuclear collapse, cytosolic vacuolation and cell shrinkage. During this time the cell is unable to respond to agonists, is immobilised and inert [132]. Apoptotic neutrophils become instantly recognisable to alveolar macrophages, which results in cell removal via efferocytosis [188]. This process can induce changes in the activation phenotype of lung macrophages, suppressing the release of proinflammatory mediators [189].
Neutrophil apoptosis is an active process that can be modulated by various mediators. Defined inflammatory stimuli such as growth factors (e.g. GM-CSF and G-CSF) cytokines (e.g. IL-1 and IL-6), chemokines (e.g. IL-8), and even bacterial products (e.g. LPS) can delay neutrophil apoptosis [190]. Conversely, TNF-α and Fas-ligand (Fas-L) can increase the rate of neutrophil apoptosis [132]. Interestingly, certain pathogenic bacteria and viruses have strong effects on inducing neutrophil apoptosis, increasing the likelihood of these organisms to evade intracellular killing and inhibiting neutrophil
33 functions. Corticosteroids delay neutrophil apoptosis, thus increasing their survival time, and influencing the persistence of neutrophilic inflammation [40].
Mechanisms inducing neutrophil apoptosis involve the activation of cellular caspases, and can occur through two main pathways. The first is the death receptor (DR) pathway, where clustering of TNF and Fas-receptors activates the caspase cascade beginning with cleavage of pro-caspase 8 [191]. The second is an intrinsic pathway that involves mitochondrial cytochrome c and members of the Bcl-2 family, which forms and apoptosome activating caspase 9. Cell stress due to exposure to ROS, DNA damage or lack of growth factors can result in apoptosis, induced by the release of cytochrome c [192]. Activated caspase 8 and 9 can then activated caspase 3, which cleaves proteins essential of cell survival [190]. There is cross-talk between the two apoptosis pathways, for example, studies have shown that the activation of caspase 8 by ligation of DRs is not enough to trigger caspase-3 activation, and that activation of caspase-8 through this mechanism cleaves Bid, a protein of the Bcl-2 family, which amplifies the release of cytochrome c from the mitochondria [193].
1.5.8 Cytokine Synthesis
The neutrophil is both a target and a source of various proinflammatory cytokines (e.g. TNF-α and IL-1), chemokines (e.g. IL-8) and growth factors (e.g. GM-CSF and G- CSF), and hence has the ability to create a positive feedback loop on its own proinflammatory functions. These cytokines can amplify several neutrophil functions, including the generation of ROS, and chemokines can promote neutrophil migration to the inflammatory site. In addition, both cytokines and chemokines may act as priming agents for neutrophils.
Neutrophils were always thought to be devoid of transcriptional activity or protein synthesis, however recent evidence proves that neutrophils are an important source of newly synthesised cytokines and growth factors [194]. Cytokine synthesis by neutrophils occurs to a lower degree when compared to monocytes [195], however this is overshadowed in vivo by both the number of circulating neutrophils being 20 times that of monocytes, and the fact that neutrophils are the first to arrive at inflammatory sites. The production of cytokines by neutrophils is increased by inflammatory stimuli,
34 bacterial endotoxin the most potent of these [194]. The secretion of cytokines is varied and dependent on the agonist, and for some cytokine production, stimulation with more than one agonist is required, such as stimulation with interferon and LPS is needed for IL-12 production [195]. Cytokine expression from neutrophils can be modulated by T cell derived cytokines, positively for Th1 cytokines e.g. IFN and negatively by Th2 cytokines e.g. IL-4 and IL-13 [194].
TNF-α is a multifunctional cytokine that is produced mainly by monocytes and macrophages, but also by other airway cells including eosinophils and epithelial cells [196]. TNF-α is a homotrimer of 17kDa subunits, and belongs to the superfamily of membrane anchored and soluble cytokines involved in T cell mediated immunity. It’s potent proinflammatory effects are mainly due to its ability to upregulate cell adhesion molecules on the endothelium, thus promoting the neutrophil adherence and migration. Neutrophils can express TNF-α mRNA upon LPS and GM-CSF stimulation [197]. TNF-α is a priming agent for neutrophils that can markedly increase their capacity for phagocytosis, degranulation and respiratory burst. These effects are mediated by two TNF-α receptors: type A of 75kDa (p75) and type B of 55kDa (p55), and these two receptors activate different signalling pathways. p75 is mainly expressed on myeloid cells and p55 is mainly expressed on epithelioid cells [86]. TNF-α has been detected in the airways in both COPD and asthma [71].
IL-1β is a multifunctional cytokine that is highly proinflammatory in nature [198-201]. IL-1β acts synergistically with TNF-α, to activate proinflammatory responses in a wide range of cells and induces the expression of many effector proteins, for example chemokines, cytokines, MMPs, and nitric oxide synthase [202]. IL-1β is released as a 31 kDa cytokine that is activated via intracellular cleavage by caspase-1 to the 17 kDa form [203]. A variety of mediators induce the production of IL-1β, including PAMPs such as LPS, TNF-α, IFN-α, IFN-β and IL-1β itself. The receptors for IL-1 cytokines (including IL-1RI for IL-1β) are structurally related to TLRs, with identical signalling pathways activated upon ligand binding.
Oncostatin M (OSM) is a 28 kDa multifunctional cytokine that belongs to the IL-6 cytokine family. OSM is produced by activated T lymphocytes [204], monocytes [205] and neutrophils [206, 207]. Peripheral blood neutrophils have a preformed stock of
35 OSM that is rapidly mobilised by degranulating agents such as phorbol myristate acetate (PMA) and GM-CSF. In addition, neutrophils can produce OSM at several hours of stimulation e.g. with LPS, and this is dependent on OSM gene transcription [206]. OSM has many functions, and is thought to act predominantly as an anti- inflammatory mediator in the lung, playing an important role in wound repair [208]. OSM is a potent inhibitor of TNF-α release following LPS treatment in vivo [209]. In vitro, OSM can inhibit IL-1β induced expression of IL-8 and GM-CSF by synovial and lung fibroblasts [207]. OSM may affect the protease/antiprotease balance in the lungs. OSM is a potent inducer of TIMP-1 expression in human lung fibroblasts and synovial fibroblasts [210], and α1-antitrypsin and α1-proteinase inhibitor in lung derived epithelial cells. In contrast to the lung, OSM induces the production of MMP-1, MMP- 8, MMP-13, and MMP-14 in human chondrocytes [211].
1.5.9 Neutrophil Priming and Activation
Excessive neutrophil activation has the ability to cause severe tissue damage [212, 213]. Products of neutrophils can induce AHR, and also increased mucus secretion and airway smooth muscle responsiveness [66, 214]. It is well recognised that tissue injury in a variety of conditions, such as acute respiratory distress syndrome (ARDS), result from uncontrolled neutrophil activation and/or reduced apoptosis limiting the clearance of neutrophils from the inflammed site. For neutrophils to be able to degranulate or undergo respiratory burst activity they must first be primed by agents such as endotoxin, TNF-α, GM-CSF, or PAF so that the cell has the capability to undergo full receptor mediated activation [132]. This priming results in a range of functional changes including the upregulation of cell surface integrin expression, cell shape change, reduced cell deformability and increased capacity for the cell to release superoxide anions [215].
Primed neutrophils can respond to activating stimuli in an exaggerated fashion, which could lead to tissue injury. Neutrophil priming and activation are distinct events supporting a ‘two hit’ model for neutrophil mediated tissue damage [216]. Resting neutrophils have a low rate of superoxide production, which increases slightly upon exposure to parts of the bacterial cell wall such as fMLP [217]. However if the cell has been exposed previously to agents such as PAF, the production of superoxide to the
36 same amount of fMLP is greatly increased [218]. Low doses of endotoxin have also been shown to induce neutrophil priming, allowing the neutrophils to adhere to the pulmonary endothelium, but cause no tissue damage at this stage. In the absence of a second stimulus the neutrophils detached from the endothelium after 12 hours. However, exposure of the cell to a second stimulus during the time that it was adherent was able to cause significant tissue damage [216]. This demonstrates that primed neutrophils adhere to the endothelium but do not damage it in the absence of a second stimulus, and neutrophils that are adherent to the endothelium and further activated can cause significant damage.
It is likely that neutrophil priming and activation are both required for efficient clearance of bacterial infection, and that this response may not lead to uncontrolled neutrophil activation and tissue damage, rather act as a control point preventing inappropriate cell activation. Neutrophil mediated tissue damage may also reflect an imbalance between appropriate degranulation and respiratory burst within the phagolysosome and inappropriate extracellular release of granule contents and ROS [132]. Priming as a control mechanism is supported by the fact that primed neutrophils can also be ‘deprimed’ and simply return to the circulation, in fact, the cell is able to undergo a full cycle of priming, depriming and repriming [219].
1.6 Neutrophil Gene Expression
Neutrophils have long been considered to be phagocytes whose main purpose is to engulf and degrade microorganisms. However, recent microarray studies have provided substantial evidence that neutrophils are capable of extensive gene expression changes that are important in the regulation of many neutrophil functions, as well as modulation of the immune response. A wide range of genes are expressed in unstimulated neutrophils, and this gene profile is dramatically changed in response to bacteria [220- 222]. Alterations in neutrophil gene expression have been reported in response to LPS exposure in vitro [220, 223-225] and in vivo [226], as well as during phagocytosis [227], and apoptosis [228-231]. Genes that are altered during exposure to bacteria include cytokines, receptors, genes involved in host defense, apoptosis-related genes, transcription factors, and chromatin-remodeling genes [220].
37 Neutrophil gene expression profiles are not only altered with stimulation due to exposure to bacteria and other agents, but gene expression also changes in disease. There is an up-regulation of proinflammatory genes (e.g. IL-8, CXCL1, CD14, TLR5, CCR1, and calgranulins A and B) and a down-regulation of anti-inflammatory genes (e.g. those of the TGF-β pathway) at rest in neutrophils isolated from subjects with chronic granulomatous disease (CGD) [232].
Studies by Kobayashi et al [230, 231] have demonstrated that neutrophil phagocytosis induces both an early response of enhanced proinflammatory activity, and a late response promoting apoptosis and resolution of inflammation. These responses are dependent on changes in gene transcription. The early response involves upregulation of proinflammatory genes including chemokines and cytokines that recruit macrophages, T cells and neutrophils [e.g. monocyte chemotactic protein (MCP)-1, macrophage inhibitory protein (MIP)-1α, MIP-1β, oncostatin M and IL-1β]. The late transcriptional response promotes neutrophil apoptosis and efferocytosis by macrophages. This stage induces the up-regulation of pro-apoptotic genes [e.g. TNF-α, TRAIL, TNFR1, TRAILR, caspase-1, and BAX]. In conjunction with an up-regulation of pro-apoptotic genes was a down-regulation of proinflammatory capacity, specifically receptors for inflammatory mediators [e.g. IL-8Rα, IL-8Rβ, IL-9R, IL-10Rα, IL-13Rα-1, IL-15Rα, IL-17R, TLR1, and TLR6].
An elegant study by Coldren and colleagues [226] demonstrated dramatic changes in gene expression of air space neutrophils compared to circulating neutrophils after bronchoscopic instillation of LPS. The altered genes included those related to inflammation and chemotaxis, as well as antiapoptotic and IKK-activating pathways. As well as alterations in gene expression, air space neutrophils induced functional changes, which included increased superoxide release, diminished apoptosis, and decreased chemotaxis in response to IL-8. This study demonstrated profound functional and gene expression changes that occur upon neutrophil migration to the lung due to endotoxin challenge [226].
Clearly active regulation of neutrophil gene expression plays an important role in determining the cells fate and function. The transcriptional program of neutrophils is
38 modified not only by exposure to infectious agents, but also through their migration to separate bodily compartments such as the airways.
1.7 Contribution of Neutrophils to the Adaptive Immune Response
Neutrophils make important contributions to adaptive immunity, through the activation and programming of antigen presenting cells (APCs) such as monocytes and dendritic cells (DCs) [233]. Neutrophil products can attract monocytes, T lymphocytes and DCs, and can influence the differentiation of monocytes into the pro- or anti-inflammatory state [234-236]. Neutrophil proteases, including cathepsin G and elastase, activate prochemerin to generate chemerin, which is a chemoattractant for both immature and plasmacytoid DCs [237]. Furthermore, neutrophils produce both interferon-γ (IFN-γ) [238] which activates macrophages and drives the differentiation of T cells, and TNF- related ligand B-lymphocyte stimulator (BLyS), which influences the differentiation and maturation of B cells [239].
Proinflammatory cytokines such as TNF-α are released after activation of neutrophils via stimulation of TLRs including TLR4 by its agonist LPS. These cytokines also play an important role in the recruitment of cells of the adaptive immune response. Therefore the function of TLRs is not only important in regulating the innate immune response, but also the development of the adaptive immune response.
Just as neutrophils can influence the differentiation of APCs at sites of inflammation, the adaptive immune system is able to control the rate of neutrophil development in the bone marrow. IL-23 released by macrophages triggers T cell subsets to release IL-17, which regulates the release of G-CSF from stromal cells [240]. G-CSF is an essential regulator of neutrophil production through stimulation of the release of MMP-9 which increases neutrophil mobilisation, and the suppression of stromal cell expression of CXCL12 which retains neutrophils in the bone marrow [241].
1.8 Neutrophil Function in Ageing
The innate immune system experiences changes with advancing age, and this is termed immune senescence. Clinically, elderly people have an impaired ability to respond to
39 infection, and as a result of this, aged individuals have a greater risk of contracting more severe infections that can last longer. Contributing to this is a decline in neutrophil function with age; particularly an impairment in the production of ROS and intracellular signalling, as well as altered rates of neutrophil apoptosis after exposure to inflammatory mediators.
With increasing age, there is a decrease in the production of ROS by neutrophils after stimulation with various agents, including fMLP, GM-CSF, and LPS [242, 243]. Impaired intracellular signalling is implicated in the reduced production of ROS from older individuals, specifically there is an impaired calcium influx [244, 245]. Actin polymerization is also significantly decreased after stimulation of neutrophils with fMLP or PMA in aged subjects compared to young subjects [246]. These differences were associated with altered cell surface marker expression [247].
Variations in neutrophil superoxide production with ageing are also dependent on the stimulus, implicating different pathways of neutrophil activation. There is suggestion that gram-positive and gram-negative bacteria have differential effects, and that this is possibly due to the fact that they function through different TLRs. Impaired functioning and expression of TLRs in ageing has been reported to occur in mouse macrophages [248]. Fulop and colleagues [249] investigated the presence of TLR2 and TLR4 in neutrophil lipid rafts, where they demonstrated that TLR4 was increased in unstimulated raft and non-raft fractions in older subjects compared to their younger counterparts. Furthermore, LPS was shown to increase the expression of TLR4 in neutrophils obtained from young subjects, in both raft and non-raft fractions, whereas there was no reorganisation of TLR4 after LPS stimulation of neutrophils from older subjects [249]. This is consistent with the inflammation in ageing theory which suggests that there is a low grade of inflammation present with increasing age [250], and the slightly stimulated state of neutrophils demonstrated from aged individuals [251].
Neutrophils from elderly subjects are not able to be primed efficiently with GM-CSF [252], which may affect their activity at inflammatory sites. The lifespan of neutrophils can be extended by exposure to a variety of factors, including proinflammatory cytokines such as IL-8 and GM-CSF, and bacterial products such as LPS. Neutrophils
40 from aged individuals cannot be rescued from apoptosis by these agents, specifically GM-CSF, G-CSF, LPS, IL-1, IL-6 and steroids [253, 254].
1.9 Neutrophilic Asthma
Subjects with neutrophilic asthma experience asthma symptoms and have airways hyperresponsiveness which is associated with the presence of airway neutrophils of greater than 61% [9]. Between 10 and 30% of adults with stable asthma have neutrophilic inflammation [9, 255, 256]. Neutrophilic asthma is associated with an increase in age [9, 255, 256] and less severe airway hyperreactivity. However, subjects with neutrophilic asthma are similar in terms of gender, atopy, smoking and lung function in comparison to other asthma subtypes [9].
Even though airway inflammation involving neutrophils occurs with increasing age, evidence suggests that both eosinophils and neutrophils are elevated in difficult asthma in children [257]. It has been recently suggested that neutrophilic airway inflammation may contribute to childhood asthma. Supporting evidence for a role for neutrophils in the development of asthma comes from observations of a low prevalence of asthma in children with autoimmune neutropaenia in contrast to healthy control children [258]. Children with mild to moderate asthma also had increased neutrophils and neutrophil proteases compared to children with intermittent asthma [259].
1.9.1 Mechanisms of Neutrophilic Asthma
A fundamental mechanism underlying neutrophilic airway inflammation is activation of innate immunity. Simpson and colleagues [255] have shown increases in several key steps in the innate immune activation pathway in neutrophilic asthma (Figure 1.6). The innate immune receptors including TLR2, TLR4, CD14 and surfactant protein A are upregulated, and there is an increase in the level of the neutrophil chemoattractant IL-8 and proinflammatory cytokine IL-1β. Activation of this pathway in neutrophilic asthma is thought to be mediated by high levels of airway endotoxin and bacterial colonisation, particularly Haemophilus influenzae [255]. In this study, airway endotoxin levels in subjects with asthma showed a significant negative correlation with airway obstruction
(FEV1% predicted and FEV1/FVC) and significant positive correlation with sputum
41 neutrophils and sputum IL-8 protein levels [255], implicating the stimulation of neutrophilic airway inflammation by endotoxin in the worsening of asthma. Therefore, neutrophilic asthma may be driven by dysfunction of distinct innate immune mechanisms.
Important neutrophil proteases are altered in neutrophilic asthma. Neutrophil elastase activity is increased in neutrophilic asthma, and present in more than 40% of subjects compared to just 5% of those with eosinophilic asthma, and this is in contrast to subjects with paucigranulocytic asthma and healthy controls where elastase activity is not detected [164]. The primary inhibitor of neutrophil elastase, α1-AT, is increased in the airways of those with neutrophilic asthma, however this appears insufficient to control elastase activity. The level of total MMP-9 and the MMP-9 inhibitor TIMP-1 measured in sputum supernatant are significantly increased in neutrophilic asthma compared to other asthma subtypes [260, 261]. However, there is decreased activity of MMP-9 in neutrophilic asthma, and MMP-9 activity is higher in eosinophilic asthma [260].
42
NEUTROPHILIC ASTHMA
INNATE PATHWAY Endotoxin
Trigger
Increased Receptors
Receptor TLR2 CD14 TLR4 SPA Signal Transduction MyD88
Transcription NF-κB Increased IL-8, IL-1β
Increased Inflammatory Cytokines NEUTROPHILS
Increased NE NEUTROPHILS
Airway Mediators Obstruction
Asthma
Figure 1.6 Innate Immune Activation Pathway in Neutrophilic Asthma
43 1.9.2 Triggers of Neutrophilic Airway Inflammation
1.9.2.1 Endotoxin
The main agent that is thought to induce the neutrophilic response seen in occupational asthma is endotoxin [44]. Endotoxin and its derivative lipopolysaccharide (LPS) are strong proinflammatory outer membrane components of gram-negative bacteria, which activate innate immunity [139, 262]. Heavy brief exposure to endotoxin is a well-known cause of acute illness with a resulting fever, cough and dyspnea. Exposure to lower concentrations over an extended period of time is implicated in the development of lung diseases such as chronic bronchitis, emphysema and asthma, but also tolerance of the acute effects [139]. Endotoxin is present in house dust, and is known to exacerbate existing asthma. Inhalation of LPS has been associated with neutrophilic inflammation with increases in IL-8 in both normal and asthmatic subjects [263].
A great deal of evidence links exposure to endotoxin to the development of asthma. Exposure to endotoxin in infancy has been thought to possibly have protective effects against the development of asthma by enhancement of the Th-1 responses, away from the Th-2 response associated with asthma and atopy. Interestingly, growing up on a farm, and in particular having contact with farm animals, has been related to a substantial decrease in risk for allergic diseases including asthma [264]. This effect is due to elevated levels of endotoxin found in homes of farmers and children with regular contact with livestock rather than non-farm children without animal contact. This view is controversial, as an increase of the risk of repeated wheeze in the first year of life has been reported in children that have early exposure to endotoxin [265], which contradicts the protective role.
Neutrophils are known to respond to LPS by production of reactive oxygen intermediates, lipid mediator and cytokine release, adhesion and phagocytosis [223, 224]. In vitro, stimulation of neutrophils with TLR agonists causes neutrophil activation, measured by upregulation of CD11b, cytokine generation and enhanced cell survival [266].
44 1.9.2.2 Respiratory Viruses
Exacerbations of asthma are frequently caused by viral infection, including influenza, RSV and rhinovirus. Asthma exacerbations in children are predominantly caused by viral infection, which accounts for up to 85% of cases. The most common viruses isolated from childhood asthma exacerbations are rhinovirus and respiratory syncitial virus (RSV) [267]. RSV infection is thought to play a role in the development of asthma, and accounts for more than 70% of all cases of infantile bronchiolitis [268].
Neutrophils play a prominent role in the innate immune response to viral infection, and these cells are recruited early due to the release of IL-8 from infected airway epithelial cells. Sputum neutrophils are increased at day 4 of a natural cold, and this correlates with the level of sputum IL-8 [269]. There is an increase in IL-8 in nasal lavage from children with natural colds. There were elevated levels of IL-8 and increased numbers of neutrophils in nasal aspirates from asthmatic children that were experimentally induced with a viral infection and, the level of neutrophil myeloperoxidase was correlated with the severity of symptoms [270].
Respiratory viruses can activate the innate immune pathway via the stimulation of TLRs. RSV proteins can stimulate TLR4 inducing the production IL-6 [271]. Interestingly, both TLR4 [272] and SP-A [273] knock out mice have delayed clearance of RSV from the airways upon infection. In the SP-A-/- mice, this delayed clearance was associated with a greater infiltration of neutrophils and an increased level of TNF-α and IL-6 in the mouse lung.
1.9.2.3 Air Pollution
Exposure to air pollution is associated with airway inflammation, where both particulate matter and ozone are of considerable importance. The inhalation of particulate matter from air pollution can cause asthma exacerbations. Inhalation of larger size particles
(PM2.5-10) induces airway inflammation, with increases in the infiltration of neutrophils. Components of ambient air pollution can stimulate TLR2 and TLR4, indicating that bacteria were associated with the particles, including LPS [274]. In alveolar macrophages, cytokine production in response to exposure to bacteria found in
45 particulate matter was inhibited by blocking CD14 and TLR4 [274, 275]. Airway epithelial cells produce IL-8 and undergo oxidative stress in response to exposure to particulate matter in vitro [276, 277]. Diesel exhaust particles promote neutrophil airway inflammation, neutrophil mobilisation from the bone marrow, and upregulation of adhesion molecules and IL-8 [278].
Ozone is a powerful oxidant and major component of urban smog that has been reported to exacerbate pre-existing asthma. Inhalation of the irritant gas ozone causes weakened lung function and tissue damage [279]. Increased neutrophil counts have been demonstrated following ozone exposure, in bronchoalveolar lavage fluid [280], and induced sputum [279, 281]. Inflammation that is caused by inhalation of diesel exhaust is further enhanced by ozone exposure. This includes a potentiation of neutrophils infiltration and activation in the airways [282].
1.10 Neutrophils and Corticosteroids
Existing treatments for asthma; e.g. corticosteroids (CS); are based on alleviating the allergic/eosinophilic response, and whilst they are effective at targeting this type of inflammation, they appear relatively ineffective in non-eosinophilic variants of the disease [33, 34]. In bronchial biopsies of moderate to severe asthmatics, corticosteroids have been shown to reduce eosinophil numbers and chemokine expression, specifically eotaxin, MCP-3 and MCP-4, whilst increasing neutrophil numbers and chemokine expression, specifically IL-8, and IP-10 [283]. In vitro experiments on the effect of corticosteroids on the survival of eosinophils and neutrophils show opposite effects. Corticosteroids induce eosinophil apoptosis, but delay neutrophil apoptosis [30, 40]. Therefore corticosteroids have the potential to worsen neutrophilic inflammation through increase in cytokine expression and neutrophil survival in the airways.
46 1.11 Aims & Hypotheses
Neutrophils are important effector cells of the innate immune system. Neutrophils are the first to migrate to inflammatory sites, where they contain and eliminate pathogenic microorganisms. Neutrophils also release cytokines and chemokines that initiate and amplify inflammatory responses. The mechanisms of neutrophilic asthma remain largely unknown; however activation of the innate immune response is implicated. This thesis examines innate immune responses of airway and circulating neutrophils, with a focus on neutrophilic asthma. Innate immune neutrophil activation consists of the release of preformed granule associated mediators such as NE, MMP-9 and OSM, new gene transcription and release of inflammatory cytokines such as IL-8, IL-1β and TNF- α, and new gene transcription of TLR2 & TLR4 which serve to amplify neutrophil responses. I hypothesise that dysregulation of innate immune neutrophil responses occurs with ageing and airway disease, particularly neutrophilic asthma and COPD.
1.12 Specific Hypotheses
¾ Neutrophils are activated by LPS, IL-8 or IL-18 to increase the release of the neutrophil proteases (MMP-9 and NE), cytokines (TNF-α) and chemokines (IL- 8) (Chapter 3). ¾ LPS increases the production of innate immune mediators from both airway and circulating neutrophils (Chapter 4). ¾ Airway and circulating neutrophil activation is altered with age (Chapter 5). ¾ Airway and circulating neutrophil activation is altered with airway disease, including asthma and COPD (Chapter 6). ¾ Airway and circulating neutrophil activation is altered with the inflammatory subtype of asthma (Chapter 7). ¾ Differential gene expression profiles will distinguish between neutrophilic and eosinophilic asthma subtypes, and implicate systemic neutrophil activation in the pathogenesis of neutrophilic asthma (Chapter 8).
47 Chapter 2: Materials and Methods
This chapter outlines methods used in this thesis and is divided into 5 sections: 1) Clinical Information 2) Cell Isolation 3) Assays 4) Molecular Methods 5) Data Analysis
2.1 Clinical Information
2.1.1 Collection of Clinical Information
Subjects were asked about their allergy history, respiratory symptoms including rhinitis specific symptoms, and medication use (See Appendix A). Subjects were excluded if they had a course of antibiotics or had increased their asthma medication to include a dose of oral steroids within one month prior to their visit. Clinical asthma pattern was determined in accordance with the GINA guidelines and subjects with chronic obstructive pulmonary disease (COPD) were assessed using the GOLD criteria. An induced sputum and blood sample were collected from each patient.
2.1.2 Spirometry
Baseline spirometry was measured using an electronic spirometer (Minato Autospiro AS-600 Minato Medical Science Co Ltd, Osaka Japan or KoKo K313100 PDS Instrumentation, Inc, Louisville CO USA) in accordance with the American Thoracic Society (ATS) guidelines [284]. Room temperature, barometric pressure and humidity were recorded and the spirometer was calibrated on a daily basis.
FEV1 and FVC were measured on each subject from a forced expiratory flow volume curve and predicted FEV1 and FVC measurements were calculated using the Knudson values [285]. Greater than 99% of all infectious particles were filtered with a bactericidal spirometry filter. Each subject was asked to use a nose clip and was seated at the time of the procedure. After tidal breathing, the subject inhaled to total lung capacity and then was asked to forcefully exhale to residual volume (minimum of 6
48 seconds). A minimum of 3 tests was performed until there were 2 acceptable results (within 200mL).
2.1.3 Saline Challenge and Sputum Induction
A sputum induction was carried out in subjects with a baseline FEV1 > 1.0L. If a subject had FEV1 > 1.0L but < 1.2L sputum induction was performed with normal (0.9%) saline. For subjects with FEV1>1.2l, sputum induction was performed with 4.5% hypertonic saline. To prevent contamination of the sputum sample with squamous cells, the subjects were asked to rinse their mouth with water before the procedure. A nose clip was applied and hypertonic saline (4.5%, room temperature) was administered using a mouthpiece and two-way valve connected to a high output ultrasonic nebuliser (DeVilbiss 2000, Oregon, PA, USA). Subjects inhaled the saline for 30 seconds first, and then doubling time periods of 1 minute, 2 minutes, 4 minutes. FEV1 was measured at 1 minute after each nebulisation period and the percentage fall of FEV1 from baseline was recorded. At each break the subject was asked to clear their throat, cough and transfer any sample into a sterile container. After 15.5 cumulative minutes on the nebulised saline, or at the subject’s request, the induction was stopped. If there was a fall in FEV1 greater than 15% of baseline during the induction, the bronchodilator salbutamol (β2-agonist, 200μg, Allen and Hanbury’s Melbourne, VIC, Australia) was given using a pressurised metered dose inhaler (pMDI) and valved holding chamber. Only if the subject’s lung function returned to normal (within 10% of baseline) could the test proceed. To determine the amount of saline supplied to the subject the nebuliser cup and tubing was weighed before and after the induction. Subjects undergoing combined hypertonic saline challenge and sputum induction were required to withhold their medications (Table 2.1). Airway hyperresponsiveness was determined by the dose of saline required to achieve a 15% fall in FEV1 (PD15) from baseline.
49 Table 2.1 Trade Names of Medication Withheld for 4.5% Saline Challenge 6 HOURS 12 HOURS 24 HOURS Ventolin Austyn Serevent Respolin Neulin Seretide Airomir Oxis Asmol Foradile Bricanyl Singulair Atrovent Theo-Dur, Neulin SR Atrovent Forte Slo-bid Combivent Symbicort Intal/Intal Forte Spiriva Tilade
Blood Collection 50mL of blood was collected from seated subjects into sodium citrate tubes by venepuncture.
2.1.4 Allergy Skin Prick Testing
A skin prick test was used to determine whether the subject was atopic or non-atopic. A positive allergy reaction was measured as the presence of at least one immediate reaction (weal>3mm) carried out using a 1 in 10 dilution of 5 common allergens (Bayer Australia Ltd, Pymble, NSW, Australia). These included Aspergilllus fumigatus, Alternaria tenius, Dust Mite (Dermatophagoides pteronyssinus), Cockroach and Grass mix (including Kentucky Blue grass, Cocksfoot, Red Top, Timothy, Sweet Vernal, Meadow fescue and Perennial Rye grass). Control solutions included histamine (10mg/mL, positive control) and saline (negative control). Subjects withheld antihistamine medications for their duration of action before testing.
50 2.1.5 Ethics
The Hunter Area Research Ethics Committee and the University of Newcastle Research Ethics Committee approved this study. Written informed consent was obtained from eligible participants who agreed to participate.
2.2 Cell Isolation
2.2.1 Isolation of Sputum Neutrophils
2.2.1.1 Induced Sputum Processing
All sputum samples were processed as soon as they were collected. Mucus plugs were selected from saliva and dispersed with 0.1% dithiothreitol (DTT, Calbiochem, La Jolla Ca USA) as described previously [286]. For every mL of sputum, 4mL of DTT was added. The tube was capped and placed on a rotating mixer (MACS mixer, Miltenyi Biotec, Gladbach, Germany) for 30 minutes at room temperature, to ensure optimal cell dispersion. Cells were washed with the same volume phosphate buffered saline (PBS) which was added (4 times the volume of sputum) and the suspension was filtered (60μm, Millipore, Australia). A leukocyte total cell count (TCC) and cell viability (trypan blue exclusion) were performed using a haemocytometer. At this stage, the sample was divided into two portions, 0.5mL was used for the preparation of cytospins for differential cell counts and the remainder of the sample used for the isolation of neutrophils via magnetic cell separation as described below. Both suspensions were centrifuged at 400g for 10 minutes at 4˚C. The supernatant from both portions was aliquoted and stored at –80˚C and the cell pellet for the cytospins was resuspended to a concentration of 1 x 106 cells/mL using phosphate buffered saline (PBS) and the slides were prepared from the resuspended cell pellet.
2.2.1.2 Sputum Differential Cell Count
Cytospins were fixed in methanol and stained with May and Grunwald stain and subsequently visualised with Giemsa. 400 non-squamous cells were counted, with the squamous cell proportion recorded separately. Cells were identified by their morphology and the differential cell count was expressed as a percentage of non-
51 squamous cells. Chromotrope 2R (C2R) staining was also performed to confirm the presence or absence of eosinophils.
2.2.2 Sputum Cell Isolation via Magnetic Cell Separation
Sputum cells were resuspended in 50uL of cold PBS (pH 7.3) supplemented with 0.1% bovine serum albumin (BSA) and 2mM EDTA (MACS buffer), and incubated with CD16 microbeads (60uL per 50 x 106cells) at 4˚C for 30 minutes. A LD column (Miltenyi Biotec, Gladbach, Germany) was prepared during this time by placing it in an LS column adapter within the magnetic field of the VarioMACS System (Miltenyi Biotec, Gladbach, Germany) and washing it with 2mL of MACS buffer. A pre- separation filter (30μm, Miltenyi Biotec, Gladbach, Germany) was rinsed with MACS buffer and placed on top of the column. After binding of the magnetic beads, the sputum cell sample volume was adjusted to 1mL with cold MACS buffer and applied to the pre- separation filter/LD column. The CD16- cell fraction was eluted with 2 x 1mL of MACS buffer. The LD column was removed from the magnetic field and the CD16+ neutrophil enriched cell fraction was eluted with 2 x 3mL of MACS buffer. Cytospins were prepared for the cell fractions, stained (May-Grunwald Giemsa) and a differential cell count obtained from 400 cells, to assess cell purity. Sputum neutrophil viability was assessed by trypan blue exclusion and found to have a mean of 67%.
2.2.3 Isolation of Peripheral Blood Granulocytes
2.2.3.1 Percoll Density Gradient
Peripheral blood (50mL) was anticoagulated by collection in sodium citrate tubes. After centrifugation at 150g for 20 minutes the platelet rich plasma was removed. The remaining pellet was diluted to 100mL with 1 x hanks buffered salt solution (HBSS, Invitrogen, Carlsbad, CA, USA), overlayed onto 3 x 15mL lots of 65% Percoll (65% Percoll, Amersham, 10% 10 x HBSS, Invitrogen, and distilled water, pH 7.3, 300mOsm). The mixture was centrifuged (400g, 30 minutes, no brake) to form 3 layers, HBSS, Percoll, and red blood cell/granulocyte pellet, with the mononuclear cells located Percoll interface. The HBSS, Percoll and monouclear cells located at the Percoll interface were removed leaving the red blood cell/granulocyte pellet. Red blood cells were lysed with isotonic ammonium chloride (15 minutes, ice cold) and centrifuged
52 (275g, 4˚C, 6 minutes). The cell pellets were resuspended in 1 x HBSS to wash, centrifuged (275g, 4˚C, 6 minutes) and this process was repeated to obtain a clean granulocyte pellet. Granulocyte viability was assessed by trypan blue exclusion and found to have a mean 99.6%.
2.2.3.2 Magnetic Cell Separation
The granulocyte pellet was resuspended in 100-200μL cold MACS buffer, and incubated with CD16 microbeads (60uL per 50 x 106cells) at 4˚C for 30 minutes. A LS column (Miltenyi Biotec, Gladbach, Germany) was prepared during this time by placing it an LS column adapter within the magnetic field of the VarioMACS System (Miltenyi Biotec, Gladbach, Germany) and washing it with 2mL of MACS buffer. A pre- separation filter (30μm Miltenyi Biotec, Gladbach, Germany) was rinsed with MACS buffer and placed on top of the column. The cells were then applied to the pre- separation filter/LS column, and the CD16- eosinophil enriched cell fraction was eluted with 3 x 3mL of MACS buffer. The LS column was removed from the magnetic field and the CD16+ neutrophil enriched cell fraction was eluted with 5mL of MACS buffer. The cell fractions were centrifuged (400g, 10 minutes, 4˚C), the pellet resuspended in 0.5-1mL of phenol red free RPMI 1640 (1% FCS) and a TCC was performed to assess cell viability (trypan blue exclusion). Cytospins were prepared for each cell fraction, they were stained (May-Grunwald Giemsa) and a differential cell count obtained from 400 cells, to assess cell purity. Blood neutrophil viability was assessed by trypan blue exclusion and found to have a mean 98%.
2.2.3.3 Cell Culture and Stimulation
Cell pellets were resuspended in phenol red free RPMI 1640 (Invitrogen, Carlsbad, CA, USA) with 10mM HEPES, 1% heat inactivated fetal calf serum and antibiotics (Penicillin/Streptomycin). The cells were cultured at concentrations of 1 x 105 cells/mL or 1 x 106 cells/mL with or without LPS (100ng/mL, E.Coli LPS, Sigma,) or IL-8 (10, 100, 200ng/mL, R & D Systems, Minneapolis MN USA) or IL-18 (10, 100ng/mL R &
D Systems, Minneapolis MN USA) at 37˚C (5% CO2). Cell cultures were collected at the maximum time of 24 hours, and cell pellets (in 350μL RLT buffer, QIAGEN, Hilden, Germany) and cell free supernatants were stored at -80˚C.
53 2.3 Assays
2.3.1 Interleukin-8 (IL-8)
IL-8 protein levels were assessed using the R & D systems (Minneapolis MN USA) Human DuoSet ELISA. This kit contains the basic components needed for the development of a sandwich ELISA to measure natural and recombinant IL-8. The standard curve for this kit ranged from 31.2pg/mL to 2000pg/mL. Assay sensitivity was determined previously to be 5.6pg/mL for the IL-8 DuoSet [9].
2.3.2 Interleukin-1β (IL-1β)
IL-1β protein levels were assessed using the R & D systems (Minneapolis MN USA) Human DuoSet ELISA. This kit contains the basic components needed for the development of a sandwich ELISA to measure natural and recombinant IL-1β. The standard curve for this kit ranged from 7.8pg/mL to 250pg/mL. Assay sensitivity was previously determined to be 1.6pg/mL for the IL-1β DuoSet [287].
2.3.3 Tumor Necrosis Factor-α (TNF-α)
TNF-α protein levels were assessed using the R & D systems (Minneapolis MN USA) Human DuoSet ELISA. This kit contains the basic components needed for the development of a sandwich ELISA to measure natural and recombinant TNF-α. The standard curve for this kit ranged from 31.3pg/mL to 1000pg/mL. Assay sensitivity was previously determined to be 6.8pg/mL for the TNF-α DuoSet [287].
2.3.4 Oncostatin M (OSM)
OSM protein levels were assessed using the R & D Systems (Minneapolis MN USA) Human DuoSet ELISA. These kits contain the basic components for the development of a sandwich ELISA to measure natural and recombinant mediator OSM. The standard curve for this kit ranged from 31.2pg/mL to 2000pg/mL Assay sensitivity of the OSM DuoSet was 1.6pg/mL.
54 2.3.5 Neutrophil Elastase
Levels of free neutrophil elastase were measured by a colourmetric assay using a chromogenic substrate specific for human neutrophil elastase, n-methoxysuccinyl-l- alanyl-prolyl-l-valyl-p-nitroanilide (Sigma, St Louis, MO, USA) as described previously [288]. The substrate was reconstituted in 1-methyl-2-pyrrolidinone (Sigma- Aldrich, Australia), with further working dilutions made up with 0.2M Tris buffer. Human neutrophil elastase (Calbiochem, La Jolla, CA, USA) was used for the construction of a standard reference curve with a range of 39ng/mL to 2500ng/mL. Cell culture supernatants were reacted with the elastase substrate at a concentration of 1mM and the absorbance was read at 405nm in 10-minute intervals for up to 1.5 hours. It was determined that the fetal calf serum (FCS) in the cell culture media had an effect on the elastase standard curve (Figure 2.1) and therefore all samples were compared to the RPMI curve.
2 1.8 1.6 y = 0.0008x - 0.0051 R2 = 0.9991 1.4 1.2 1 O.D. 0.8 y = 0.0004x + 0.0132 2 0.6 R = 0.9996 0.4 0.2 0 0 500 1000 1500 2000 2500 3000 Active NE (ng/mL)
0.2M Tris RPMI 1% FCS Linear (0.2M Tris) Linear (RPMI 1% FCS)
Figure 2.1 Neutrophil Elastase Standard Curve
2.3.6 Matrix Metalloproteinase-9 (MMP-9)
Total and Active MMP-9 was assessed using the R & D systems (Minneapolis MN USA) Fluorokine E active MMP-9 assay. The standard curve for this assay ranges from
55 0.25ng/mL to 16ng/ml. A monoclonal antibody that captures all three forms of MMP-9 (92, 82 and 65kDa) is used in this assay to measure human active MMP-9. A fluorogenic substrate was added once the sample has bound to the antibody and the resulting fluorescent signal was measured by a fluorometer (FLUOstar Optima, BMG Labtech, VIC Australia). This kit can also measure total MMP-9 by the addition of an activating reagent (4-aminophenylmercuric, APMA) to the sample wells, as APMA activates all forms of MMP-9. The sensitivity of this assay is reported by the manufacturer to be 0.005ng/mL.
2.3.7 Zymography of MMP-9 Activity
Samples were mixed with 2x sample buffer (0.125M Tris-Cl pH 6.8, 4% SDS, 20% glycerol, 0.04% bromophenol blue) and were separated on 10% SDS PAGE gel containing 10% gelatin Type A (Sigma St Louis MO USA). Samples were compared to known reference standards for MMP-9 and MMP-2 (Chemicon International Inc., Temecula, Ca USA). Gels were then incubated in 2.5% Triton-X 100 for 45 minutes and then in development buffer (0.05M Tris-Cl pH 8.8, 5mM CaCl2, 0.02% NaN3) for 18 hours at 37˚C. 0.1% Coomassie Brilliant Blue-250 (Bio-Rad, Reagents Park, NSW Australia) was used to stain the gels and subsequently 45% methanol, 10% acetic acid was used to destain the gel. Clear bands against a blue background indicated gelatinolytic activity.
2.4 Molecular Methods
2.4.1 RNA Extraction
Sputum and blood cell pellets were stored in 350μL of Buffer RLT (QIAGEN, Hilden, Germany) at –80˚C. Buffer RLT contains guanidine isothiocanate, which immediately inactivates RNases to make sure there is isolation of intact RNA. Blood cell sample RNA was extracted using the QIAGEN RNeasy Mini Kit as per manufacturers instructions. Briefly, the sample was thawed and ethanol was added and mixed well. The sample was then applied to a spin column and the eluate discarded. The column was washed and the RNA eluted with 40μL of RNase free water. The extracted RNA was stored at –80˚C.
56 Sputum cell culture sample RNA was extracted using the QIAGEN Rneasy MinElute Cleanup Kit as per manufacturers instructions. Briefly, the sample was thawed and ethanol was added and mixed well. The sample was then applied to a spin column and the eluate discarded. The column was washed and the RNA eluted with 14μL of RNase free water. The extracted RNA was stored at –80˚C.
2.4.2 RNA Quantitation for Real-Time PCR
RNA was diluted in TE buffer (Tris EDTA) buffer and the ratio of sample absorbance at 260nm to 280nm was measured (Cary 50: Varian Bio, Palo Alto, CA USA). Samples with a ratio of less than 1.7 were considered low quality and discarded. The concentration of RNA in the sample was determined from the optical density of the sample at 260nm.
2.4.3 Reverse Transcription for Real-Time PCR
DNAse (Invitrogen, Carlsbad, CA, USA) was added to each RNA sample before the reverse transcription to remove any contaminating genomic DNA. To do this, the following master mix (10μL) was prepared and incubated at room temperature for 15 minutes. VOLUME PER REACTION COMPONENT 1μL 10 x DNAse reaction buffer 1μL DNAse I, Amp Grade, 1U/μL XμL RNA sample (500ng) (8-X)μL RNAse free water
EDTA (1μL, Invitrogen, Carlsbad, CA, USA) was added and the sample was incubated at 65˚C for 10 minutes to stop the reaction. A quick pulse spin was performed and the reaction chilled rapidly on ice.
The following master mix (2.5μL) was added to the DNAse treated RNA for random primer reverse transcription.
57 VOLUME PER REACTION COMPONENT 0.1μL Random Primers (3μg/μL) 1μL 10mM dNTP mix 1.4μL RNAse free water
The reaction was incubated for 5 minutes at 65˚C, pulse spun and chilled on ice. A second master mix (7μL) was prepared and added to the reaction. VOLUME PER REACTION COMPONENT 4μL 5 x SuperScript II buffer 2μL 100mM DTT 1μL RNAse OUT (40U/μL)
The reaction was incubated at 25˚C for 10 minutes followed by 42˚C for 2 minutes. Superscript II reverse transcriptase (100U, Invitrogen) was added and incubated at 42˚C for 50 minutes followed by 70˚C for 15 minutes. The resultant cDNA was stored at – 20˚C.
2.4.3.1 Primers and Probe Sequences
Real time PCR primers and probes for 18S rRNA, IL-8, IL-1β, TNF-α, TLR-4, TLR-2 and OSM were purchased as proprietary preoptimised reagents (Taqman Gene Expression Assays, Applied Biosystems, Forster City, CA, USA). See table for probe sequence reference.
TARGET NCBI ACCESSION
18S X03205 IL-8 NM_000584 IL-1β NM_000576 TNF-α NM_000594 OSM NM_020530 TLR2 NM_003264 TLR4 NM_138554
58 2.4.4 Real-Time PCR
Duplex PCR was performed to amplify the endogenous control gene 18S and the target gene simultaneously using the commercial RealMaster Mix (Eppendorf, Hamburg, Germany) and an ABI 7500 Real Time PCR System (Applied Biosystems). The following master mix (25μL) was prepared for each real-time PCR reaction.
VOLUME PER REACTION COMPONENT 2μL cDNA sample 1.25μL 18S Probe 1.25μL Target Probe 10μL RealMaster Mix Probe (Eppendorf) 10.5μL Nuclease free water
The target fluorogenic probes incorporated the 5’ reporter dye FAM and the 3’ quencher dye MGB. All reactions were run under the following conditions: 95˚C for 2 minutes, and then 40 cycles of 95˚C for 15 seconds and 60˚C for 1 minute.
2.4.4.1 Analysis of Relative Real-Time PCR results
The amount of target present was calculated as below, and normalised to the endogenous control (18S rRNA), expressed relative to the calibrator. 2-∆∆Ct Where: ∆∆Ct = ∆Ct (sample) - ∆Ct (calibrator) ∆Ct is the difference in the threshold cycles between the target and the endogenous control.
2.4.4.2 PCR Controls
2.4.4.2.1 Endogenous Control 18S ribosomal RNA was used as the endogenous control in all of the real-time PCR reactions.
59 2.4.4.2.2 Specific Target Positive Controls (Calibrators)
2.4.4.2.2.1 IL-8 mRNA The positive calibrator used for IL-8 real-time PCR was a universal human reference RNA (ALLPOS, Stratagene).
2.4.4.2.2.2 IL-1β and TNF-α mRNA The positive calibrator used for IL-1β and TNF-α real-time PCR was PHA stimulated peripheral blood monocytes (PBMCs). Briefly, blood (20mL, EDTA tubes) was diluted with HBSS (50mL) and layered onto Lymphoprep solution (Nycomed, Oslo, Norway), and centrifuged (400g, 20 minutes, RT, no brake). The PBMC layer was carefully removed and diluted to 50mL with HBSS and centrifuged (400g, 10 minutes, RT). The pellet was resuspended to 1mL (HBSS) and a TCC performed. The cell concentration was adjusted to 1 x 106cells/mL and 5μg/mL of PHA was added (Sigma-Aldrich, Australia). The stimulated cells were cultured in 24 well plates (37˚C, 5% CO) for 6 hours, and the cell pellet stored in RLT buffer (QIAGEN, Hilden, Germany) at –80˚C until RNA was extracted.
2.4.4.2.2.3 TLR2, TLR4, and OSM mRNA The positive calibrator used for TLR2 and TLR4 mRNA was LPS stimulated peripheral blood granulocytes. Granulocytes were isolated as per section 2.2.3.1 of this thesis. The cell concentration was adjusted to 1 x 106cells/mL and stimulated with LPS (100ng/mL, Sigma-Aldrich, Australia) for 2-4 hours. Cell pellets were stored in RLT buffer (QIAGEN, Hilden, Germany) at –80˚C until RNA was extracted.
2.4.5 Gene Expression Studies with Illumina BeadArrays
2.4.5.1 RNA Quantitation for BeadArrays
RNA was quantitated using the Quant-iT RiboGreen RNA Quantitation Assay and Kit (Molecular Probes Inc, Invitrogen, Eugene, OR, USA). An RNA standard curve was constructed with a range from 0 – 200ng/μL. 1μL of sample was added to 99μL of 1 x TE (Tris EDTA) in a 96 well plate format. Black plates were used so as to minimise contact of RiboGreen with light. Fluorescence was measured at wavelengths 485nm for
60 excitation and 520nm for emission with a fluorometer (FLUOstar Optima, BMG Labtech, VIC Australia).
2.4.5.2 RNA Amplification for BeadArrays
RNA was amplified using the Illumina TotalPrep RNA Amplification Kit (Ambion, Texas, USA) as per manufacturers instructions. This kit is a complete system for generating biotinylated, amplified RNA for hybridisation with Illumina Sentrix arrays (Illumina Inc, San Diego, CA, USA).
2.4.5.2.1 Reverse Transcription to Synthesise First Strand cDNA
500ng of RNA was adjusted to a volume of 11μL and mixed with 9μL of reverse transcription master mix (See Table). This was incubated at 42˚C for 2 hours.
Table: Reverse Transcription Master Mix VOLUME PER REACTION KIT COMPONENT 1μL T7 Oligo (dT) Primer 2μL 10 x First Strand Buffer 4μL dNTP Mix 1μL RNase Inhibitor 1μL ArrayScript
2.4.5.2.2 Second Strand cDNA Synthesis
The reactions were placed on ice, and 80μL of Second Strand master mix (see table) was added to the reaction, mixed and placed at 16˚C for another 2 hours. Table: Second Strand Master Mix VOLUME PER REACTION KIT COMPONENT 63μL Nuclease-free Water 10μL 10 x Second Strand Buffer 4μL dNTP Mix 2μL DNA Polymerase 1μL RNase H
61 2.4.5.2.3 cDNA Purification The reactions were placed on ice and 250μL of cDNA binding buffer was added. The mix was then applied to a filter cartridge inside a wash tube and centrifuged for 1 minute at 10,000g. The eluate was discarded, the filter washed and the purified cDNA eluted in 19μL of nuclease-free water warmed to 50-55˚C.
2.4.5.2.4 In Vitro Transcription (IVT) IVT was performed to synthesise biotinylated cRNA for hybridisation to Illumina BeadChips. 7.5μL of IVT master mix (see table) was added to purified cDNA and incubated at 37˚C overnight.
Table: IVT Master Mix VOLUME PER REACTION KIT COMPONENT 2.5μL T7 10x Reaction Buffer 2.5μL T7 Enzyme Mix 2.5μL Biotin-NTP Mix
75μL of nuclease free water was added to stop the reaction.
2.4.5.2.5 cRNA Purification
It is necessary to purify cRNA to remove any enzymes, buffers and salts from the previous reaction. 350μL of cRNA binding buffer was added to the sample and mixed, and another 250μL of ethanol was subsequently added. The mix was applied to a cRNA filter cartridge and centrifuged at 10,000g for 1 minute. The eluate was discarded, the filter washed and cRNA eluted with 100μL of nuclease free water warmed to 50-60˚C.
2.4.5.3 Illumina BeadChip Protocol
RNA samples were hybridised to Sentrix HumanRef-8 Expression BeadChips, which allowed the testing of 8 samples per chip. Hybridisation, washing and scanning of the BeadChip was performed as per Illumina protocol (Illumina Inc., San Diego, CA, USA).
62 2.4.5.3.1 Hybridisation
Briefly, cRNA samples were dried down on a rotary evaporator (DNA Speed Vac DNA 110, Savant, Ramsay, Minnesota, USA) if necessary to achieve a concentration of 0.85μg RNA in a volume of 11.3μL. 125μL of Hyb E1 buffer was combined with 75μL of formamide (Deionized, Ambion, Texas, USA) and mixed well. 22.7μL of this mix was added to each cRNA sample and heated for 5 minutes at 65˚C. Each sample was added to the BeadChip within a hybridisation chamber and this was incubated whilst rotating at 55˚C for 16-20 hours (Illumina Hybridisation Oven, Illumina, San Diego, CA, USA).
2.4.5.3.2 Washing
The BeadChip was removed from its hybridisation chamber and subject to the following series of washes: room temperature E1BC wash buffer (Illumina Inc., San Diego, CA, USA), high temperature wash (55˚C, 10 minutes, Illumina Inc., San Diego, CA, USA), E1BC wash (RT, 5 minutes, orbital shaker), 100% ethanol wash (10 minutes, orbital shaker), E1BC wash (RT, 2 minutes, orbital shaker). The BeadChip was then blocked (Block E1 buffer, 10 minutes, Illumina) and stained with Cy3 Streptavidin (Block E1 buffer with 2μL of 1 mg/mL Cy3 stock, Amersham Pharmacia Biotech). Following staining was a final wash (E1BC buffer, 5 minutes, orbital shaker). The chip was centrifuged dry at 275g for 4 minutes.
2.4.5.3.3 Scanning
Each BeadChip was scanned using the Illumina BeadArray Reader (Illumina, San Diego, CA, USA) and captured using BeadScan 3.5.11 (Illumina, San Diego, CA, USA).
2.5 Data Analysis
Data was analysed using Stata 9 (Stata Corporation, College Station, Texas USA) and GraphPad Prism 4.0 (GraphPad Software Inc, San Diego, CA, USA). Unless otherwise stated data was non-parametric and median (25th-75th percentile) is reported. Non- parametric analyses were performed using the two-sample Wilcoxon Rank Sum, and the
63 Kruskal-Wallis test for comparison of 3 or more samples. Associations between non- parametric data were determined using the Spearman rank correlation. Results were considered statistically significant when p<0.05.
2.5.1 Microarray Data Analysis
Alterations in gene expression of peripheral blood neutrophils were measured using the Illumina HumanRef-8 Gene Expression BeadChips (Illumina, San Diego, CA, USA). Average fluorescent signals were normalised by cubic spline in BeadStudio 2 (Illumina) software.
2.5.1.1 Resting versus Stimulated Neutrophils
Data was exported to GeneSpring 5.0 (Silicon Genetics, USA) and a list was created of genes present in all samples. From those genes detected in all samples, a statistically significant gene list was created comparing resting neutrophils to those stimulated with LPS using a non-parametric test (Wilcoxon-Mann-Whitney) where a p value of <0.05 was considered significant. Using this list a dendogram (Experiment tree) was created using standard correlation to reveal significant relationships between the expression profiles of all samples. A second dendogram (Gene tree) was created to show relationships between the expression levels of genes across all samples.
2.5.1.2 Resting Neutrophils in Asthma Subtypes
Data was exported to GeneSpring 5.0 (Silicon Genetics, USA) and a list was created of genes present in all resting neutrophil samples. From those genes detected in all control samples, a statistically significant gene list was created comparing baseline neutrophil activity in neutrophilic asthma to eosinophilic asthma using a non-parametric test (Wilcoxon-Mann-Whitney) where a p value of <0.05 was considered significant. Using this list a dendogram (Experiment tree) was created using standard correlation to reveal significant relationships between the expression profiles of resting neutrophil samples. A second dendogram (Gene tree) was created to show relationships between the expression levels of genes across all samples.
64 2.5.1.3 LPS Stimulated Neutrophils Asthma Subtypes
Data was exported to GeneSpring 5.0 (Silicon Genetics, USA) and a list was created of genes present in all LPS stimulated neutrophil samples. From those genes detected in all LPS samples, a statistically significant gene list was created comparing LPS responses in neutrophilic asthma to eosinophilic asthma using a non-parametric test (Wilcoxon- Mann-Whitney) where a p value of <0.05 was considered significant. Using this list a dendogram (Experiment tree) was created using standard correlation to reveal significant relationships between the expression profiles of all LPS samples. A second dendogram (Gene tree) was created to show relationships between the expression levels of genes across all samples.
65 Chapter 3: Activation of Blood Granulocytes
3.1 Introduction
Neutrophils are the primary cells of the innate immune system that arrive first at an inflammatory site, protecting the host from infection by microorganisms. Neutrophils have been implicated in the pathogenesis of a number of inflammatory diseases of the airways, including chronic obstructive pulmonary disease (COPD) and asthma. Activation of neutrophils in the bloodstream leads to their migration to the inflammatory site, e.g. the airways, where the cells contribute to significant tissue damage through the release of proteases, reactive oxygen species, cytokines and chemokines.
Neutrophilic airway inflammation is associated with the accumulation of high levels of the potent neutrophil chemoattractant IL-8, the cytokines TNF-α and IL-1β, and the proteolytic enzymes MMP-9 and NE [164, 255]. NE is a serine protease that is found in high concentrations in neutrophil azurophilic granules. It is important in the remodelling of the extracellular matrix as well as cell migration, angiogenesis, vasculogenesis, and apoptosis [161].
MMPs are a family of zinc containing proteinases that are produced by phagocytes and are thought to be involved in cell migration and tissue remodelling [168]. The association of high levels of these mediators with neutrophilic airway inflammation could be due to the influx of large numbers of neutrophils into the airways; however neutrophils themselves are an important and perhaps somewhat undervalued source of these mediators. The activation of the release of innate immune mediators from neutrophils is likely to be important in the regulation of neutrophilic inflammation.
The reactivity of neutrophils is modulated by many cytokines including IL-1, TNF-α, GM-CSF and the chemokine IL-8, which are activators of many neutrophil functions [120, 289, 290]. IL-8 is a potent chemotactic factor for neutrophils and is proinflammatory in its effects [119, 120]. IL-8 is initially synthesised as a 99 amino acid precursor, which undergoes N-terminal cleavage to either 72 or 69 amino acid forms [120]. IL-8 is a member of the CXC subfamily, and is produced by several cell
66 types, including epithelial cells, alveolar macrophages, lymphocytes, fibroblasts, and endothelial cells. Interestingly, IL-8 is one of the main cytokines produced by neutrophils [195, 291], and in this way the cells may maintain their own activation by providing feedback in an autocrine manner. IL-8 is released upon proinflammatory stimulation [124] and has long been recognised to regulate neutrophil recruitment, including the shedding of L-selectin, upregulation of β2 integrins (CD11b/CD18), adhesion to the endothelium and migration [292]. Furthermore, IL-8 can have priming effects on various functions of the neutrophil, including degranulation, oxidative burst and phagocytosis of bacteria [125, 126, 293].
IL-18 is a member of the IL-1 cytokine family that is expressed by many cell types but primarily macrophages/monocytes [294]. IL-18 is an early mediator for the Th1 response. It is a co-stimulus important for the release of interferon (IFN)-γ, which acts in concert with IL-12 and LPS or other bacterial products [294]. The precursor form of IL-18 (26kDa) is cleaved by IL-1 converting enzyme (ICE, also called caspase 1) to a functionally active form (18.3kDa) [295]. A proinflammatory role for IL-18 has recently been implicated in asthma [296-298], acute lung inflammation [299], and rheumatoid arthritis [300], diseases also known to involve neutrophilic inflammation. Leung et al [301] has recently proposed a role for IL-18 in neutrophil activation and the promotion of early innate immune responses.
Neutrophils become activated when they encounter bacteria and their products, in particular lipopolysaccharide (LPS) present on gram-negative bacteria. Airway inflammation can be associated with infection by gram negative bacteria or the presence of endotoxin (LPS) and is still a major cause of severe airway disease [139, 302]. Our airways are constantly faced with endotoxin in the environment (e.g. dust, air pollution, tobacco smoke). Inhalation of LPS has been associated with neutrophilic inflammation with increases in IL-8 in both normal and asthmatic subjects [263]. Neutrophils are known to respond to LPS by production of reactive oxygen intermediates, lipid mediator and cytokine release, adhesion and phagocytosis [223, 224].
The mechanism behind the activation and accumulation of neutrophils in COPD and asthma remains to be fully elucidated. Bacterial products such as lipopolysaccharide (LPS), chemokines such as interleukin-8 (IL-8), and cytokines such as interleukin-18
67 (IL-18) play a role in neutrophil migration and activation. This chapter investigates the activation of neutrophils by LPS, IL-8 and IL-18. The hypothesis being tested is that addition of these mediators will increase the release of neutrophil proteases (MMP-9 and NE), cytokines (TNF-α) and chemokines (IL-8). A suitable model of neutrophil activation is established in this chapter, and will be the subject of further study in subsequent chapters.
3.2 Methods
Blood samples were collected from healthy control volunteers (n=10). Granulocytes were isolated using percoll density gradient and erythrocyte lysis. Isolated granulocytes were cultured in RPMI 1640 (1% FCS) with or without stimulation with the following neutrophil activating agents: LPS (100ng/mL), IL-8 (10, 100, 200ng/mL), or IL-18 (10, 100ng/mL). Cells were cultured at a concentration 1 x 106 cells/mL. Cells and cell free supernatants were collected at the specified time point and stored at –80˚C until analysis. IL-8 and TNF-α were measured using a commercial sandwich ELISA. MMP-9 was measured using a commercial fluorescent ELISA. NE was measured using a colourmetric assay with the substrate n-methoxysuccinyl-l-alanyl-prolyl-l-valyl-p- nitroanilide. RNA was extracted and mRNA quantified by real-time PCR with commercial primers and probes for TLR4, IL-1β, and TNF-α. Data was analysed by Stata 9, all data is non-parametric and reported as medians (IQR). Significant differences (p<0.05) were detected with the 2-sample test Wilcoxon rank sum. For multiple group comparisons, the non-parametric Kruskal Wallis test was used. For more detailed methods please refer to Chapter 2 of this thesis.
3.3 Results
Granulocytes were isolated from blood via Percoll density gradient. This resulted in a cell fraction that contained an average of 93% neutrophils and 6% eosinophils, with a small number (1%) of contaminating lymphocytes and monocytes. Neutrophils are short-lived cells, so it was necessary to monitor their viability over the 24-time period. As expected, viability of granulocytes significantly decreased over time (Table 3.1), however at 24 hours approximately 96% of cells were still alive. Cell viability did not significantly differ between the different treatments.
68
Time (hours) Viability (%) 0 99.4 (98.6-99.5) 0.5 98.9 (96.8-100) 2 99.6 (98.4-100) 18 97.1 (94.4-99.1) * 24 95.5 (88-97.9) **#
Table 3.1 Cell Viability of Granulocytes at Specified Time Points Total cell counts were performed and cell viability determine by trypan blue exclusion (n=10). Data is reported as median (IQR) and analysed using the Kruskil Wallis multiple comparison test for non-parametric data. *p<0.00001 compared to 2 hours, **p<0.00001 compared to 0.5 and 2 hours. #p=0.0002 compared to 0 hours.
Neutrophils synthesise cytokines and chemokines such as TNF-α and IL-8 de novo, under various conditions. Within their cytoplasmic granules they also contain proteases such as NE and MMP-9, which can be released upon stimulation without transcription and translation.
The ability of the neutrophil activating agents LPS, IL-8 and IL-18 to induce the release of total MMP-9, TNF-α, IL-8 and NE from blood granulocytes in vitro was assessed. Isolated granulocytes were cultured for time periods ranging from 30 minutes to 24 hours and total MMP-9, TNF-α, IL-8 and NE levels were measured in the cell-free supernatants. Levels of TNF-α, TLR4 and IL-1β mRNA were measured using real time PCR.
3.3.2 Protease Release
Degranulation in response to LPS, IL-8 and IL-18 was evaluated by measuring the release of NE from azurophilic granules and MMP-9 from specific/gelatinase granules. Out of the neutrophil stimulants assessed in this study, only LPS (100ng/mL) induced the release of MMP-9 from isolated granulocytes. LPS induced the release of total MMP-9 to a maximum of 517 (379-591) pg/mL in comparison to 161 (158-169) pg/mL
69 released from unstimulated granulocytes at 24 hours of culture (Figure 3.1). Neither IL- 8 nor IL-18 induced total MMP-9 release at the concentrations studied. LPS significantly increased the quantity of total MMP-9 in 2 hours, indicating the release from preformed granules. Total MMP-9 protein was increased up to its maximum level at 24 hours (Figure 3.2A). NE was not significantly enhanced by any of the inflammatory mediators tested compared to the unstimulated control, however NE release significantly increased over the 24 hour period (Figure 3.2B).
700 *
600
500
400
300
200 Total MMP-9 (pg/ml) MMP-9 Total
100
0 Cntrl LPS 10ng IL-8 100ng IL-8 200ng IL-8 10ng IL-18 100ng IL-18 Treatments
Figure 3.1 Total MMP-9 release from stimulated blood granulocytes at 24 hours The presence of LPS significantly increased the release of total MMP-9 from isolated granulocytes (Unstimulated n=10, LPS n=10). IL-8 (10, 100, 200ng/mL, n=5) and IL-18 (10, 100ng/mL, n=5) had no effect on total MMP-9 release. *p<0.01 compared to granulocyte control.
70 A B * # 700 ** 350 600 * 300 500 250
400 200 300 150 200 100 Active NE (ng/ml) NE Active Total MMP-9Total (pg/mL) 100 50 0 0 0.5 2 18 24 0.5 2 18 24 Treatment Time (hours) Treatment Time (hours)
Figure 3.2 Total MMP-9 release from LPS stimulated granulocytes (A) and NE release from control granulocytes (B) over 24 hours (n=10) MMP-9 and NE release increased over time to reach a maximum at 24 hours after stimulation. *p<0.001, **p<0.0001 compared to 0.5 hours, #p<0.01 compared to 2 hours.
3.3.3 Chemokine and Cytokine Release
LPS stimulation also induced the release of large amounts of IL-8 [6368 (6111-9673) pg/mL] compared to only low amounts released from the control and IL-18 stimulated granulocytes [Control: 64(50-70) pg/mL, 100ng/mL IL-18: 78(77-122) pg/mL)] (Figure 3.3). Release of IL-8 was not at detectable levels in any sample at 30 minutes, however IL-8 release increased over time to a maximum level detected at 24 hours of culture (Figure 3.4).
71 * 24000
14000
4000
200 IL-8 (pg/ml) IL-8 100
0 Control LPS 10ng IL-18 100ng IL-18
Figure 3.3 IL-8 release from granulocytes at 24 hours IL-8 release from granulocytes is significantly increased with LPS stimulation. IL-18 (10, 100ng/mL) had no effect on IL-8 release. *p<0.01 versus to granulocyte control (n=5).
* 17500
15000 12500 * 10000 7500
IL-8 (pg/mL) IL-8 5000
2500 0 2hr 18hr 24hr
Figure 3.4 IL-8 release from LPS stimulated granulocytes over 24 hours IL-8 release increased in LPS stimulated granulocytes over the 24-hour period. *p<0.001 versus 2 hours (n=5).
TNF-α protein release reached detectable levels only in LPS stimulated granulocyte samples, and only at the later time points (2, 18, 24 hours), consistent with the transcript and translation of protein. The level of TNF-α released was highest at 24 hours, where it reached 70 (42-220) pg/mL, however this was not significantly more than that released at 18 hours 56 (34-164) pg/mL (Figure 3.5A).
72
Neutrophils have only recently been shown to be transcriptionally active, adding to their traditional role as phagocytes. TNF-α mRNA was assessed by real time PCR at all time points. LPS significantly induced the transcription of TNF-α mRNA (Figure 3.6), the level being highest at 2 hours after stimulation (Figure 3.5B). Although mRNA levels were lower at later time points, significant differences between control and LPS stimulated granulocytes were still detected up to 24 hours after activation of the cells (Figure 3.5B). mRNA levels of TLR4 and IL-1β were also measured in a selection of samples. At 2 hours, TLR4 mRNA levels remained unchanged, whilst IL-1β was significantly increased with LPS stimulation (Table 3.2).
TNF-a release over time TNF-a mRNA over time Resting A B LPS *** # 240 4.5 4 ** 200 *** 3.5 160 3 2.5 120 2 * 80 1.5 * TNF-a mRNA TNF-a TNF-a (pg/mL) 1 40 0.5 0 0 0.5 2 18 24 0.5 2 18 24 Time (hours) Time (hrs)
Figure 3.5 Kinetics of TNF-α mRNA and protein levels over time Data not normally distributed, so medians values are plotted with the interquartile range as the error bars. TNF-α protein release (A) increases in 24 hours of LPS stimulation. TNF-α mRNA (B) peaks at 2 hours, but stays significantly elevated compared to the unstimulated controls up to 24 hours after LPS stimulation. ***p<0.0001 compared to 0.5 hours, #p<0.05 compared to 2 hours (A). **p<0.001, *p<0.01 compared to unstimulated control (B) (n=5).
73 5.5 * 5.0 4.5 4.0 3.5 3.0 2.5
TNF-a mRNA 2.0 1.5 1.0 0.5 0.0 Control LPS 10ng IL-8 100ng IL-8 200ng IL-8 10ng IL-18 100ng IL-18
Figure 3.6 TNF-α mRNA levels at 2 hours LPS (100ng/mL, n=10) significantly increased the levels of TNF-α mRNA. IL-8 (10, 100, 200ng/mL, n=5) and IL-18 (10, 100ng/mL, n=5) had no effect on TNF-α mRNA levels. *p<0.00001 versus control granulocytes.
TLR4 mRNA IL-1β mRNA Control 0.27 (0.12-0.32) 0.17 (0.12-0.33) 100ng/mL LPS 0.39 (0.15-0.39) 2.32 (1.97-2.6) * 100ng/mL IL-8 0.3 (0.17-0.49) 0.35 (0.31-0.53) 100ng/mL IL-18 0.43 (0.27-0.76) 0.41 (0.24-0.51) Table 3.2 Levels of TLR4 and IL-1β mRNA at 2 hours LPS stimulation significantly increases the level of IL-1β mRNA, however TLR4 mRNA remains unchanged. * p<0.005 kwallis2 p value versus granulocyte control. Control and LPS n=10, IL-8 and IL-18 n=5.
3.4 Discussion
Neutrophils play a key role in the innate immune response to infection. The factors that recruit and activate neutrophils are of major significance in the sequestration of neutrophils to the lung, where they can cause the airway inflammation and remodelling associated with the respiratory diseases asthma and COPD. This chapter examines the
74 capacity of LPS, IL-8 and IL-18 to activate neutrophils as measured by the release of inflammatory mediators NE, MMP-9, TNF-α and IL-8, known to be involved in the pathogenesis of asthma and COPD.
The release of azurophilic granules was determined by the measurement of NE, however neither LPS, IL-8 nor IL-18 induced the release NE above baseline levels. The level of total MMP-9 in the cell culture supernatant determined the release of specific/gelatinase granules. LPS, but not IL-8 or IL-18 significantly increased the release of total MMP-9. Significant increases of total MMP-9 were observed with LPS stimulation at 2, 18 and a maximum level at 24 hours of cell culture. MMP-9 was spontaneously released from resting neutrophils at all time points. Pugin et al [168] have previously reported the release of MMP-9 by stimulation of whole blood with LPS at as early as 20 minutes after stimulation. They found that release of MMP-9 due to LPS stimulation was dependent on the interaction of LPS with CD14, involved in the binding of LPS to its receptor (TLR4).
Other inflammatory mediators such as IL-8, TNF-α, GM-CSF and G-CSF have been demonstrated to induce the release of MMP-9 [168, 169]. This study was unable to confirm the release of MMP-9 induced by IL-8 (10, 100, 200ng/mL) in isolated granulocytes above that of the unstimulated control. This could be due to the longer time period (24 hours) assessed, as the release of MMP-9 increased over time, and this may have masked a small increase in MMP-9 release due to IL-8 stimulation.
The ability of neutrophils to synthesise and release cytokines, in particular TNF-α, after stimulation with LPS, IL-18 or IL-8 was determined by ELISA and real-time PCR. LPS significantly induced the release of TNF-α protein at the earliest time of 2 hours, and increased up to 24 hours. Resting granulocytes and those stimulated with IL-8 or IL-18 did not release TNF-α to detectable amounts. Significant transcription of TNF-α mRNA was seen at 2 hours after LPS stimulation; and this remained significantly upregulated at both 18 and 24 hours compared to unstimulated granulocytes. The chemokine IL-8 was also measured in the cell supernatants of LPS and IL-18 stimulated granulocytes. Resting granulocytes spontaneously released low levels of IL-8 over 24 hours. IL-18 did not increase the release of IL-8 from resting granulocytes.
75 The current results are confirmed by previous studies that have demonstrated LPS stimulation of granulocytes resulting in the production of IL-1β and TNF-α, and IL-8, [303]. The kinetics of chemokine production in neutrophils responding to LPS involve two phases, the first phase resulting from direct stimulation of the cell with LPS which can last for some hours, and the second phase which is sustained and appears to be due to the endogenous action of IL-1β and TNF-α [303, 304]. This study is in agreement with these findings, as the majority of TNF-α was released from the cells by 18 hours, however the release of IL-8 continued to higher amounts at up to 24 hours.
LPS stimulates neutrophils through the activation of TLR4 and the NF-κB pathway. There are a number of proteins that facilitate this interaction, including LPS binding protein, which transfers LPS to CD14, a receptor present on the cell surface. This leads to the association of CD14 with the TLR4-MD-2 complex, inducing the dimerisation of TLR4. Activation of TLR4 induces a signalling cascade beginning with the adapter protein MyD88, associated with IL-1 receptor associated kinase (IRAK). Phophorylation of IRAK leads to the activation of TNF-associated factor 6 (TRAF-6). This leads to the activation of mitogen activated protein kinase kinase kinase (MAP3K) resulting in NF-κB translocation to the nucleus and inflammatory gene transcription [136].
IL-18 exerts its effects in part by the induction of IFN-γ, and in this way IL-18 has proinflammatory properties. IL-18 has been shown also to activate NF-κB in murine T cells [305], and induces the release of IL-8, TNF-α and IL-1β from human blood mononuclear cells [306]. Neutralisation of IL-18 using IL-18 antibodies protected mice from lethal endotoxemia, thought to be in part due to a reduction in neutrophil mediated tissue injury and cytokine and chemokine release [307]. Leung et al [301] demonstrated the effects of IL-18 on neutrophils to include cytokine and chemokine release, upregulation of CD11b, degranulation and an increase in respiratory burst following fMLP exposure. In the current study however, IL-18 had no effect on cytokine and chemokine release or degranulation of neutrophils. IL-18 by itself does not induce IFN- γ production, however it greatly enhances IFN-γ production in the presence of other stimuli including IL-12, and bacterial products such as LPS [308]. IL-18 requires processing by ICE to its active form. In the absence of ICE, there is no IFN-γ production despite the presence of IL-12 [309]. LPS activates ICE in peripheral blood
76 monocytes as well as the monocyte cell line THP-1 [310]. Responses to IL-18 are also modulated by IL-18 binding protein [311]. It is clear that many factors determine the activity of IL-18, and it is possible that this study may lack a number of these factors, explaining the absence of neutrophil activation.
IL-8 has been shown to be a potent inducer of degranulation for both specific and azurophilic granules from human neutrophils [312, 313]. It is not clear why the current study did not result in the cells activation by IL-8; however, it is possible that higher concentrations of IL-8 are needed to activate neutrophil to release elastase and MMP-9 under these in vitro conditions. In fact, human neutrophils have been shown to degranulate when exposed in vitro to doses of IL-8 that are 10 to 100 fold higher than those required for chemotaxis [313]. Higher amounts of IL-8 (1000ng/mL) were needed to cause degranulation in the study by Djeu et al [314]. Perhaps the concentrations of IL-8 used in this study (10, 100, 200ng/mL) alone were not high enough to caiuse neutrophil degranulation, however these concentrations are of more relevance physiologically.
The migration of neutrophils to the airways is a complex process, and requires adhesion to the endothelium, which may accelerate the cells’ activation. Neutrophils that are adherent to the endothelium show increased inflammatory responses as compared to suspended neutrophils, in particular, they release larger amounts of oxygen radicals and NE, suggesting that the binding of the cells’ to the endothelium increases their activation [315, 316]. NF-κB is a transcription factor that controls gene expression during inflammation, and is activated via several stimuli including cytokines such as IL- 8. Interestingly, Kettritz and colleagues [315] have shown that IL-8 did not activate NF- kB in suspended neutrophils, but that IL-8 rapidly activated the NF-κB under adherent conditions on a matrix (fibronectin), which provides a strong co-stimulatory signal for NF-κB activation. It is probable that both bacterial and host derived interactions and neutrophil and endothelial cell interactions are involved in neutrophil mediated tissue injury.
At inflammatory sites in vivo, mediators may interact to promote the activation of neutrophils. As an example of this, synergistic effects for TNF-α and platelet activating factor (PAF) [317] have been described. IL-8 also primes the neutrophils for enhanced
77 superoxide production rather than directly activating them [318], in contrast to what was initially thought [312]. IL-8 also readily induces NE release from IL-1β primed neutrophils [319]. Priming refers to a process whereby the response of neutrophils to an activating stimulus is potentiated by prior exposure to a priming agent. The phenomenon of neutrophil priming has been suggested to be critical in maximising tissue injury by enhancing oxidase dependent and independent cytotoxic function [320]. Functional consequences of priming include enhancement of the respiratory burst activity, shape change and deformability, that is, polarisation of the cell (representing chemotaxis), upregulation of adhesion molecule expression needed for trans-endothelial migration (e.g. CD11b/CD18), inhibition of neutrophil apoptosis, degranulation and lipid mediator release [321].
The neutrophil is a pivotal effector cell that is able to respond to mediators in its environment and generate cytokines. The generation of cytokines may be important for elicitation of additional neutrophils or to orchestrate the immune response at sites of inflammation. The secretory products of neutrophils, including proteolytic enzymes such as elastase, can be a major source of damage to inflamed tissues. This chapter clearly demonstrates that LPS significantly activates neutrophils in vitro to release MMP-9, IL-8, and TNF-α. This stimulation reached a maximum at 24 hours for protein release, and 2 hours for mRNA levels, however significant differences in mRNA were also detected as late as 24 hours. In conclusion, LPS stimulation of neutrophils can be used as an appropriate in vitro model allowing the evaluation of the neutrophils capacity to release important innate immune mediators.
78 Chapter 4: Differences Between Airway and Systemic Innate Immune Function
4.1 Introduction
The effector mechanisms of innate immunity are activated immediately after infection and rapidly control the replication of the pathogen via antimicrobial peptides, phagocytes and the alternative complement pathway [136]. The innate immune response in the airways relies on the recognition and differentiation between potentially pathogenic and non-pathogenic particles that enter the respiratory tract through inspiration. This response involves the detection of pathogen associated molecular patterns (PAMPs) present on microbes, by pattern recognition receptors (PRRs) on the cell surface. A model of innate immunity is the response to LPS from gram negative bacteria, however other examples PAMPs include peptidoglycan and lipoteichoic acids, present on gram positive bacteria, which stimulate Toll like receptor (TLR) 2 [135].
Lipopolysaccharide (LPS) stimulates neutrophils through the activation of TLR4 and the NF-κB pathway. There are a number of proteins that facilitate this interaction, including LPS binding protein, which is produced by the liver and circulates in the blood where it recognizes the lipid A moiety of LPS. LBP initially binds with LPS and forms a complex with CD14, enabling LPS to be transferred to the receptor complex TLR4 and MD-2. CD14 is present in two forms, including soluble CD14, which assists in LPS signalling in cells lacking membrane CD14 e.g. epithelial and endothelial cells, and membrane bound CD14 attached to the cell surface of myeloid cells. The binding of LPS to TLR4 and MD-2 assisted by LBP and CD14 induces the homodimerization of the receptor.
Signalling pathways activated through TLR4 are either MyD88 dependent or independent [140]. Early LPS signalling leads to the activation of NF-κB, IRF3 and MAPK pathways, which are mediated by MyD88 and Mal. Following the phosphorylation of IRAK, TRAF-6 is activated which leads to the production of proinflammatory genes. The later response to LPS is mediated by TRIF and TRAM and results in the activation of TRAF-6 and TANK-binding protein (TBK)-1 and finally NF- κB and IRF3 and the induction of cytokines, chemokines and other transcription factors [140].
79
Inhalation of endotoxin (LPS) is known to be an effective model of airway disease. LPS may cause airway dysfunction, due to its wide spectrum of acute and chronic airway symptoms and its extensive and common presence in everything from household pets and carpet, to grain dust, tobacco smoke and air pollution [139, 322]. However, studies have shown evidence that LPS exposure in early life may be protective against the development of allergy and asthma [264]. The heterogeneity of the response to LPS is thought to be related to both the dose and timing of exposure as well as genetic influences [302]. LPS installation into the airways results in recruitment of macrophages and neutrophils as well as increased levels of inflammatory mediators (especially IL-1, TNF-α, IL-6 and IL-8) in bronchoaveolar lavage fluid (BAL) [323] and neutrophilic inflammation relating to increased IL-8 levels in sputum [263].
Neutrophils migrate rapidly to sites of inflammation including the airways, where these motile phagocytic cells become activated by microbial products such as LPS, or cytokines secreted by T cells, macrophages and other structural cells. Neutrophils express TLR2 and TLR4 on their cell surface and these receptors can detect the presence of bacteria [142, 324]. Activation of TLR2 and TLR4 regulates important neutrophil functions, including recruitment, e.g. regulation of adhesion molecule expression (CD11b); release of chemokines, e.g. IL-8; generation of reactive oxygen species; and regulation of the neutrophils life span, e.g. stimulation of TLR4 activates the NF-κB pathway which promotes neutrophil survival [325]. Expression of TLR2 but not TLR4 is increased by proinflammatory cytokines [142] and exposure to a range of different bacteria [326]. In this study we have used commercially available LPS which can be viewed as a dual TLR2 and TLR4 agonist due to contamination with bacterial lipopeptide [327].
LPS stimulation of blood neutrophils in vitro results in the production of IL-8, and this is both increased and sustained in the presence of IL-1β and TNF-α, also induced by LPS [81]. Through the production of these mediators, neutrophils further influence their recruitment by positive feedback. Previous studies have been carried out mainly in blood neutrophils, and the results from these experiments may not reflect the activity of neutrophils in the airways. Under normal conditions the peripheral blood is sterile and therefore responds to pathogen exposure with a vigorous inflammatory response. In
80 contrast to this, the airways are continually exposed to sources of LPS, including bacteria, air pollution, and tobacco smoke. Neutrophils also undergo vast changes upon migration from the blood to the airways, which has the capacity to change the passive circulating neutrophil into an active effector cell of innate immunity. The response of airway cells must also be tightly regulated to identify the presence of pathogens and distinguish their presence from harmless exposure. The mechanisms by which this occurs remain largely unknown but involve innate immunity, and therefore it is essential to investigate airway neutrophil activation as it is likely to be different to that of blood neutrophil responses.
Airway inflammation is an important feature of asthma, however very little is known regarding airway neutrophil responses, and how they compare to those of blood neutrophils. Cytokines and mediators produced by airway neutrophils could provide a local mechanism by which neutrophils could modulate ongoing inflammation. Induced sputum is being increasingly used as a non-invasive method for the direct study of airway inflammation, however this has been limited because of the small number of cells recovered, and the differing cell types and composition of the sample. In order to clarify the role of neutrophils in the airways and circulation, these cells were isolated and the production of innate immune mediators was investigated in resting and LPS stimulated states. The hypothesis for this chapter is that LPS will increase the production of innate immune mediators from both airway and circulating neutrophils.
4.2 Methods
Induced sputum and blood samples were collected from a total of 35 participants including subjects with asthma (n=17), COPD (n=5) and healthy controls (n=13). Neutrophils were isolated via magnetic cell separation with CD16 microbeads. Isolated blood and sputum neutrophils were cultured at cell concentrations of with 1 x 105 or 1 x 106 cells/mL in RPMI 1640, with or without LPS stimulation (100ng/mL). Cells and cell free supernatants were collected at 24 hours and stored at –80˚C until analysis. IL-8, IL-1β, TNF-α and OSM were measured using a commercial sandwich ELISA. MMP-9 was measured using a commercial fluorescent ELISA and zymography. RNA was extracted and mRNA quantified by real-time PCR with commercial primers and probes for TLR2, TLR4, IL-8, IL-1β, TNF-α and OSM. Data was analysed by Stata 9, all data
81 are non-parametric and reported as medians (25th-75th percentile). Significant differences were detected with the 2-sample test Wilcoxon rank sum, with a p value of <0.05 considered significant.
Neutrophils were isolated from sputum or blood using magnetic cell separation (MACS) with CD16 microbeads. This technique involved the sample being incubated with CD16 microbeads, and passed through a column surrounded by a magnetic field. Microbeads are superparamagnetic particles approximately 50nm in diameter, and coupled to a CD16 antibody. They have no known effect on cell structure, function or activity status of labeled cells and do not interfere with subsequent experiments (See http://www.miltenyibiotec.com/en/NN_21_MACS_Cell_Separation.aspx for more information). The labelled cells (neutrophils) are retained in the column whilst the unlabelled cells are eluted to a collection vial. The column is then removed from the magnetic field and the neutrophils are eluted into a separate collection vial. CD16 is present on the surface of neutrophils as well as a proportion of macrophages. For more detailed methods please refer to Chapter 2 of this thesis.
4.3 Results
4.3.1 Magnetic Cell Separation of Neutrophils
For the blood isolation, granulocytes were separated from the blood by percoll density gradient centrifugation and erythrocyte lysis. CD16 microbeads were used to label CD16 positive cells (neutrophils) allowing the depletion of peripheral blood eosinophils. This technique resulted in the isolation of highly pure blood neutrophils (99.8%, (96.8-100%)) (Figure 4.1B, Table 4.1) and blood eosinophils (94.3% (92.4- 96.5%)) (Figure 4.1A, Table 4.1). Contaminating cells in the neutrophil fraction were eosinophils, and in the eosinophil fraction were neutrophils, lymphocytes or monocytes. The cells were resuspended in RPMI 1640 1% FCS to a concentration of 1 x 105 and 1 x 106 cells/mL and cultured for 24 hours.
82
CD16- Fraction CD16+ Fraction p value Neutrophils (%) 0.5 (0-1.6) 99.8 (96.8-100) <0.00001 Eosinophils (%) 94.3 (92.4-96.5) 0.2 (0-1.5) <0.00001 Other (%) 4.5 (2.3-6.1) 0 (0-0.5) <0.00001 Table 4.1 Proportions of neutrophils, eosinophils and other (lymphocytes or monocytes) of the CD16- and CD16+ blood cell fractions created using MACS
83 Figure 4.1 Magnetic cell separation of peripheral blood eosinophils (A) and neutrophils (B) using CD16 microbeads, magnification 400x A)
B)
84 For the sputum isolation method, the sputum sample was dispersed with 0.1% DTT/PBS, filtered and centrifuged to obtain a cell pellet. The sputum cells were labelled with CD16 microbeads and separated into CD16 negative and CD16 positive cell fractions. This technique resulted in the creation of a non-eosinophilic (CD16 positive) cell fraction that consisted mainly of neutrophils (59.2% (30-78.1%), Figure 4.2, Table 4.2) as well as a population of macrophages (35.3% (21.6-56.8%)). The cells were resuspended in RPMI 1640 1% FCS to a concentration of 1 x 105 cells/mL and cultured for 24 hours.
CD16- Fraction CD16+ Fraction p value Neutrophils (%) 1.2 (0.3-3.3) 59.2 (30-78.1) <0.00001 Macrophages (%) 60 (39-73.2) 35.3 (21.6-56.8) 0.0034 Eosinophils (%) 11 (0.7-24.3) 0 (0-0.7) <0.00001 Table 4.2 Proportions of neutrophils, macrophages and eosinophils in the CD16- and CD16+ sputum cell fractions created using MACS
Figure 4.2 Isolated Sputum Neutrophils, Magnification 1000x
Cell viability was monitored before and after cell separation. Viability of blood eosinophils and neutrophils decreased after magnetic cell separation (Refer to Table 4.3), however 98% of the cells remained viable. Sputum viability was significantly lower than blood [Blood: 99.6% (99.2-99.8); Sputum: 75.8% (67.2-81.4); p<0.00001], but was not affected by magnetic cell separation. Cell viability of blood neutrophils decreased over 24 hours of culture [0hrs: 98% (97-99), 24hrs: 94%(89-96) p<0.0001].
85 Cell viability did not significantly differ between control and LPS stimulated blood neutrophils at 24 hours [Control: 94%(89-96) LPS: 95% (88-97)].
Sample Pre MACS (%) CD16- Fraction (%) CD16+ Fraction (%) p
Peripheral Blood 99.6 (99.2-99.8) 98.4 (97.7-99.5)* 98 (97.1-99.1)* 0.0001
Induced Sputum 75.8 (67.2-81.4) 82.3 (61.9-88.9) 66.7 (57.1-82.1) 0.14 Table 4.3 Cell viability before and after MACS
4.3.2 Effects of LPS Stimulation on Peripheral Blood Neutrophils
Expression and release of cytokines from blood neutrophils and eosinophils was dependent on the presence of LPS. LPS significantly increased the production of IL-8, IL-1β, TNF-α, and OSM from blood neutrophils (Figure 4.3), with corresponding increases in mRNA transcripts (Table 4.4). Levels of mRNA significantly correlated with protein levels for all cytokines (Figure 4.4).
86 Resting LPS
9000 **40 7500 30 6000
4500 20 (pg/mL) β 3000 IL-8 (pgmL)
IL-1 10 1500
0 0
45 * 70 * 40 60 35 50 30 25 40
(pg/mL) 20 α 30 15 20 OSM (pg/mL) OSM
TNF- 10 5 10 0 0
Figure 4.3 Chemokine (A: IL-8) and cytokine (B: IL-1β, C: TNF-α, D: OSM) release from blood neutrophils (1 x 106 cell/mL) *p<0.00001 versus control
Resting 100ng/mL LPS IL-8 2.28 (1.22-3.57) 44.94 (19.03-79.34) * IL-1β 0.0032 (0.001-0.007) 0.085 (0.028-0.14) * TNF-α 0.0023 (0.0012-0.0056) 0.11 (0.056-0.17) * OSM 0.12 (0.07-0.18) 0.65 (0.41-0.84) * Table 4.4 Levels of chemokine and cytokine mRNA expression in isolated blood neutrophils (1 x 106 cells/mL) at rest and with LPS stimulation *p<0.00001 versus control
87 r=0.81 200 r=0.74 20000 p<0.0001 p<0.0001 150 15000
100 10000
IL-8 (pg/mL) IL-8 50
5000 IL-1beta (pg/mL)
0 0 0 100 200 300 400 0.0 0.5 1.0 1.5 2.0 2.5 3.0 IL-8 mRNA IL-1beta mRNA
1000 350 300 r=0.56 r=0.64 p<0.0001 750 p<0.0001 250 200 500 150 100 OSM (pg/mL)
TNF-a (pg/mL) 250 50 0 0 0.00 0.25 0.50 0.75 1.00 1.25 0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 TNF-a mRNA OSM mRNA
Figure 4.4 mRNA expression positively correlates with protein release
LPS stimulation significantly increased the release of total MMP-9 from isolated neutrophils but did not change the activity of the protease (Table 4.5, Figure 4.5).
Resting 100ng LPS Total MMP-9 285.7 (215-381.1) 584.8 (432.8-690.3) * Active MMP-9 3.5 (2.2-4.8) 3.9 (2.8-6.6) Table 4.5 Total and active MMP-9 (pg/mL) release from isolated peripheral blood neutrophils *p<0.00001 versus resting neutrophils
88 L 1 2 3 4
MMP-9 Homodimer MMP-9 High Molecular Weight Form (MMP-9/Lipocalin complex)
MMP-9 Latent Form
MMP-2 Latent Form
Figure 4.5 Zymography gel confirms the presence of MMP-9 Lanes 1 (Control) and 2 (LPS) contain peripheral blood eosinophil samples, which show little MMP-9 activity. Lanes 3 (Control) and 4 (LPS) contain peripheral blood neutrophil samples, which show large amounts of MMP-9 in all the 3 forms (homodimer, high molecular weight form, and the latent form). LPS clearly increases the release of the latent form of MMP-9 from peripheral blood neutrophils. Bands are also present for eosinophil and neutrophil fractions corresponding to MMP-2.
The level of TLR expression was examined in isolated blood neutrophils. Both TLR4 (Figure 4.6A) and TLR2 (Table 4.6B) mRNA expression was significantly increased with LPS stimulation.
Resting LPS A B ** 0.0125 0.4
0.0100 0.3
0.0075 0.2 0.0050 TLR2 mRNA TLR2 TLR4 mRNA 0.1 0.0025
0.0000 0.0 Figure 4.6 TLR4 and TLR2 mRNA expression in isolated blood neutrophils (1 x 106 cells/mL) * p<0.0001 versus resting neutrophils
89
4.3.3 Effects of LPS Stimulation on Sputum Neutrophils
To determine whether sputum neutrophils are a source of cytokines, isolated sputum neutrophils were cultured with or without LPS stimulation (100ng/ml) for 24 hours. As seen in Figure 4.7, sputum neutrophil samples spontaneously released high levels of IL- 8, IL-1β and TNF-α protein, with no further increase after LPS exposure. Levels of IL- 8, IL-1β and TNF-α mRNA were also elevated in resting sputum cells, and LPS stimulation had no additional stimulating effect on these levels (Table 4.6). Sputum neutrophils did not release detectable levels of OSM.
Resting LPS
A B C 3000 45 175 40 150 35 125 2000 30 25 100 20 75 1000 15 IL-8 (pg/mL) IL-8 50 10 IL-1beta (pg/mL) IL-1beta 5 TNF-alpha (pg/mL) 25 0 0 0 Figure 4.7 Chemokine (A: IL-8) and cytokine (B: IL-1β and C: TNF-α) release from sputum neutrophils (1 x 105 cell/mL)
Resting 100ng/mL LPS IL-8 50.5 (28.2-79.5) 62.2 (44.1-104) IL-1β 0.27 (0.094-0.79) 0.26 (0.16-0.78) TNF-α 0.16 (0.77-0.38) 0.2 (0.09-0.37) Table 4.6 Levels of chemokine and cytokine mRNA expression in sputum neutrophils (1 x 105 cells/mL) at rest and with LPS stimulation
The expression of TLR2 & 4 was examined in sputum neutrophils. TLR4 and TLR2 mRNA was present in sputum cells at baseline, and LPS exposure did not change the level of TLR4 (Figure 4.8A) or TLR2 (Figure 4.8B). Sputum neutrophils released very low levels of total MMP-9 [Sputum: 2.0 pg/mL (1.8-2.1)].
90 Resting LPS
A B
0.6 0.005 0.5 0.004 0.4 0.003 0.3 0.002 0.2 TLR2 mRNA TLR4 mRNA 0.001 0.1 0.000 0.0 Figure 4.8 TLR4 and TLR2 mRNA expression in sputum neutrophils (1 x 105 cells/mL)
4.3.4 Comparison of Sputum to Blood Neutrophils
Cell viability is significantly higher in peripheral blood samples before and after separation compared to induced sputum samples (Table 4.7). The purity of CD16+ isolated cells is also different between sputum and blood fractions. Isolated blood neutrophils were highly pure, whereas the sputum neutrophil fraction contained a population of CD16 positive macrophages. There are also a number of differences between the isolation procedures for neutrophils from the blood versus the sputum. Sputum samples were dispersed using DTT, and blood neutrophils were isolated from granulocytes after Percoll density gradient. Cell viability of blood granulocytes was decreased by magnetic cell separation, in contrast to sputum cells where no difference of cell viability was observed between isolated cell fractions (Table 4.3).
Peripheral Blood Induced Sputum p value Pre MACS 99.6 (99.2-99.8) 75.8 (67.2-81.4) <0.00001 CD16- Fraction (%) 98.4 (97.7-99.5) 82.3 (61.9-88.9) <0.00001 CD16+ Fraction (%) 98 (97.1-99.1) 66.7 (57.1-82.1) <0.00001 Table 4.7 Cell viability of blood compared to sputum samples before and after MACS
The expression of cytokines from blood neutrophils is dependent on the presence of LPS, and this differs to sputum neutrophils, as cytokine release from these cells is not dependent on LPS. Resting Sputum Neutrophils are considerably more active and
91 release higher levels of innate immune cytokines compared to blood neutrophils (Figure 4.9), and have higher levels of cytokine mRNA (Table 4.8). Resting sputum neutrophils have significantly higher levels of TLR2, but similar levels of TLR4 mRNA expression compared to blood neutrophils (Figure 4.10). LPS stimulated sputum neutrophils still release significantly more cytokines compared to LPS stimulated blood neutrophils (Figure 4.10), with this also corresponding to a higher level of mRNA (Table 4.9). LPS stimulated blood neutrophils have higher levels of TLR4 and similar levels of TLR2 mRNA expression compared to sputum neutrophils (Figure 4.12). Sputum neutrophils release only low levels of total MMP-9 compared to blood neutrophils [Blood: 34.8pg/mL (27.4-44.8) Sputum: 2pg/mL (1.8-2.1) p=0.019] and do not release OSM, another granule associated neutrophil product.
Sputum Blood A * B * C * 3000 45 175 40 150 35 125 2000 30 25 100 20 75 1000 15
IL-8 (pg/mL) IL-8 50 10 IL-1beta (pg/mL) 5 TNF-alpha (pg/mL) 25 0 0 0 Figure 4.9 Chemokine (A: IL-8) and cytokine (B: IL-1β and C: TNF-α) release from resting sputum neutrophils compared to blood neutrophils (1 x 105 cell/mL) *p<0.0001 versus blood
Sputum Blood IL-8 mRNA 50.5 (28.2-79.5) * 1.4 (0.7-2.0) IL-1β mRNA 0.27 (0.09-0.79) * 0.0013 (0.0002-0.004) TNF-α mRNA 0.16 (0.08-0.38) * 0.0036 (0.002-0.008) Table 4.8 Levels of chemokine and cytokine mRNA expression in resting sputum and blood neutrophils (1 x 105 cell/mL) *p<0.0001 versus blood
92 Sputum Blood A B 0.6 * 0.005 0.5 0.004 0.4 0.003 0.3 0.002 0.2 TLR2 mRNA TLR2 TLR4 mRNA 0.001 0.1 0.000 0.0 Figure 4.10 Levels of TLR4 (A) and TLR2 (B) mRNA expression in resting sputum and blood neutrophils (1 x 105 cell/mL) *p<0.00001 versus blood
Sputum Blood
A B C 3000 # 35 * 175 * 30 150 25 125 2000 20 100 15 75 1000
IL-8 (pg/mL) IL-8 10 50 IL-1beta (pg/mL) IL-1beta
5 TNF-alpha (pg/mL) 25 0 0 0 Figure 4.11 Chemokine (A: IL-8) and Cytokine (B: IL-1β and C: TNF-α) release from LPS stimulated sputum neutrophils compared to LPS stimulated blood neutrophils (1 x 105 cell/mL) *p<0.0001, #p<0.01 versus blood
Sputum Blood IL-8 mRNA 62.2 (44-104) # 28.9 (17.8-51.4) IL-1β mRNA 0.26 (0.16-0.78) * 0.05 (0.01-0.11) TNF-α mRNA 0.2 (0.09-0.37) # 0.07 (0.03-0.18) Table 4.9 Levels of chemokine and cytokine mRNA expression in LPS stimulated sputum and blood neutrophils (1 x 105 cell/mL) #p<0.05 *p<0.00001 versus blood
93 Sputum Blood A * B 0.007 0.35 0.006 0.30 0.005 0.25 0.004 0.20 0.003 0.15 TLR2 mRNATLR2 TLR4 mRNA TLR4 0.002 0.10 0.001 0.05 0.000 0.00 Figure 4.12 Levels of TLR4 (A) and TLR2 (B) mRNA expression in LPS stimulated sputum and blood neutrophils (1 x 105 cell/mL) *p<0.05 versus sputum
4.3.5 Effects of DTT Treatment of Peripheral Blood Neutrophils
To assess whether DTT treatment had any effects on cytokine production and LPS responsiveness, isolated blood neutrophils were treated with DTT for 30 minutes and then the cells were cultured with or without LPS stimulation for 24 hours. Using IL-8 release as a marker, it was clear that DTT treatment had no effect on baseline or LPS stimulated IL-8 release (Figure 4.13).
2500
2000
1500
1000 IL-8 (pg/mL) IL-8 500
0
l S S o rol LP ont Contr C T + LP T TT D D
Figure 4.13 DTT treatment of neutrophils had no effect on IL-8 production
94 4.4 Discussion
These experiments have isolated neutrophils from blood and sputum using a magnetic cell separation technique with CD16 microbeads. This technique successfully isolates blood neutrophils from blood eosinophils, resulting in highly pure neutrophil and eosinophil fractions. The use of magnetic cell separation in sputum samples using CD16 microbeads removes CD16- cells, including eosinophils, lymphocytes, epithelial cells, squamous cells and most macrophages from sputum, enriching a CD16+ fraction that comprised approximately 59% neutrophils and 35% macrophages. Since a proportion of macrophages in the airways are CD16+, this method was unable to separate these cell populations. However the methods result in a ‘non-eosinophilic’ neutrophil enriched cell fraction, which enables the study of these cells in the mechanisms of non- eosinophilic asthma.
These results demonstrate that activation of blood neutrophils is dependent on the presence of LPS. LPS stimulation of peripheral blood neutrophils increased the release of IL-8, IL-1β, TNF-α, and total MMP-9, in keeping with other data [168, 303]. LPS stimulation also increased the level of IL-8, IL-1β, and TNF-α mRNA expression. LPS stimulation increased OSM mRNA expression and protein release from isolated blood neutrophils, which is consistent with other reports of increased OSM release from macrophages and dendritic cells after LPS stimulation [328, 329]. Indeed, the release of IL-8, IL-1β, TNF-α and OSM protein was significantly correlated with the level of mRNA suggesting that the production of these cytokines was dependent on de novo synthesis rather than the release from stored granules.
LPS is known to signal through the TLR4 pathway, whereas TLR2 is the receptor for a range of microbial ligands, for example, gram positive bacteria, peptidoglycan and yeast zymosan [330]. TLR4 and TLR2 expression levels are of importance in facilitating optimal LPS responsiveness. This chapter demonstrates that LPS stimulation increases the levels of TLR2 and TLR4 mRNA in isolated blood neutrophils, which may enhance the functional response of neutrophils to TLR2 & 4 ligands. The upregulation of TLR2 mRNA is consistent with other studies in which neutrophil TLR2 cell surface expression has been shown to be increased by several stimuli including GM-CSF, G- CSF and LPS [142, 331]. TLR4 expression is known to be regulated by LPS [332-334].
95 In isolated blood neutrophils, TLR4 cell surface levels are unchanged after LPS stimulation [331], however at the mRNA level expression was upregulated with LPS stimulation [333], which is consistent with the results of this chapter. These responses are teleologically appropriate for a normally sterile compartment such as the blood circulation. They demonstrate rapid activation by bacterial products, with additional amplification mechanisms also involved to further enhance activation and promote a vigorous inflammatory response that will eradicate invading pathogens, maintain sterility and resolve infection. A different response was observed in the airway compartment however.
Studies using induced sputum so far have mainly focused on the assessment of sputum inflammatory cell counts, properties of sputum supernatant, and staining of sputum cells for activation markers. Only recently has sputum been shown to be a useful source of airway cells for assessment of cellular function by cell culture and cytokine production [335-339]. The majority of studies investigating cytokine release from sputum cells have looked at the global sputum cell populations. This approach is problematic, as only speculations can be made about the cell type involved in the cytokine production, and differences between disease and control states, since all sputum samples are comprised of different proportions of these cells. A further limitation is that using global cell populations limits the ability to investigate the heterogeneity of the inflammatory response in the airway.
This chapter adds to the current knowledge of the activity of sputum cells, specifically neutrophils, and is the first report of CD16 positive magnetic cell separation of neutrophils from induced sputum samples. Sputum neutrophils are a significant source of cytokines, with spontaneous release of high levels of IL-8, IL-1β and TNF-α. However, sputum neutrophils were refractory to further stimulation by LPS. LPS unresponsiveness of sputum cells has been reported previously for TNF-α production from patients with COPD [336], IL-8, IL-1β and TNF-α production from patients with chronic bronchial sepsis [340] and IL-8 production from patients with CF [341]. My data builds on these results to show that sputum cells in asthma as well as healthy people are also non-responsive to LPS stimulation. Subsequent chapters will explore these observations in more detail. Sputum cells have been reported to be responsive to other stimulants including PHA [337] and fMLP [337, 339], suggesting that the
96 unresponsiveness of sputum neutrophils reported in this chapter may be specific for LPS stimulation.
The insensitivity of sputum neutrophils to LPS stimulation resembles the phenomenon of endotoxin tolerance, where activation of the cells with LPS renders the same cells refractory to further challenges with LPS [342]. Endotoxin tolerance is known to occur in monocytes, and the mechanisms of this relate to a decrease in transcription factor activation [332], in the release of some but not all proinflammatory cytokines [343], and the downregulation of components of the TLR4 signalling pathway [344]. Endotoxin tolerance has also been demonstrated to occur in neutrophils, which may represent a reprogramming of neutrophil cell function as a means of adaptation to bacterial infection [345]. This is biologically plausible, as levels of endotoxin in airway samples have been reported previously [255].
This study demonstrates important differences between neutrophil activity in the airways and circulation. Airway neutrophils release high levels of IL-8, IL-1β and TNF- α at rest and have higher levels of IL-8, IL-1β and TNF-α mRNA compared to resting blood neutrophils. Airway neutrophils also differed in their expression of TLR2 mRNA, with airway neutrophils having higher levels of TLR2 mRNA compared to blood, but similar levels of TLR4 mRNA expression. Neutrophils isolated from sputum still release more IL-8, IL-1β and TNF-α compared to LPS stimulated blood neutrophils. This suggests that these cells have come into contact with various stimuli when they were recruited to the airways through the migratory process, and that the neutrophil response is much more complex than simply contact with LPS. The adhesion and migration of neutrophils to the airways involves several steps, which expose neutrophils to additional factors in vivo, including exposure to other cell types, growth factors, adhesion and migration through the endothelium, extracellular matrix and epithelium, which presumably prepares the cell for an effective response to pathogens.
Both resting and LPS stimulated blood neutrophils release a significant amount of total MMP-9, in contrast to sputum neutrophils, which release very low amounts of the protease. This may represent the phenomenon of ‘neutrophil exhaustion’, whereby the total contents of the cells granules have already been released, which may have occurred through the process of the cells migrating to the airways.
97
It is thought that production of IL-1β in sputum neutrophils could enhance the production of IL-8 in an autocrine fashion [303, 340]. This chapter demonstrates that both IL-8 and IL-1β are enhanced in sputum neutrophils compared to blood neutrophils. Therefore it is possible that local mechanisms within the airways can further amplify neutrophil recruitment and activation through cytokine-mediated positive feedback loops. This may be an important mechanism by which there is amplification of neutrophil activation in the airways, which is different to LPS stimulation of peripheral blood neutrophils in vitro.
Interpreting the differences between sputum and blood neutrophils is limited in this study due to a number of factors. Firstly, dithiothreitol (DTT) is widely used to disperse induced sputum samples. There are a number of studies that show that DTT does not affect cell morphology and differential cell counts [346, 347]. Cytokine levels in the sputum supernatant are also unaffected by DTT treatment [348]. However, DTT can affect the expression of cell surface molecules on lymphocytes, neutrophils and eosinophils [349]. Specifically, the expression of CD11a was decreased, CD11b was increased and CD16 was unchanged by DTT treatment of peripheral blood neutrophils [349]. My data demostrates that DTT treatment of blood neutrophils did not affect IL-8 release at rest or with LPS stimulation, and therefore it is unlikely that DTT treatment accounts for the differences between airway and circulating neutrophils in this study.
Secondly, magnetic cell separation of blood neutrophils resulted in a highly pure neutrophil fraction, however the isolation of sputum neutrophils was more troublesome. The CD16 postive fraction of sputum cells contained a proportion of macrophages. It is known that the presence of peripheral blood monocytes can influence the activation of peripheral blood neutrophil preparations. Specifically, the presence of monocytes in peripheral blood neutrophil preparations can modulate the neutrophils response to LPS stimulation, particularly this can cause an increase in the expression of CD11b and the shedding of L-selectin, however the presence of contaminating monocytes does not affect the level of IL-8 mRNA induced by LPS stimulation [350]. This shows that neutrophil signaling was still active to induce a cytokine response which was not changed due to monocytes, however the presence of cell surface adhesion molecules could be modulated, probably due to the production of inflammatory mediators from the
98 small number of contaminating LPS stimulated monocytes. Neutrophil survival induced by LPS is also amplified by the presence of low levels of monocytes, however in the absence of stimuli, neutrophil survival was not altered [350]. The presence of macrophages in the neutrophil enriched sputum cell fraction may manipulate the responses of these neutrophils. The effects found by Sabroe et al were only shown in response to LPS, and did not involve the cytokine, protease responses or TLR mRNA expression of neutrophils as reported in this thesis. I have shown that the sputum neutrophil cell fraction is refractory to LPS stimulation, therefore it is unlikely that the presence of monocytes has influenced the LPS responses. This unresponsiveness is a result of the cells being in the airway, not the lack of purity.
A criticism of positive cell selection is that it the cross linking of cell surface antigens has the potential to result in cellular activation, which has led to the endorsement of negative selection to provide a population of ‘untouched cells’. However there is little evidence to support the advantages of negative selection, and studies are now emerging which have addressed the issue of cell activation after positive selection. Lyons et al [351] demonstrated that there were no significant changes in gene expression that are due to positive selection of cells. Furthermore, negative selection gave far inferior results, thought to be due to reduced purity of the cell fractions. It is unlikely that positive selection has activated the airway and circulating neutrophils in this study, as resting blood neutrophils did not release detectable TNF-α or IL-1β and only low levels of IL-8 in comparison to stimulated cells.
This chapter shows clear and important differences between innate immune activation in the airways compared to the circulation. We can conclude that while circulating neutrophils are responsive to and dependent on activation by LPS, sputum neutrophils remain refractive to LPS stimulation. This unresponsiveness of sputum neutrophils seems specific to LPS stimulation and probably due to a level of tolerance. Sputum neutrophils are activated without stimulation, representing the modulation of the neutrophil phenotype after its migration to the airways. LPS stimulation alone was not sufficient to create a ‘sputum like’ neutrophil, demonstrating the complexities in vivo of the neutrophils’ journey from the blood to the airways.
99 Chapter 5: Innate Immune Responses of Neutrophils in Ageing
5.1 Introduction
It is well known that during healthy ageing, there is a decrease in the immune response, termed immunosenescence. Both innate and adaptive immune system functions decline with advancing age, which leads to an increased susceptibility to infections, cancer, and autoimmune disorders. Ageing is associated with decreased resistance or an impaired ability to respond to bacterial infections, and an increase in systemic inflammation, including the circulating plasma levels of inflammatory mediators including TNF-α [352] and IL-6 [353]. Infections of the respiratory tract, particularly pneumonia, are common in the elderly and a significant cause of morbidity and mortality.
Alterations of the innate immune system in ageing include a decline in neutrophil function, specifically chemotaxis, free radical production and survival/apoptosis, resulting from changes in signal transduction. There is a decrease in the production of ROS by neutrophils after stimulation with various agents, including fMLP, GM-CSF, and LPS [242, 243]. Impaired intracellular signalling is implicated in the reduced production of ROS from older individuals, specifically there is impaired calcium influx [244, 245]. Actin polymerization is also significantly decreased after stimulation of neutrophils with fMLP or PMA in aged subjects compared to younger ones [246]. These differences were associated with altered cell surface marker expression [247].
Neutrophils from elderly subjects are not able to be primed efficiently with GM-CSF [252], which may affect their activity at inflammatory sites. The lifespan of neutrophils can be extended by exposure to a variety of factors, including proinflammatory cytokines such as IL-8 and GM-CSF, and bacterial products such as LPS. Neutrophils from aged individuals cannot be rescued from apoptosis by these agents, specifically GM-CSF, G-CSF, LPS, IL-1, IL-6 and steroids [253, 254]. There is also a significant reduction in neutrophil chemotaxis associated with ageing, however this is specific to the stimulus, and observed with GM-CSF and fMLP but not LPS [249]. When neutrophils were pre-treated with GM-CSF, chemotaxis of neutrophils from young subjects increased significantly, whereas it did not alter in elderly subjects [249].
100 An effect of ageing on innate immune responses of airway neutrophils is not known, but is suggested by the age related increase in BAL and sputum neutrophils. Results in the literature so far suggest that there is age related impairment in response to innate immune stimuli, such as LPS stimulation of whole blood [354] and peripheral blood monocytes [355]. The effect of age on neutrophil activation and expression of innate immune receptors is not known. Specifically, alterations in the secretion of important innate immune mediators, such as IL-8, TNF-α, IL-1β, OSM and MMP-9 from neutrophils have not been investigated in ageing. Furthermore, very little is known in regards to changes in airway neutrophil function and the expression of innate immune receptors with ageing. This study tested the hypothesis that airway and circulating innate immune neutrophil responses, including cytokine, chemokine and protease release and TLR expression is altered with age.
5.2 Methods
Induced sputum and peripheral blood samples were collected from healthy non-smoking control subjects aged between 26 and 77 years. Induced sputum was dispersed and neutrophils were isolated using magnetic cell separation and CD16 microbeads. Neutrophils were isolated from peripheral blood samples via percoll density gradient, erythrocyte lysis and magnetic cell separation using CD16 microbeads. Either the sputum neutrophil enriched fraction [53 (31-62)% neutrophils; 46 (32-68)% macrophages] or highly pure blood neutrophils (100% (96-100%)) were resuspended in phenol red free RPMI 1640(1% FCS) with or without LPS stimulation (E.Coli, 100ng/mL). Cells were cultured at a concentration 1 x 105 and/or 1 x 106 cells/mL. Cells and cell free supernatants were collected at 24 hours and stored at –80˚C until analysis. IL-8, IL-1β, TNF-α and OSM were measured using a commercial sandwich ELISA. MMP-9 was measured using a commercial fluorescent ELISA. RNA was extracted and mRNA quantified by real-time PCR with commercial primers and probes for TLR4, TLR2, IL-8, IL-1β, OSM and TNF-α, and expressed as 2-∆∆Ct. Data was analysed by Stata 9. All data, unless otherwise stated, is non-parametric and reported as the median
(Q1-Q3). In the case of age, FEV1% predicted and FEV1/FVC data is reported as mean (SD) and significant differences were determined using Student’s t test. For all other data significant differences (p<0.05) were detected with the 2-sample test Wilcoxon rank sum or the multiple sample Kruskal Wallis test. For categorical data (Gender and
101 Atopy) Fischer’s exact test was applied (p<0.05 considered significant). For more detailed methods please refer to Chapter 2 of this thesis.
5.3 Results
Samples were successfully collected from 13 healthy control subjects with an age range of 26 to 77 years. Blood samples were collected from all participants, and sputum samples were successfully collected from 11 participants. Neutrophils and eosinophils were isolated from all blood samples (n=13), and neutrophils were successfully isolated from sputum for 69% (n=9) of the samples. For the analysis the subjects were divided into 2 groups above and below the mean age of the subjects (55 years).
5.3.1 Clinical Features
The characteristics of the study participants are summarised in Table 5.1. The younger group had a mean age of 35 years, and the older group had a mean age of 68 years. Younger subjects tended to be more atopic than the older group (80% versus 38%). All subjects had lung function within the normal range.
Under 55 Over 55 p
n 5 8
Age years, mean (SD) 35 (8) 68 (8) <0.00001
Gender M | F 3 | 2 3 | 5 0.59
Atopy n (%) 4 (80) 3 (38) 0.27
FEV1 % predicted 101 (16) 99 (18) 0.86
FEV1/FVC % 82 (7) 77 (7) 0.22 Table 5.1 Clinical characteristics of healthy control participants under and over 55 years of age
102 5.3.2 Inflammatory Cells
Total and differential cell counts obtained from induced sputum samples are shown in Table 5.2. There were no significant differences in the sputum differential cell counts between the younger and older groups.
Under 55 Over 55 p* n 5 5 Total cell count x 106/mL 3.4 (2.2-3.9) 3.2 (2.4-5.6) 0.86 Viability (%) 80 (79.2-81.4) 77.9 (73.9-87.8) 0.93 Neutrophils, % 30.5 (29.3-36.5) 34.8 (14.5-58.5) 0.47 Neutrophils 104/mL 65.9 (65.7-100.2) 135.3 (49.1-179.8) 0.36 Eosinophils, % 0 (0-0.5) 0 (0-0.2) 0.53 Eosinophils 104/mL 0 (0-1.6) 0 (0-0.6) 0.53 Macrophage, % 66.1 (58.4-66.8) 58.7 (35-83.3) 0.72 Macrophages 104/mL 226 (142.8-228.3) 196.7 (131.7-307.4) 0.72 Lymphocytes, % 1 (0-2) 3 (1.3-3.7) 0.14 Lymphocytes 104/mL 3.7 (0-4.2) 8.1 (2.6-17.9) 0.27 Columnar epithelial cells, % 1.3 (1.2-11.3) 1.4 (1.2-3.3) 0.85 Columnar epithelial cells 104/mL 4.6 (4.3-20.3) 7.3 (3.9-11.4) 0.72 Squamous cells, % 16.5 (5.1-16.6) 5.1 (1.8-18.1) 0.58 Table 5.2 Induced sputum inflammatory cell counts healthy control participants under and over 55 years of age
5.3.3 Sputum Supernatant IL-8
Levels of IL-8 were measured in the sputum supernatant by ELISA. There was no difference between the level of IL-8 detected in the sputum supernatant between younger and older healthy controls (Figure 5.1).
103 Under 55 years Over 55 years
7500
5000
2500 IL-8 (pg/mL) IL-8
0 Figure 5.1 Level of IL-8 detected in sputum supernatant from participants under 55 (n=5) and over 55 (n=6)
5.3.4 Chemokine and Cytokine Production from Sputum Neutrophils
Neutrophils isolated from sputum samples did not respond to LPS stimulation with an increase in chemokine and cytokine production (See Chapter 4), therefore the results reported are from sputum neutrophils at rest. Resting sputum neutrophils isolated from participants aged over 55 released significantly more TNF-α compared to those aged under 55 years (Figure 5.2C). The level of IL-8 and IL-1β were also higher in the older group; however this did not reach statistical significance (Figure 5.2A&B). Gene expression of IL-8, IL-1β and TNF-α was significantly increased in participants that were over 55 compared to those aged under 55 years (Table 5.3).
104 Under 55 years
A B Over 55 years 9000 250
7500 200
6000 150 4500 100 3000 IL-8 (pg/mL) IL-8
IL-1beta (pg/mL) 50 1500 0 0 C * 1400 1200 1000 800 600 400
TNF-alpha (pg/mL) 200 0 Figure 5.2 Chemokine (A: IL-8) and Cytokine (B: IL-1β, C: TNF-α) release from resting sputum neutrophils (105 cells/mL) isolated from participants under (n=5) and over 55 (n=4) *p<0.05 versus under 55
Under 55 Over 55 p IL-8 79.5 (49-82.3) 196.7 (186.1-347.3) 0.034 IL-1β 0.79 (0.36-0.92) 2.65 (1.8-10) 0.034 TNF-α 0.22 (0.11-0.35) 0.54 (0.5-1.9) 0.034 Table 5.3 Levels of chemokine and cytokine mRNA expression in resting sputum neutrophils from healthy control participants under (n=5) and over (n=4) 55 years of age
5.3.5 Chemokine and Cytokine Production from Peripheral Blood Neutrophils
Isolated blood neutrophils were cultured with or without LPS stimulation (100ng/mL). The cells and cell culture supernatants were collected after 24 hours and stored at -80°C until analysis. Resting blood neutrophils did not release detectable IL-1β, TNF-α or OSM. All resting blood neutrophil samples released detectable levels of IL-8, however
105 the amount of IL-8 released did not differ between the different age groups [Under 55: 83.4pg/mL (71.4-116.1) Over 55: 114.7pg/mL (64.5-197.4) p= 0.66].
The level of IL-8, IL-1β, TNF-α and OSM gene expression was also examined in resting neutrophils (Table 5.4). There was no difference in cytokine gene expression in resting blood neutrophils between healthy controls under 55 compared with those subjects over 55.
Under 55 Over 55 p IL-8 2.2 (1.6-2.3) 3.2 (1.9-4.3) 0.57 IL-1β 0.003 (0.002-0.006) 0.008 (0.006-0.02) 0.17 TNF-α 0.003 (0.002-0.004) 0.002 (0.001-0.006) 0.57 OSM 0.08 (0.05-0.09) 0.2 (0.1-0.2) 0.29 Table 5.4 Levels of chemokine and cytokine mRNA expression in resting blood neutrophils isolated from participants under (n=5) and over (n=8) years of age
Isolated LPS stimulated blood neutrophils from older healthy control participants released significantly less IL-1β, TNF-α and OSM compared to younger healthy control participants (Figure 5.3). There was also a trend for lower levels of IL-8 released from older versus younger healthy control participants (p=0.057, Figure 5.3A). No difference was found between younger and older healthy controls when gene expression for these cytokines was analysed (Table 5.5).
106 Under 55 years A B Over 55 years 17500 90 80 15000 70 12500 60 10000 50 7500 40 * 30
IL-8 (pg/mL) IL-8 5000 20 IL-1beta (pg/mL) 2500 10 0 0 C D 120 140 100 120
80 100 80 60 * * 60 40 40 OSM (pg/mL)OSM
TNF-alpha (pg/mL) 20 20 0 0 Figure 5.3 Chemokine (A: IL-8) and Cytokine (B: IL-1β, C: TNF-α D: OSM) release from LPS stimulated blood neutrophils (106 cells/mL) isolated from participants under (n=5) and over (n=8) years of age *p<0.05 versus Under 55
Under 55 Over 55 p IL-8 68.6 (52.5-95.3) 48.3 (12.5-79.3) 0.37 IL-1β 0.12 (0.08-0.14) 0.18 (0.06-0.25) 0.22 TNF-α 0.1 (0.05-0.12) 0.13 (0.06-0.17) 0.46 OSM 0.84 (0.64-0.98) 0.66 (0.22-0.84) 0.25 Table 5.5 Levels of chemokine and cytokine mRNA expression in LPS stimulated blood neutrophils isolated from participants under (n=5) and over (n=8) years of age
5.3.6 Total MMP-9 Release from Peripheral Blood Neutrophils
Resting blood neutrophils isolated from older healthy control participants released significantly more total MMP-9 compared to younger healthy controls (Table 5.6). Once the cells were stimulated with LPS there was no difference in the amount of total MMP-9 released into the culture supernatant.
107
Under 55 Over 55 P value Neutrophils 176 (145-262) 484 (374-589) 0.0054 Neutrophils + LPS 452 (408-545) 692 (544-820) 0.14 Table 5.6 Total MMP-9 release from blood neutrophils (106 cells/mL) isolated from participants under (n=5) and over (n=8) years of age
5.3.7 Neutrophil TLR Expression
The levels of TLR4 and TLR2 mRNA were measured in resting sputum neutrophils and resting and LPS stimulated blood neutrophils. Older healthy control participants had a similar level of TLR4 mRNA detected in their sputum neutrophils in comparison to younger healthy controls. However, older healthy controls had significantly higher TLR4 mRNA expression in resting blood neutrophils compared to younger healthy controls. The level of TLR4 mRNA did not differ in LPS stimulated blood neutrophils (Table 5.7). Gene expression for TLR2 was significantly increased in sputum neutrophils isolated from older healthy controls compared to younger healthy controls. The level of TLR2 gene expression did not differ with age for both resting and LPS stimulated blood neutrophils (Table 5.8).
Under 55 Over 55 P value Sputum Neutrophils 0.0041 (0.0027-0.0042) 0.014 (0.004-0.026) 0.21 Resting Blood 0.0021 (0.002-0.003) 0.0063 (0.005-0.008) 0.042 Neutrophils LPS Stimulated 0.006 (0.005-0.01) 0.012 (0.009-0.02) 0.19 Blood Neutrophils Table 5.7 Relative mRNA levels of TLR4 in sputum and blood neutrophils isolated from participants under (n=5) and over (n=8) years of age
108
Under 55 Over 55 p Sputum Neutrophils 0.13 (0.01-0.26) 0.69 (0.44-1.21) 0.0495 Resting Blood 0.028 (0.009-0.054) 0.066 (0.032-0.18) 0.17 Neutrophils LPS Stimulated 0.34 (0.16-0.39) 0.43 (0.25-0.88) 0.17 Blood Neutrophils Table 5.8 Relative mRNA levels of TLR2 in sputum and blood neutrophils isolated from participants under (n=5) and over (n=8) years of age
Results are summarised in Table 5.9.
Airway Resting Blood LPS Stimulated
Neutrophils Neutrophils Blood Neutrophils IL-8 protein Higher Unchanged Lower IL-8 mRNA Higher Unchanged Unchanged IL-1β protein Higher Not Detected Lower IL-1β mRNA Higher Unchanged Unchanged TNF-α protein Higher Not Detected Lower TNF-α mRNA Higher Unchanged Unchanged OSM protein Not Detected Not Detected Lower OSM mRNA N/A Unchanged Unchanged Total MMP-9 N/A Higher Unchanged TLR4 mRNA Unchanged Higher Unchanged TLR2 mRNA Higher Unchanged Unchanged Table 5.9 Summary of results for the comparison between older versus younger healthy control participants
Significantly lower in older healthy volunteers versus younger healthy volunteers Trend for lower levels in older healthy volunteers versus younger healthy volunteers Significantly higher in older healthy volunteers versus younger healthy volunteers Trend for higher levels in older healthy volunteers versus younger healthy volunteers N/A means not measured
109 5.4 Discussion
The major findings of this chapter demonstrate that there are distinct alterations in neutrophil function that occur with age. Airway neutrophils from older subjects release higher levels TNF-α, and IL-8 and IL-1β. The higher levels of these innate immune cytokines were paralleled by a significant up regulation of the corresponding mRNA transcripts. There was also an up regulation of TLR2 gene expression in airway neutrophils isolated from older subjects compared to their younger counterparts. Resting blood neutrophils from older subjects release significantly higher levels of MMP-9 and have higher levels of TLR4 gene expression compared to the younger group. Blood neutrophils from older subjects had a decreased response to LPS stimulation in terms of cytokine release, however there was no difference in the mRNA levels for each cytokine tested, suggesting the involvement of post-transcriptional mechanisms.
The results support an age-related enhancement of innate immune activation responses in airway neutrophils. Although there is a considerable amount of information regarding systemic immune responses in ageing, there is relatively little that is known about compartmentalised innate immune responses in the lung, especially those involving airway neutrophils. There are increases in the number of both lymphocytes and neutrophils in BAL fluid of healthy elderly subjects who had never smoked compared to younger individuals [356-358]. This is associated with a shift in T cell subsets with an increase in the numbers of CD4+ lymphocytes which demonstrated an increased expression of cell surface activation markers HLA-DR and CD69 [357]. There also is an increase in BAL levels of IL-6 [358], IL-8 and neutrophil elastase bound to α1- antiprotease [356], as well as an increase in superoxide release from BAL cells stimulated with PMA [358]. Many of these findings suggest the presence of a low-grade mucosal inflammation that may be present in the ageing healthy human lung, as well as increase in activity in infiltrating lung cells. This supports the increased activity of airway neutrophils in older subjects found in this study. These changes may be beneficial for immune responses to infections, however they could also represent age- associated immune dysregulation, altered responses to environmental triggers, structural changes of the lung, or a decline in mucociliary clearance [359].
110 It is possible that ageing alters the balance of pro and anti-inflammatory cytokines released from inflammatory cells including neutrophils. This balance is crucial during an immune response and for the resolution of inflammation. Although we have not measured the concentrations of anti-inflammatory cytokines released from airway neutrophils in this study, we have found an increase in the release of proinflammatory cytokines with age, which could be associated with a decrease in anti-inflammatory cytokine release. Impaired production of IL-10 has been demonstrated in alveolar macrophages from aged rats, and correlated with an accentuated inflammatory response in the lungs of these rats following challenge with carrageenan, a pharmacological inducer of local inflammation, in comparison to young rats [360]. Lung inflammation in the old rats in this study was predominantly neutrophilic with increases in neutrophil infiltration and activation as measured by an increased level of myeloperoxidase [360].
During airway infection or inflammation, neutrophils migrate from the blood to the airways where they play a crucial role in innate immune defence. In the pulmonary circulation, neutrophil migration occurs through at least two pathways that are either CD11b/CD18-dependent or -independent. In vitro models have demonstrated that the bacterial derived chemoattractant fMLP stimulates CD18-dependent neutrophil migration, whereas the host derived chemoattractants IL-8 and LTB4 stimulate CD18- independent neutrophil migration [111]. It is unclear if one pathway is preferentially used over the other in relation to lung neutrophil influx. However, a recent study in which cystic fibrosis sputum, which is a cocktail of both bacterial and host derived chemoattractants, was added to neutrophils in vitro there was preferential induction of CD18-independent migration [112]. Tortorella et al [361] demonstrated a larger contribution of CD18-dependent versus CD18-independent pathways for neutrophil adhesion to fibronectin (FN) that occurred in neutrophils of older subjects, regardless of the stimulus. In this study, neutrophils of aged individuals show a significant impairment of FN primed superoxide release as a result of GM-CSF or fMLP, but not TNF-α or PMA stimulation. FN primed superoxide release was suppressed to a greater degree in TNF-α compared to GM-CSF stimulated neutrophils following incubation with anti-CD18 monoclonal antibody [361]. Different pathways of cell migration may impact on the activity of the cells once they reach the inflammatory site such as the airways.
111 Matrix Metalloproteinases (MMPs) are a family of zinc containing enzymes that are capable of degrading proteins of the extracellular matrix (ECM), and modulating chemokine and cytokine function. MMPs facilitate leukocyte recruitment through remodelling the ECM, the conversion of inactive cytokines/chemokines to their active form and the establishment of chemotactic gradients that guide leukocyte chemotaxis across endothelial and epithelial barriers [362]. Resting blood neutrophils from older individuals release significantly more total MMP-9 when compared to their younger counterparts. This may reflect a greater degree of neutrophil activation. Several proinflammatory mediators including TNF-α stimulate the release of MMP-9 from neutrophils [363], and TNF-α is known to be increased in ageing plasma [352].
Resting blood neutrophils from the older group had significantly increased TLR4 gene expression compared to the younger group. Impaired functioning and expression of TLRs in ageing has been reported to occur in mouse macrophages [248]. Fulop and colleagues [249] investigated the presence of TLR2 and TLR4 in neutrophil lipid rafts, and demonstrated that TLR4 was increased in unstimulated raft and non-raft fractions in older subjects compared to younger counterparts. Furthermore, LPS was shown to increase the expression of TLR4 in neutrophils obtained from young subjects, in both raft and non-raft fractions, whereas there was no reorganisation of TLR4 after LPS stimulation of neutrophils from older subjects [249].
The increased levels of TLR4 mRNA and total MMP-9 released from neutrophils isolated from older individuals is consistent with the ‘inflamm-ageing’ theory which suggests that there is a low grade of inflammation present with increasing age [250], and the slightly stimulated state of neutrophils demonstrated from aged individuals [251].
My results show a decreased cytokine response to LPS of circulating neutrophils in older subjects. In support of this are other studies demonstrating decreased LPS-induced TNF-α and IL-1β production in whole blood [354] and isolated peripheral blood monocytes [355]. Decreased production of these cytokines from monocytes after LPS stimulation has also been reported in animal studies [364, 365]. Other LPS induced neutrophil functions are altered in ageing, including the inability of LPS to rescue
112 neutrophils from apoptosis, and the decreased production of superoxide in response to LPS stimulation.
Altered signal transduction pathways may cause the deficient TLR induced cytokine release of circulating neutrophils in this study. A number of negative signalling regulators such as IRAK-M, SOCS-1, A20 and Tollip have been shown to play an important role in TLR signal transduction [366]. Anti-inflammatory cytokines may also be important in limiting cytokine responses in ageing. There is a known increase in anti- inflammatory factors particularly prostaglandin E2 (PGE2) with ageing in both mice and humans, which is due to an enhanced expression of cyclooxygenase [367, 368]. The anti-inflammatory properties of PGE2 include the suppression of IL-12, enhancement of IL-10 production and decreased class II MHC expression on antigen presenting cells.
Low levels of proinflammatory cytokines in response to stimulation may cause a weak local inflammatory response, which may play a role in the age-associated increased incidence of severe and invasive infections. This could be due to depressed inflammatory responses, and reduced recruitment and/or activation of effector cells at the site of inflammation.
This chapter has described the effects of age on the innate immune responses of airway and circulating neutrophils, particularly the secretion of cytokines (IL-1β, TNF-α and OSM), chemokines (IL-8) and proteases (MMP-9) and the expression of TLR2 and TLR4. With advancing age, there are alterations in the innate immune responses of neutrophils characterised by enhanced spontaneous activation of both circulating and airway neutrophils and a decreased response of circulating neutrophils to LPS. Increased expression of TLR2 is a potential mechanism for the increased cytokine response in airway neutrophils from older individuals.
113 Chapter 6: Innate Immune Responses of Neutrophils in Airway Disease
6.1 Introduction
Migration of neutrophils into the airways is an important function of innate immunity and neutrophils are commonly found in the airways of healthy people. However, increased infiltration of neutrophils into the airways occurs in both acute and chronic respiratory diseases including asthma and COPD, where these cells contribute significantly to inflammation through the release of proinflammatory mediators including cytokines, chemokines and proteases.
Neutrophils are a hallmark feature of COPD with their increased presence in the airways of affected patients [69, 70]. The presence of neutrophils in the airways of COPD patients is associated with increased levels of neutrophilic inflammatory mediators including IL-8, TNF-α and MMP-9 [71-73]. Markers of airway neutrophilic inflammation are correlated with COPD disease progression [74], and the degree of neutrophilic inflammation is also correlated with clinical severity [75]. Neutrophilic airway inflammatory markers are further increased with COPD exacerbations [76]. Peripheral blood neutrophils from subjects with COPD show altered activity in both stable disease and exacerbations, including increased presence of cell surface adhesion molecule expression [77-79], upregulation of genes relating to inflammation [80] and enhanced respiratory burst [78].
A role for neutrophils in the pathogenesis of asthma is being increasingly recognised. Neutrophils are increased in more severe forms of asthma including in subjects with severe asthma requiring intubation [41], sudden onset fatal asthma and life threatening asthma [49-51]. Neutrophilic inflammation is increased with increasing severity of asthma [53], as well as with exacerbations of asthma [48]. More recently reported are a subgroup of patients with stable asthma that have persistent neutrophilic inflammation [9, 32, 33, 53]. In addition, peripheral blood neutrophils from subjects with allergic asthma release increased amounts of myeloperoxidase upon stimulation with fMLP [54], which is thought to be mediated by activation of IgE receptors [55]. There is also increased production of reactive oxygen species from peripheral blood neutrophils obtained from subjects with asthma both spontaneously [56] and with stimulation [57].
114
While it is clear that neutrophils play an important role in promoting airway inflammation in both asthma and COPD, the precise mechanisms underlying this remain unclear. It is possible that abnormal neutrophil function would contribute to the enhanced airway inflammation seen in asthma and COPD. To further investigate the involvement of both airway and circulating neutrophils in asthma and COPD, we have investigated the release of innate immune mediators and the expression of Toll-like receptors 2 and 4 at both resting and stimulated states. We hypothesised that airway and circulating neutrophils from participants with asthma and COPD would have enhanced activation which would result in increased release of innate immune mediators compared to age matched healthy controls.
6.2 Methods
Induced sputum and peripheral blood samples were collected from subjects with asthma, COPD and healthy control volunteers. The group of healthy controls was selected to be of similar age and gender as the participants with asthma. Induced sputum was dispersed and neutrophils were isolated using magnetic cell separation and CD16 microbeads. Neutrophils were isolated from peripheral blood samples via percoll density gradient, erythrocyte lysis and magnetic cell separation using CD16 microbeads. Isolated neutrophils were cultured in RPMI 1640 (1% FCS) with or without LPS stimulation (E.Coli, 100ng/mL). Cells were cultured at a concentration 1 x 105 or 1 x 106 cells/mL. Cells and cell free supernatants were collected at 24 hours and stored at – 80˚C until analysis. IL-8, IL-1β, TNF-α and OSM were measured using a commercial sandwich ELISA. MMP-9 was measured using a commercial fluorescent ELISA, and zymography. RNA was extracted and mRNA quantified by real-time PCR with commercial primers and probes for TLR4, TLR2, IL-8, IL-1β, OSM and TNF-α. Data was analysed by Stata 9. All data, unless otherwise stated, is non-parametric and reported as the median (Q1-Q3), or in the case of figures the median as the bar and the upper quartile as the error bars. In the case of age, FEV1% predicted and FEV1/FVC data is reported as mean (SD) and significant differences were determined using Student’s t test. For categorical data (Gender and Atopy) Fischer’s exact test was applied (p<0.05 considered significant). For all other data, significant differences (p<0.05) were detected with the 2-sample Wilcoxon rank sum test. To adjust for
115 multiple group comparisons, post-hoc analysis was conducted using kwallis2 (Stata 9). For more detailed methods please refer to Chapter 2 of this thesis.
6.3 Results
6.3.1 Comparison of Asthma and Healthy Controls
Samples were collected from 17 participants with asthma, and 11 age and gender matched healthy controls. Blood samples were successfully collected from all participants, and sputum samples were successfully obtained from 93% (n=26) of participants. Neutrophils and eosinophils were isolated from all blood samples (n=28), and neutrophils were isolated from 81% (n=21) of sputum samples.
6.3.1.1 Clinical Features
The characteristics of the study participants are summarised in Table 6.1. Subjects with asthma tended to be more atopic (85% versus 55%) in comparison to healthy controls. The healthy control group had pulmonary function parameters within the normal range, and this was significantly reduced in participants with asthma (FEV1% predicted and
FEV1/FVC: Table 1). All subjects with asthma were on inhaled corticosteroids in the medium dosage range.
116 Healthy Asthma p
n 11 17
Age years, mean (SD) 56 (18) 59 (16) 0.65
Gender M | F 5 | 6 8 | 9 1.0
Atopy n (%) 6 (55) 14 (82) 0.2
FEV1 % predicted 98 (17) 71 (19) 0.0007
FEV1/FVC % 77 (7) 67 (10) 0.0086 ICS dose (μg) ‡‡ - 1000 (400-2000) Table 6.1 Clinical characteristics of participants with asthma and healthy control subjects ‡‡Inhaled corticosteroid (ICS) dose is calculated as 1μg of beclomethasone = 1μg budesonide = 0.5μg fluticasone.
6.3.1.2 Inflammatory Cells
Total and differential cell counts obtained from induced sputum samples are shown in Table 6.2. Subjects with asthma had a significantly increased proportion and absolute number of sputum eosinophils in comparison to healthy controls (p<0.05). The proportion of macrophages was lower in subjects with asthma, however absolute numbers did not differ between groups.
117 Healthy Asthma p n 9 17 Total cell count x 106/mL 3.7 (2.4-5.6) 6.9 (3.3-10.6) 0.11 Viability (%) 79.2 (73.9-84.2) 69.2 (49.7-82.9) 0.21 Neutrophils, % 30.5 (14.5-37.4) 40.5 (23.8-58.5) 0.19 Neutrophils 104/mL 90.9 (49.1-150.7) 185.9 (74.5-589.7) 0.09 Eosinophils, % 0 (0-0.3) 1.5 (0.3-3.9) 0.006 Eosinophils 104/mL 0 (0-1.4) 10.6 (0.8-23.5) 0.006 Macrophages, % 66.1 (58.4-82.4) 48.3 (40.3-54) 0.009 Macrophages 104/mL 226 (172.8-307.4) 240.1 (148.8-402.7) 0.69 Lymphocytes, % 2.0 (1.0-3.2) 1.2 (0.7-1.9) 0.28 Lymphocytes 104/mL 4.2 (2.6-9.4) 6.9 (3.4-10.1) 0.77 Columnar epithelial cells, % 1.3 (1.2-3.3) 4.3 (1.9-13.1) 0.15 Columnar epithelial cells 104/mL 6.7 (4.3-11.4) 50.7 (7.4-79) 0.09 Squamous cells, % 5.1 (1.8-16.5) 6.8 (1.7-11.1) 0.61 Table 6.2 Induced sputum inflammatory cell counts from participants with asthma compared to healthy controls
6.3.1.3 Sputum Supernatant IL-8
Subjects with asthma had significantly increased levels of sputum supernatant IL-8 (Figure 6.1). Healthy * 30 Asthma
20
10 IL-8 (ng/mL) IL-8
0 Figure 6.1 Levels of IL-8 detected in sputum supernatant
118
6.3.1.4 Chemokine and Cytokine Production from Sputum Neutrophils
In order to analyse local chemokine and cytokine production, isolated sputum neutrophils were cultured at a concentration of 1 x 105 cells/mL with or without LPS (100ng/mL). LPS stimulation did not alter cytokine or chemokine production from sputum neutrophils (See Chapter 4), and therefore differences between healthy control and asthma responses were examined in resting sputum neutrophils. Sputum neutrophils isolated from subjects with asthma released significantly less IL-8 (Figure 6.2A), IL-1β (Figure 6.2B) and TNF-α (Figure 6.2C) protein when compared to healthy controls.
AB 7000 100 6000 75 5000 4000 50 (pg/mL)
3000 * β *
IL-8 (pg/ml) IL-8 2000 IL-1 25 1000 0 0
C Healthy 600 Asthma 500
400
(pg/mL) 300 α 200 * TNF- 100
0
Figure 6.2 Chemokine (A: IL-8) and Cytokine (B: IL-1β, C: TNF-α) release from resting sputum neutrophils (105 cells/mL) *p<0.05 versus healthy controls
Messenger RNA was extracted from isolated and cultured sputum neutrophils. Relative levels of mRNA for IL-8, IL-1β and TNF-α were significantly decreased in participants with asthma compared to healthy controls (Table 6.3).
119
Healthy Asthma p IL-8 134.4 (77.2-196.7) 39.3 (29.6-58.2) 0.022 IL-1β 1.34 (0.68-2.65) 0.23 (0.09-0.34) 0.025 TNF-α 0.47 (0.16-0.54) 0.14 (0.06-0.25) 0.035 Table 6.3 Levels of chemokine and cytokine mRNA expression in resting sputum neutrophils from participants with asthma compared to healthy controls
6.3.1.5 Chemokine and Cytokine Production from Peripheral Blood Neutrophils
Resting blood neutrophils did not release detectable levels of TNF-α (<15.6pg/mL) or IL-1β (<3.3pg/mL). 93% of resting blood neutrophils did not release detectable levels of OSM (<32pg/mL). All resting blood neutrophils released detectable levels of IL-8, however the amount of IL-8 released did not differ between asthma and control groups [Healthy: 83.4 (57.9-145.1) pg/mL, Asthma: 81.1 (56.5-178.9) pg/mL; p=0.869]. Levels of the corresponding mRNA transcripts were also measured in resting neutrophils. Levels of mRNA for IL-8 and TNF-α did not differ between healthy controls and participants with asthma. However, subjects with asthma had significantly lower levels of IL-1β mRNA in blood neutrophils at rest in comparison to healthy controls (Table 6.4).
Healthy Asthma p IL-8 2.27 (1.59-4.26) 2.41 (1.28-3.66) 0.80 IL-1β 0.007 (0.003-0.019) 0.002 (0.0007-0.004) 0.026 TNF-α 0.002 (0.001-0.006) 0.002 (0.002-0.009) 0.48 OSM 0.18 (0.07-0.24) 0.13 (0.1-0.17) 0.6 Table 6.4 Levels of chemokine and cytokine mRNA expression in resting neutrophils from subjects with asthma and healthy controls
LPS stimulation induced cytokine and chemokine protein release in both participants with asthma and healthy controls. LPS stimulated blood neutrophils from subjects with asthma released similar amounts of IL-8, IL-1β, and TNF-α when compared to healthy control samples (Figure 6.3A, B, C respectively). Production of OSM from isolated
120 blood neutrophils was less in participants with asthma compared to healthy controls, however this did not reach statistical significance (p=0.059, Figure 6.3D).
Healthy ABAsthma
9000 40 8000 7000 30 6000 5000 20 4000 (pg/mL) β 3000 IL-8 (pg/mL) IL-8 2000 IL-1 10 1000 0 0
CD 45 75 40 35 30 50 25 (pg/mL) 20 α 15 25 OSM (pg/mL) OSM
TNF- 10 5 0 0
Figure 6.3 Chemokine (A: IL-8) and Cytokine (B: IL-1β, C: TNF-α D: OSM) release from LPS stimulated blood neutrophils (106 cells/mL)
Levels of the corresponding mRNA transcripts were also measured in LPS stimulated blood neutrophils. Levels of mRNA for IL-8, TNF-α, and OSM in LPS stimulated blood neutrophils did not differ between healthy controls and participants with asthma. However, subjects with asthma had significantly lower levels of IL-1β mRNA in LPS stimulated blood neutrophils in comparison to healthy controls (Table 6.5).
Healthy Asthma p IL-8 54.2 (40.6-95.3) 36.8 (12.6-64) 0.15 IL-1β 0.14 (0.06-0.2) 0.06 (0.01-0.1) 0.018 TNF-α 0.12 (0.06-0.17) 0.11 (0.05-0.17) 0.72 OSM 0.66 (0.35-0.84) 0.61 (0.36-0.78) 0.6 Table 6.5 Levels of chemokine and cytokine mRNA expression in LPS stimulated neutrophils from participants with asthma compared with healthy controls
121
6.3.1.6 Total MMP-9 Release from Peripheral Blood Neutrophils
Both resting and LPS stimulated blood neutrophils release high levels of total MMP-9 into the culture supernatant. Levels of Total MMP-9 released from neutrophils were similar in subjects with asthma and healthy controls (Table 6.6).
Healthy Asthma p Neutrophils 369 (176-554) 250 (213-348) 0.46 Neutrophils + LPS 577 (452-857) 516 (432-637) 0.3 Table 6.6 Total MMP-9 release from blood neutrophils (106 cells/mL) isolated from participants with asthma compared with healthy controls
Resting and stimulated blood neutrophils released low levels of active MMP-9 that did not differ between healthy controls and participants with asthma (Table 6.7).
Healthy Asthma p Neutrophils 3.3 (1.9-4.1) 3.3 (2.7-6.1) 0.38 Neutrophils + LPS 4 (2.4-7.2) 4.7 (3-6.7) 0.61 Table 6.7 Active MMP-9 release from blood neutrophils (106 cells/mL) isolated from participants with asthma compared with healthy controls
6.3.1.7 Neutrophil TLR Expression
Levels of TLR4 and TLR2 mRNA were measured in resting sputum neutrophils and resting and LPS stimulated blood neutrophils. Participants with asthma had significantly lower levels of TLR4 mRNA detected in sputum neutrophils in comparison to healthy controls (Table 6.8). Resting and LPS stimulated blood neutrophils from participants with asthma had similar levels of TLR4 mRNA in comparison to healthy controls (Table 6.8). Participants with asthma had similar levels of sputum neutrophil TLR2 mRNA expression compared with healthy controls (Table 6.9). However, participants
122 with asthma had significantly increased TLR2 mRNA in both resting and LPS stimulated blood neutrophils compared to healthy controls (Table 6.9).
Healthy Asthma p Sputum Neutrophils 0.004 (0.004-0.014) 0.002 (0.001-0.003) 0.0348 Resting Blood 0.005 (0.002-0.008) 0.003 (0.002 –0.004) 0.31 Neutrophils LPS Stimulated Blood 0.009 (0.005-0.01) 0.007 (0.005-0.01) 0.56 Neutrophils Table 6.8 Relative mRNA levels of TLR4 in sputum and blood neutrophils from participants with asthma compared with healthy controls
Healthy Asthma p Sputum Neutrophils 0.44 (0.13-0.69) 0.21 (0.07-0.46) 0.61 Resting Blood 0.057 (0.03-0.09) 0.02 (0.01-0.03) 0.008 Neutrophils LPS Stimulated Blood 0.39 (0.25-0.84) 0.21 (0.12-0.31) 0.0055 Neutrophils Table 6.9 Relative mRNA levels of TLR2 in sputum and blood neutrophils from participants with asthma compared with healthy controls
Results of this section are summarised in Table 6.10.
123 Airway Resting Blood LPS Stimulated
Neutrophils Neutrophils Blood Neutrophils IL-8 protein Lower Unchanged Unchanged IL-8 mRNA Lower Unchanged Unchanged IL-1β protein Lower Not Detected Unchanged IL-1β mRNA Lower Lower Lower TNF-α protein Lower Not Detected Unchanged TNF-α mRNA Lower Unchanged Unchanged OSM protein Not Detected Not Detected Unchanged OSM mRNA N/A Unchanged Unchanged Total MMP-9 N/A Unchanged Unchanged Active MMP-9 N/A Unchanged Unchanged TLR4 mRNA Lower Unchanged Unchanged TLR2 mRNA Unchanged Lower Lower Table 6.10 Summary of results for the comparison between participants with asthma and age matched healthy controls
Significantly lower in subjects with asthma versus healthy controls N/A means not measured as very low amounts detected in a small sample group
124 6.3.2 Comparison of COPD and Healthy Controls
Samples were collected from 5 subjects with COPD, and 8 age matched healthy controls. Blood samples were successfully collected from all participants, and sputum samples were successfully obtained from 85% (n=11) of subjects. Neutrophils and eosinophils were isolated from all blood samples (n=13), and neutrophils were isolated for 69% (n=9) sputum samples.
6.3.2.1 Clinical Features
The characteristics of the study participants are summarised in Table 6.11. The healthy control group had pulmonary function parameters within the normal range, and this was significantly reduced in participants with COPD (FEV1% predicted and FEV1/FVC: Table 11).
Healthy COPD p n 8 5
Age years, mean (SD) 68 (8) 68 (7) 0.97
Sex M | F 3 | 5 3 | 2 0.59
Atopy n (%) 3 (38) 3 (60) 0.59
FEV1 % predicted 99 (18) 69 (14) 0.01
FEV1/FVC % 77 (8) 66 (6) 0.023 Table 6.11 Clinical characteristics of participants with COPD compared to healthy controls
6.3.2.2 Inflammatory Cells
There were no significant differences in the inflammatory cell counts between participants with COPD and healthy controls (Table 6.12).
125 Healthy COPD p n 6 5 Total cell count x 106/mL 3.2 (2.4-5.6) 3.7 (3.2-6.5) 0.41 Viability (%) 77.9 (73.9-87.8) 77.8 (69.2-79.5) 0.58 Neutrophils, % 34.8 (14.5-58.8) 55.8 (43.0-60.8) 0.36 Neutrophils 104/mL 135.3 (49.1-179.8) 158.5 (130.6-454.2) 0.47 Eosinophils, % 0 (0-0.2) 0.2 (0-4.2) 0.32 Eosinophils 104/mL 0 (0-0.7) 1.7 (0-27.2) 0.16 Macrophages, % 58.7 (35-83.3) 36.1 (35.7-40.1) 0.36 Macrophages 104/mL 196.7 (131.7-307.4) 133.4 (132.2-230.3) 0.72 Lymphocytes, % 3 (1.3-3.7) 1 (0.7-2.1) 0.20 Lymphocytes 104/mL 8.1 (2.6-17.9) 3.2 (2.3-10.9) 0.58 Columnar epithelial cells, % 1.4 (1.2-3.3) 1.6 (1.2-3.1) 0.78 Columnar epithelial cells 104/mL 7.3 (3.9-11.4) 7.2 (6.4-9.0) 0.72 Squamous cells, % 5.1 (1.8-18.1) 8 (2.1-9.9) 1.00 Table 6.12 Induced sputum inflammatory cell counts for participants with COPD compared with healthy controls
6.3.2.3 Sputum Supernatant IL-8
There was no significant difference (p=0.08) between the levels of IL-8 measured in the sputum supernatant (Figure 6.4), however there was a trend for levels of IL-8 to be higher in subjects with COPD.
40
Healthy 30 COPD
20
IL-8 (ng/mL) IL-8 10
0 Figure 6.4 Level of IL-8 in sputum supernatant from subjects with COPD compared to healthy controls
126 6.3.2.4 Chemokine and Cytokine Production from Sputum Neutrophils
Although not reaching significance, resting sputum neutrophils from subjects with COPD released lower levels of IL-8, IL-1β and TNF-α (Table 6.13). The level of mRNA for IL-8, IL-1β and TNF-α in resting sputum neutrophils also did not reach significance but was lower in subjects with COPD compared to healthy controls (Table 6.14).
Healthy COPD p n 4 5 IL-8 3112.1 (1909.3-5313) 689.5 (554.2-1687) 0.22 IL-1β 97.2 (40.8-233) 8.5 (6.6-27.1) 0.086 TNF-α 669.9 (373.9-1377.3) 20.7 (11.3-30.4) 0.086 Table 6.13 Levels of chemokine and cytokine protein release from sputum neutrophils isolated from participants with COPD compared to healthy controls
Healthy COPD p IL-8 196.7 (186.1-347.3) 27.3 (24.8-68.1) 0.18 IL-1β 0.54 (0.5-1.88) 0.11 (0.08-0.3) 0.1 TNF-α 2.65 (1.78-10.02) 0.18 (0.08-0.22) 0.053 Table 6.14 Levels of chemokine and cytokine mRNA expression in sputum neutrophils isolated from participants with COPD compared to healthy controls
6.3.2.5 Chemokine and Cytokine Production from Peripheral Blood Neutrophils
Isolated blood neutrophils were cultured with or without LPS stimulation (100ng/ml). Resting blood neutrophils did not release detectable levels of IL-1β, TNF-α, or OSM. All resting blood neutrophils released detectable levels of IL-8, however the amount of IL-8 released was comparable between the 2 groups [Healthy: 114.7 (64.5-197.4) pg/mL, COPD: 80.5 (78.8-234.3) pg/mL, p= 0.249]. The level of mRNA for TNF-α did not differ between healthy controls and participants with COPD. However, subjects with COPD had lower levels of IL-8, IL-1β and OSM mRNA in blood neutrophils at
127 rest in comparison to healthy controls, although this only reached statistical significance for OSM (Table 6.15).
Healthy COPD p IL-8 3.2 (1.9-4.26) 1.2 (0.99-1.9) 0.06 IL-1β 0.008 (0.006-0.019) 0.001 (0.0007-0.002) 0.09 TNF-α 0.002 (0.001-0.006) 0.001 (0.0005-0.002) 0.22 OSM 0.2 (0.15-0.24) 0.1 (0.008-0.13) 0.04 Table 6.15 Levels of chemokine and cytokine mRNA in resting blood neutrophils isolated from participants with COPD and healthy controls
LPS stimulation induced cytokine and chemokine protein release in both participants with COPD and healthy controls. Isolated blood neutrophils stimulated with LPS from participants with COPD released higher levels of IL-8 and TNF-α, and lower levels IL- 1β compared to healthy controls, however this did not reach statistical significance (IL- 8: p=0.079, IL-1β: p=0.057, TNF-α: p=0.057, Figure 6.5).
128 ABHealthy 30 17500 COPD 15000 12500 20 10000 (pg/mL)
7500 β 10 IL-8 (pg/mL) IL-8
5000 IL-1 2500 0 0
CD 90 100 80 70 75 60 50
(pg/mL) 50 40 α 30 OSM (pg/mL)OSM TNF- 20 25 10 0 0
Figure 6.5 Chemokine (A: IL-8) and Cytokine (B: IL-1β, C: TNF-α D: OSM) release from LPS stimulated blood neutrophils (106 cells/mL) isolated from participants with COPD compared with healthy controls.
Levels of mRNA for IL-8, IL-1β, TNF-α and OSM were measured in LPS stimulated neutrophils. Levels of mRNA for IL-8, IL-1β and TNF-α from LPS stimulated blood neutrophils did not significantly differ between healthy controls and participants with COPD (Table 6.16).
Healthy COPD p IL-8 48.3 (12.5-79.3) 22 (19-91.8) 0.94 IL-1β 0.18 (0.06-0.25) 0.09 (0.01-0.1) 0.22 TNF-α 0.13 (0.06-0.17) 0.11 (0.07-0.31) 0.94 OSM 0.66 (0.22-0.69) 0.81 (0.44-1.28) 0.22 Table 6.16 Levels of chemokine and cytokine mRNA expression in LPS stimulated neutrophils from participants with COPD compared with healthy controls
129 6.3.2.6 Total MMP-9 Release from Peripheral Blood Neutrophils
Both resting and LPS stimulated blood neutrophils released high levels of total MMP-9 into the culture supernatant. Levels of Total MMP-9 released from resting neutrophils were lower from participants with COPD compared to healthy controls (p=0.01, Figure 6.6A). The release of total MMP-9 from LPS stimulated blood neutrophils was similar in COPD patients and healthy controls (Figure 6.6B). Healthy A B COPD 600 900 800 500 * 700 400 600 500 300 400 200 300 200 100
Total MMP-9(pg/mL) Total MMP-9(pg/mL) 100 0 0 Figure 6.6 Total MMP-9 release from isolated blood neutrophils at rest (A) and stimulated with 100ng/mL LPS (B) in COPD compared with healthy controls *p=0.01 versus healthy controls
Resting and stimulated blood neutrophils released low levels of active MMP-9 that did not differ between healthy controls and COPD patients (Table 6.17).
Healthy COPD p Neutrophils 1.9 (1.9-3.8) 3.5 (2-4.8) 0.62 Neutrophils + LPS 2.6 (0.9-3.6) 2.9 (2.6-3.6) 0.56 Table 6.17 Active MMP-9 release from blood neutrophils (106 cells/mL) isolated from participants with COPD compared with healthy controls
6.3.2.7 Neutrophil TLR Expression
Levels of both TLR4 and TLR2 mRNA expression were examined in sputum and blood neutrophils. Sputum neutrophils from participants with COPD had lower TLR4 mRNA levels compared to healthy control sputum neutrophils, although this did not reach statistical significance (p=0.053, Table 6.18). However, resting blood neutrophils from subjects with COPD had significantly lower levels of TLR4 mRNA expression
130 compared to healthy controls (p=0.023, Table 6.18). These levels did not differ between the patient groups when the cells were stimulated with LPS. Resting blood neutrophils from subjects with COPD had significantly lower TLR2 gene expression compared to healthy controls (p=0.012, Table 6.28)
Healthy COPD p Sputum Neutrophils 0.014 (0.004-0.026) 0.0023 (0.0014-0.0024) 0.053 Resting Blood 0.006 (0.005-0.008) 0.0016 (0.0013-0.0017) 0.023 Neutrophils LPS Stimulated Blood 0.01 (0.009-0.02) 0.006 (0.006-0.008) 0.12 Neutrophils Table 6.18 Relative mRNA levels of TLR4 in sputum and blood neutrophils from participants with COPD compared with healthy controls
Healthy COPD p Sputum Neutrophils 0.69 (0.44-1.21) 0.05 (0.05-0.16) 0.099 Resting Blood 0.06 (0.03-0.09) 0.02 (0.005-0.03) 0.012 Neutrophils LPS Stimulated Blood 0.43 (0.25-0.88) 0.22 (0.21-0.4) 0.18 Neutrophils Table 6.19 Relative mRNA levels of TLR2 in sputum and blood neutrophils from participants with COPD compared with healthy controls
Results of this section are summarised in Table 6.20.
131 Airway Resting Blood LPS Stimulated
Neutrophils Neutrophils Blood Neutrophils IL-8 protein Lower Unchanged Higher IL-8 mRNA Lower Lower Unchanged IL-1β protein Lower Not Detected Lower IL-1β mRNA Lower Lower Unchanged TNF-α protein Lower Not Detected Higher TNF-α mRNA Lower Unchanged Unchanged OSM protein Not Detected Not Detected Unchanged OSM mRNA N/A Lower Unchanged Total MMP-9 N/A Lower Unchanged Active MMP-9 N/A Unchanged Unchanged TLR4 mRNA Lower Lower Unchanged TLR2 mRNA Unchanged Lower Lower Table 6.20 Summary of results for the comparison between participants with COPD and age matched healthy controls Significantly lower in participants with COPD versus healthy controls. Trend for lower levels of the mediator in participants with COPD versus healthy controls. Trend for higher levels of the mediator in participants with COPD versus healthy controls N/A means not measured as very low amounts detected in a small sample group
132 6.3.3 Comparison of Asthma and COPD
6.3.3.1 Clinical Features
For this comparison, care was taken to age match the participants with asthma to those with COPD so that we could determine significant differences in neutrophil function specifically due to either disease groups. Results for this section are summarised in Table 6.29. There were no significant differences between the clinical parameters measured in either asthma or COPD (Table 6.21).
Asthma COPD p
N 12 5
Age years, mean (SD) 68 (6) 68 (7) 0.91
Sex M | F 7 | 5 3 | 2 1.0
Atopy n (%) 10 (83) 3 (60) 0.54
FEV1 % predicted 63 (16) 69 (14) 0.47
FEV1/FVC % 63 (7) 66 (6) 0.42 Table 6.21 Clinical characteristics of participants with asthma compared to those with COPD
6.3.3.2 Inflammatory Cells
There were no significant differences in the inflammatory cell counts between participants with asthma and COPD (Table 6.22).
133 Asthma COPD p n 12 5 Total cell count x 106/mL 8.6 (4.7-16) 3.7 (3.2-6.5) 0.17 Viability (%) 69.5 (56.4-86.6) 77.8 (69.2-79.5) 0.64 Neutrophils, % 46.3 (28.6-71.8) 55.8 (43.0-60.8) 0.92 Neutrophils 104/mL 496.1 (67.2-951.6) 158.5 (130.6-454.2) 0.34 Eosinophils, % 1.4 (0.4-5.1) 0.2 (0-4.2) 0.67 Eosinophils 104/mL 12.2 (2.6-26.9) 1.7 (0-27.2) 0.67 Macrophages, % 43.3 (22.8-52.4) 36.1 (35.7-40.1) 0.67 Macrophages 104/mL 288.1 (109.7-405.5) 133.4 (132.2-230.3) 0.29 Lymphocytes, % 1 (0.6-1.6) 1 (0.7-2.1) 0.43 Lymphocytes 104/mL 5.3 (2.9-11.5) 3.2 (2.3-10.9) 0.75 Columnar epithelial cells, % 3.9 (1.3-6.5) 1.6 (1.2-3.1) 0.46 Columnar epithelial cells 104/mL 35 (4.8-75.5) 7.2 (6.4-9.0) 0.25 Squamous cells, % 7 (1.5-11.2) 8(2.1-9.9)) 0.67 Table 6.22 Induced sputum inflammatory cell counts for participants with asthma compared with COPD
6.3.3.3 Sputum supernatant IL-8
There was no significant difference between the level of IL-8 detected in the sputum supernatant [Asthma: 11.7ng/mL (6.1-39), COPD: 11.9ng/mL (10.4-16.5) p=0.92].
6.3.3.4 Chemokine and Cytokine Production from Sputum Neutrophils
The level of IL-8, IL-1β and TNF-α protein and mRNA measured in resting sputum neutrophils was similar between subjects with asthma and COPD (Table 6.23).
134 Asthma COPD p IL-8 412.5 (191.4–1890.3) 689.5 (554.2-1687) 0.46 IL-8 mRNA 39.3 (29-57.3) 27.3 (24.8-68.1) 0.9 IL-1β 5.9 (3.1-9.3) 8.5 (6.6-27.1) 0.16 IL-1β mRNA 0.18 (0.082-0.28) 0.11 (0.08-0.3) 0.9 TNF-α 7.1 (0-48.2) 20.7 (11.3-30.4) 0.33 TNF-α mRNA 0.11 (0.03-0.16) 0.18 (0.08-0.22) 0.54 Table 6.23 Chemokine and cytokine production of sputum neutrophils isolated from participants with asthma compared with COPD
6.3.3.5 Chemokine and Cytokine Production from Peripheral Blood Neutrophils
Resting blood neutrophils did not release detectable IL-1β or TNF-α. The level of IL-8 released from resting blood neutrophils did not differ between participants with asthma or COPD [Asthma: 70.1ng/mL (51.3-147.8, COPD: 80.5ng/mL (78.8-234.3), p=0.75]. Levels of mRNA for IL-8, IL-1β, TNF-α or OSM did not differ between participants with asthma and COPD for resting blood neutrophils (Table 6.24).
Asthma COPD p IL-8 2.4 (0.82-3.43) 1.2 (0.99-1.9) 0.34 IL-1β 0.0013 (0.0007-0.004) 0.001 (0.0007-0.002) 0.69 TNF-α 0.004 (0.002-0.01) 0.001 (0.0005-0.002) 0.09 OSM 0.1 (0.08-0.16) 0.1 (0.008-0.13) 0.56 Table 6.24 Levels of chemokine and cytokine mRNA expression in resting blood neutrophils isolated from participants with asthma compared with COPD
LPS stimulated neutrophils isolated from subjects with COPD released a significantly higher level of IL-8 (Figure 6.7A). There was also a trend (p=0.09, Figure 6.7C) for an increased TNF-α release from COPD neutrophils with LPS stimulation. LPS stimulated blood neutrophils released similar amounts of IL-1β and OSM between the asthma and COPD groups (Figure 6.7B & D). Interestingly, the level of IL-8, IL-1β, TNF-α and
135 OSM gene expression was not different between asthma and COPD patient groups (Table 6.25).
Asthma A B * 35 COPD 17500 30 15000 12500 25 10000 20
(pg/mL) 15 7500 β 10 IL-8 (pg/mL) IL-8
5000 IL-1 2500 5 0 0
CD 90 100 80 70 75 60 50
(pg/mL) 50 40 α 30 OSM (pg/mL) OSM TNF- 20 25 10 0 0
Figure 6.7 Chemokine (A:IL-8) and Cytokine (B: IL-1β, C: TNF-α, D: OSM) release from LPS stimulated blood neutrophils (106cells/mL) isolated from participants with asthma compared with COPD *p=0.018 versus asthma
Asthma COPD p IL-8 28.9 (11.8-44.9) 22 (19-91.8) 0.53 IL-1β 0.02 (0.01-0.09) 0.09 (0.01-0.1) 0.16 TNF-α 0.12 (0.04-0.2) 0.11 (0.07-0.31) 0.6 OSM 0.56 (0.33-0.75) 0.81 (0.44-1.28) 0.21 Table 6.25 Levels of chemokine and cytokine mRNA expression in LPS stimulated neutrophils isolated from participants with asthma or COPD
6.3.3.6 Total MMP-9 Release from Peripheral Blood Neutrophils
Total MMP-9 released from both resting and LPS stimulated neutrophils did not differ between asthma and COPD groups (Table 6.26).
136
Asthma COPD p
Neutrophils 293 (215-381) 279 (256-326) 0.87
Neutrophils + LPS 609 (430-645) 606 (592-636) 0.78
Table 6.26 Total MMP-9 release from blood neutrophils (106cells/mL) isolated from participants with asthma compared with COPD
6.3.3.7 Neutrophil TLR Expression
TLR4 (Table 6.27) and TLR2 (Table 6.28) gene expression was not significantly different between participants with asthma and COPD.
Asthma COPD p Sputum Neutrophils 0.002 (0.002-0.003) 0.0023 (0.0014-0.0024) 0.95 Resting Blood 0.003 (0.002-0.004) 0.0016 (0.0013-0.0017) 0.19 Neutrophils LPS Stimulated Blood 0.006 (0.004-0.01) 0.006 (0.006-0.008) 0.67 Neutrophils Table 6.27 Relative mRNA levels of TLR4 in sputum and blood neutrophils from participants with asthma and COPD
Asthma COPD p Sputum Neutrophils 0.2 (0.08-0.46) 0.05 (0.05-0.16) 0.12 Resting Blood 0.02 (0.009-0.03) 0.02 (0.005-0.03) 0.67 Neutrophils LPS Stimulated Blood 0.18 (0.1-0.3) 0.22 (0.21-0.4) 0.15 Neutrophils Table 6.28 Relative mRNA levels of TLR2 in sputum and blood neutrophils from participants with asthma and COPD
Results of this section are summarised in Table 6.29.
137 Airway Resting Blood LPS Stimulated Neutrophils Neutrophils Blood Neutrophils IL-8 protein Unchanged Unchanged COPD Higher IL-8 mRNA Unchanged Unchanged Unchanged IL-1β protein Unchanged Not Detected Unchanged IL-1β mRNA Unchanged Unchanged Unchanged TNF-α protein Unchanged Not Detected COPD Higher TNF-α mRNA Unchanged Unchanged Unchanged OSM protein Not Detected Not Detected Unchanged OSM mRNA N/A Unchanged Unchanged Total MMP-9 N/A Unchanged Unchanged Active MMP-9 N/A Unchanged Unchanged TLR4 mRNA Unchanged Unchanged Unchanged TLR2 mRNA Unchanged Unchanged Unchanged Table 6.29 Summary of results for the comparison between participants with asthma and COPD Significantly higher in subjects with COPD versus asthma. Trend for higher levels of the mediator in subjects with COPD versus asthma. N/A means not measured as very low amounts detected in a small sample group
138 6.4 Discussion
This study has shown that there is a decreased activation of airway neutrophils from subjects with asthma compared to airway neutrophils isolated from age matched healthy controls. Airway neutrophils from subjects with asthma released significantly less IL-8, IL-1β and TNF-α and have significantly lower levels of IL-8, IL-1β, TNF-α and TLR4 gene expression. Both resting and LPS stimulated blood neutrophils have similar innate immune mediator release in asthma and healthy controls. The only difference between blood neutrophil responses in asthma compared to healthy controls is a decrease in IL- 1β and TLR2 gene expression in both resting and LPS stimulated blood neutrophils.
For subjects with COPD, there was a similar decreased activation of airway neutrophils compared to airway neutrophils isolated from age matched healthy controls. Airway neutrophils from subjects with COPD released lower levels of IL-8, IL-1β and TNF-α and had lower levels of IL-8, IL-1β, and TNF-α gene expression. There was also a decreased activation of resting blood neutrophils in COPD. Resting blood neutrophils from subjects with COPD released significantly less total MMP-9 and had significantly lower levels of OSM, TLR2 and TLR4 gene expression. However when isolated blood neutrophils were stimulated with LPS, subjects with COPD had enhanced activation in the form of increased release of IL-8 and TNF-α protein. This enhanced activation of blood neutrophils stimulated with LPS was also enhanced in comparison to participants with asthma, indicating that it is specific to COPD responses.
Interesting findings in this study relate to airway neutrophil activity in asthma and COPD, where the neutrophil is thought to play an important pathogenic role. Unexpectedly, we have shown a deficient innate immune response of neutrophils in asthma and COPD compared to healthy age matched controls. To our knowledge, the activation of airway neutrophils to release innate immune mediators has not been studied in asthma or COPD. Our data, that suggests suppression of the inflammatory potential in asthma and COPD are also supported by previous reports of decreased TNF-α release from airway cells in COPD [369], and reduced cytokine production by primary bronchial epithelial cells of COPD patients [370].
139 The innate immune response is critical in controlling infections by bacteria and viruses, a process that cytokines such as TNF-α and IL-1β and chemokines such as IL-8 play a central role by directly activating both neutrophils and macrophages. Reduced release of these mediators from airway neutrophils could contribute to impaired local defence, which could lead to an increased susceptibility of subjects with asthma and COPD to infections. On the other hand, high levels of these cytokines are known to lead to significant tissue damage in airway diseases.
Neutrophils are also responsive to the cytokine composition of the surrounding extracellular microenvironment, which is known to be different between subjects with airway diseases such as asthma and COPD compared to that of healthy people. Higher levels of cytokines and chemokines in the airways of subjects with asthma and COPD, for example the increased level of sputum supernatant IL-8 in this study, may provide feedback to downregulate innate immune responses. Further studies are needed to elucidate the underlying mechanism and clinical implication of reduced cytokine release from airway neutrophils ex vivo.
Corticosteroids are widely used for the reduction of inflammation in chronic inflammatory diseases including asthma and COPD. These drugs suppress the expression of proinflammatory genes through direct inhibitory action on the transcription factors NF-κB and AP-1 [371, 372]. Corticosteroids are also known to inhibit neutrophil apoptosis [40]. Subjects with asthma in this study were on varying doses of inhaled corticosteroids, which may have had an impact on the reduction of cultured airway neutrophil cytokine responses seen in this study. However subjects with COPD were not on any corticosteroids and still had reduced release of innate immune cytokines from cultured airway neutrophils ex vivo. Therefore, it seems unlikely that inhaled corticosteroid use is completely to blame for the deficient airway neutrophil response in asthma and COPD.
TLR4 is important in the recognition of the gram-negative bacterial component LPS. TLR4 is crucial in recognising pulmonary pathogens such as Haemophilus influenzae [373] and respiratory syncytial virus [271]. We have demonstrated decreased gene expression of TLR4 in airway neutrophils from subjects with asthma compared to healthy controls. Previous reports of decreased TLR4 mRNA expression have been
140 associated with neutrophilic inflammation and bacterial colonisation in children with chronic cough [374]. There is also a decreased expression of both TLR4 and TLR2 in the bronchial epithelium of subjects with cystic fibrosis, another airway disease characterised by airway bacterial colonisation [375]. This evidence suggests that loss of TLR4 expression may contribute to impaired defence against LPS, which may be related to reduced bacterial clearance [376].
Importantly, TLR4 mRNA levels are decreased in alveolar macrophages after inhalation of LPS by healthy human volunteers [377]. Considerable evidence suggests that endotoxin can both cause and prevent asthma [378], and this depends on the timing, dose and host genetics, but it can also exacerbate asthma if it already exists [265]. Both asthma severity and frequency of asthma symptoms have been associated with endotoxin concentrations in house dust [379, 380]. It is feasible that higher levels of endotoxin in asthmatic airways may induce a level of tolerance, and result in the downregulation of TLR4. These airway neutrophils may be dysfunctional and unable to successfully defend against invading pathogenic bacteria and viruses that signal through TLR4.
This study also reports an increase in the release of IL-8 and TNF-α from LPS stimulated blood neutrophils isolated from COPD patients compared to both age matched healthy controls and subjects with asthma. This study failed to show any difference between mRNA levels, suggesting this was due to post-translational differences. Enhanced response of COPD blood neutrophils to LPS could contribute significantly to the systemic inflammation seen in COPD patients. Other studies have shown an enhanced activation of systemic neutrophils in COPD, including increased presence of cell surface adhesion molecule expression [77-79], upregulation of genes relating to inflammation [80] and enhanced respiratory burst [78], however this is the first report of increased response to LPS in COPD to our knowledge.
This chapter has demonstrated important differences between the innate immune response in asthma and COPD in comparison to healthy controls. Cytokine production from airway neutrophils in asthma and COPD is impaired, which may result in a defective local defence to invading pathogens. Blood neutrophils from subjects with
141 COPD had an enhanced response to LPS, which may impact on the systemic inflammation that occurs in this condition.
142 Chapter 7: Innate Immune Responses of Neutrophils in Asthma Subtypes
7.1 Introduction
The current definition of asthma states that it is a chronic inflammatory disease of the airways that involves many cells and cellular elements [1]. A well-characterised pathway of inflammation implicated in asthma pathogenesis involves the inhalation of allergens, which induce activation of Th2 lymphocytes, IL-5 production and eosinophil influx. Despite effective treatments targeting this inflammatory process, asthma symptoms and heightened airway responsiveness are known to persist in the absence of eosinophilia [32-36]. This inflammatory subtype is termed non-eosinophilic asthma [37].
Non-eosinophilic asthma may occur in all grades of asthma severity, including mild asthma [34], persistent asthma [32] and severe refractory asthma [36]. Non-eosinophilic asthma is associated with corticosteroid unresponsiveness [34]. Corticosteroid treatments have little impact on non-eosinophilic inflammation, and they potentially promote neutrophilic inflammation by reducing neutrophil apoptosis [40]. Recently, the use of induced sputum inflammatory cell counts has suggested the presence of four distinct subtypes of asthma based on sputum eosinophil and neutrophil proportions. These subtypes are eosinophilic, neutrophilic, mixed granulocytic and paucigranulocytic asthma [9].
The mechanisms underlying non-eosinophilic inflammation in asthma are unclear, however studies of non-eosinophilic asthma have identified increased numbers of airway neutrophils and elevated levels of the neutrophil chemoattractant IL-8 [32, 36, 41]. Furthermore, neutrophilic asthma is associated with innate immune activation, specifically increases in the expression of the receptors TLR2, TLR4, and CD14, as well as the proinflammatory cytokines IL-8 and IL-1β [255]. The levels of these innate immune mediators measured in the sputum were correlated with the number of neutrophils in the airways, implicating a role for neutrophils in the local production of these mediators.
143 It is unclear whether the increased innate immune activation seen in neutrophilic asthma is associated with up-regulation of this pathway in isolated neutrophils on a per cell basis, or whether this is simply related to the large number of neutrophils present in the airways of these subjects. This study investigates whether the elevated innate immune mediators found in the sputum from subjects with neutrophilic asthma was linked to an enhanced activation of isolated blood and sputum neutrophils.
The hypothesis of this study is that isolated sputum and peripheral blood neutrophils from subjects with non-eosinophilic asthma release more innate immune mediators in comparison with subjects with eosinophilic asthma, and healthy controls, at rest and with LPS stimulation. Furthermore sputum and blood neutrophils from subjects with neutrophilic asthma release more innate immune mediators in comparison to eosinophilic asthma and paucigranulocytic asthma. This excessive release of inflammatory mediators would promote further neutrophilic inflammation that would contribute significantly to airway inflammation.
7.2 Methods
Induced sputum and peripheral blood samples were collected from subjects with asthma and healthy control volunteers. Subjects with asthma were classified into inflammatory subtypes based on their induced sputum cell counts (See 7.2.1). Induced sputum was dispersed and neutrophils were isolated using magnetic cell separation. Neutrophils were isolated from peripheral blood samples via percoll density gradient, erythrocyte lysis and magnetic cell separation using CD16 microbeads. Isolated neutrophils were cultured in RPMI 1640 (1% FCS) with or without LPS stimulation (E.Coli, 100ng/mL). Cells were cultured at a concentration 1 x 105 or 1 x 106 cells/mL. Cells and cell free supernatants were collected at 24 hours and stored at –80˚C until analysis. IL-8, IL-1β, TNF-α and OSM were measured using a commercial sandwich ELISA. MMP-9 was measured using a commercial fluorescent ELISA, and zymography. RNA was extracted and mRNA quantified by real-time PCR with commercial primers and probes for TLR4, TLR2, IL-8, IL-1β, OSM and TNF-α, and expressed as 2-∆∆Ct. Data was analysed by Stata 9. All data, unless otherwise stated, is non-parametric and reported as the median (Q1-Q3), or in the case of figures the median as the bar and the upper quartile as the error bars. In the case of age, FEV1% predicted and FEV1/FVC data is reported as mean
144 (SD) and significant differences were determined using Student’s t test. For categorical data (Gender and Atopy) Fischer’s exact test was applied (p<0.05 considered significant). For all other data, significant differences (p<0.05) were detected with the 2- sample Wilcoxon rank sum test. To adjust for multiple group comparisons, post-hoc analysis was conducted using kwallis2 (Stata 9). For more detailed methods please refer to Chapter 2 of this thesis.
7.2.1 Asthma Subtype Classification
Based on previous studies [9], the upper limit of normal sputum eosinophil and neutrophil counts was 1% and 61%, respectively, and was determined by the 95th percentile of a healthy control population studied. Subjects with a sputum eosinophil count of >1% in the absence of an increase in sputum neutrophils were classified as eosinophilic asthma, and the remaining subjects were classified as non-eosinophilic asthma. Non-eosinophilic asthma was further divided into 2 groups, neutrophilic asthma if subjects had >61% sputum neutrophils and paucigranulocytic asthma if subjects had normal levels of sputum neutrophils and eosinophils. Subjects with both increased neutrophils and eosinophils were classified as neutrophilic asthma, since in prior examinations the clinical and inflammatory mediator profiles of these subjects resembled neutrophilic asthma [9].
7.3 Results
7.3.1 Comparison of Eosinophilic and Non-Eosinophilic Asthma
7.3.1.1 Clinical Features
Eight (47%) of the 17 subjects had eosinophilic asthma, and the remaining 9 (53%) had non-eosinophilic asthma. Eleven age matched healthy controls were also collected as a reference group. Subjects with non-eosinophilic asthma had significantly worse lung function in comparison to healthy controls (FEV1% predicted and FEV/FVC: Table 7.1).
145
7.3.1.2 Inflammatory Cells
Total and differential cells counts obtained from induced sputum samples are shown in Table 7.2. Subjects with eosinophilic asthma had an increased proportion of eosinophils compared to non-eosinophilic asthma and healthy controls, and an increased absolute number of eosinophils compared to healthy controls. There was a trend for subjects with non-eosinophilic asthma to have an increased proportion and absolute number of neutrophils. Subjects with non-eosinophilic asthma had significantly decreased proportion of macrophages compared to healthy controls. Absolute numbers of macrophages were similar between the groups and therefore the reduced proportion of macrophages was due to an increased proportion of neutrophils and expressing the result as a percentage.
146 Healthy Controls Eosinophilic Asthma Non-Eosinophilic Asthma p n 11 8 9 Age years, mean (SD) 56 (18) 53 (20) 65 (9) 0.35 Sex M | F 5 | 6 4 | 4 4 | 5 1.0 Atopy n (%) 6 (55) 7 (88) 7 (78) 0.29 FEV1 % predicted 98 (17) 77 (19) 66 (18) ‡ 0.002 FEV/FVC % 77 (7) 70 (9) 65 (10) ‡ 0.01 ICS ‡‡ dose (μg) median (IQR) - 750 (400-1500) 1000 (500-2000) 0.24 Table 7.1 Clinical characteristics of eosinophilic asthma, non-eosinophilic asthma and healthy controls ‡p<0.008 versus healthy controls ‡‡ ICS dose is calculated 1μg of beclomethasone = 1μg of budesonide = 0.5μg of fluticasone.
147 Healthy Eosinophilic Asthma Non-Eosinophilic Asthma p n 9 8 9 Total cell count x 106/mL 3.7 (2.4-5.6) 5.3 (2.8-8.3) 10.1 (4.4-17.6) 0.13 Viability (%) 79.2 (73.9-84.2) 64.7 (49.8-71.1) 82.9 (49.7-87.5) 0.14 Neutrophils, % 30.5 (14.5-37.4) 31.2 (16-41.1) 58.5 (24.5-72) 0.08 Neutrophils 104/mL 90.9 (49.1-150.7) 104.7 (64.4-218.7) 589.7 (81.6-1043.3) 0.06 Eosinophils, % 0 (0-0.3) 5.1 (2.1-8.6) ‡║ 0.2 (0-0.8) 0.0003 Eosinophils 104/mL 0 (0-1.4) 16.1 (8.8-47.7)‡ 0.8 (0-13.2) 0.002 Macrophages, % 66.1 (58.4-82.4) 52.6 (49.8-62.8) 40.5 (22.6-48.3)‡ 0.009 Macrophages 104/mL 226 (172.8-307.4) 241.5 (189.1-437.3) 238.1 (120.6-402.7) 0.82 Lymphocytes, % 2.0 (1.0-3.2) 1.4 (0.8-2.5) 1 (0.3-1.7) 0.37 Lymphocytes 104/mL 4.2 (2.6-9.4) 6.4 (3.7-7.6) 9.9 (2.4-12.8) 0.92 Columnar epithelial cells, % 1.3 (1.2-3.3) 6.5 (3.1-13.2) 3.5 (0.8-5.2) 0.18 Columnar epithelial cells 104/mL 6.7 (4.3-11.4) 48.1 (10.8-101.9) 50.7 (7.4-74.6) 0.19 Squamous cells, % 5.1 (1.8-16.5) 7.3 (4.3-12) 2.9 (0-11.1) 0.54 Table 7.2 Inflammatory cell counts for subjects with eosinophilic asthma, non-eosinophilic asthma and healthy controls p<0.008 ‡ versus healthy controls, ║ versus non-eosinophilic asthma
148 7.3.1.3 Sputum Supernatant IL-8
There was a non-significant trend (p=0.097) for subjects with non-eosinophilic asthma to have an increased level of IL-8 in their sputum supernatant in comparison to subjects with eosinophilic asthma and healthy controls (Figure 7.1).
Healthy
60 Eosinophilic Non Eosinophilic 50
40
30
IL-8 (ng/mL) IL-8 20
10
0 Figure 7.1 Level of IL-8 detected in sputum supernatant
7.3.1.4 Chemokine and Cytokine Production from Sputum Neutrophils
Local chemokine and cytokine production was examined in isolated sputum neutrophils. IL-8 and IL-1β were released from sputum neutrophils however levels from subjects with non-eosinophilic asthma and eosinophilic asthma tended to be lower compared to that released from healthy controls [IL-8: p=0.13; Figure 7.2A and IL-1β: p=0.07 Figure 7.2B]. Sputum neutrophils from subjects with non-eosinophilic asthma released significantly less TNF-α compared with healthy controls [p<0.008, Figure 7.2C].
149 Healthy Eosinophilic A B 7000 100 Non Eosinophilic 6000 75 5000 4000 50 (pg/mL)
3000 β
IL-8 (pg/mL) IL-8 2000 IL-1 25 1000 0 0 C 600
500
400
(pg/mL) 300 α 200 * TNF- 100
0
Figure 7.2 Chemokine (A: IL-8) and Cytokine (B: IL-1β, C: TNF-α) release from resting sputum neutrophils isolated from subjects with non-eosinophilic asthma, eosinophilic asthma and healthy controls *p<0.008 versus healthy controls
The levels of mRNA for IL-8, IL-1β and TNF-α were also examined in sputum neutrophils. IL-8, IL-1β and TNF-α mRNA expression were lower in both eosinophilic and non-eosinophilic asthma, however this did not reach significance (Table 7.3).
Eosinophilic Non-Eosinophilic Healthy p Asthma Asthma IL-8 134.4 (77.2-196.7) 40.5 (38.1-43.7) 36.9 (21.8-59.1) 0.06 IL-1β 1.34 (0.68-2.65) 0.17 (0.08-0.19) 0.28 (0.1-0.39) 0.067 TNF-α 0.47 (0.16-0.54) 0.098 (0.08-0.15) 0.16 (0.03-0.36) 0.086 Table 7.3 Levels of chemokine and cytokine mRNA expression in resting sputum neutrophils from participants with eosinophilic asthma, and non-eosinophilic asthma compared to healthy controls
150
7.3.1.5 Chemokine and Cytokine Production from Blood Neutrophils
Resting peripheral blood neutrophils did not release detectable levels of TNF-α in any of the samples. 91% (n=20) of resting neutrophil samples had undetectable levels of IL- 1β and OSM. All resting neutrophil samples released IL-8 at detectable levels. Resting neutrophils from subjects with non-eosinophilic asthma released significantly more IL-8 compared to subjects with eosinophilic asthma (Figure 7.3).
Healthy Eosinophilic 450 * Non Eosinophilic 400 350 300 250 200 150 IL-8 (pg/mL) IL-8 100 50 0 Figure 7.3 Enhanced IL-8 release from neutrophils in non-eosinophilic asthma *p<0.008 versus eosinophilic asthma
IL-8, IL-1β, TNF-α and OSM mRNA expression was measured using real-time PCR and results are shown in Table 4. Levels of IL-8 mRNA expression was highest in participants with non-eosinophilic asthma, however this was not significantly different between the groups. IL-1β mRNA expression was lower in resting neutrophils from participants with eosinophilic asthma, however this did not reach significance (p=0.067). TNF-α mRNA expression in resting neutrophils was not different between healthy controls and participants with eosinophilic or non-eosinophilic asthma.
151 Eosinophilic Non-Eosinophilic Healthy p Asthma Asthma IL-8 2.27 (1.59-4.26) 1.07 (0.71-3.63) 3.07 (2.4-3.66) 0.17 IL-1β 0.007 (0.003-0.019) 0.001 (0.0007-0.004) 0.003 (0.002-0.004) 0.068 TNF-α 0.002 (0.001-0.006) 0.002 (0.001-0.003) 0.008 (0.002-0.01) 0.23 OSM 0.18 (0.07-0.24) 0.1 (0.07-0.15) 0.15 (0.13-0.18) 0.29 Table 7.4 Levels of chemokine and cytokine mRNA expression in resting blood neutrophils from participants with eosinophilic asthma and non-eosinophilic asthma compared to healthy controls
LPS stimulation induced blood neutrophil to release IL-8, IL-1β, TNF-α and OSM to detectable levels. There was no difference in the neutrophils response to LPS when comparing subjects with non-eosinophilic or eosinophilic asthma to healthy controls (Figure 7.4). LPS stimulated neutrophils from subjects with eosinophilic asthma released significantly less OSM compared healthy controls (Figure 7.4D). Healthy
Eosinophilic Non Eosinophilic AB50 9000 8000 40 7000 6000 30 5000 (pg/mL)
4000 β 20 3000 IL-8 (pg/mL) IL-8 IL-1 2000 10 1000 0 0
CD 50 90 80 40 70 60 30 50
(pg/mL) 40 α 20 * 30 OSM (pg/mL) TNF- 10 20 10 0 0
Figure 7.4 Chemokine (A: IL-8) and Cytokine (B: IL-1β, C: TNF-α) release from LPS stimulated blood neutrophils (106 cells/mL) from subjects with non- eosinophilic asthma, eosinophilic asthma and healthy controls *p<0.008 versus healthy controls
152
IL-8, IL-1β, TNF-α and OSM mRNA expression is shown in Table 7.5. IL-8 and TNF-α mRNA expression was not different between healthy controls and participants with eosinophilic and non-eosinophilic asthma. However, IL-1β mRNA expression was significantly lower in LPS stimulated neutrophils from participants with eosinophilic asthma compared to healthy controls.
Eosinophilic Non-Eosinophilic Healthy p Asthma Asthma IL-8 54.2 (40.6-95.3) 26.4 (10.9-56.4) 44.6 (29.7-84.4) 0.23 IL-1β 0.14 (0.06-0.2) 0.04 (0.01-0.075) * 0.07 (0.02-0.13) 0.03 TNF-α 0.12 (0.06-0.17) 0.15 (0.04-0.19) 0.1 (0.06-0.15) 0.83 OSM 0.66 (0.35-0.84) 0.64 (0.26-1.09) 0.56 (0.46-0.72) 0.87 Table 7.5 Relative messenger RNA levels of IL-8, IL-1β and TNF-a of LPS stimulated blood neutrophils from participants with asthma compared to healthy controls *p<0.008 versus healthy controls
7.3.1.6 MMP-9 Release from Peripheral Blood Neutrophils
Both resting and stimulated blood neutrophils release high levels of total MMP-9, but low levels of active MMP-9 into the cell culture supernatant. Similar amounts of both total and active MMP-9 were released from resting and LPS stimulated blood neutrophils between healthy controls and participants with eosinophilic and non- eosinophilic asthma (Table 7.6, Table 7.7).
Eosinophilic Non Eosinophilic Sample Healthy p Asthma Asthma Neutrophils 369 (176-554) 242 (213-283) 299 (212-408) 0.53 Neutrophils +LPS 577 (452-857) 504 (433-629) 613 (389-668) 0.55 Table 7.6 Total MMP-9 release from blood neutrophils (106 cells/mL) isolated from participants with non-eosinophilic asthma, eosinophilic asthma and healthy controls
153
Eosinophilic Non-Eosinophilic Sample Healthy p Asthma Asthma Neutrophils 3.3 (1.9-4.1) 2.8 (2.6-6.1) 3.8 (3.2-9) 0.51 Neutrophils +LPS 4 (2.4-7.2) 4.7 (2.5-8) 4.7 (3-6.6) 0.88
Table 7.7 Active MMP-9 release from blood neutrophils (106 cells/mL) isolated from participants with non-eosinophilic asthma, eosinophilic asthma and healthy controls
7.3.1.7 Neutrophil TLR Expression
TLR4 and TLR2 mRNA expression were measured in both resting sputum neutrophils and resting and LPS stimulated blood neutrophils. There was no significant difference between healthy controls and participants with eosinophilic or non-eosinophilic asthma for the levels of TLR4 mRNA (Table 7.8). Resting neutrophils had significantly lower levels of TLR2 mRNA in eosinophilic asthma compared to healthy controls (Table 7.9). LPS stimulated neutrophils had significantly lower levels of TLR2 mRNA in non- eosinophilic asthma compared to healthy controls (Table 7.9).
154 Healthy Controls Eosinophilic Asthma Non-Eosinophilic Asthma p Sputum Neutrophils 0.004 (0.004-0.014) 0.0022 (0.0021-0.0024) 0.002 (0.001-0.004) 0.11 Resting Blood 0.005 (0.002-0.008) 0.002 (0.0016-0.0054) 0.004 (0.003-0.004) 0.48 Neutrophils LPS Stimulated 0.009 (0.005-0.01) 0.01 (0.005-0.013) 0.006 (0.005-0.01) 0.66 Blood Neutrophils Table 7.8 Relative mRNA levels of TLR4 in sputum and blood neutrophils from subjects with non-eosinophilic asthma, eosinophilic asthma and healthy controls
Healthy Controls Eosinophilic Asthma Non-Eosinophilic Asthma p Sputum Neutrophils 0.44 (0.13-0.69) 0.2 (0.07-0.29) 0.29 (0.08-0.46) 0.79 Resting Blood 0.057 (0.03-0.09) 0.014 (0.006-0.04) * 0.02 (0.02-0.03) 0.02 Neutrophils LPS Stimulated 0.39 (0.25-0.84) 0.21 (0.16-0.34) 0.23 (0.11-0.29) * 0.02 Blood Neutrophils Table 7.9 Relative mRNA levels of TLR2 in sputum and blood neutrophils from subjects with non-eosinophilic asthma, eosinophilic asthma and healthy controls *p<0.008 versus healthy controls
155
LPS Stimulated Airway Resting Blood Blood Neutrophils Neutrophils Neutrophils Lower in EA & IL-8 protein Higher in NEA Unchanged NEA Lower in EA & IL-8 mRNA Higher in NEA Unchanged NEA Lower in EA & IL-1β protein Not Detected Lower in EA NEA Lower in EA & IL-1β mRNA Lower in EA Lower in EA NEA TNF-α protein Lower in NEA Not Detected Unchanged Lower in EA & TNF-α mRNA Unchanged Unchanged NEA OSM protein Not Measured Not Detected Lower in EA OSM mRNA Not Measured Unchanged Unchanged Total MMP-9 Not Measured Unchanged Unchanged Active MMP-9 Not Measured Unchanged Unchanged TLR4 mRNA Unchanged Unchanged Unchanged Lower in EA & Lower in EA & TLR2 mRNA Unchanged NEA NEA Table 7.10 Summary of results for the comparison between participants with non- eosinophilic asthma, eosinophilic asthma and healthy controls Significantly lower in subjects with EA or NEA versus healthy controls Trend for lower levels in subjects with EA or NEA versus healthy controls Significantly higher in subjects with EA or NEA versus healthy controls Trend for higher levels in subjects with EA or NEA versus healthy controls
156 7.3.2 Comparison of Eosinophilic, Neutrophilic and Paucigranulocytic Asthma
7.3.2.1 Clinical Features
Four (24%) of the 17 subjects with asthma had neutrophilic asthma, 8 (47%) had eosinophilic asthma and 5 (29%) had paucigranulocytic asthma. Two subjects with neutrophilic asthma had a mixed granulocytic pattern of inflammation with increased proportions of neutrophils and eosinophils. Subjects with neutrophilic asthma had significantly worse lung function (Table 7.11).
7.3.2.2 Inflammatory Cells
Subjects with eosinophilic asthma had an increased proportion of eosinophils compared to healthy controls and paucigranulocytic asthma, and an increased absolute number of eosinophils compared to healthy controls (Table 7.12). There was a non-significant trend (p=0.06) for subjects with neutrophilic asthma to have an increased total cell count. Subjects with neutrophilic asthma had an increased proportion and absolute number of neutrophils, and a decreased proportion of macrophages compared to eosinophilic asthma and healthy controls. Absolute numbers of macrophages were similar between the groups and therefore the reduced proportion of macrophages was due to an increased proportion of neutrophils and expressing the result as a percentage.
157 Healthy Eosinophilic Neutrophilic Paucigranulocytic p Controls Asthma Asthma Asthma n 11 8 4 5
Age years, mean (SD) 56 (18) 53 (20) 69 (7) 61 (10) 0.45
Sex M | F 5 | 6 4 | 4 2 | 2 2 | 3 1.0
Atopy n (%) 6 (55) 7 (88) 3 (75) 4 (80) 0.48
‡ FEV1 % predicted mean (SD) 98 (17) 77 (19) 53 (15) 76 (13) 0.0008
‡ FEV1/FVC % mean (SD) 77 (7) 70 (9) 58 (9) 70 (9) 0.005
ICS dose ‡‡ (μg) median (IQR) - 750 (400-1500) 2000 (1200-2500) 800 (500-1000) 0.4 Table 7.11 Clinical characteristics of neutrophilic asthma, eosinophilic asthma, paucigranulocytic asthma and healthy controls ‡p<0.004 versus healthy controls ‡‡ ICS dose is calculated 1μg of beclomethasone = 1μg of budesonide = 0.5μg of fluticasone.
158 Healthy Eosinophilic Asthma Neutrophilic Asthma Paucigranulocytic Asthma p n 9 8 4 5 Total cell count x 106/mL 3.7 (2.4-5.6) 5.3 (2.8-8.3) 16.2 (10.7-18.3) 4.4 (3.2-10.1) 0.055 Viability (%) 79.2 (73.9-84.2) 64.7 (49.8-71.1) 77.7 (59.7-91.2) 82.9 (43.8-87.5) 0.25 Neutrophils, % 30.5 (14.5-37.4) 31.2 (16-41.1) 74.3 (71.8-84.3)‡║ 24.5 (23.8-49) 0.018 Neutrophils 104/mL 90.9 (49.1-150.7) 104.7 (64.4-218.7) 1159.6 (786.7-1503.2) ‡ 81.6 (75-589.7) 0.021 Eosinophils, % 0 (0-0.3) 5.1 (2.1-8.6) ‡ * 1.2 (0.4-1.7) 0 (0-0.3) 0.0005 Eosinophils 104/mL 0 (0-1.4) 16.1 (8.8-47.7)‡ 17.7 (2.6-31.6) 0 (0-0.8) 0.003 Macrophages, % 66.1 (58.4-82.4) 52.6 (49.8-62.8) 20 (11.3-22.8) ‡ 48.3 (46-54) 0.005 Macrophages 104/mL 226 (172.8-307.4) 241.5 (189.1-437.3) 226.9 (109.7-368) 238.1 (152.1-408.2) 0.85 Lymphocytes, % 2.0 (1.0-3.2) 1.4 (0.8-2.5) 1.1 (0.5-1.7) 1 (0-1.7) 0.57 Lymphocytes 104/mL 4.2 (2.6-9.4) 6.4 (3.7-7.6) 13 (8.8-17.5) 2.4 (0-9.9) 0.28 Columnar epithelial cells, % 1.3 (1.2-3.3) 6.5 (3.1-13.2) 1.9 (0.5-3.3) 5.2 (4.3-25.3) 0.09 Columnar epithelial cells 104/mL 6.7 (4.3-11.4) 48.1 (10.8-101.9) 32.4 (8-53.2) 74.6 (7.4-87.6) 0.33 Squamous cells, % 5.1 (1.8-16.5) 7.3 (4.3-12) 1.5 (0-7.2) 6.1 (2.9-11.1) 0.53 Table 7.12 Table 12: Inflammatory cell counts for subjects with eosinophilic asthma, non-eosinophilic asthma and healthy controls. ‡ p<0.004 versus healthy controls ║ p<0.004 versus eosinophilic asthma * p<0.004 versus paucigranulocytic asthma
159
7.3.2.3 Sputum Supernatant IL-8
Subjects with neutrophilic asthma had significantly elevated levels of sputum supernatant IL-8 in comparison to paucigranulocytic asthma and healthy controls (Figure 7.5). Healthy Eosinophilic * # 80 Neutrophilic Paucigranulocytic 70
60
50
40
30 IL-8 (pg/mL) IL-8 20
10
0 Figure 7.5 Subjects with neutrophilic asthma have significantly higher levels of sputum supernatant IL-8 p<0.004 *versus healthy controls, # versus paucigranulocytic asthma
7.3.2.4 Chemokine and Cytokine Production from Sputum Neutrophils
Levels of IL-1β release from sputum neutrophils were not significantly different between asthma subtypes and healthy controls (Figure 7.6B). Although not reaching significance sputum neutrophils from participants with neutrophilic asthma subjects release less IL-8 (p=0.055, Figure 7.6A). Sputum neutrophils from participants with neutrophilic asthma did not release detectable amounts of TNF-α, which was significantly less than healthy controls (Figure 7.6C).
160 Healthy Neutrophilic Eosinophilic Paucigranulocytic
AB 7000 100 6000 75 5000 4000 50 (pg/mL)
3000 β
IL-8 (pg/mL) IL-8 2000 IL-1 25 1000 0 0
C 600
500
400
(pg/mL) 300 α 200 TNF- 100 * 0
Figure 7.6 Chemokine (A: IL-8) and cytokine (B: IL-1β, C: TNF-α) release from resting sputum neutrophils (105 cells/mL) asthma subtypes and healthy controls *p<0.004 versus healthy controls
Levels of IL-8, IL-1β and TNF-α mRNA were measured in sputum neutrophils and are shown in Table 7.13. IL-8 mRNA levels were significantly less in subjects with neutrophilic asthma compared to healthy controls. Although not quite reaching significance, IL-1β and TNF-α mRNA expression was lower in sputum neutrophils isolated from participants with neutrophilic asthma (Table 7.13).
161
Eosinophilic Neutrophilic Paucigranulocytic Healthy p Asthma Asthma Asthma IL-8 134.4 (77.2-196.7) 40.5 (38.1-43.7) 21.7 (12.6-29) * 58.2 (47.1-61.7) 0.03 IL-1β 1.34 (0.68-2.65) 0.17 (0.08-0.19) 0.11 (0.05-0.28) 0.33 (0.27-0.51) 0.08 TNF-α 0.47 (0.16-0.54) 0.098 (0.08-0.15) 0.03 (0.02-0.16) 0.34 (0.22-0.38) 0.049 Table 7.13 Relative messenger RNA levels of IL-8, IL-1β and TNF-α of resting sputum neutrophils in asthma subtypes and healthy controls *p<0.004 versus healthy controls
162
7.3.2.5 MMP-9 Release from Sputum Neutrophils
MMP-9 levels were measured in a selection of sputum neutrophil samples, however the levels were at undetectable levels. Zymography (Figure 7.7) showed minor bands for total MMP-9 and MMP-2, however there was little MMP-9 present and no obvious differences in the amounts released from sputum neutrophils across the groups.
Lane 1 2 3 4 5 6 7
MMP-9 Latent Form MMP-2 Latent Form
Figure 7.7 Zymography to assess levels of MMP-9 in culture supernatants of sputum neutrophils (105 cells/mL) at 24 hours (1: PGA, 2: COPD, 3: EA, 4: NA, 5: HC, 6: Ladder, 7: Zymography Control)
7.3.2.6 Chemokine and Cytokine Production from Blood Neutrophils
The enhanced IL-8 release from resting blood neutrophils in non-eosinophilic asthma (Figure 7.3) was not significant when the results were divided into neutrophilic and paucigranulocytic asthma subtypes, however there was a trend (p=0.06) for neutrophils from subjects with paucigranulocytic asthma to release more IL-8 (Figure 7.8).
163
Healthy 500 Eosinophilic Neutrophilic 400 Paucigranulocytic 300
200 IL-8 (pg/mL) IL-8 100
0 Figure 7.8 IL-8 release from resting blood neutrophils in asthma subtypes
Expression of IL-8, IL-1β and TNF-α mRNA did not differ between asthma subtypes and healthy controls in resting neutrophils (Table 7.14).
164
Eosinophilic Neutrophilic Paucigranulocytic Healthy p Asthma Asthma Asthma IL-8 2.27 (1.59-4.26) 1.07 (0.71-3.63) 2.7 (1.9-3.4) 3.2 (2.5-5.4) 0.25 IL-1β 0.007 (0.003-0.019) 0.001 (0.0007-0.004) 0.002 (0.002-0.003) 0.004 (0.002-0.006) 0.13 TNF-α 0.002 (0.001-0.006) 0.002 (0.001-0.003) 0.007 (0.003-0.009) 0.009 (0.002-0.01) 0.4 OSM 0.18 (0.07-0.24) 0.1 (0.07-0.15) 0.18 (0.14-0.49) 0.15 (0.13-0.15) 0.38 Table 7.14 Levels of chemokine and cytokine mRNA expression in resting blood neutrophils in asthma subtypes and healthy controls
165
Release of IL-8, IL-1β and TNF-α from LPS stimulated neutrophils was not different between asthma subtypes and healthy controls (Figure 7.9). LPS stimulated neutrophils from subjects with eosinophilic asthma released significantly less OSM compared healthy controls (Figure 7.9).
Healthy Neutrophilic Eosinophilic Paucigranulocytic
AB 70 12500 60 10000 50
7500 40
(pg/mL) 30 5000 β
IL-8 (pg/mL) 20 IL-1 2500 10 0 0
CD 75 200
150 50
(pg/mL) 100 α 25 OSM (pg/mL) OSM TNF- 50 *
0 0
Figure 7.9 Chemokine (A: IL-8) and Cytokine (B: IL-1β, C: TNF-α) release from LPS stimulated blood neutrophils (106 cells/mL) in asthma subtypes and healthy controls. *p<0.004 versus healthy controls
IL-8, IL-1β, TNF-α and OSM mRNA expression was measured in LPS stimulated blood neutrophils and the results are reported in Table 7.15.
166
Healthy Eosinophilic Neutrophilic Paucigranulocytic p Controls Asthma Asthma Asthma IL-8 54.2 (40.6-95.3) 26.4 (10.9-56.4) 28.4 (11.8-66.3) 45.3 (36.8-84.4) 0.28 IL-1β 0.14 (0.06-0.2) 0.04 (0.01-0.075) 0.05 (0.02-0.1) 0.1 (0.04-0.2) 0.051 TNF-α 0.12 (0.06-0.17) 0.15 (0.04-0.19) 0.06 (0.04-0.13) 0.1 (0.1-0.15) 0.66 OSM 0.66 (0.35-0.84) 0.64 (0.26-1.09) 0.64 (0.43-0.75) 0.56 (0.46-0.61) 0.96 Table 7.15 Levels of chemokine and cytokine mRNA expression in LPS stimulated blood neutrophils in asthma subtypes and healthy controls.
167 7.3.2.7 MMP-9 release from blood neutrophils
Levels of both total and active MMP-9 released from blood neutrophils did not differ between the asthma subtypes (Figure 7.10, Table 7.16, Table 7.17).
Lane 1 2 3 4 5 6 7
MMP-9 Latent Form
MMP-2 Latent Form
Figure 7.10 Zymography to assess levels of MMP-9 in culture supernatants of isolated blood neutrophils (106 cells/mL) at 24 hours (1: Zymography Control, 2: HC, 3: COPD, 4: EA, 5: NA, 6: PGA, 7: Ladder).
Sample Healthy EA NA PGA p Neutrophils 369 (176-554) 242 (213-283) 293 (138-381) 304 (240-435) 0.63 Neutrophils +LPS 577 (452-857) 498 (460-578) 609 (394-690) 617 (384-645) 0.75 Table 7.16 Total MMP-9 released from blood neutrophils (106 cells/mL) in asthma subtypes and healthy controls
Sample Healthy EA NA PGA p Neutrophils 3.3 (1.9-4.1) 2.8 (2.6-6.1) 4.3 (3.2-9) 3.3 (2.2-9) 0.64 Neutrophils +LPS 4 (2.4-7.2) 4.7 (2.5-8) 6.6 (4.7-8.5) 3.4 (2.9-5.1) 0.52 Table 7.17 Active MMP-9 released from blood neutrophils (106 cells/mL) in asthma subtypes and healthy controls
168 7.3.2.8 Neutrophil TLR Expression
Expression of TLR4 and TLR2 was examined in resting and stimulated sputum and blood neutrophils. Levels of sputum neutrophil TLR4 mRNA was lowest in subjects with neutrophilic asthma; however this did not reach statistical significance. TLR4 mRNA expression in resting and LPS stimulated blood neutrophils was not different in asthma subtypes compared to healthy controls (Table 7.18). The expression of sputum neutrophil TLR2 mRNA was the lowest in the subjects with neutrophilic asthma, however this did not reach significance. The expression of TLR2 did not differ in resting blood neutrophils; however subjects with neutrophilic asthma had significantly lower TLR2 mRNA in LPS stimulated neutrophils (Table 7.19).
169 Healthy Controls Eosinophilic Asthma Neutrophilic Asthma Paucigranulocytic Asthma p Sputum 0.004 (0.004-0.014) 0.0022 (0.0021-0.0024) 0.001 (0.0008-0.002) 0.003 (0.002-0.01) 0.096 Neutrophils Resting Blood 0.005 (0.002-0.008) 0.002 (0.0016-0.0054) 0.004 (0.003-0.005) 0.004 (0.002-0.004) 0.59 Neutrophils LPS Stimulated 0.009 (0.005-0.01) 0.01 (0.005-0.013) 0.005 (0.004-0.009) 0.007 (0.005-0.01) 0.69 Blood Neutrophils Table 7.18 Relative mRNA levels of TLR4 in sputum and blood neutrophils in asthma subtypes and healthy controls
Healthy Controls Eosinophilic Asthma Neutrophilic Asthma Paucigranulocytic Asthma p Sputum 0.44 (0.13-0.69) 0.2 (0.07-0.29) 0.15 (0.08-0.21) 0.41 (0.19-0.83) 0.78 Neutrophils Resting Blood 0.057 (0.03-0.09) 0.014 (0.006-0.04) 0.024 (0.017-0.035) 0.02 (0.02-0.029) 0.055 Neutrophils LPS Stimulated 0.39 (0.25-0.84) 0.21 (0.16-0.34) 0.13 (0.1-0.19) # 0.29 (0.23-0.32) 0.025 Blood Neutrophils Table 7.19 Relative mRNA levels of TLR2 in sputum and blood neutrophils in asthma subtypes and healthy controls
170
LPS Stimulated Airway Resting Blood Blood Neutrophils Neutrophils Neutrophils IL-8 protein Lower in NA Higher in PGA Unchanged IL-8 mRNA Lower in NA Unchanged Unchanged Lower in NA & IL-1β protein Not Detected Unchanged EA Lower in NA & IL-1β mRNA Unchanged Lower in EA EA TNF-α protein Lower in NA Not Detected Unchanged TNF-α mRNA Lower in NA Unchanged Unchanged OSM protein N/A Not Detected Lower in EA OSM mRNA N/A Unchanged Unchanged Total MMP-9 N/A Unchanged Unchanged Active MMP-9 N/A Unchanged Unchanged TLR4 mRNA Lower in NA Unchanged Unchanged TLR2 mRNA Lower in NA Lower in EA Lower in NA Table 7.20 Summary of results for the comparison between EA, NA, PGA versus healthy controls Significantly lower in subjects with EA, NA, or PGA versus healthy controls Trend for lower levels in subjects with EA, NA, or PGA versus healthy controls Significantly higher in subjects with EA, NA, or PGA versus healthy controls Trend for higher levels in subjects with EA, NA, or PGA versus healthy controls
7.3.3 Associations
FEV1 % predicted and FEV1/FVC were positively associated with sputum neutrophil cytokine and chemokine production but not with the expression of TLR2 or 4. However ICS dose was positively associated with sputum neutrophil expression of both TLR2 and TLR4, but not with cytokine production (Table 7.21).
171 FEV1 % Pred FEV1/FVC ICS Dose (μg) IL-8 protein 0.539* 0.531* -0.309 IL-8 mRNA 0.556* 0.434 0.025 IL-1β protein 0.422 0.521* -0.45 IL-1β mRNA 0.491* 0.402 -0.178 TNF-α protein 0.656* 0.658* -0.183 TNF-α mRNA 0.397 0.389 -0.477 TLR4 mRNA 0.011 -0.176 0.758* TLR2 mRNA 0.131 0.1 0.689* Table 7.21 Spearman correlations of innate immune mediators measured in resting sputum neutrophils and clinical parameters (*p<0.05)
7.4 Discussion
This study investigated activation of the innate immune pathway in isolated airway and circulating neutrophils relating this to previously defined inflammatory subtypes of asthma. We have shown that there is a decreased pro-inflammatory cytokine response of airway neutrophils in non-eosinophilic asthma. Airway neutrophil cytokine production was the lowest in subjects with neutrophilic asthma, including lower levels of IL-8 and IL-1β and an absence of detectable TNF-α protein, which was highly significant. Gene expression for IL-8 (-6 fold), IL-1β (-12 fold) and TNF-α (-16 fold) was also much lower in airway neutrophils in neutrophilic asthma compared to healthy controls, however only the decrease in IL-8 mRNA reached statistical significance. Consistent with the lower airway responses were the trends for decreased TLR gene expression; with TLR4 mRNA decreased by 4 fold and TLR2 mRNA decreased 2.9 fold in neutrophilic asthma compared to healthy controls. These data demonstrate that innate immune responses in airway neutrophils are relatively suppressed in their inflammatory cytokine response and innate immune signalling receptor expression. The reduced TLR- induced cytokine secretion may also be attributable to direct cytokine downregulation resulting from chronic stimulation with exogenous or endogenous ligands.
Resting blood neutrophils isolated from subjects with non-eosinophilic asthma had an enhanced release of IL-8 compared to subjects with eosinophilic asthma, suggesting that
172 the cells are partially activated or ‘primed’ for an enhanced response. LPS stimulated blood neutrophils had significantly lower TLR2 expression in subjects with non- eosinophilic asthma and particularly neutrophilic asthma. Neutrophil TLR2 expression has been linked to neutrophil apoptosis, and these findings in neutrophilic asthma are consistent with circulating neutrophils in NEA being in a partially activated state, and ready to actively participate in tissue inflammation responses if recruited to the tissue. Peripheral blood neutrophils isolated from subjects with eosinophilic asthma had decreased responses to LPS, including decreased OSM and IL-1β production.
Although there is growing evidence implicating the involvement of neutrophils and innate immune activation in the airways of subjects with neutrophilic asthma, very little is known regarding the state of activation of airway and circulating neutrophils in asthma subtypes. In this study, subjects with neutrophilic asthma had significantly increased neutrophil proportion but also approximately 13 times the absolute number of neutrophils, and 6 times the level of sputum supernatant IL-8 compared to healthy controls. This dramatic accumulation of neutrophils strongly suggests a role for these cells in the perpetuation of abnormal airway inflammation. The high levels of innate immune mediators generated in the airways in neutrophilic asthma could be due to an enhanced activation of the neutrophils, or accummulation of a large number of cells that exhibit a lesser degree of activation. It would be logical to assume enhanced activation of the cells, however evidence from this study suggests the opposite. I have found a relative reduction in the release of TNF-α, and level of TNF-α mRNA in airway neutrophils in neutrophilic asthma. The level of TNF-α released from isolated sputum neutrophils was positively associated with FEV1% predicted (Spearman r=0.61, p=0.0006). These observations suggest that it is the increased influx and accumulation of partially activated cells that is responsible for the high cytokine levels seen in vivo in neutrophilic asthma.
There is little data in the literature regarding sputum cell cytokine production in asthma and asthma subtypes. One recent report by Quaedvlieg and colleagues [338] showed that cytokine production from whole sputum cells in eosinophilic asthma differed in comparison to non-eosinophilic asthma and healthy controls, however non-eosinophilic asthma was similar to healthy controls in the cytokines measured. Sputum cells from subjects with eosinophilic asthma released significantly more IL-4 and less TNF-α
173 compared to healthy controls. The study by Quaedvlieg et al is limited compared to data collected for this thesis as the investigators did not divide the subjects based on abnormal sputum neutrophil counts, and therefore subjects with high neutrophils were evenly spread in both the eosinophilic and non-eosinophilic asthma groups depending on the presence of abnormal levels of sputum eosinophils. Results from this chapter provide additional knowledge on the activation status of sputum neutrophils in asthma subtypes that has not been previously investigated.
Cytokine production by airway neutrophils has been examined in vitro in both cystic fibrosis [341] and chronic bronchial sepsis [340]. Airway neutrophils release significantly more IL-8 in subjects with cystic fibrosis compared to subjects with dyskinetic cilia syndrome [341]. Airway neutrophils from subjects with chronic bronchial sepsis release significantly more IL-8, IL-1 and TNF-α compared to blood neutrophils [340]. Only sputum samples from subjects that had a high number of neutrophils were collected in these studies, therefore no samples from healthy subjects were obtained. In these studies, sputum samples were dispersed using Trypsin and neutrophils were isolated using Percoll density gradient [340] or immunomagnetic depletion with the HLA class II coated magnetic beads [341], methods that differed from those of this thesis. From these studies it is impossible to ascertain what a ‘normal’ amount of cytokine released from airway neutrophils is, and how this would compare to airway neutrophils from patients with neutrophilic airway diseases. One would expect that neutrophils migrated to the airways in response to airway triggers such as infection would be actively producing cytokine to regulate the inflammatory response to pathogens, through the activation of the innate immune response. This chapter adds to current knowledge on cytokine production and TLR expression of airway neutrophils in healthy controls, and how this may be impaired in neutrophilic airway diseases.
A study of airway cells cultured from both healthy subjects and subjects with COPD reported a decrease in the production of TNF-α [369]. This observation is consistent with suppression of the inflammatory response in this airway disease, and shows similarities to our observation of decreased TNF-α release from airway neutrophils ex vivo in neutrophilic asthma. In this sense, neutrophilic asthma is similar to COPD. Future studies into the mechanisms of neutrophilic airway disease should examine the mechanisms of neutrophil accumulation in the airways. Relevent processes in airway
174 neutrophil accumulation involve dysregulation of neutrophil recruitment or clearance from the airways.
Dysregulation of apoptosis could in fact lead to the persistence of neutrophils at the inflammatory site and the development of chronic inflammation. TNF-α is mainly proinflammatory in its functions; however TNF-α can induce neutrophil apoptosis [132]. Therefore lower levels of TNF-α production by sputum neutrophils in neutrophilic asthma may encourage the survival of the cells. Inflammatory stimuli such as growth factors (e.g. GM-CSF and G-CSF) cytokines (e.g. IL-1 and IL-6), chemokines (e.g. IL-8), and even bacterial products (e.g. LPS) can delay neutrophil apoptosis [190]. Corticosteroids also inhibit neutrophil apoptosis [40]. Subjects with neutrophilic asthma have high levels of airway IL-8, increased levels of endotoxin (LPS), and often these subjects are on high doses of inhaled corticosteroid, all of which may influence the persistence of neutrophilic inflammation through delaying neutrophil apoptosis [40].
The local environment can modulate functional properties of neutrophils. It is possible that the high level of IL-8 present in the airways provides feedback, which down regulates its own production from airway neutrophils. Reduced responsiveness of neutrophils to IL-8 has been reported in cystic fibrosis, where it is associated with exposure to high levels of IL-8, which induces receptor desensitisation [381]. IL-8 receptors (CXCR1 and CXCR2) are downregulated in sputum neutrophils versus blood neutrophils, and this modulation is greater in subjects with asthma compared to healthy controls [382].
Non-infective exposure to LPS present in the environment in both domestic and occupational settings is associated with the progression of airway inflammatory diseases including asthma [139, 265, 383]. LPS inhalation in humans activates effector cells, including airway epithelial cells and macrophages, which leads to the production of proinflammatory cytokines and chemokines that cause neutrophil influx [384].
Simpson et al [255] reported that neutrophilic asthma is characterised by innate immune activation of the airways, where higher levels of endotoxin in sputum supernatant and increased frequency of chronic bacterial colonisation are thought to be responsible for
175 the ongoing neutrophilic airway inflammation. The presence of these TLR agonists within the airways in neutrophilic asthma suggests that there is continual exposure to these agents, which is likely to have an effect on neutrophil function. Importantly, prolonged exposure of neutrophils to LPS is known to induce endotoxin tolerance where there is selective reprogramming of neutrophil function [385]. Tolerant neutrophils exhibit a reduction in respiratory burst activity and surface expression of TLR4, but a similar level of IL-8 release [385]. This suggests that whilst there is downregulation of TLR4 signalling during prolonged exposure of neutrophils to LPS, they continue to release IL-8 recruiting further neutrophils. In neutrophilic asthma, TLR4 gene expression is downregulated in airway neutrophils, however the cells still release IL-8 protein, albeit at a lower level in comparison to healthy controls. With such large numbers of neutrophils that are accumulated in the airways in neutrophilic asthma, this amount of IL-8 is significant and will promote further neutrophilic inflammation.
Corticosteroids are widely used for the reduction of inflammation in chronic inflammatory diseases including asthma and COPD. Non-eosinophilic asthma is associated with corticosteroid unresponsiveness [34]. Subjects with neutrophilic asthma in this study were on higher doses of inhaled corticosteroids, which may have had an impact on the reduction of cultured airway neutrophil cytokine responses seen in this study. However, this is not likely for several reasons. Corticosteroid dose was positively associated with TLR2 (Spearman r=0.76, p=0.0068) and TLR4 (Spearman r=0.69, p=0.019) mRNA expression, suggesting that expression of the innate signalling receptors was greater when subjects were using more corticosteroid. The non-response was not consistent across cytokines tested, for example levels of TNF-α were undetectable in neutrophilic asthma however IL-8 was still produced. This indicates a differential cytokine effect, which is not explained by corticosteroid exposure, and cytokine levels were different between phenotypes, but these differences did not follow differences in inhaled corticosteroid (ICS) doses. For example, while subjects with neutrophilic asthma had a higher dose of ICS than subjects with eosinophilic asthma, the IL-8 and IL-1β cytokine levels were similar in these 2 groups, and less than paucigranulocytic asthma.
The activation of the peripheral blood neutrophil results in their intravascular margination, adhesion to the endothelium, and migration to the site of inflammation e.g.
176 the airways. This process is regulated by chemokines, the most potent for neutrophils being IL-8 [119, 120, 126]. IL-8 is the main chemoattractant in the lung, since blocking of IL-8 with a neutralising antibody resulted in a 75-98% inhibition of its chemotactic activity [121]. IL-8 induces shedding of L-selectin and upregulation of the adhesion molecule Mac-1, which promotes attachment of neutrophils to the endothelium. Furthermore, IL-8 can activate various functions of the neutrophils, including degranulation and respiratory burst [125, 126]. IL-8 also causes the secretion of important neutrophil proteases such as elastase and MMP-9, which are capable of causing tissue damage [127]. IL-8 stimulates the bone marrow to release neutrophils into the circulation [98], and these can include immature neutrophils, which preferentially sequester in the lung [96].
Importantly, this study has shown that resting blood neutrophils isolated from subjects with non-eosinophilic asthma had an enhanced release of IL-8. Enhanced release of IL- 8 has previously been reported in blood neutrophils isolated from patients with cystic fibrosis [341]. Enhanced IL-8 release by peripheral blood neutrophils will provide autocrine feedback to prime the cells for migration to the airways. Enhanced IL-8 release may be due to positive feedback from the leakage of inflammatory mediators such as IL-8 from the airways, release of immature neutrophils from the bone marrow, or genetic differences such as IL-8 gene polymorphisms, however further investigation is needed to elucidate this.
Peripheral blood neutrophils from subjects with non-eosinophilic asthma, particularly neutrophilic asthma had a decreased expression of TLR2 both at rest and with LPS stimulation. TLR2 is important in the detection of gram-positive bacterial infections, therefore in this way lower TLR2 may predispose these subjects to more severe infections. TLR2 also plays an important role in the regulation of neutrophil apoptosis, as increased expression of TLR2 is associated with increased spontaneous neutrophil apoptosis [386].
This study shows that isolated blood neutrophils from subjects with eosinophilic asthma have a decreased response to LPS stimulation. Large variability in the inflammatory responses to inhalation of LPS in humans has previously been reported [387], and this is thought to be related to genetic variation of the genes in the TLR4 pathway. TLR4
177 polymorphisms have been linked with blunted airway [388] and systemic inflammatory responses to endotoxin [389]. Furthermore, the hyporesponsiveness to endotoxin caused by the presence of TLR4 polymorphisms is associated with an increase in the severity of atopy in subjects with asthma [390].
There is an intense neutrophilic inflammation in the airways of subjects with neutrophilic asthma demonstrated by massive numbers of neutrophils and levels of IL-8 in the sputum supernatant in affected individuals. This study reports enhanced IL-8 release from resting blood neutrophils in non-eosinophilic asthma, which is likely to perpetuate the migration of neutrophils to the airways, whilst airway neutrophils have impaired innate immune responses that may contribute to enhanced susceptibility to infection.
178 Chapter 8: Molecular Analysis of Neutrophils in Asthma Subtypes
8.1 Introduction
Neutrophils have long been considered to be phagocytes whose main purpose is to engulf and degrade microorganisms. However, recent microarray studies have provided substantial evidence that neutrophils are capable of extensive gene expression changes that are important in the regulation of many neutrophil functions, as well as modulation of the immune response. A wide range of genes are expressed in unstimulated neutrophils, and this gene profile is dramatically changed in response to bacterial exposure [220-222], transmigration to the airways [226], and neutrophilic mediated diseases [232].
An experimental model of neutrophil response to infection involves exposure of the cells to soluble LPS. Alterations in neutrophil gene expression have been reported in response to LPS exposure in vitro [220, 223-225] and in vivo [226], as well as during phagocytosis [227], and apoptosis [228-231]. Alterations in gene expression during exposure to bacteria include genes that encode for cytokines, receptors, genes involved in host defense, apoptosis-related genes, transcription factors, and chromatin- remodeling genes [220].
Due to their potent inflammatory nature, neutrophils have been implicated in the pathogenesis of airway diseases including asthma. Recently, subtypes of asthma have been defined based on induced sputum inflammatory cell counts, illustrating the heterogeneous nature of the disease. These inflammatory subtypes include eosinophilic asthma (increased sputum eosinophils), neutrophilic asthma (increased sputum neutrophils) and paucigranulocytic asthma (normal levels of sputum eosinophils and neutrophils)[9].
Neutrophils are present in increased numbers in neutrophilic asthma; however the precise mechanisms of their recruitment and accumulation remain largely unknown. Microarray technology is very useful and can uncover gene expression profiles, as well as novel genes and pathways that are associated with disease pathogenesis. Whole genome gene expression analysis has not been widely used to investigate the molecular
179 mechanisms underlying asthma, but could provide useful information relating to the heterogeneity of disease. To address the complex nature of the asthma pathogenesis and explore a potential role for neutrophils, we have investigated differential gene expression of isolated peripheral blood neutrophils in relation to asthma subtype. We hypothesise that neutrophil differential gene expression profiles will distinguish between neutrophilic and eosinophilic asthma subtypes, and implicate systemic neutrophil activation in the pathogenesis of neutrophilic asthma.
8.2 Methods
Induced sputum and peripheral blood samples were collected from subjects with asthma. Induced sputum was dispersed and differential and total cell counts were performed. Asthma was classified as eosinophilic if subjects had a sputum eosinophil count of >1% alone. Asthma was classified as neutrophilic if the subjects had >61% sputum neutrophils. Neutrophils were isolated from peripheral blood samples via percoll density gradient, erythrocyte lysis and magnetic cell separation using CD16 microbeads. Isolated neutrophils were cultured at 1 x 106 cells/mL in RPMI 1640 (1% FCS) with or without LPS stimulation (E.Coli, 100ng/mL) for 24 hours. RNA was extracted using Qiagen RNeasy columns and quantified using RiboGreen. 500ng of RNA was amplified using the Illumina TotalPrep RNA Amplification Kit (Ambion, Texas, USA) as per manufacturer’s instructions and hybridised to Sentrix HumanRef-8 Expression BeadChips overnight. The arrays were washed, stained with Cy3, and scanned using the Illumina BeadStation. For further details on methods please see Chapter 2 of this thesis.
8.2.1 Data Analysis
Data was normalised using cubic spline in Illumina’s BeadStudio 2.0 software and exported to GeneSpring 5.0 for further analysis. Genes were judged to be differentially regulated in resting and LPS stimulated neutrophils only when 1) the gene was present in all samples studied, 2) the difference in expression was >1.5 fold; and 3) the extent of difference in expression was statistically significant (p<0.05 Wilcoxon-Mann-Whitney test). The dominant themes of gene expression changes were determined by using the Protein Analysis Through Evolutionary Relationships (PANTHER) algorithm
180 (http://www.pantherdb.org/tools/). PANTHER categorises differentially expressed genes based on biological processes that can be associated with over and under- represented functional categories. This analysis is similar to Gene Ontology, but greatly simplified for high throughput analysis.
8.3 Results
8.3.1 Clinical Features Gene expression profiles were successfully generated for resting and LPS stimulated peripheral blood neutrophils isolated from 4 subjects with neutrophilic asthma and 5 subjects with eosinophilic asthma. Clinical characteristics for these subjects are shown in Table 8.1. Two subjects with neutrophilic asthma had a mixed granulocytic pattern of inflammation with an increased proportion of neutrophils and eosinophils.
Neutrophilic Asthma Eosinophilic Asthma p n 4 5 Age years, median (IQR) 70 (64-75) 63 (55-69) 0.33 Sex M | F 2 | 2 3 | 2 1.00 Atopy n (%) 3 (75) 4 (80) 1.00
FEV1 % predicted 53 (15) 66 (15) 0.21
FEV1/FVC % 71 (9) 80 (7) 0.15 ICS ‡‡ dose (μg) median (IQR) 2000 (1200-2500) 1000 (500-2000) 0.3 Table 8.1 Clinical characteristics of neutrophilic and eosinophilic asthma subjects ‡‡ ICS dose is calculated 1μg of beclomethasone = 1μg of budesonide = 0.5μg of fluticasone.
8.3.2 Induced Sputum Inflammatory Cells
Subjects with eosinophilic asthma had an increased proportion of sputum eosinophils compared to those with neutrophilic asthma (Table 8.2). Subjects with neutrophilic asthma had a significantly increased sputum total cell count, proportion and absolute number of neutrophils, and a decreased proportion of macrophages compared to eosinophilic asthma. Absolute numbers of macrophages were similar between the two
181 groups and therefore the reduced proportion of macrophages was due to an increased proportion of neutrophils and expressing the result as a percentage. Subjects with neutrophilic asthma had similar proportions but increased absolute numbers of lymphocytes, most likely due to an increased total cell count in these subjects.
Neutrophilic Asthma Eosinophilic Asthma p n 4 5 Total cell count x 106/mL 16.2 (10.7-18.3) 3.3 (2.3-6) 0.03 Viability (%) 77.7 (59.7-91.2) 67.2 (62.2-69.2) 0.33 Neutrophils, % 74.3 (71.8-84. 29.7 (17.8-32.7) 0.01 Neutrophils 104/mL 1159.6 (786.7-1503.2) 69.5 (59.3-75.2) 0.01 Eosinophils, % 1.2 (0.4-1.7) 6.3 (3.9-9.6) 0.01 Eosinophils 104/mL 17.7 (2.6-31.6) 20.9 (11.2-23.5) 0.62 Macrophages, % 20 (11.3-22.8) 52.8 (51.9-63.6) 0.01 Macrophages 104/mL 226.9 (109.7-368) 229.4 (148.8-243) 0.81 Lymphocytes, % 1.1 (0.5-1.7) 1.2 (1-1.9) 0.54 Lymphocytes 104/mL 13 (8.8-17.5) 4 (3.4-5.8) 0.05 Columnar epithelial cells, % 1.9 (0.5-3.3) 5.8 (1.9-13.1) 0.14 Columnar epithelial cells 104/mL 32.4 (8-53.2) 19.3 (2.2-79) 0.62 Squamous cells, % 1.5 (0-7.2) 10.8 (7.1-13.2) 0.09 Table 8.2 Induced sputum inflammatory cell counts for subjects with eosinophilic asthma, non-eosinophilic asthma and healthy controls
8.3.3 Altered Neutrophil Gene Expression with LPS Stimulation
Dramatic changes in gene expression were apparent between resting and LPS stimulated circulating neutrophils isolated from subjects with asthma. Using the Wilcoxon-Mann- Whitney test, 1080 genes were identified with a mean ratio of expression that was significantly different comparing resting to LPS stimulated neutrophils. Construction of a dendogram containing these 1080 genes showed that the gene expression profiles from resting neutrophils are closely related, but significantly different to the LPS stimulated neutrophils (Figure 8.1). As expected, the LPS stimulated gene profile represented a proinflammatory state of neutrophil activation with increases in cytokines
182 (e.g. OSM), chemokines (e.g. IL-8, CCL3L1, CXCL1), signalling molecules (e.g. IRAK1, IRAK3), receptors (e.g. TLR2, CXCR4, CCR1), molecules regulating apoptosis (e.g. GADD45B, SGK, CEBPB) and components of the NF-κB pathway (e.g. NFKB1, RIPK2, TNFRSF14). The LPS regulated genes OSM, TLR2 and IL-8 were confirmed to be upregulated via real time PCR (Table 8.3).
183 all...
...
Figure 8.1 Gene expression profiles of resting versus LPS stimulated neutrophils Columns represent gene expression for both resting and LPS stimulated neutrophils from each of the subjects with asthma. Down-regulation is represented as green, and up- regulation is represented as red. The dendrogram at the top of the figure represents the relationship between resting and LPS stimulated neutrophils (blue branches: resting neutrophils; and red branches: LPS stimulated neutrophils). The dendrogram on the side represents the relationship between the expression levels of the gene across all the samples.
184 Gene Symbol Array Results Real-Time Results
TLR2 + 5.8 + 5.6
OSM + 2.7 + 6.8
IL-8 + 6.3 + 22.2
Table 8.3 OSM, TLR2 and IL-8 fold change increase with LPS stimulation from baseline for microarrays and real-time PCR
8.3.4 Altered Gene Expression Profiles in Asthma Subtypes
To investigate changes in gene expression profiles of resting and LPS stimulated neutrophils in asthma subtypes, the subjects were divided according to the presence of either high sputum neutrophils or high sputum eosinophils.
8.3.4.1 Resting Blood Neutrophils
Using the Wilcoxon-Mann-Whitney test, 317 genes were identified as having significantly different levels of expression across the asthma subtypes, for resting neutrophils. Construction of a dendogram containing these 317 genes showed that the gene expression profiles from subjects with neutrophilic asthma were closely related, but significantly different to the subjects with eosinophilic asthma (Figure 8.2). Of these 317 genes that were significantly altered in resting neutrophils from subjects with neutrophilic asthma versus eosinophilic asthma, 121 were upregulated more than 1.5 fold, and 182 genes were downregulated more than 1.5 fold.
When comparing the genes that were differentially regulated by LPS treatment and in resting neutrophils in neutrophilic asthma it became apparent that a significant number of genes that were regulated in resting neutrophils in neutrophilic asthma were also regulated by LPS. This is demonstrated in Figure 8.3, which shows the amount of genes in common between the two gene lists. There were 171 from 317 (54%) genes altered in resting neutrophils in neutrophilic asthma that were also altered by LPS treatment in vitro. These genes are likely to play a role in neutrophil activation, which suggests that
185 the circulating neutrophils in neutrophilic asthma have increased activation in comparison to neutrophils isolated from subjects with eosinophilic asthma. Table 8.4 lists 47 genes that were upregulated in resting neutrophils in neutrophilic asthma that were also upregulated by in vitro LPS stimulation in this study.
Genes involved in signal transduction that were upregulated in neutrophilic asthma are listed in Table 8.5. Analysis of genes that were upregulated in neutrophilic asthma revealed an increased number involved in cell motility (Table 8.6), apoptosis (Table 8.7) and the NF-κB cascade (Table 8.8). Other selected genes with immune related functions that were altered in resting neutrophils from subjects with neutrophilic asthma in comparison to eosinophilic asthma are shown in Table 8.9. There were a significant number of genes that were of unknown function, of which 38% were upregulated and 51% of downregulated.
186 asthma...
...
Figure 8.2 Gene expression profiles of resting neutrophils from subjects with eosinophilic asthma versus those with neutrophilic asthma Columns represent gene expression for resting neutrophils from each of the subjects with asthma. Down-regulation is represented as green, and up-regulation is represented as red. The dendrogram at the top of the figure represents the relationship between subjects with neutrophilic and eosinophilic asthma (Experiment Tree: red branches=eosinophilic asthma; and blue branches=neutrophilic asthma). The dendrogram on the side represents the relationship between the expression levels of each gene across all the samples (Gene Tree).
187
909 171 146
Genes altered in resting neutrophils in neutrophilic asthma (n=317) Genes altered by LPS stimulation (n=1080) Figure 8.3 Schematic representation of genes altered both by LPS and in neutrophilic asthma The genes altered by LPS are represented in the blue circle and the genes altered in resting neutrophils in neutrophilic asthma are represented in the red circle. The area where these circles overlap represents genes in common, which equates to 171 out of 317 or 54% of genes.
188 Table 8.4 LPS regulated genes that are also upregulated in resting neutrophils from subjects with neutrophilic asthma versus eosinophilic asthma GenBank Symbol p value Fold Change NM 001456.1 FLNA Filamin A, α 0.04 + 5.1 NM_007199.1 IRAK3 Interleukin-1 receptor associated kinase-3 0.04 + 4.3 XM_370714.2 FTHL7 Ferritin, heavy polypeptide like 7 0.01 + 4.2 NM_018370.1 DRAM Damage regulated autophagy modulator 0.003 + 3.8 NM_003965.3 CCRL2 Chemokine CC motif receptor like 2 0.04 + 3.4 NM_003131.1 SRF Serum response factor 0.01 + 3.2 NM_002638.2 PI3 Protease Inhibitor 3, skin-derived (SKALP) 0.01 + 3.2 NM_015675.1 GADD45B Growth arrest and DNA damage inducible β 0.003 + 3.1 NM_004251.3 RAB9A RAB9A, member RAS oncogene family 0.01 + 3.1 NM_006018.1 HM74 G protein coupled receptor 109B 0.01 + 3.0 NM 002964.3 S100A8 S100 calcium binding protein A8 0.04 + 2.6 NM_003821.4 RIPK2 Receptor (TNFRSF) interacting serine- 0.003 + 2.6 threonine kinase 2 NM_005340.2 HINT1 Histidine triad nucleotide binding protein 1 0.003 + 2.6 NM_006755.1 TALDO1 Transaldolase 1 0.003 + 2.4 NM_022044.1 SDF2L1 Stromal cell derived factor 2-like protein 0.01 + 2.4 NM_024067.2 C7orf26 Chromosome 7 open reading frame 26 0.01 + 2.4 NM_013439.2 PILRA Paired immunoglobulin-like type 2 receptor 0.04 + 2.4 alpha NM_031419.1 MAIL Molecule possessing ankyrin repeats induced 0.04 + 2.4 by lipopolysaccharide NM_000584.2 IL8 Interleukin-8 0.04 + 2.4 NM_014330.2 PPP1R15A Protein phosphatase 1, regulatory (inhibitor) 0.04 + 2.4 subunit 15A NM_007267.5 EVER1 Transmembrane channel like 6 0.04 + 2.3 NM_016154.2 RAB4B RAB4B, member RAS oncogene family 0.003 + 2.3 NM_015266.1 SLC9A8 Solute carrier 9 0.003 + 2.3 NM_000146.2 FTL Ferritin light polypeptide 0.01 + 2.2 NM_021009.2 UBC Ubiquitin C 0.04 + 2.2 NM_004048.2 B2M Beta 2 microglobulin 0.01 + 2.2 NM_003820.2 TNFRSF14 Tumor necrosis factor receptor superfamily, 0.003 + 2.2 member 14 NM_003375.2 VDAC2 Voltage dependent anion channel 2 0.003 + 2.2 NM_006520.1 TCTE1L T-complex associated -testis-expressed 1 like 0.04 + 2.1 NM_006460.1 HIS1 Hematopoietic insertion site 1 0.01 + 2.1 NM_001693.2 ATP6V1B2 Vacuolar H+ATPase B2 0.01 + 2.0 NM_004547.4 NDUFB4 NADH dehydrogenase (ubiquinone) 1 beta 0.003 + 2.0 subcomplex 4 NM_005706.2 TSSC4 Tumor suppressing subchromosomal 0.003 + 2.0 transferable fragment 4 NM_182608.2 DKFZp686O168 Ankyrin repeat domain 33 0.04 + 2.0 NM_003370.1 VASP Vasodilator stimulated phosphoprotein 0.04 + 2.0 NM_005627.2 SGK Serum/glucocorticoid regulate kinase 0.01 + 2.0 NM_000433.1 NCF2 Neutrophilic cytosolic factor 2 0.04 + 2.0 NM_001183.3 ATP6AP1 ATPase, H+ transporting lysosomal accessory 0.04 + 1.9 protein NM_025205.1 EG1 Endothelial derived gene 1 0.04 + 1.9 NM_003830.1 SIGLEC5 Siliac acid binding Ig-like lectin 5 0.01 + 1.9 NM_001861.2 COX4I1 Cytochrome c oxidase subunit IV isoform 1 0.04 + 1.8 NM_016202.1 LOC51157 Zinc finger protein 580 0.04 + 1.7 NM_005729.3 PPIF Peptidylprolyl isomerase F (cyclophilin F) 0.04 + 1.7 NM_005194.2 CEBPB CCAAT/enhancer binding protein (C/EBP), 0.01 + 1.7 beta NM_017548.2 H41 CDV3 homolog (mouse) (CDV3) 0.01 + 1.7 NM_000086.1 CLN3 Ceroid lipofuscinosis, neuronal 3, juvenile 0.04 + 1.6 NM_004637.5 RAB7 RAB7, member RAS oncogene family 0.04 + 1.5
189 Table 8.5 Upregulated genes in resting neutrophils from subjects with neutrophilic asthma versus eosinophilic asthma involved in signal transduction GenBank Symbol p value Fold Change NM 007199.1 IRAK3 Interleukin-1 receptor associated kinase-3 0.04 + 4.3 NM_003965.3 CCRL2 Chemokine CC motif receptor like 2 0.04 + 3.4 NM_015675.1 GADD45B Growth arrest and DNA damage inducible β 0.003 + 3.1 NM_004251.3 RAB9A RAB9A, member RAS oncogene family 0.01 + 3.1 NM_006018.1 HM74 G-protein coupled receptor 109B 0.01 + 3.0 NM_003821.4 RIPK2 Receptor (TNFRSF) interacting serine- 0.003 + 2.6 threonine kinase 2 NM_153023.1 SPATA13 Spermatogenesis associated 13 0.01 + 2.4 NM_000584.2 IL-8 Interleukin-8 0.04 + 2.4 NM_016154.2 RAB4B RAB4B, member RAS oncogene family 0.003 + 2.3 NM_000228.1 LAMB3 Laminin beta 0.04 + 2.3 NM_003820.2 TNFRSF14 Tumor necrosis factor receptor superfamily, 0.003 + 2.2 member 14 NM_005627.2 SGK Serum/glucocorticoid regulate kinase 0.01 + 2.0 NM 004309.3 ARHGDIA Rho GDP dissociation inhibitor GDI alpha 0.04 + 1.9 NM_003830.1 SIGLEC5 Siliac acid binding Ig-like lectin 5 0.01 + 1.9 NM_020680.2 SCYL1 SCY1-like 1 (S.cerevisiae) 0.04 + 1.7 NM_025144.2 LAK Alpha kinase 1 0.01 + 1.7 NM_004637.5 RAB7 RAB7, member RAS oncogene family 0.04 + 1.5
Table 8.6 Differentially regulated genes in resting neutrophils from subjects with neutrophilic asthma versus eosinophilic asthma related to cell motility GenBank Symbol p value Fold Change NM 001456.1 FLNA Filamin A, α 0.04 + 5.1 NM_003965.3 CCRL2 Chemokine CC motif receptor like 2 0.04 + 3.4 NM_003131.1 SRF Serum response factor 0.01 + 3.2 NM_002964.3 S100A8 S100 calcium binding protein A8 0.04 + 2.6 NM_000584.2 IL-8 Interleukin-8 0.04 + 2.4 NM_006000.1 TUBA1 Tubulin alpha 0.04 + 2.1 NM_003370.1 VASP Vasodilator stimulated phosphoprotein 0.04 + 2.0
190 Table 8.7 Genes that were altered in resting neutrophils from subjects with neutrophilic asthma versus eosinophilic asthma involved in apoptosis GenBank Symbol p value Fold Change NM 015675.1 GADD45B Growth arrest and DNA damage inducible 0.003 + 3.1 NM_003883.2 HDAC3 Histone deacetylase 3 0.003 + 2.7 NM_003821.4 RIPK2 Receptor (TNFRSF) interacting serine- 0.003 + 2.6 threonine kinase 2 NM_139205.1 HDAC5 Histone deacetylase 5 0.01 + 2.5 NM_014330.2 PPP1R15A Protein phosphatase 1, regulatory 0.04 + 2.4 (inhibitor) subunit 15A NM_003820.2 TNFRSF14 Tumor necrosis factor receptor 0.003 + 2.2 superfamily, member 14 NM_005627.2 SGK Serum/glucocorticoid regulate kinase 0.01 + 2.0 NM_005194.2 CEBPB CCAAT/enhancer binding protein 0.01 + 1.7 (C/EBP), beta NM_005118.2 TNFSF15 Tumor necrosis factor (ligand) 0.01 - 2.1 superfamily member 15
Table 8.8 Genes that were upregulated in resting neutrophils from subjects with neutrophilic asthma versus eosinophilic asthma involved in the NF-κB cascade GenBank Symbol p value Fold Change NM_015675 GADD45B Growth arrest and DNA damage inducible β 0.003 + 3.1 NM_003821.4 RIPK2 Receptor (TNFRSF) interacting serine- 0.003 + 2.6 threonine kinase 2 NM_000584.2 IL-8 Interleukin-8 0.04 + 2.4 NM_003820.2 TNFRSF14 Tumor necrosis factor receptor superfamily, 0.003 + 2.2 member 14
191 Table 8.9 Selected genes with immune related function that were altered in resting neutrophils from subjects with neutrophilic asthma compared to eosinophilic asthma GenBank Symbol PANTHER Biological Process p value Fold Change Upregulated NM_014015.3 DEXI Dexamethasone-induced transcript Unclassified 0.04 + 4.9 NM_012092.2 ICOS Inducible T cell co-stimulator Unclassified 0.04 + 3.1 NM_000247.1 MICA MHC class I polypeptide related sequence A MHCI mediated immunity 0.01 + 2.8 NM_002117.3 HLA-C MHCI mediated immunity 0.003 + 2.3 NM_004604.3 STX4A Syntaxin 4A Regulated exocytosis 0.003 + 1.8 NM_005516.3 HLA-E Major histocompatibility complex class I, E MHCI mediated immunity 0.01 + 1.8 NM_001613.1 ACTA2 Actin α2 Cell Structure 0.04 + 1.8 Downregulated NM_002697.2 POU2F1 POU domain class 2 transcription factor 1 mRNA transcription regulation 0.003 - 1.5 NM_133280.1 FCAR Fc fragment of IgA, receptor for Immunity and defence 0.04 - 1.6 NM_000896.1 CYP4F3 Cytochrome P450 family 4 subfamily F polypeptide 3 Steroid metabolism 0.01 - 1.6 NM_003298.2 NR2C2 Nuclear receptor subfamily 2, group C, member 2 mRNA transcription regulation 0.003 - 1.7 NM_031483.3 ITCH Itchy homologue E3 ubiquitin protein ligase Proteolysis 0.003 - 1.7 NM_177536.1 SULT1A1 Sulfotransferase family, cytosolic, 1A, phenol preferring, member 1 Steroid hormone metabolism 0.04 - 1.7 USP14 Ubiquitin specific peptidase 14 Proteolysis 0.04 - 1.8 NM_147134.1 NFX1 Nuclear transcription factor, X-box binding 1 mRNA transcription regulation 0.04 - 1.9 NM_178509.3 STXBP4 Syntaxin binding protein 4 Unclassified 0.04 - 1.9 NM_147191.1 MMP21 Matrix metalloproteinase 21 Proteolysis 0.01 - 1.9 NM_000437.2 PAFAH2 Platelet activating factor acetylhydrolase 2, 40kDa Immunity and defence 0.003 - 1.9 NM_006885.2 ATBF1 AT-binding transcription factor mRNA transcription regulation 0.04 - 1.9 NM_016459.2 PACAP Proapoptotic caspase adapter protein Unclassified 0.003 - 2.1 NM_152878.1 MAFF v-maf musculoaponeurotic fibrosarcoma oncogene homologue F mRNA transcription regulation 0.04 - 2.2 NM_006896.2 HOXA7 Homeobox A7 mRNA transcription regulation 0.01 - 2.2 NM_001074.1 UGT2B7 UDP glucuronosyltransferase 2 family, polypeptide B7 Steroid hormone metabolism 0.04 - 2.6 NM_001075.2 UGT2B10 UDP glucuronosyltransferase 2 family, polypeptide B10 Steroid hormone metabolism 0.003 - 3.1 NM_198257.1 E2F6 E2F transcription factor mRNA transcription regulation 0.01 - 3.1 NM_007038.1 ADAMTS5 A disintegrin like and metallopeptidase with thrombospondin type 1 Proteolysis 0.003 - 3.3 NM_001073.1 UGT2B11 UDP glucuronosyltransferase 2 family, polypeptide B11 Steroid hormone metabolism 0.003 - 3.5 NM_080764.2 SUHW2 Suppressor of hairy wing homologue 2 mRNA transcription regulation 0.01 - 9.9
192 8.3.4.2 LPS Stimulated Blood Neutrophils
Using the Wilcoxon-Mann-Whitney test, 221 genes were identified with a mean ratio of expression that was significantly different across the asthma subtypes for LPS stimulated neutrophils. Construction of a dendrogram containing these 221 genes showed that the gene expression profiles from subjects with neutrophilic asthma are closely related, but significantly different to the subjects with eosinophilic asthma (Figure 8.4). Selected genes with immune related functions that were altered in LPS stimulated neutrophils from subjects with neutrophilic asthma compared to subjects with eosinophilic asthma are listed in Table 8.10.
193 asthma lps...
...
Figure 8.4 Gene expression profiles of LPS stimulated neutrophils from subjects with neutrophilic asthma versus those with eosinophilic asthma Columns represent gene expression for LPS stimulated neutrophils from each of the subjects with asthma. Down-regulation is represented as green, and up-regulation is represented as red. The dendrogram at the top of the figure represents the relationship between subjects with neutrophilic and eosinophilic asthma (Experiment Tree: red branches=eosinophilic asthma; and blue branches=neutrophilic asthma). The dendrogram on the side represents the relationship between the expression levels of the gene across all the samples (Gene Tree).
194 Table 8.10 Selected genes with immune related function that were altered in resting neutrophils from subjects with neutrophilic asthma compared to eosinophilic asthma GenBank Symbol Name PANTHER Biological Process p value Fold Change Upregulated NM_002658.1 PLAU Plasminogen activator, urokinase Proteolysis, Cell Motility 0.04 + 3.4 NM_012342.1 BAMBI BMP and activin membrane bound Unclassified 0.01 + 3.1 NM_001618.2 PARP1 Poly (ADP-ribose) polymerase family member 1 DNA damage 0.01 + 2.5 NM_006904.6 PRKDC Protein kinase, DNA activated, catalytic polypeptide DNA damage 0.003 + 2.4 NM_000181.1 GUSB Glucuronidase β Carbohydrate metabolism 0.04 + 2.3 NM_005723.2 TM4SF9 Tetraspanin 5 Cell Adhesion 0.04 + 2.3 NM_005746.1 PBEF Pre B cell colony enhancing factor Immunity and defence 0.04 + 1.9 NM_000877.2 IL1R1 Interleukin-1 receptor type 1 Cytokine/Chemokine mediated pathway 0.01 + 1.9 NM_006290.2 TNFAIP3 Tumor necrosis factor alpha induced protein 3 Unclassified 0.003 + 1.8 NM_005063.3 SCD Stearoyl-CoA desaturase (delta-9-desaturase) Fatty acid metabolism 0.04 + 1.7 NM_033405.2 PRIC285 Peroxysomal proliferator activated receptor A, interacting complex 285 mRNA transcription regulation 0.04 + 1.6 NM_006534.2 NCOA3 Nuclear receptor coactivator 3 mRNA transcription regulation 0.04 + 1.6 NM_002514.2 NOV Nephroblastoma overexpressed gene Cell communication 0.04 + 1.6 NM_004333.2 BRAF v-raf murine sarcoma viral oncogene homologue B1 Apoptosis 0.04 + 1.5 NM_021738.1 SVIL Supervillan Cell Structure 0.04 + 1.5 NM_003153.3 STAT6 Signal transducer and activator of transcription-6 JAK-STAT cascade 0.04 + 1.5 Downregulated NM_024535.1 FLJ22021 Coronin 7 Cell motility 0.04 - 1.5 NM_005981.3 TSPAN31 Tetraspanin 31 Cell proliferentiation and differentiation 0.04 - 1.7 NM_016150.3 ASB2 Ankyrin repeat and SOCS box containing 2 Unclassified 0.04 - 1.7 NM_198291.1 SRC V-src sarcoma (Schmidt-Ruppin A-2) viral oncogene homologue Immunity and defence 0.003 - 2.4 NM_058171.2 ING2 Inhibitor of growth family, member 2 Apoptosis 0.04 - 2.8 NM_003265.2 TLR3 Toll-like receptor 3 Cytokine/Chemokine mediated signalling 0.04 - 2.9 NM_145898.1 CCL23 Chemokine (CC motif) ligand 23 Cytokine/Chemokine mediated pathway 0.003 - 5.9
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8.4 Discussion
There is a substantial degree of genetic heterogeneity in resting neutrophils from subjects with neutrophilic and eosinophilic asthma, reflecting the complex nature of asthma pathogenesis. This study highlights the ability of microarray technology to define inflammatory gene profiles associated with asthma, and is the first study to investigate neutrophil gene expression in asthma in relation to neutrophilic and eosinophilic subtypes. In this study, novel and distinct gene expression profiles were created from resting and LPS stimulated peripheral blood neutrophils using Illumina BeadArrays that were used to identify gene expression profiles relating to asthma subtype. These data also implicate systemic neutrophil activation in the pathogenesis of neutrophilic asthma, in particular, the enhancement of neutrophil chemotaxis, survival, and activation of the NF-κB cascade.
The development of high-throughput screening and genome wide gene expression by microarrays has allowed many diseases to be characterised into groups by gene expression profiling. Analysis of the current data suggests that the type of airway inflammation present can separate asthma into subgroups based on altered systemic neutrophil gene expression profiles. Although relatively small groups were studied here, significant differences (p<0.05) in gene expression and distinct dendrograms were observed. In addition, genes in peripheral blood neutrophils from asthma subtypes with known immune related functions were identified and 3 known LPS regulated genes IL-8, TLR2 and OSM were confirmed via real-time PCR.
It is well documented that LPS stimulation of neutrophils generates a robust transcriptional response [220, 223-225], especially for cytokines, indicating that the neutrophil contributes much more than the release of pre-formed mediators and bactericidal agents. However, previous studies have been limited, as they have not examined gene expression across the whole genome. This study has reported 1080 genes that were significantly altered by LPS treatment in vitro across the >24,000 genes tested. These included the up-regulation of many genes that are important in regulating the immune response.
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Key insights were gained into the pathogenesis of neutrophilic asthma through the analysis of altered genes from resting neutrophils once the asthma subtypes were compared. These analyses showed that many genes (54%) that were upregulated in resting neutrophils in neutrophilic asthma were also further upregulated by LPS stimulation, indicating that these genes play a role in neutrophil activation. Importantly, genes relating to cell motility, apoptosis and the NF-κB cascade were also among those upregulated.
Large numbers of neutrophils are present in the airways of subjects with neutrophilic asthma. Increased accumulation of neutrophils in the airways could be due to either enhanced chemotaxis from the blood and/or enhanced survival of these cells. Here we have shown that peripheral blood neutrophils have increased expression of genes relating to enhanced cell motility and survival. Genes relating to cell motility that were upregulated in neutrophilic asthma include, most importantly, IL-8, S100A8, CCRL2 and SRF, suggesting that these cells are primed systemically for migration to the airways.
IL-8 is a potent chemotactic factor for neutrophils and is proinflammatory in its effects [119, 120]. It is the main chemoattractant in the lung, since blocking of IL-8 with a neutralising antibody resulted in a 75-98% inhibition of chemotaxis [121]. IL-8 can delay neutrophil apoptosis [391] and stimulate the bone marrow to release neutrophils into the circulation [392]. We have also reported down-regulation of POU2F1, a transcription factor important in the regulation of IL-8 expression. Down-regulation of POU2F1 has been reported to increase IL-8 gene expression [393].
S100A8 belongs to a family of low molecular weight calcium binding proteins that are involved in regulating intracellular calcium dependent processes [394]. S100A8 is also involved in neutrophil chemotaxis, adhesion and other proinflammatory activities [395, 396]. S100A8 forms a heterodimer with S100A9, and interestingly this heterodimer has been shown to induce NF-kB activation in macrophages [397], as well as IL-8 production in airway epithelial cells [398]. S100A8 is reported to be associated with LPS-induced neutrophilic lung inflammation that was steroid resistant in a mouse model [399].
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A number of genes encoding proteins with receptor activity were also upregulated in neutrophilic asthma, including the orphan chemokine receptor CCRL2. CCRL2 gene expression has previously been reported in neutrophils and its expression was increased with LPS stimulation in vitro [400]. CCRL2 expression is also upregulated on synovial neutrophils in rheumatoid arthritis [400], and alveolar neutrophils after endotoxin instillation in the lung [226].
Serum response factor (SRF) plays a crucial role in cellular migration and normal actin cytoskeleton and contractile biology. SRF is a transcription factor that regulates an expanding number of genes that encode for actin-cytoskeleton and contractile proteins [401]. SRF expression has been previously reported in human neutrophils, and is increased with both LPS and IL-8 stimulation in vitro [402].
Neutrophils that do not undergo apoptosis undergo secondary necrosis, where they release their toxic cytosolic contents, which can cause significant tissue damage. A large number of the changes in apoptosis related genes are associated with increased neutrophil survival. Particular examples of genes that were increased in neutrophilic asthma and relate to a delay in apoptosis include GADD45β, HDAC3, HDAC5, SGK, and CEBPB.
The NF-κB pathway plays an important role in the control of neutrophil apoptosis, and there is increased gene expression of several members of this pathway in neutrophils isolated from subjects with neutrophilic asthma. GADD45β has recently been identified as a novel antiapoptotic gene whose expression depends on NF-κB. GADD45B is a pivotal mediator of the NF-κB protective activity [403, 404] that mediates cell survival via the inhibition of JNK signalling through the TNF receptor [403]. RIPK2 is another important signalling molecule involved in the activation of NF-κB through stimulation of numerous innate immune receptors including TLR2, TLR4, NOD proteins, IL-1R, and IL-18R [405]. Serum/glucocorticoid-regulated kinase (SGK) is involved in signal transduction and its gene expression is crucial in promoting neutrophil survival. SGK expression is decreased during spontaneous apoptosis and increased with GM-CSF treatment [406]. The mechanism by which SGK influences cell survival is also thought to involve the NF-κB
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pathway [407]. IRAK-M (IRAK3), a negative regulator of TLR signalling [408] was upregulated in resting neutrophils in neutrophilic asthma. IRAK-M has recently been linked with the pathogenesis of early-onset persistent asthma [409].
There is a considerable amount of literature demonstrating that cell fate is regulated at the level of gene expression, and that these changes are important in the resolution of inflammatory processes [227-231]. Although data from this study strongly suggests that peripheral blood neutrophils have an increased survival rate in neutrophilic asthma compared to eosinophilic asthma, it is important to clarify whether these changes in gene expression are related to changes in neutrophil survival at the functional level in future studies.
Activation of the innate immune response in the form of increases in the expression of the key receptors TLR4, TLR2, CD14 and SP-A and the proinflammatory cytokines IL-8 and IL-1β has been demonstrated in the airways of subjects with neutrophilic asthma. This study provides evidence that the activation of neutrophils begins in the circulation in these subjects, which could both promote the development of neutrophilic airway inflammation, and further potentiate existing neutrophilic airway inflammation. Many genes that were increased with LPS stimulation were also increased in resting neutrophils from subjects with neutrophilic asthma. Abraham et al [410] demonstrated that there is a significant correlation that exists between peripheral blood neutrophil phenotypes and the pulmonary response to endotoxin, that is, the accumulation of neutrophils and the intensity of the immune response in the airways to endotoxin challenge is directly associated with the activation state of circulating neutrophils.
Differences in the response to LPS may also play a role in the innate immune defence against invading microorganisms, and may contribute to airway inflammation. This study demonstrated significant alterations in gene expression after LPS stimulation in neutrophilic asthma compared to eosinophilic asthma. The genes that were altered suggested that there was a potentiation of LPS responses in neutrophilic asthma (e.g PLAU
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and IL-1R1), and further increases in genes relating to cell survival (e.g. PBEF, TNFAIP3, BRAF, PRKDC, SVIL).
Plasminogen activator (PLAU) is a serine protease that catalyses the conversion of plasminogen to plasmin. PLAU was upregulated in LPS stimulated neutrophils in neutrophilic asthma, and has been shown to potentiate LPS induced neutrophil activation [411]. Supervillin (SVIL) is an F-actin binding protein that associates with both actin filaments and the plasma membrane [412]. SVIL could cause morphological changes in the cell cytoskeleton that could lead to altered neutrophil functions including random migration and chemotaxis [413]. Pre-B cell colony enhancing factor (PBEF) is an adipokine that is involved in the inhibition of neutrophil apoptosis and IL-1β, OSM and LPS regulate its expression [414]. PBEF is also implicated in the pathogenesis of rheumatoid arthritis [415]. Also interesting is the 6-fold down-regulation of CCL23 in LPS stimulated neutrophils in neutrophilic asthma. CCL23 suppresses neutrophil and monocyte progenitors, decreases pool size and slows their turnover in the bone marrow [416].
These findings support the notion that neutrophils are transcriptionally active cells that are responsive to environmental stimuli and capable of a complex series of late transcriptional changes. This is the first study to examine whole genome gene expression of peripheral blood neutrophils relating to asthma subtype. We have identified specific gene profiles associated with neutrophilic and eosinophilic asthma, providing further validation that these subtypes of asthma involve very different molecular mechanisms of disease pathogenesis at the systemic level. This study has important implications for treatment strategies and highlights the importance of neutrophils in the pathogenesis of neutrophilic asthma.
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Chapter 9: General Discussion
It is clear that neutrophils are an important part of innate immune defense against infections, however dysfunctional neutrophil responses could contribute extensively to the pathogenesis of a number of inflammatory lung diseases including neutrophilic asthma. This thesis has examined innate immune responses of both airway and circulating neutrophils, with a focus on the contribution of neutrophils to inflammation in asthma and subtypes of asthma. Comparison of innate immune responses of airway and blood neutrophils identified distinct differences, illustrating that airway neutrophils have a unique phenotype, characterised by a high level of spontaneous activation but tolerance to further activation by an innate stimulus. It was found that there are modifications to the innate immune responses of neutrophils that occur during healthy ageing both in the airways and in response to LPS stimulation, which are likely to have an impact on both the development and the persistence of neutrophilic airway disease.
Neutrophils are present in large numbers in the airways in airway diseases such as neutrophilic asthma and COPD. Whole genome gene expression analysis of isolated peripheral blood neutrophils revealed distinct profiles corresponding to neutrophilic and eosinophilic asthma subtypes, and implicated systemic neutrophil activation in the pathogenesis of neutrophilic asthma. This is also supported by in vitro studies showing a relative increase in circulating neutrophil activation in asthma and COPD. These mechanisms would be expected to increase neutrophil accumulation in the airways. It is not known whether the accumulation of these cells in the airway is associated with excess cell activation. This thesis has demonstrated that airway diseases; particularly neutrophilic asthma and COPD are associated with an impaired neutrophil response in the airways in comparison to healthy volunteers, and these findings add to increasing evidence that suggests that increased presence of airway neutrophils does not correspond to increased cellular activity. A summary of the results presented in this thesis can be seen in Table 9.1.
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Systemic Airway LPS Age Asthma COPD NA V’s Blood Age Asthma COPD NA Cytokines ↑ ↓ = ↑ ↑* ↑ ↑ ↓ ↓ ↓ Degranulation ↑ ↑* = = = ↓ = = = = TLR4 ↑ ↑* = = = = = ↓ ↓ ↓ TLR2 ↑ = ↓ ↓ ↓ ↑ ↑ = ↓ ↓
Table 9.1 Summary of Results. * Resting neutrophils only. ↑ Increase, ↓ Decrease, = Unchanged. Systemic Responses: When peripheral blood neutrophils are stimulated with LPS, there are increases in the production of cytokines, the degranulation of gelatinase granules and the expression of TLR2 and TLR4. Airway Responses: Resting airway neutrophils are highly activated and have an increased production of cytokines, and expression of TLR2 compared to resting and LPS stimulated blood neutrophils, however they no longer have the capacity to degranulate and release MMP-9. Ageing: During healthy ageing, peripheral blood neutrophils have a decreased production of cytokines in response to LPS, however they have an increased degranulation and expression of TLR4 at rest. Airway neutrophils from older subjects have an increased production of cytokines and expression of TLR2. Airway Disease: There is a decreased expression of TLR2 in LPS stimulated blood neutrophils in asthma, COPD and neutrophilic asthma, an increase in the production of IL-8 from resting neutrophils in non- eosinophilic asthma, and an increased production of cytokines in response to LPS from COPD neutrophils. Airway neutrophils have a decreased release of cytokines and expression of TLR4 in asthma, particularly neutrophilic asthma and COPD. In addition airway neutrophils from subjects with neutrophilic asthma or COPD have a decreased expression of TLR2.
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9.2 Relative Cytokine Production from Isolated Neutrophils
This thesis reports a decreased proinflammatory cytokine release from airway neutrophils in neutrophilic asthma, of interest however is how this compares to the amount released from blood neutrophils at rest and under stimulation. Using IL-8 release as an example, the baseline level of activation of airway neutrophils in neutrophilic asthma is similar to LPS stimulated blood neutrophils, but significantly more than resting blood neutrophils (Figure 9.2). So in asthma and neutrophilic asthma, airway neutrophils are relatively activated compared to resting blood neutrophils, but relatively suppressed compared to airway neutrophils from healthy people (Figure 9.1). The level of IL-8 produced from airway neutrophils in neutrophilic asthma was similar to that produced by LPS activated peripheral blood neutrophils cultured at the same cell concentration.
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150
100
50 IL-8 Fold Change
0 LPS NA Asthma Healthy
Blood neutrophils Airway neutrophils Figure 9.1 Relative cytokine production (IL-8 protein expressed as average fold change from resting blood neutrophils, 1 x 105 cells/mL) IL-8 production from LPS stimulated blood neutrophils is similar to the spontaneous IL-8 production from airway neutrophils in neutrophilic asthma. IL-8 production from airway neutrophils is less in asthma, particularly neutrophilic asthma compared to healthy controls. For complete data see Chapter 4 and Chapter 7.
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There are several implications of these results. The spontaneous cytokine release from airway neutrophils in neutrophilic asthma, coupled with the increase in cell number that occurs in neutrophilic asthma can explain the marked increase in cytokines seen when whole airway samples are examined [255]. The relative suppression of cytokine release from individual airway neutrophils in neutrophilic asthma compared to healthy controls suggests that there are adaptive mechanisms operating in neutrophilic asthma in an effort to control the inflammatory response.
9.3 Distinct differences between airway and blood neutrophils
The analysis of innate immune activation of airway neutrophils revealed an inflammatory profile that was different to that of blood neutrophils in a number of ways. Firstly, the amount of spontaneously released proinflammatory cytokines and expression of TLR2 was significantly higher than that of blood neutrophils. Secondly, addition of LPS failed to further enhance the secretion of proinflammatory cytokines and total MMP-9 or the expression of TLR2 or TLR4 in airway neutrophils. Thirdly, airway neutrophils release only low levels of total MMP-9 compared to blood neutrophils and do not release OSM, another granule associated neutrophil product. This suggests that upon migration of neutrophils to the airways, the contents of gelatinase granules is released, and therefore airway neutrophils are deficient of further release from gelatinase granules. These differences in the innate immune response of airway and circulating neutrophils were not specific to the disease group, as they also occurred in the population of healthy volunteers. Clearly the responsiveness of neutrophils is different in the airway compared to the circulation, and it is likely the local cellular environment modifies the functional properties of neutrophils.
There is now growing evidence that neutrophils display distinct functional capacities that depend on their local environment. Pang et al [340] investigated the production of IL-8, IL- 1 and TNF-α by sputum neutrophils in chronic bronchial sepsis. In these subjects the spontaneous production of cytokines from sputum neutrophils was higher compared to blood neutrophils, and sputum neutrophils were not responsive to LPS stimulation. Corvol et al [341] examined IL-8 and IL-1-receptor antagonist (IL-1ra) release from sputum and
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blood neutrophils isolated from subjects with cystic fibrosis (CF). Similarly, they showed increased spontaneous cytokine production of sputum neutrophils compared to blood, with LPS failing to further enhance this cytokine production. Therefore irrespective of the presence of airway disease there are clear and distinct differences in the production of cytokines and responsiveness to LPS stimulation in the airways compared to circulating neutrophils. These data support the view that there are specific mechanisms that regulate the activation of neutrophils in the airways and that these are different compared to the systemic circulation.
In vitro studies of the effects of dexamethasone on sputum neutrophils obtained from CF patients showed a difference in the responsiveness of airway neutrophils compared to blood neutrophils [341]. When used at concentrations that were sufficient to suppress IL-8 production from peripheral blood neutrophils, dexamethasone was unable to suppress the production of IL-8 from airway neutrophils. Higher concentrations of dexamethasone were needed to obtain the same effect.
9.4 Endotoxin Tolerance
The inability of sputum cells to respond to stimulation appears to be specific for LPS as sputum cells have been reported to respond to PHA [337] and fMLP [339] stimulation in vitro. The inability of airway neutrophils to respond to LPS described in this thesis and other studies resembles the in vitro phenomenon of endotoxin tolerance, whereby cells are refractory to secondary LPS exposure [342]. Studies of endotoxin tolerance of monocytes have determined that tolerance results in a decrease in the activation of transcription factors [332], decreased release of some cytokines [343], and downregulation of the TLR4 pathway [344].
Endotoxin tolerance has been demonstrated to induce important changes in the neutrophil phenotype [345]. Intracellular signalling pathways and the respiratory burst are affected by endotoxin tolerance of isolated blood neutrophils. In this study by Parker et al, initial LPS exposure resulted in the phosphorylation of MAPKs and the activation of NF-κB, however the secondary LPS dose induced no such effect. Neutrophils that are tolerant retained the
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ability to respond to survival factors with a delay in apoptosis, and the cells continued to release IL-8, the level of IL-8 released was maintained from the initial LPS exposure but not further increased upon additional exposure [345]. It appears that a single activating stimulus is all that is needed to transform the neutrophil into a pro-inflammatory cytokine producing cell, and this cytokine production persists even when the inflammatory stimulus is withdrawn.
Tolerance to endotoxin is associated with a loss of TLR4 expression on the neutrophils cell surface in vitro [345]. Inhalation of endotoxin by healthy volunteers also decreases the expression of TLR4 on alveolar macrophages [377]. We have observed a decreased gene expression of TLR4 in airway neutrophils in asthma, particularly neutrophilic asthma and COPD. Neutrophilic asthma has been associated with increased levels of endotoxin and increased frequency of bacterial infection, particularly with Haemophilis Influenzae [255]. Bacterial airway infections e.g. H.influenzae are also commonly associated with COPD, during stable disease as well as exacerbations [417]. The effect of continual exposure to endotoxin and microbial products on neutrophil function in airway disease is not known. Our data suggests that this continual exposure results in a downregulation of innate immune responses of neutrophils, however further investigations are warranted. The combination of systemically activated neutrophils with airway LPS exposure could induce the phenotype of a spontaneously activated, endotoxin tolerant cell that was seen in this study.
9.5 Impact of Ageing
Subjects with neutrophilic asthma are often older than subjects from the other asthma inflammatory phenotypes [9]. Sputum neutrophilia is generally accepted as a feature of COPD, which also occurs in an ageing population. Our study did not detect differences in sputum neutrophil percentages between subjects with COPD and healthy controls. This is most likely due to the low number of patients studied. However we took care to match our healthy control subjects with respect to age since it has been reported that sputum neutrophil proportions are correlated with age in healthy non-smoking control subjects [418]. In addition, the effect of age on neutrophil activation was investigated.
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The neutrophil phenotype in relation to increasing age is characterised by an enhanced spontaneous activation of both circulating and airway neutrophils, and an impaired response of circulating neutrophils to LPS. The enhanced spontaneous activation may represent a low grade underlying inflammation that exists at the airway mucosal surface as well as systemically, which occurs during healthy ageing. An impaired reponse to bacterial stimuli such as LPS may predispose older people to more severe and prolonged infections that may lead to chronic inflammation and airway remodeling. This is likely to greatly impact on subjects with underlying respiratory conditions such as asthma and COPD. Whilst evidence is accumulating that reveals changes in neutrophil function with ageing, there is a need for more research into how these alterations affect the ageing population of subjects that have underlying respiratory conditions. Investigating the mechanisms of the age related changes in neutrophil function are also needed.
9.6 Innate Immune Dysfunction in Airway Disease
This thesis has described downregulation in the innate immune activity of airway neutrophils that occurs in airway disease, which is most pronounced in neutrophilic asthma and COPD. There was a relationship between cytokine release and lung function, in that reduced lung function was associated with lower levels of pro-inflammatory cytokine release. Mechanisms of this dysfunction of the innate immune system involving neutrophils in asthma have not yet been described. Since neutrophils are important effector cells in innate immunity to microorganisms including bacteria, any dysfunction of these cells may impact on susceptibility to and severity of infections.
Our findings of reduced neutrophil cytokine release and TLR expression in asthma and COPD is in agreement with a number of emerging studies that have reported a reduction in neutrophil function in airway disease, including a decreased neutrophil chemotaxis in COPD and chronic bronchitis [419, 420], and reduced phagocytosis and killing of Candida albicans in chronic bronchitis [419]. Our data also adds to growing evidence that in neutrophilic airway disease there is a discrepancy between individual cell activation and the
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cumulative effects of increased numbers of airway neutrophils. The global increase in neutrophil burden leads to over all increase in cytokine levels, however this is not necessarily associated with more inflammatory cell activity in the airways. There have now been a number of observations of reduced activity of inflammatory cells in airway disease. These include a decreased TNF-α production from sputum cells in COPD [369], TLR2 expression on alveolar macrophages from subjects with COPD [421], and pro- inflammatory cytokine production including TNF-α and IL-8 from bronchial epithelial cells from COPD patients [422]. The results of this thesis therefore need to be interpreted from the perspective of neutrophil accumulation in the airways. Then the impact of the changes in cellular activation can be related to other observations.
9.7 Accumulation of Neutrophils in the Airways in Neutrophilic Asthma
This thesis has shown that neutrophils have a decreased innate immune activation in neutrophilic asthma and COPD. Even though there is a relative suppression of the innate immune pro-inflammatory responses of airway neutrophils in neutrophilic asthma there is still a massive increase in the numbers of neutrophils, which means there is ongoing neutrophil recruitment leading to their accumulation in the airways. Accumulation of neutrophils in the airways in neutrophilic asthma is associated with high concentrations of neutrophil derived mediators including IL-8. The mechanisms by which neutrophils accumulate in the airways in neutrophilic asthma are unknown, however it is likely to involve either enhanced neutrophil migration to the airways, or delayed clearance of neutrophils. Delayed clearance of neutrophils could be due to either delayed apoptosis or impaired efferocytosis, which may lead to secondary necrosis and tissue damage.
9.7.1 Increased Migration
Very little is known regarding the mechanisms of neutrophil accumulation in neutrophilic asthma, however increased migration of neutrophils from the blood to the airways may play a role. Gene expression analysis of peripheral blood neutrophils in neutrophilic and eosinophilic asthma in this study demonstrated that there is upregulation of genes relating
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to cell motility and inhibition of apoptosis in neutrophilic asthma. Among those cell motility genes were IL-8 and S100A8, which are involved in neutrophil migration. This suggests that programming of neutrophils to migrate to the airways begins in the circulation. Feedback from possible leakage of inflammatory mediators (e.g. IL-8) from the airways back to the blood could influence the activity of blood neutrophils and act to prime these cells for increased migration, survival, and further release from the bone marrow [98].
Peripheral blood neutrophils isolated from subjects with non-eosinophilic asthma also released significantly more IL-8 protein at rest. The combination of these two sources of IL-8 would have the ability to perpetuate an ongoing cycle of neutrophil production and release from the bone marrow, priming and migration to the airways (Figure 9.2).
Enhanced Neutrophil Chemotaxis and Survival
BLOOD IL-8 AIRWAYS
? Leakage of Inflammatory Mediators ? Effects on the Bone Marrow Figure 9.2 Cycle of neutrophilic airway inflammation in neutrophilic asthma
Several studies have described an increased expression of adhesion molecules on the cell surface of peripheral blood neutrophils in subjects with COPD, which would facilitate their migration to the airways [77, 78]. The expression of Mac-1 was differentially expressed depending on the clinical condition of the patient, in that there was increased cell surface expression during stable disease, but decreased cell surface expressed during exacerbations [77]. These results, along with the data from this thesis suggest that systemic alterations in
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neutrophil activation, including the presence of adhesion molecules, the release of IL-8, and alterations in gene expression, can directly influence airway inflammation.
Other work on neutrophil chemotactic responsiveness and chemotactic activity of airway secretions in COPD gives mixed results. Yoshikawa and colleagues [420] have just reported reduced chemotactic activity of blood neutrophils from subjects with moderate to severe COPD in response to fMLP and IL-8 when compared to blood neutrophils from healthy nonsmokers and subjects with mild COPD. Increased neutrophil migration to fMLP has been reported in chronic bronchitis and emphysema [423]. However, future work is needed to investigate neutrophil chemotaxis in neutrophilic asthma.
9.7.2 Delayed Apoptosis
There is significant evidence to suggest that removal of neutrophils from sites of infection by apoptosis is essential for the regulation of the inflammatory response. Dysregulation of neutrophil apoptosis by bacterial infections and secreted products may influence disease processes, as premature apoptosis potentially results in failure of host defence [424], and delay of apoptosis results in neutrophil cell death via secondary necrosis and leads to damage to host tissue [425]. In fact, H.influenzae, a relevant respiratory bacterial pathogen in neutrophilic airway disease, induces neutrophil necrosis with the infiltrating neutrophils failing to kill the bacteria, rather releasing their damaging contents into the lung [417].
Importantly recent studies have linked delayed apoptosis with accumulation of neutrophils in inflammatory diseases [426]. The increased neutrophil survival was associated with reduced levels of Bax, which is a pro-apoptotic member of the Bcl-2 family. This Bax deficiency was induced by cytokines that are present at sites of neutrophilic inflammation, including G-CSF and GM-CSF [426]. Neutrophil apoptosis can be inhibited by a variety of mediators and environmental factors; this functioning to extend the time that the cell is able to exert it’s antimicrobial effect. There are a number of factors including high airway IL-8, endotoxin and inhaled corticosteroid treatment that are associated with neutrophilic asthma that would prolong survival of neutrophils in the airways.
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The role of neutrophil apoptosis in airway disease remains unclear. During acute exacerbations of COPD, peripheral blood neutrophil apoptosis is significantly suppressed. This defect reverses progressively after treatment and returns to normal healthy values on recovery [427]. However, during stable COPD there is no difference in the proportion of apoptotic neutrophils in sputum, or the in vitro survival of peripheral blood neutrophils cultured with sputum supernatant from COPD patients [428], or the rate of spontaneous apoptosis of peripheral blood neutrophils [79]. Apoptotic peripheral blood neutrophils from subjects with COPD have an increased expression of adhesion molecules compared to those from healthy controls, which may represent increased activation of these cells during apoptosis [79]. Bronchiectasis is associated with a lower percentage of neutrophils undergoing apoptosis, which was thought to be associated with the inhibition of apoptosis by inflammatory mediators [429]. In subjects with allergic asthma, the presence of IgE delayed spontaneous neutrophil apoptosis in vitro [430].
TLR4 and TLR2 play a significant role in the regulation of neutrophil apoptosis. Whilst activation of TLR4 by LPS delays apoptosis, activation of TLR2 in monocytes can result in the activation of both pro-survival and pro-death pathways through the activation of caspases [325]. Increased TLR2 expression has been associated with increased rate of spontaneous neutrophil apoptosis [386]. Our data shows a reduced expression of TLR2 in peripheral blood and airway neutrophils that is associated with airway disease. This may implicate modulation of neutrophil apoptosis, however further investigations are needed.
9.7.3 Impaired Efferocytosis
Removal of apoptotic cells occurs by a form of phagocytosis termed efferocytosis. This process can be carried out by structural cells, such as fibroblasts, epithelial cells and endothelial cells, as well as by “professional phagocytes”, such as macrophages and dendritic cells. Efferocytosis is essential in the resolution of inflammation as it results in the physical removal of apoptotic neutrophils before membrane permeability occurs, which prevents the release of harmful inflammatory mediators, proteases and oxidants. In order for apoptotic cells to be recognised, changes in the cell membrane occur that distinguish
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them from viable cells. A key element of this recognition is the externalisation of phosphatidylserine (PS), which is normally located on the inner membrane leaflet [431]. Calreticulin is also important in enhancing efferocytosis as it associates with PS on the cell surface [432].
The interaction between apoptotic cells and phagocytes also has important functions in modulating the innate immune response. Efferocytosis directly induces phagocytes to produce anti-inflammatory mediators, including TGF–β, IL-10, and PGE2, and these mediators act in an autocrine/paracrine manner to suppress the production of pro- inflammatory mediators [189, 433]. In comparison, efferocytosis of lysed neutrophils is much less effective at inducing the production of TGF-β and does not suppress the release of IL-8 or MIP-2, which can attract further neutrophils to the inflammatory site [434]. Efferocytosis may also be important in maintaining the protease/antiprotease balance, as it results in the production of secretory leukoprotease inhibitor (SLPI) an anti-protease for neutrophil elastase and cathepsin G [435].
Under normal conditions, this process occurs so efficiently that apoptotic cells are rarely seen in the non-diseased human lung. In contrast, apoptotic cells are far more prevalent in chronic inflammatory diseases of the lung including CF, non-CF bronchiectasis, COPD, emphysema and asthma [436, 437]. It is not clear whether this increased appearance is due to an enhanced induction of apoptosis or impaired efferocytosis, however increasing evidence suggests the latter. Alveolar macrophages from subjects with COPD, have an impaired ability to phagocytose apoptotic airway epithelial cells [438].
Imbalance of the protease/antiprotease ratio is now implicated as a key factor causing impaired efferocytosis in airways of subjects with CF. It was shown that airway fluid from subjects with CF or bronchietasis directly inhibited apoptotic cell removal by alveolar macrophages. This was thought to be due to cleavage of the PS receptor by neutrophil elastase [436]. One of the features of neutrophilic asthma is the high level of neutrophil elastase activity [255]. It is possible that the presence of elastase may impact on the persistence of apoptotic neutrophils in the airways in neutrophilic asthma. Further
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investigations of the rate of apoptosis and efferocytosis are warranted in neutrophilic asthma.
9.8 Complications of Reporting Neutrophil Data: Volume or Cell Number?
Neutrophilic asthma has been reported to be associated with an increase in innate immune activation [255], with increased levels of innate immune cytokines and receptors measured in induced sputum samples. These results are calculated by volume of sputum samples, but not corrected for the number of cells present in the sample. Subjects with neutrophilic asthma have increased total cell counts, which are mainly due to the high numbers of neutrophils that have accumulated in the airways. This global increase in innate immune activation markers contrasts with the results of this thesis and other published work showing reduced activation of airway inflammatory cells in neutrophilic airway diseases, and once the global sputum results are corrected for the amount of neutrophils in the sample, very different results are observed (Table 9.2). Using IL-8 as an example, there was significantly less IL-8 measured per neutrophil in neutrophilic asthma in comparison with paucigranulocytic asthma. This indicates that the high level of IL-8 measured in the sputum supernatant simply reflects the number of neutrophils that have accumulated rather than an excessive activation of these cells. This provides further evidence that neutrophil migration and accumulation in the airway are key processes contributing to neutrophilic airway diseases such as neutrophilic asthma.
NA EA PGA p N 11 16 20
Neutrophils x 104 664 (254-1328)*# 80.2 (46.0-206.1) 42.1 (33.5-116) 0.0001
IL-8 ng/mL 10.8 (7.4-28.9) 4.1 (2.8-8.3) 5.9 (2.2-11.1) 0.08 IL-8 ng/104 neutrophils 0.020 (0.013-0.049)* 0.056 (0.031-0.086) 0.078 (0.03-0.15) 0.006 Table 9.2 Level of IL-8 measured in sputum supernatant corrected for the number of neutrophils present in the sample (Reanalysed data via personal communication), *p<0.008 versus PGA, #p<0.008 versus EA
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9.9 Conclusions
The work embodied in this thesis demonstrates that neutrophils are able to influence the development and persistence of airway inflammation in asthma. LPS stimulation of neutrophils in vitro is an appropriate model to evalulate neutrophil activation and the capacity of cells to release important innate immune mediators. Whilst circulating neutrophils are responsive to and dependent on activation by LPS, airway neutrophils have very different properties. Airway neutrophils are highly activated at rest, but remain refractory to LPS stimulation. This may be due to previous LPS exposure and a level of endotoxin tolerance. Normal healthy aging has an impact on neutrophil function that is associated with increased spontaneous activation of airway and circulating neutrophils, but a reduced cytokine response of circulating neutrophils to LPS stimulation.
Neutrophil function is altered with airway disease, including asthma and COPD. Airway neutrophils have a relative impairment of cytokine responses in asthma, particularly neutrophilic asthma and COPD, which is linked with a downregulation of TLR4 and TLR2. Circulating neutrophil cytokine responses to LPS remain unchanged in asthma, but are increased in COPD. Whilst airway neutrophil responses in neutrophilic asthma are impaired, circulating neutrophils have gene expression profiles that implicate enhanced cell motililty and survival, which is likely to influence the accumulation of neutrophils in the airways. These results can help guide further research into mechanisms and treatments of neutrophilic airway diseases. Future work could usefully investigate the intracellular mechanisms of these changes, and may identify ways to reduce neutrophil accumulation. Neutrophil accumulation in the airway emerges as an important therapeutic target for patients with neutrophilic airway diseases such as neutrophilic asthma.
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