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PhD Thesis

Epithelial barrier integrity and function in paediatric asthma

Kevin Looi B.Sc. (Hons)

This thesis is presented for the Degree of Doctor of Philosophy at the University of

Western Australia, School of Paediatrics and Child Health

2015

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Declaration

DECLARATION FOR THESES CONTAINING PUBLISHED WORK AND/OR

WORK PREPARED FOR PUBLICATION

This thesis contains published work and/or work prepared for publication, some of

which has been co-authored. The bibliographical details of the work and where it

appears in the thesis are outlined below.

Signed___

Kevin Looi

Date 27 March 2015

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1. Looi, K., Sutanto, E.N., Banerjee, B., Garratt, L.W., Ling, K-M., Foo, C.J.,

Stick, S.M. & Kicic, A. (2011). Bronchial brushings for investigating airway

inflammation and remodelling. Respirology 16: 725–737.

This manuscript constitutes aspects of the Literature Review of this thesis. K Looi

(80%) was involved in manuscript preparation, literature collation, writing and editing.

EN Sutanto, B Banerjee, LW Garratt, KM Ling, CJ Foo assisted in manuscript

preparation, literature collation and editing. SM Stick and A Kicic were involved in the

initial manuscript concept and design, drafting and editing of the manuscript.

Signed Date 27 March 2015

Sunalene Devadason (Co-ordinating Supervisor)

Signed__ Date 27 March 2015

Anthony Kicic (Senior Author)

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2. Looi, K., Buckley, A.G., Rigby, P., Garratt, L.W., Iosifidis, T., Knight, D.A.,

Bosco, A., Troy, N.M., Ling, K-M., Martinovich, K.M., Kicic-Starcevich, E.,

Shaw, N.C., Sutanto, E.N., Kicic, A. & Stick, S.M. (2015). Human rhinovirus

infection of airway epithelium triggers tight junction protein disassembly

resulting in increased transepithelial permeability. (In preparation)

This manuscript constitutes Chapter 4 of this thesis. K Looi (80%) was involved in the

initial manuscript concept and experimental design, sample collection, performing

experimental assays, data analysis and manuscript writing. AG Buckley and P Rigby

provided technical expertise in confocal microscopy and image data analysis. LW

Garratt, T Iosifidis, K-M Ling, KM Martinovich, E Kicic-Starcevich, NC Shaw and EN

Sutanto assisted in manuscript preparation and editing. NM Troy and A Bosco provided

technical expertise in Ingenuity Pathway Analysis. DA Knight assisted in data

interpretation and critical revision of the manuscript. SM Stick and A Kicic were

involved in manuscript concept and design, data interpretation, drafting and critical

revision of the manuscript.

Signed_ Date 27 March 2015




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3. Looi, K., Buckley, A.G., Rigby, P., Garratt, L.W., Iosifidis, T., Knight, D.A.,

Zosky, G.R., Larcombe, A.N., Lannigan, F.J., Ling, K-M., Martinovich, K.M.,

Kicic-Starcevich, E., Shaw, N.C., Sutanto, E.N., Kicic, A. & Stick, S.M. (2015).

Effects of human rhinovirus on epithelial barrier integrity and function in

childhood asthma. (In preparation)

This manuscript constitutes Chapter 5, 6 and 7 of this thesis. K Looi (80%) was

involved in the initial manuscript concept and design, sample collection, performing

experimental assays, data analysis and manuscript writing. AG Buckley and P Rigby

provided technical expertise in confocal microscopy and image data analysis. LW

Garratt, T Iosifidis, GR Zosky, AN Larcombe, K-M Ling, KM Martinovich, NC Shaw

and EN Sutanto assisted in manuscript preparation and editing. E Kicic-Starcevich was

involved in co-ordinating patient recruitment. FJ Lannigan was the Otolaryngologist

involved in patient sample collection. DA Knight assisted in data interpretation and

manuscript preparation. SM Stick, GR Zosky, AN Larcombe and A Kicic initiated the

study. SM Stick and A Kicic were involved in manuscript concept, data interpretation,

drafting and critical revision of the manuscript.




Stephen M. Stick (Senior Author)

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4. Garratt, L.W., Sutanto, E.N., Ling, K-M., Looi, K., Iosifidis, T., Martinovich,

K.M., Shaw, N.C., Kicic-Starcevich, E., Knight, D.A., Ranganathan, S., Stick,

S.M. & Kicic, A. on behalf of AREST CF (2015) MMP activation by free NE

contributes to bronchiectasis progression in early CF. European Respirology

Journal (In Press)

This publication is not directly related to the content of this thesis or its chapters. K

Looi provided feedback and assisted in critical revision of the manuscript.



Signed__ Date 27 March 2015


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5. Garratt, L.W., Sutanto, E.N., Ling, K-M., Looi, K., Iosifidis, T., Martinovich,

K.M., Shaw, N.C., Buckley, A.G., Kicic-Starcevich, E., Lannigan, F.J., Knight,

D.A., Stick, S.M. & Kicic, A. on behalf of AREST CF (2015) Alpha 1-

antitrypsin mitigates the inhibition of primary airway epithelial cell repair by

elastase (Under Review)


Looi was involved in sample collection and processing for the study and critical

revision of the manuscript.





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6. Garratt, L.W., Sutanto, E.N., Foo, C.J., Ling, K-M., Looi, K., Kicic-Starcevich,

E., Iosifidis, T., Martinovich, K.M., Lannigan, F.J., Stick, S.M. & Kicic, A. on

behalf of AREST CF (2014) Determinants of culture success in an airway

epithelium sampling program of young children with cystic fibrosis.

Experimental Lung Research. Early Online 1-13.


Looi provided feedback and assisted in critical revision of the manuscript.





Signed___

Kevin Looi

Date 27 March 2015

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Abstract

The airway epithelium is the primary contact point with the inhaled air and provides a

physical barrier defence against injurious stimuli. This barrier is achieved and

maintained through a myriad of junctional complexes, however, of key interest, are the

tight junctional complex proteins. Tight junctions (TJ) are located at the terminal end of

the epithelial cell layer and serve to maintain structural integrity as well as regulate

transepithelial permeability. Although studies have demonstrated TJ abnormalities in

adults with asthma, few studies have addressed whether these abnormalities are intrinsic

to asthma, are consequences of chronic inflammation, or due to atopy rather than

asthma. In addition, it is unknown whether these abnormalities can be detected early in

the disease progression and are correlated with disease severity.

Evidence has also demonstrated a close association between respiratory viral infections,

in particular, human rhinoviruses (HRVs) and resulting asthma exacerbations. Although

it has been previously shown that there is disassembly of specific TJ proteins, in

particular, zonula occludens-1 (ZO-1) following HRV infection, there still remains a

paucity of data on the susceptibility of the asthmatic epithelium to TJ disassembly

following HRV infection, especially within a paediatric population. This presents

significant rationale for the re-evaluation of the concept of epithelial barrier dysfunction

in asthma as well as the effects of HRV infection on barrier integrity. Therefore, this

study was conducted to study epithelial barrier function in non-asthmatic and asthmatic

individuals to test the following hypotheses. Firstly, the epithelial barrier function is

defective in children with asthma. Secondly, a defective barrier function in asthma is

independent of atopy. Thirdly, epithelial integrity and barrier function is further

compromised by HRVs in asthmatic airways compared to non-asthmatics.

In order to address these hypotheses, this project initially adapted, designed and

optimised two methodologies for the in vitro assessment of barrier integrity. A novel,

quantitative immunofluorescence assay termed In-Cell™ Western (ICW) was used to

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quantify multiple membrane TJ proteins simultaneously from small sample sizes. Here,

different parameters including concentration of primary antibodies, length of primary

antibodies incubation and concentration of secondary antibodies were all optimised

using modified human AECs namely NuLi-1, which was subsequently corroborated in

primary AECs (pAECs). Results obtained indicated a 1:200 dilution of primary

antibodies and an incubation temperature and time of 4°C overnight, followed by

secondary antibody detection at a 1:800 dilution was the most appropriate for assessing

membrane TJ protein expression in pAECs. In addition, a transepithelial permeability

assay was also established. Here, fluorescently labelled dextran molecules of two

different molecular weights (4 and 20 kDa) were used and experiments performed to

ascertain the effects of varying sampling durations (4 and 6 hours) on absorbance

measurements and calculated permeability coefficients. Data generated reports the

successful establishment of this assay and demonstrated that absorbance values

continually increased over time and reached a maximum value at 6 h. Furthermore, the

additional sampling time period provided a more accurate measure of calculated

permeability coefficients than a shorter sampling period.

In addition, it was also essential to establish a direct correlation between HRV infection,

altered TJ expression and transepithelial permeability. This was performed using

modified healthy AECs, namely NuLi-1. However, prior to elucidating the potential

pathways and regulatory effects HRV has on epithelial TJ proteins, it was first

necessary to assess the susceptibility of NuLi-1 cells to the virus. Cells were initially

infected with HRV-1B at 50% Tissue Culture Infectivity Dose (TCID50) 2.5, 10 and 20

x 104 TCID50/ml for 24 h and cell viability, apoptosis and viral replication all

subsequently assessed. Membrane TJ protein disassembly and the resulting permeability

was assessed using previously optimised ICW and transepithelial permeability assay.

Finally, focused arrays on human TJs were performed to identify potential pathways

and regulatory effects of HRV infection on epithelial TJ proteins. Results confirmed a

typical loss of viability (p<0.05), induction of apoptosis following infection at a titre of

20x104 TCID50/ml HRV-1B for 24 h (p<0.05) and the presence of high viral RNA copy

number following HRV-1B infection for 24h (p<0.05). Focused qPCR array data

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demonstrated a down-regulation of 58 key genes with an up-regulation of 26 key genes

encoding for epithelial barrier junction proteins following infection with HRV-1B,

providing additional insights and better understanding of the molecular mechanisms

behind TJ-mediated cell biology. Membrane TJ disassembly in response to HRV-1B

infection was observed following infection at a titre of 20x104 TCID50/ml HRV-1B for

24 h. Although this was non-significant, when assessed functionally, a significant

increase in transepithelial permeability was observed (p<0.05).

To address the first hypothesis that epithelial barrier function is defective in children

with asthma, pAECs derived from non-asthmatic and asthmatic children were assessed

for the gene and protein expression of the TJs, claudin-1, occludin and ZO-1, via RT-

PCR and immunocytochemistry. Data demonstrated a significant increase in claudin-1

and occludin gene expression within the asthmatic cohorts. In contrast, ex vivo protein

expression for all three TJ was observed to be markedly decreased within the asthmatic

cohorts. Furthermore, assessment of TJ protein expression in vitro further corroborated

this decrease with significant reduction in membrane expression of all three TJ proteins

within the asthmatic cohorts. (p<0.05). Using submerged monolayer cultures, basal

transepithelial permeability was observed to be higher within the asthmatic cohorts

compared to the non-asthmatics. Collectively, these findings demonstrate that paediatric

asthmatic epithelium, despite higher TJ gene expression, has lower protein expression

and subsequently, an increase in basal transepithelial permeability compared to their

non-asthmatic counterpart. The discordance between increased TJ gene expression and

decreased TJ protein expression suggests post-transcriptional regulation or a

compensatory effect by other junctional proteins.

In order to test the second hypothesis that defective barrier function in asthma is

independent of atopy, pAECs were sub-categorised according to confirmed allergen

sensitisation, resulting in the following phenotypes; healthy non-atopic (HNA), healthy

atopic (HA), non-atopic asthmatic (NAA) and atopic asthmatic (AA) cohorts. Ex vivo

TJ gene and protein expression were assessed using RT-PCR and

immunocytochemistry. In vitro assessment of TJ protein expression and barrier integrity

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was achieved via ICW and transepithelial permeability assays. Significant differences in

ex vivo claudin-1 and occludin TJ gene expression was observed between the HNA and

HA cohorts (p<0.05). Marked differences in ex vivo TJ protein expression was also

similarly observed between the HNA and HA cohorts. Moreover, in vitro analysis

demonstrated significantly lower membrane TJ protein expression within the HA

cohorts when compared to HNA (p<0.05). However, the lower protein expression did

not translate into a significant increase in transepithelial permeability within the HA

cohorts. Due to the limited availability of paediatric derived pAECNAA samples, no

statistical analysis could be performed on this cohort.

To test the third hypothesis that barrier function and epithelial integrity is further

compromised by HRV in asthmatic compared to non-asthmatic airways, asthmatic and

non-asthmatic paediatric derived pAECs were infected with viral titres of 2.5x104

TCID50/ml and 20x104 TCID50/ml of HRV-1B for 24 and 48 h. Membrane TJ protein

expression and barrier function was assessed using the ICW and transepithelial

permeability assay respectively. Data demonstrated significant disassembly of

membrane claudin-1, occludin and ZO-1 proteins in non-asthmatic cohort following

infection with viral titres of 20x104 TCID50/ml (p<0.05) of HRV-1B for 24 h was

observed, while, significant disassembly of all 3 TJ proteins was similarly observed in

the asthmatic cohort (p<0.05). Interestingly, a restoration towards non-infected levels

was observed for all TJ membrane proteins in non-asthmatic cohort following 48 h

infection while sustained disassembly occurred in the asthmatic cohort. When barrier

integrity was assessed functionally, a significant increase in transepithelial permeability

was only observed in the non-asthmatic cohorts (p<0.05) while increased transepithelial

permeability was maintained within the asthmatic cohorts post infection. Collectively,

these findings indicate that maintenance of increased permeability within the asthmatic

cohorts could be attributed to the low basal membrane TJ protein expression and that

HRV infection did not increase transepithelial permeability significantly in an already

permeable state. When assessing the effects of atopy on TJ protein expression in

response to HRV infection, a significant decrease (p<0.05) in all 3 TJ protein

expression was observed in pAECHNA, pAECHA and pAECAA after infection with viral

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titres of 2.5 and 20x104 TCID50/ml of HRV-1B for 24 h. Interestingly, restoration of all

3 TJ protein expression towards non-infected levels was only observed in pAECHNA

while sustained decreased expression was observed in pAECHA, pAECNAA and pAECAA

cohorts. Despite this significant decrease in membrane TJ protein expression in all

cohorts, no significant differences in transepithelial permeability was observed between

submerged monolayer cultures of pAECHNA, pAECHA, pAECNAA and pAECAA.

Findings were then recapitulated in a physiologically more representative culture model

of the airway, using paediatric derived non-asthmatic and asthmatic pAECs. Cells were

cultured on inserts and once confluent, were grown at air-liquid interface (ALI) and

allowed to terminally differentiate. Transepithelial electrical resistance (TEER)

measurements and transepithelial permeability assays were performed to assess barrier

integrity and function respectively. Cells were then infected with viral titres of 10x104

TCID50/ml of HRV-1B for 24 h at 33°C, TEER and permeability to different sized inert

molecules subsequently re-assessed. The results showed lower basal membrane TJ

protein expression within the asthmatic epithelium compared to non-asthmatic

counterpart. Elevated basal transepithelial permeability, concomitant with reduced RT

values, strongly suggests that asthmatic epithelial cells have an intrinsically altered TJ

expression, hence leading to impairment of barrier functionality. Collectively, these

observations demonstrate potential relationship between atopy and asthma in altering

barrier integrity as well as the effects of HRV infection on epithelial integrity and

functionality in paediatric derived non-asthmatic and asthmatic pAEC ALI cultures.

In conclusion, this project has demonstrated an association between TJ disassembly and

the subsequent increase in transepithelial permeability. The data have also demonstrated

significantly lower membrane TJ protein and significantly higher permeability towards

small sized molecules in the asthmatic cohorts, which supports the notion that intrinsic

differences in basal TJ protein expression exist between non-asthmatic and asthmatic

epithelium. This may partly explain the increased susceptibility to aeroallergen

sensitisation and pathogenic challenges in children with asthma. Interestingly, this

project also demonstrated significant differences between non-atopic and atopic

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phenotypes within the non-asthmatic cohort. This suggests that the presence of atopy

might contribute to an increased predisposition towards membrane TJ protein

disassembly. However, the precise role of atopy, whether causative or co-contributing to

TJ disassembly, warrants further detailed analysis. Following HRV infection,

significant disassembly of all three membrane TJ protein within the non-asthmatic

cohort which is concomitant with elevated permeability may explain the increased

vulnerability of the non-asthmatic epithelium to successive aeroallergen sensitisation or

pathogenic challenges. However, despite significant disassembly of membrane occludin

within the asthmatic cohort following infection, maintenance of elevated permeability

was observed, suggesting that this intrinsically higher level of permeability could

facilitate trafficking of aeroallergens or pathogens into the sub-epithelial space,

resulting in an endless cycle of aeroallergen sensitisation, pathogenic challenges and

asthma exacerbations.

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Contents

Abstract ......................................................................................................................... viii

List of Figures .............................................................................................................. xxii

List of Tables ............................................................................................................. xxvii

List of Abbreviations ............................................................................................... xxviii

Publications arising from this project .................................................................... xxxiii

Presentations arising from this project .................................................................. xxxiv

International Conference Paper .............................................................................. xxxiv

National Conference Paper ..................................................................................... xxxv

Local Conference Paper ........................................................................................ xxxvii

Awards ...................................................................................................................... xxxix

Australian Postgraduate Award (APA) .................................................................. xxxix

Princess Margaret Hospital Foundation PhD Top-Up Scholarship ....................... xxxix

New Investigator Awards ....................................................................................... xxxix

Best Poster Prize .................................................................................................... xxxix

Travel Awards .............................................................................................................. xl

Acknowledgements ........................................................................................................ xli

CHAPTER 1: Literature Review ................................................................................... 1

1. Asthma ................................................................................................................... 1

1.2 Recent advances in the pathophysiology of asthma ........................................... 6

1.3 Respiratory epithelium ....................................................................................... 8

1.3.1 Form and function of the epithelium ........................................................... 8

1.3.2 Airway responses in the asthmatic epithelium .......................................... 14

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1.3.3 Intrinsic abnormalities of the asthmatic epithelium .................................. 17

1.4 Respiratory viruses and asthma ........................................................................ 20

1.4.1 Influenza virus ........................................................................................... 21

1.4.2 Respiratory syncytial virus (RSV) ............................................................ 23

1.4.3 Human rhinovirus ..................................................................................... 25

1.5 Assessing airway integrity ................................................................................ 29

1.5.1 Animal models .......................................................................................... 29

1.5.2 Cell culture models ................................................................................... 31

1.6 Summary .......................................................................................................... 34

1.7 Hypotheses and research aims .......................................................................... 36

CHAPTER 2: General Materials and Methods ......................................................... 37

2.1 General Materials ............................................................................................. 37

2.2 Antibodies ............................................................................................................. 40

2.2.1 Primary antibodies .................................................................................... 40

2.2.2 Secondary antibodies ................................................................................ 40

2.3 General Equipment................................................................................................ 41

2.3.1 Autoclave .................................................................................................. 41

2.3.2 Balances .................................................................................................... 41

2.3.3 Bronchoscope ............................................................................................ 41

2.3.4 Bronchial brush ......................................................................................... 41

2.3.5 Centrifuges ................................................................................................ 41

2.3.6 Glassware .................................................................................................. 42

2.3.7 Heating devices ......................................................................................... 42

2.3.8 Incubators .................................................................................................. 42

2.3.9 Infrared scanner ......................................................................................... 42

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2.3.10 Laminar flow cabinets ............................................................................... 42

2.3.11 Microscope ................................................................................................ 43

2.3.12 pH meter .................................................................................................... 43

2.3.13 Pipettes ...................................................................................................... 43

2.3.14 Plate readers .............................................................................................. 43

2.3.15 Real Time Quantitative PCR (RT-qPCR) ................................................. 44

2.3.16 Semi-dry Western Blot Transfer ............................................................... 44

2.3.17 Spectrophotometer .................................................................................... 44

2.3.18 Stirrer, shakers and rockers ....................................................................... 44

2.3.19 Tissue culture and general plastic ware .................................................... 44

2.3.20 Water bath ................................................................................................. 45

2.4 General Buffers and Solutions.......................................................................... 45

2.4.1 General purpose ........................................................................................ 45

2.4.2 Cell culture ................................................................................................ 48

2.4.3 Assays and associated solutions ................................................................ 53

2.5 General Methodology ....................................................................................... 56

2.5.1 Cell line types ............................................................................................ 56

2.5.2 Immortalised cell line culture, sub-culture and cryopreservation ............. 59

2.5.3 Ethics approval .......................................................................................... 60

2.5.4 Primary paediatric airway epithelial cells ................................................. 60

2.5.5 Plasma and buffy coat isolation ................................................................ 63

2.5.6 Cytospin preparation ................................................................................. 63

2.5.7 Human Rhinovirus .................................................................................... 63

2.5.8 Immunocytochemistry............................................................................... 65

2.5.9 Immunohistochemistry .............................................................................. 66

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2.5.10 In-Cell™ Western ...................................................................................... 67

2.5.11 Transepithelial permeability...................................................................... 67

2.5.12 Transepithelial electrical resistance (TEER) ............................................. 68

2.5.13 Reverse Transcriptase-Polymerase Chain Reaction (RT-PCR) and Real

Time quantitative Polymerase Chain Reaction (RT-qPCR) ................................... 69

2.5.14 Total cellular protein extraction ................................................................ 69

2.5.15 Total cellular protein quantification .......................................................... 70

2.5.16 Western Blot.............................................................................................. 70

2.5.17 Statistical analysis ..................................................................................... 71

CHAPTER 3: Optimisation of In Cell™ Western and Transepithelial permeability

assays .............................................................................................................................. 72

3.1 Introduction ...................................................................................................... 72

3.2 In Cell™ Western assay .................................................................................... 75

3.2.1 Materials .................................................................................................... 75

3.2.2 Methods ..................................................................................................... 76

3.2.3 Results / Discussion .................................................................................. 77

3.2.4 Conclusion ................................................................................................ 82

3.3 Transepithelial permeability assay ................................................................... 83

3.3.1 Materials .................................................................................................... 83

3.3.2 Methods ..................................................................................................... 84

3.3.3 Results / Discussion .................................................................................. 85

3.3.4 Conclusion ................................................................................................ 87

CHAPTER 4: Effect of human rhinovirus infection on tight junction disassembly

and the subsequent changes to barrier function ........................................................ 88

4.1 Introduction ...................................................................................................... 88

4.2 Materials and Methods ..................................................................................... 89

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4.2.1 Cell culture ................................................................................................ 89

4.2.2 Human rhinovirus and titrations................................................................ 89

4.2.3 Human tight junction Polymerase Chain Reaction (PCR) arrays ............. 89

4.2.4 Infection of cell cultures............................................................................ 90

4.2.5 In Cell™ Western assay ............................................................................. 90

4.2.6 MTS cell viability assay ............................................................................ 90

4.2.7 Quantification of Human Rhinovirus viral copy number ......................... 91

4.2.8 Real-Time Quantitative Polymerase Chain Reaction (RT-qPCR) ............ 91

4.2.9 Single-stranded DNA (ssDNA) apoptosis assay ....................................... 91

4.2.10 Transepithelial permeability assay ............................................................ 92

4.2.11 Statistical analysis ..................................................................................... 93

4.3 Results .............................................................................................................. 93

4.3.1 Effect of human rhinoviral infection on NuLi-1 cell viability .................. 93

4.3.2 Apoptotic response and viral replication following infection with HRV-1B

94

4.3.3 Effect of HRV-1B infection on mRNA expression of tight junction

complexes ................................................................................................................ 95

4.3.4 Effect of human rhinovirus infection on membrane tight junction

disassembly ............................................................................................................. 95

4.3.5 Effect of human rhinovirus infection on transepithelial permeability ...... 96

4.4 Discussion ........................................................................................................ 97

4.5 Conclusion ........................................................................................................ 99

CHAPTER 5: Epithelial barrier integrity and function in paediatric asthma ..... 101

5.1 Introduction .................................................................................................... 101

5.2 Materials and Methods ................................................................................... 103

5.2.1 Patient Demographics ............................................................................. 103

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5.2.2 Cell culture .............................................................................................. 103

5.2.3 In Cell™ Western ..................................................................................... 103

5.2.4 Immunocytochemistry............................................................................. 103

5.2.5 Real-Time Quantitative Polymerase Chain Reaction (RT-qPCR) .......... 104

5.2.6 Transepithelial permeability assay .......................................................... 104

5.2.7 Statistical analysis ................................................................................... 104

5.3 Results ............................................................................................................ 104

5.3.1 Basal tight junction gene expression ....................................................... 104

5.3.2 Basal tight junction protein expression ................................................... 106

5.3.3 In vitro tight junction protein expression ................................................ 108

5.3.4 In vitro transepithelial permeability ........................................................ 110

5.4 Discussion ...................................................................................................... 112

5.5 Conclusion ...................................................................................................... 117

CHAPTER 6: Effects of human rhinovirus on epithelial barrier integrity and

function in vitro and its role in paediatric asthma ................................................... 118

6.1 Introduction .................................................................................................... 118



6.2.2 Cell culture .............................................................................................. 120

6.2.3 Human rhinovirus and titrations.............................................................. 120

6.2.4 Infection of cell cultures.......................................................................... 120

6.2.5 In Cell™ Western ..................................................................................... 120



6.3 Results ............................................................................................................ 121

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6.3.1 Effect of human rhinovirus infection on membrane tight junction protein

expression after 24 and 48 h.................................................................................. 121

6.3.2 Effect of human rhinovirus infection on in vitro transepithelial

permeability........................................................................................................... 134

6.4 Discussion ...................................................................................................... 136

6.5 Conclusion ...................................................................................................... 141

CHAPTER 7: Effects of human rhinovirus on epithelial barrier integrity and

function in well-differentiated air-liquid interface cultures ................................... 142

7.1 Introduction .................................................................................................... 142



7.2.2 Cell culture .............................................................................................. 144

7.2.3 Establishment of air-liquid interface (ALI) cultures ............................... 144

7.2.4 Immunohistochemistry for visualisation of ciliated and goblet cells...... 144

7.2.5 Immunofluorescence and confocal microscopy ...................................... 145

7.2.6 Stereological analysis and quantification of tight junction expression ... 145

7.2.7 Human rhinovirus and titrations.............................................................. 146

7.2.8 Infection of cell cultures.......................................................................... 146

7.2.9 Transepithelial electrical resistance measurement .................................. 146



7.3 Results ............................................................................................................ 147

7.3.1 Establishment of air-liquid interface cultures ......................................... 147

7.3.2 Physical properties of pAEC derived ALI cell cultures of non-asthmatic

and asthmatic cohorts ............................................................................................ 148

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expression in pAEC derived well differentiated air-liquid interface (ALI) cultures

149

7.3.4 Effect of human rhinovirus infection on transepithelial electrical resistance

(TEER) and permeability in pAEC derived well differentiated air-liquid interface

(ALI) cultures ........................................................................................................ 152

7.4 Discussion ...................................................................................................... 155

7.5 Conclusion ...................................................................................................... 159

Chapter 8: General Discussion .................................................................................. 160

References .................................................................................................................... 169

Appendix A .................................................................................................................. 190

Appendix B .................................................................................................................. 192

Appendix C .................................................................................................................. 194

Appendix D .................................................................................................................. 196

Appendix E .................................................................................................................. 197

Appendix F ................................................................................................................... 198

Appendix G .................................................................................................................. 199

Appendix H .................................................................................................................. 200

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List of Figures

Chapter 1: Literature Review

Fig 1.1: Schematic diagram of the factors contributing to the heterogeneity and

complexity in asthma pathogenesis and diagnosis.

Fig 1.2: Schematic of major cell types lining the respiratory airways.

Fig 1.3: Illustration of the complexity between protein-protein interactions at the tight

junction complex.

Fig 1.4: Schematic of air-liquid interface culture process.

Chapter 3: Optimisation of In-Cell™ Western and Transepithelial permeability

assay

Fig 3.1: Effect of primary antibody concentration on NuLi-1 TJ signal intensity.

Fig 3.2: Effect of primary antibody concentration on paediatric derived pAECHNA

TJ signal intensity.

Fig 3.3: Effect of incubation temperature of primary antibody at 25°C on NuLi-1 TJ

signal intensity.

Fig 3.4: Effect of incubation temperature of primary antibody at 4°C on NuLi-1 TJ

signal intensity.

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Fig 3.5: Effect of incubation temperature of primary antibody at 25°C on paediatric

derived pAECHNA TJ signal intensity.

Fig 3.6: Effect of incubation temperature of primary antibody at 4°C on paediatric

derived pAECHNA TJ signal intensity.

Fig 3.7: Effect of secondary antibody concentration on NuLi-1 TJ signal intensity.

Fig 3.8: Effect of secondary antibody concentration on paediatric derived pAECHNA

TJ signal intensity.

Fig 3.9: Methodology of In-Cell™ Western (ICW) assay

Fig 3.10: Effect of sampling time on FITC-dextran molecules across cell

monolayers.

Fig 3.11: Methodology of Transepithelial permeability assay

Chapter 4: Effect of human rhinovirus infection on tight junction disassembly and

the subsequent changes to barrier function

Fig 4.1: Effect of HRV-1B on cellular viability in NuLi-1 over time.

Fig 4.2: Effect of HRV-1B on apoptosis and viral replication in NuLi-1.

Fig 4.3: Effect of HRV-1B on mRNA expression of TJ in NuLi-1.

Fig 4.4: Effect of HRV-1B infection on membrane TJ protein expression in NuLi-1.

Fig 4.5: Effect of HRV-1B infection on transepithelial permeability in NuLi-1.

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Chapter 5: Epithelial barrier integrity and function in paediatric asthma

Fig 5.1: Ex vivo mRNA expression of TJs from pAECs of non-asthmatic and asthmatic

cohorts.

Fig 5.2: Ex vivo mRNA expression of TJs in pAECs of non-asthmatic and asthmatic

cohorts with each cohort further categorised based on atopy.

Fig 5.3: Ex vivo membrane protein expression of TJs from pAECs of non-asthmatic and

asthmatic cohorts.

Fig 5.4: Ex vivo membrane protein expression of TJs of pAECs from non-asthmatic and

asthmatic cohorts with each cohort further categorised based on atopy.

Fig 5.5: Basal membrane TJ protein expression in pAECs from non-asthmatic and

asthmatic cohorts.

Fig 5.6: Basal membrane TJ protein expression in pAECs from non-asthmatic and

asthmatic cohorts with each cohort further categorised based on atopy.

Fig 5.7: Basal transepithelial permeability in pAECs from non-asthmatic and asthmatic

cohorts and with each cohort further categorised based on atopy.

Chapter 6: Effects of human rhinovirus on epithelial barrier integrity and function

in vitro and its role in paediatric asthma

Fig 6.1: Membrane TJ protein expression over time in pAECs from non-asthmatic and

asthmatic cohorts following viral infection.

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Fig 6.2: Membrane TJ protein expression over time in pAECs from non-asthmatic and

asthmatic cohorts following viral infection with each cohort further categorised based

on atopy.

Fig 6.3: Transepithelial permeability in pAECs from non-asthmatic and asthmatic

cohorts following viral infection.

Fig 6.4: Transepithelial permeability in pAECs from non-asthmatic and asthmatic

cohorts following viral infection with each cohort further categorised based on atopy.


in well-differentiated air-liquid interface cultures

Fig 7.1: Generation of ALI cultures from pAECs of non-asthmatic and asthmatic

cohorts.

Fig 7.2: Generation of ALI cultures from pAECs of non-asthmatic and asthmatic

cohorts with each cohort further categorised based on atopy.

Fig 7.3: Membrane TJ protein expression in ALI cultures generated from pAECs of

non-asthmatic cohorts following viral infection.

Fig 7.4: Membrane TJ protein expression in ALI cultures generated from pAECs of

asthmatic cohorts following viral infection.

Fig 7.5: Expression of membrane TJ protein in ALI cultures from pAECs of non-

asthmatic and asthmatic cohorts following viral infection.

Fig 7.6: Membrane TJ protein expression in ALI cultures generated from pAECHNA


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Fig 7.7: Membrane TJ protein expression in ALI cultures generated from pAECHA


Fig 7.8: Membrane TJ protein expression in ALI cultures generated from pAECAA


Fig 7.9: Expression of membrane TJ protein in ALI cultures from pAECs of non-

asthmatic and asthmatic cohorts with each cohort further categorised based on atopy.

Fig 7.10: Transepithelial electrical resistance (RT) and permeability in ALI cultures

from pAECs of non-asthmatic and asthmatic cohorts.

Fig 7.11: Transepithelial electrical resistance (RT) and permeability in ALI cultures

from pAECs of non-asthmatic and asthmatic cohorts with each cohort further

categorised based on atopy.

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List of Tables

Chapter 1: Literature Review

Table 1: Asthma susceptibility genes with the associated functions and pathway.

Chapter 2: General Materials and Methods

Table 2: Radioallergosorbent test (RAST) to a panel of common allergens for

determination of atopy.

Chapter 5: Epithelial barrier integrity and function in paediatric asthma

Table 5.1: Demographic of patient cohort categorised according to atopy.


in vitro and its role in paediatric asthma



in well-differentiated air-liquid interface cultures


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List of Abbreviations

°C degrees Celsius

AA atopic asthmatic

AEC airway epithelial cell

AHR airway hyper-responsiveness

AJ adherens junction

ALI air liquid interface

ASL airway surface liquid

ASM airway smooth muscle

ATCC American type culture collection

ATS American Thoracic Society

AU Arbitrary Units

BAL bronchoalveolar lavage

BCA bicinchoninic acid

BEBM bronchial epithelial basal medium

BPE bovine pituitary extract

BSA bovine serum albumin

CaCl2 calcium chloride

CEB cell protein extraction buffer

CF cystic fibrosis

cDNA complementary deoxyribonucleotide acid

CLDN claudin

cm2 squared centimetre

CO2 carbon dioxide

DAPI 4′,6-diamidino-2-phenylindole

DC dendritic cell

ddH2O double deionised water

DMEM dulbecco’s modified eagle medium

DMSO dimethyl sulfoxide

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DNA deoxyribonucleotide acid

dNTP deoxyribonucleotide triphosphates

ECACC European collection of cell cultures

ECM extracellular matrix

EDTA ethylenediamine tetraacetic acid

EGF epidermal growth factor

EMEM earl’s modified essential medium

FCS foetal calf serum

FEV1 forced expiratory volume in 1 second

FITC fluorescein isothiocynate

g grams

g gravitational force

GWAS genome wide association studies

h hour

HA healthy atopic

HBSS hank’s balanced salt solution

HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid

HCl hydrochloric acid

HNA healthy non-atopic

HRV human rhinovirus

ICC immunocytochemistry

ICW In Cell™ Western

Ig immunoglobulin

IHC immunohistochemistry

I.I integrated intensity

IL interlukin

iNOS inducible nitric oxide synthase

IQR interquartile range

ISAAC International Study of Asthma and Allergies in Childhood

JAMs junctional adhesion molecules

KCl potassium chloride

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kDa kilodaltons

KH2PO4 potassium dihydrogen orthophosphate

l litre

LDS lithium dodecyl sulphate

M molar

MEM minimum essential media

MES-SDS 2-(N-morpholino)ethanesulfonic acid sodium dodecyl sulphate

mg milligram

MgCl2 magnesium chloride

MHC major histocompatibility complex

min minutes

ml millilitres

mM millimolar

MOI multiplicity of infection

mRNA messenger ribonucleic acid

MW molecular weight

NAA non-atopic asthmatic

NaCl sodium chloride

Na2CO3 sodium carbonate

Na2HPO4 sodium phosphate dibasic

NaF sodium fluoride

NaHCO3 sodium bicarbonate

NaH2PO4 sodium dihydrogen orthophosphate

NaOH sodium hydroxide

Na3VO4 sodium orthovanadate

NBF neutral buffered formalin

ng nanogram

nm nanometre

OCLN occludin

pAEC paediatric-derived primary airway epithelial cells

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pAECAA atopic asthmatic paediatric-derived primary airway epithelial

cells

pAECHA healthy atopic paediatric-derived primary airway epithelial cells

pAECHNA healthy non-atopic paediatric-derived primary airway epithelial

cells

pAECNAA non-atopic asthmatic paediatric-derived primary airway epithelial

cells

PAGE polyacrylamide gel electrophoresis

Papp Apparent permeability coefficient

PBS phosphate buffered saline

PCR polymerase chain reaction

pH -log [H+]

PPIA peptidylprolyl isomerase A

PVDF polyvinylidene fluoride

qPCR quantitative polymerase chain reaction

RANTES regulated upon activation, normal T-cell expressed and secreted

RAST radioallergosorbent test

RLT ribonucleic acid lysis buffer

RNA ribonucleic acid

rpm revolutions per minute

RPMI Roswell Park Memorial Institute

RSV respiratory syncytial virus

RT room temperature

RT-PCR reverse transcriptase-polymerase chain reaction

RT-qPCR real time quantitative polymerase chain reaction

SD standard deviation

SDS sodium dodecyl sulfate

SE standard error

SEM standard error of mean

TBS tris buffered saline

TCID50 50% tissue culture infective dose

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TEER transepithelial electrical resistance

TH T helper cell

TJ tight junction

UV ultraviolet

v/v volume per volume

w/v weight per volume

ZO-1 zonula occludens-1

µg microgram

µl microlitre

µM micromolar

% percent

Ω ohms

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Publications arising from this project

1. Looi K., Sutanto E.N., Banerjee B., Garratt L.W., Ling K-M., Foo C.J., Stick

S.M. & Kicic A. (2011). Bronchial brushings for investigating airway

inflammation and remodelling. Respirology 16: 725–737.

2. Looi K., Buckley A.G., Rigby P., Garratt L.W., Iosifidis T., Knight, D.A.,

Bosco A., Troy N.M., Ling K-M., Martinovich K.M., Kicic-Starcevich E., Shaw

N.C., Sutanto E.N., Kicic A. & Stick S. M. (2015). Human rhinovirus infection

of airway epithelium triggers tight junction protein disassembly resulting in

increased transepithelial permeability. (In preparation)

3. Looi K., Buckley A.G., Rigby P., Garratt L.W., Iosifidis T., Knight D.A., Zosky

G.R., Larcombe A.N., Lannigan F.J., Ling K-M., Martinovich K.M., Kicic-

Starcevich E., Shaw N.C., Sutanto E.N., Kicic A. & Stick S. M. (2015). Effects

of human rhinovirus on epithelial barrier integrity and function in childhood

asthma. (In preparation)

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Presentations arising from this project

International Conference Paper

Oral Presentation

Looi K., Larcombe A.N., Zosky G.R, Rigby P., Knight D.A., Stick S.M., Kicic

A. (2013). Human rhinovirus infection of asthmatic airway epithelial cells

causes tight junction disassembly resulting in increased permeability.

Respirology 18(4): 16

Full oral presentation at the 18th Congress of the Asian Pacific Society of Respirology

(APSR), Yokohama, Japan (2013).

Poster Presentation

1. Looi K., Buckley A.G., Rigby P., Garratt L.W., Iosifidis T., Lannigan F.J.,

Knight D.A., Zosky G.R., Larcombe A.N., Ling K-M., Martinovich K.M.,

Kicic-Starcevich E., Shaw N.C., Sutanto E.N., Kicic A. & Stick S.M. (2015).

Intrinsic differences of epithelial tight junction in asthmatic airway epithelium

and barrier compromisation following human rhinoviral insult.

Poster presentation at the American Thoracic Society (ATS) annual scientific meeting,

Denver, USA (2015).

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2. Looi K., Ling K-M., Sutanto E.N., Larcombe A.N., Foong R., Knight D.A.,

Rigby P., Stick S.M., Kicic A. (2011). Barrier integrity compromisation as an

intrinsically abnormal process in asthmatic epithelium independent of atopy. Am

J Respir Crit Care Med 183: A2062

Poster presentation at the American Thoracic Society (ATS) annual scientific meeting,

Denver, USA (2011).

National Conference Paper

Oral Presentation

Looi K., Garratt L.W., Iosifidis T., Lannigan F.J., Ling K-M., Martinovich

K.M., Kicic-Starcevich E., Sutanto E.N., Kicic A. & Stick S.M. (2014). Tight

junction disassembly following human rhinovirus infection results in airway

epithelial permeability changes. Respirology 19 (S2): 56

Oral presentation at Thoracic Society of Australia and New Zealand (TSANZ) Annual

Scientific Meeting, Adelaide, Australia (2014).

Looi 2015

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Poster Presentation

1. Looi K., Buckley, A.G., Rigby P, Garratt L.W., Iosifidis T, Lannigan F.J.,

Knight D.A., Zosky G.R., Larcombe A.N., Ling K-M, Martinovich K.M., Kicic-

Starcevich E, Shaw N.C., Sutanto E.N., Stick S.M. & Kicic A (2015)

Disassembly of epithelial tight junctions in well-differentiated air-liquid

interface (ALI) cultures following human rhinovirus infection results in airway

epithelial permeability changes.

Poster presentation at Thoracic Society of Australia and New Zealand (TSANZ)

Annual Scientific Meeting, Gold Coast, Australia (2015).

2. Looi K., Larcombe A.N., Zosky G.R., Rigby P, Knight D.A., Stick S.M. &

Kicic A (2013) Human rhinovirus infection initiates airway epithelial tight

junction disassembly resulting in barrier function disruption. Respirology 18

(S2): 49


Annual Scientific Meeting, Darwin, Australia (2013).

3. Looi K., Ling K-M, Sutanto E.N., Larcombe A.N., Foong R. Rigby P, Stick

S.M. & Kicic A (2011) Airway epithelium barrier integrity compromisation

following rhinoviral insult. Respirology 16 (S1): 47


Annual Scientific Meeting, Perth, Australia (2011).

Looi 2015

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Local Conference Paper

Oral Presentation

1. Looi K., Larcombe A.N., Zosky G.R., Rigby P, Knight D.A., Stick S.M. &

Kicic A (2013) Disassembly of asthmatic airway epithelial cells tight junctions

following rhinoviral insult results in increased permeability.

Oral presentation at the Child and Adolescent Health Service Research and Advances

Seminar, Princess Margaret Hospital, Perth, WA (2013)

2. Looi K., Buckley A.G., Rigby P, Knight D.A., Zosky G.R., Larcombe A.N.,

Stick S.M. & Kicic A (2013) Change in airway epithelial permeability following

human rhinovirus infection is associated with disassembly of tight junctions in

well-differentiated air-liquid interface (ALI) cultures.

Oral presentation at the Respiratory Medicine Annual Scientific Meeting, Perth, WA

(2013) New Investigator Finalist.

Poster Presentation

1 Looi K.,, Garratt L.W., Iosifidis T, Lannigan F.J., Ling K-M, Martinovich K.M.,

Kicic-Starcevich E, Sutanto E.N., Stick S.M. & Kicic A (2013) Tight junction

expression in healthy and asthmatic airway epithelium: Are they intrinsically

different?

Poster presentation at Thoracic Society of Australia and New Zealand (TSANZ) WA

Branch Annual Scientific Meeting, Perth, Australia (2013)

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2 Looi K., Stick S.M. & Kicic A (2012) Evaluating changes on epithelial tight

junction expression in response to rhinoviral insult to determine potential

regulatory mechanisms.



3 Looi K., Stick S.M. & Kicic A (2012) Assessing the effects of rhinoviral

infection on epithelial tight junction expression to elucidate potential regulatory

mechanisms.

Poster presentation at the Child and Adolescent Health Service Research and Advances


4 Looi K., Ling K-M, Sutanto E.N., Larcombe A.N., Foong R. Rigby P, Stick

S.M. & Kicic A (2010) Changes in the airway epithelium barrier integrity in

response to viral insult.



5 Looi K., Ling K-M, Sutanto E.N., Larcombe A.N., Foong R. Rigby P, Stick

S.M. & Kicic A (2010) The effect of rhinoviral insult on airway epithelium

barrier integrity.

Poster presentation at the Child and Adolescent Health Service Research and Advances


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Awards

Australian Postgraduate Award (APA)

A three year scholarship was awarded during the PhD candidature by the University of

Western Australia, Perth, Western Australia, Australia (2010)

Princess Margaret Hospital Foundation PhD Top-Up Scholarship

A three year top-up scholarship was awarded during the PhD candidature by the

Princess Margaret Hospital Foundation, Perth, Western Australia (2010)

New Investigator Awards

Recipient of the Distinguished Young Scientist Award inclusive of conference

registration, accommodation and travel reimbursement from the European Respiratory

Society at Yokohama, Japan (2013)

Finalist for the Young Investigator Award at the Respiratory Medicine Annual

Scientific Meeting, Perth, Western Australia (2013)

Best Poster Prize

A conference travel award of $500 for best poster at the annual Child and Adolescent

Health Services Research and Advances Seminar, Perth, WA (2010)

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Travel Awards

Winner of the Lung Institute of Western Australia (LIWA) Young Scientist Award of

$1750 conference travel reimbursement for attendance at the American Thoracic

Society Annual Scientific Meeting at Denver, Colorado, USA (2011)

Conference travel awards of $500 from the Thoracic Society of Australia and New

Zealand (TSANZ) to attend the annual scientific conference in Darwin, Adelaide and

Gold Coast (2013, 2014, 2015)

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Acknowledgements

I would like to thank all the study patients and their families who have supported and

participated in this study. I would also like to thank all the funding bodies who have

provided financial support over the years. I would like to acknowledge the contribution

of all the doctors and recruitment officers at the Department of Respiratory Medicine at

Princess Margaret Hospital and St John of God Hospital Subiaco, who assisted in

patient recruitment and sample collection, in particular, Dr Desmond Cox, Dr Srinivas

Poreddy, Dr Francis Lannigan and Ms Cindy Bailey. I would especially like to thank Dr

Elizabeth Kicic-Starcevich for her dedication in recruiting and co-ordinating the

collection of samples, without whom, this study would not have been possible. I would

also like to thank all past and present members of the Epithelial Research Group for

their assistance in the laboratory, particularly, Ms Kak-Ming Ling, for her patience in

teaching and guidance.

For their aspiring guidance, I would like to thank my supervisors, Professor Stephen

Stick and Associate Professor Sunalene Devadason for their constant support, feedback,

constructive criticisms and their truthful and illuminating views on the various issues

related to this project. I am especially thankful to my supervisor, Associate Professor

Anthony Kicic, for his continuous support, guidance and patience on a daily basis. You

have been a tremendous mentor for me, encouraging my research and allowing me to

grow as a research scientist.

A special thanks to my family. Words alone cannot adequately express how grateful I

am to my parents and parents-in-laws for all the sacrifices that you’ve made on my

behalf. I would also like to thank my friends who supported me and encouraged me to

strive towards my goal. At the very end, I would like to express my special appreciation

to Erika, my beloved wife and mother of our two beautiful children, Evelyn and Kenzo,

who endured many a sleepless nights with and was always my pillar of strength in those

moments when all seemed impossible.

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In loving memory of my late Grandfather,

Looi Kim Thean

3rd July 2007

When the journey ahead seems arduous,

And all light has faded and failed,

Your words of “Never stop learning”,

Kept me persevering,

And so, against all odds, I have prevailed.

Thank you for being such an inspiration to me.

Looi 2015

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CHAPTER 1: Literature Review

1. Asthma

Asthma is among the most common chronic condition worldwide, affecting both

children and adults. The report by Masoli and colleagues (2004) estimated that 300

million people worldwide has a history of asthma and that this number would increase

to 400 million by 2025 as cities becoming increasingly urbanised (Masoli et al. 2004).

Asthma is the most common chronic illness in childhood and adolescence (GINA

2011), imparting physical, mental as well as an impaired quality of life burden to the

patients and their care-givers. Studies have shown that the quality of life experienced by

the asthmatic individual is closely associated with the severity of symptoms

(Warschburger et al. 2003; Merikallio et al. 2005) and it has been reported that

presentation of severe asthmatic symptoms has been closely linked to anxiety or

depression disorders (Richardson et al. 2006). In addition to an impaired quality of life

in patients, asthmatic episodes can also result in the increased need for treatment,

hospitalisation and emergency department services. Care-givers for asthma patients can

also experience a loss of personal time, absenteeism from work resulting in lost

productivity as well as the need to cope with the mental and physical demands.

Collectively, these requirements can then place further strain on the overall healthcare

system and social costs (Kenny et al. 2005; Simonella et al. 2006; Watson et al. 2007;

Ivanova et al. 2012). The total cost of asthma for the 2008-09 fiscal year in Australia

was $655 million, which represented 0.9% of the total health expenditure of that year

(AIHW 2011). Despite comprehensive analyses of the prevalence and burden of asthma,

attempts at advancing the understanding of the pathophysiology of asthma remain

elusive.

Historically, the earliest concept of asthma was derived from detailed clinical

observations that eventually contributed to the development of the bronchoconstrictor

paradigm. The Belgian physician, Jean van Helmont, who suffered from asthma,

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suggested one of the first pathophysiologic mechanisms of asthma in 1662, stating that:

“the lungs are contracted or otherwise drawn together” (Keeney 1964; Sakula 1988). In

1698, the English physician, Sir John Floyer, who developed asthma following a

respiratory infection, provided detailed accounts of asthma signs and symptoms,

treatment, prevention and prognosis. He also described a hereditary component of

asthma as well as other contributing factors to asthma exacerbations such as air

pollution, infection, cold air, exercise, psychological stress and tobacco smoke. These

observations aided in the clear distinction of asthma from other respiratory disorders.

By early 1900s, Brodie and Dixon proposed that the physiologic process of bronchial

narrowing due to constriction of the airway smooth muscle (ASM) resulted in asthma.

Subsequent observations from studies performed on individuals with asthma further

documented an exaggerated bronchoconstrictor response to various agents (Boushey et

al. 1980), which led to a general consensus that airway hyper-reactivity played a central

role in the pathogenesis of asthma. However, as airway hyper-reactivity could not be

consistently associated with ASM behaviour between healthy individuals and

individuals with asthma, it was postulated that non-muscle factors may contribute to the

development of airway hyper-reactivity in asthma as well as other chronic inflammatory

airway diseases (Solway and Fredberg 1997). Furthermore, the development of airway

hyper-reactivity in acute or chronic bronchitis has resulted in the postulation of a

common pathogenic mechanism between airway inflammatory diseases and asthma

(Bleecker 2004; Postma and Boezen 2004). Recent findings have documented

differences in ASM cells of animal models and individuals with asthma (Stephens et al.

2003; Roth et al. 2004). This accumulated evidence suggests the possibility of intrinsic

differences within the smooth muscles of individuals with asthma potentially

contributing to an increased level of bronchoconstriction. Although the importance of

ASM cells in asthma has long been recognised, the precise nature of its involvement in

the pathogenesis of airway hyper-reactivity remains unclear. Studies performed within

the past decade have challenged the well-established concept that ASM cells were

primarily an effector, while airway inflammation has been generally considered to be

the causal pathophysiological mechanism underlying airway hyper-reactivity and

ultimately, asthma pathogenesis. However, several studies have indicated that airway

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hyper-reactivity and airway inflammation can occur independently following specific

interventions at the inflammatory mediator level (Johnson et al. 2004; Kariyawasam et

al. 2007; Fattouh et al. 2008) .

Another suggestion was that an abnormal nervous system resulted in bronchial

constriction (Simonsson et al. 1967), thereby extending the bronchoconstriction

paradigm. In the past century, respiratory neurobiologists have shown that the lungs are

innervated with sympathetic, parasympathetic as well as non-adrenergic non-cholinergic

(NANC) nervous systems. Although recent studies have shown the influence of the

NANC nervous systems on airway behaviour, the primary role of NANC nervous

systems continues to be largely undefined and remains of interest to researchers

particularly in relation to cough associated with airway inflammation.

A dominant concept arose from observations of the many similarities between asthma

and allergic conditions (Stolkind 1933; Persson 1985) such as the high incidence of

allergen skin test reactivity and tissue eosinophilia (Powell and Hartley 1911; Cooke

and Vander Veer 1916). However, further examination indicated that certain individuals

with asthma have an allergic diathesis while some others do not. An early study of over

600 individuals with asthma categorised individuals as having “intrinsic” or “extrinsic”

asthma, largely dependent on the nature of the primary attributed factor (Rackemann

1921). Intrinsic asthma was thought to be precipitated by a factor within the individual

and was often associated with either respiratory bacterial or viral infection, a reflex of

upper airway irritation or stress and anxiety. Alternatively, extrinsic asthma was

observed to occur within hypersensitive individuals and asthma exacerbations were

induced by exposure to external allergens (Rackemann 1921). The commonly

associated link between allergy and asthma has been the major focus for asthma

research for at least 4 decades. Although much has been learned about the nature of the

association, very few new and effective therapeutic strategies have resulted. However,

the importance of airway inflammation has emerged as an almost ubiquitous factor in

asthma regardless of apparent phenotype.

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The presence of airway inflammation in individuals with asthma was evident in a series

of autopsies performed on individuals who died from status asthmaticus in 1922.

Evidence from these autopsies described the clinical presentation and classic

histopathological features of asthma which includes bronchial glands and muscle

hypertrophy, basement membrane thickening, injury to the AEC layer, mucous cell

metaplasia and accumulation of inflammatory cells such as eosinophils, neutrophils,

lymphocytes and macrophages into the sub-epithelial and epithelial layer (Huber and

Koessler 1922). However, it was also noted by Huber and Koessler that not all

individuals exhibited the classical eosinophilic infiltration of the airways typical of

allergic asthma, thus postulating the existence of other non-allergic subtypes of asthma.

Nonetheless, the inflammatory concept of asthma pathogenesis emerged as the most

widely regarded model of asthma pathogenesis in the early 1980s in which a pioneer

study quantified the severity of experimental airway inflammation and suggested that

non-allergic inflammation directed by AECs could also be responsible for airway hyper-

reactivity (Holtzman et al. 1983). Advancement of tissue sampling methods has enabled

studies to be performed utilising samples obtained via fibre-optic bronchoscopy for the

confirmation of previously obtained findings from autopsies demonstrating the presence

of mucosal inflammation within individuals with asthma (Djukanović et al. 1990;

Bousquet et al. 2000). Furthermore, other studies have shown a correlation between the

level of disease severity and the extent of inflammation and that an increase in

inflammatory cell infiltration was closely associated with the state of the disease

activity (De Monchy et al. 1985; Metzger et al. 1987; Laitinen et al. 1991; Vignola et

al. 1998).

An approach which links the “allergic” asthma phenotype with airway inflammation

stems from the observations of adaptive immune responses in murine models, in which

the inflammatory patterns are characterised by the development of distinct subsets of

CD4+ T cells designated as T-helper type-1 (TH1) or type-2 (TH2) (Mosmann et al.

1986). Although the distinction is not as clear in humans, similar differentiation of cells

in humans have been described and counter-regulation between TH1 and TH2 cells have

been shown to occur (Fiorentino et al. 1989; de Waal Malefyt et al. 1993; Manetti et al.

1993; Trinchieri 1994; Wenner et al. 1996) while the interplay of TH1 and TH2 with

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other cells have been shown to regulate the release of mediators responsible for the

inflammatory responses (Wills-Karp 1999). Moreover, considerable evidence obtained

from individuals with asthma, which demonstrated a TH2 inflammatory profile, led to

the belief that TH2 cells contribute towards asthma pathogenesis (Robinson et al. 1992).

However, observations have also shown that the onset and progression of asthma is not

entirely attributed to the allergic pathway and that other pathways exists that could also

result in the development of asthma (Johnston et al. 2007; Kim et al. 2008; Pichavant et

al. 2008). This subgroup of asthma is classically termed non-TH2 mediated asthma and

unlike TH2 mediated asthma, very little is known about this subpopulation of asthma,

the phenotypes underlying it as well as the molecular aspects which regulate it.

Observations such as those that demonstrated eosinophilic and neutrophilic asthma

phenotypes (Hastie et al. 2010; Bourgeois et al. 2011) and data that have challenged

whether the T cell paradigm can be extrapolated to humans from murine models argue

that the current perception is likely to be more complex than a simple dichotomy of

innate or adaptive immune developmental processes and responses.

As history has demonstrated, countless models and concepts have been proposed to

explain the pathophysiological abnormalities of asthma and also to explain the

complexity of asthma as a disease. However, a unifying explanation for asthma and its

heterogeneity remains elusive. One acceptable description is of a heterogeneous disease

with multiple phenotypes and endotypes, involving a plethora of susceptibility genes

(Table 1) in combination with multiple environmental stimuli and associated with

inflammation that can either be TH2-driven or non-TH2 mediated (Figure 1.1).

Therefore, given such a complex description, it is not surprising that the many models

and concepts have also become increasingly complex in their attempt to describe asthma

pathogenesis. Despite the considerable progress that has been achieved, the inability to

define a cause of asthma highlights the need to investigate alternative concepts that

might contribute to a better comprehension of disease pathogenesis.

Table 1: Asthma susceptibility genes with the associated functions and pathway

Susceptibility gene Associated function / pathway References

GSTM1 Environmental and oxidative stress — detoxification

Hatsushika, K. et al.(2007)

FLG Epithelial barrier integrity Palmer, C.N. et al (2006)

IL10 Immuno-regulation Guglielmi, L. et al (2007)

CTLA4 T-cell-response inhibition and immune-regulation Jones, G. et al.(2006)

IL13 TH2 effector functions Maier, L.M. et al (2006)

IL4 TH2 differentiation and IgE induction Imboden, M. et al. (2006)

CD14 Innate immunity — microbial recognition

Bernstein, D.I. et al. (2006)

SPINK5 Epithelial serine protease inhibitor Hubiche, T. et al. (2007)

ADRB2 Bronchial smooth-muscle relaxation Leung, T.F. et al.(2007)

HAVCR1 T-cell-response regulation — HAV receptor

Page, N.S. et al (2006)

LTA Inflammation Mak, J.C. et al. (2007)

TNF Inflammation Munthe-Kaas, M.C. et al. (2007)

GPRA Regulation of cell growth and neural mechanisms

Booth, M. et al. (2006)

NAT2 Detoxification of drugs and carcinogens Batra, J. et al. (2006)

FCERIB High-affinity Fc receptor for IgE Kim, Y.K. et al. (2007)

CC16 Epithelium-derived anti-inflammatory protein

Yang, K.D. et al. (2007)

IL18 Induction of IFNγ and TNF Imboden, M. et al.(2006)

STAT6 IL-4 and IL-13 signalling Yabiku, K. et al. (2007)

NOS1 Nitric oxide synthesis — cell–cell communication

Martinez, B. et al. (2007)

IL4R α-chain of the IL-4 and IL-13 receptors Yabiku, K. et al. (2007)

CCL11 Epithelium-derived eosinophil chemoattractant

Raby, B.A. et al.(2006)

ACE Monocyte, T-cell and eosinophil chemoattractant

Tanaka, K. et al. (2006)

TBXA2R Inactivation of inflammatory mediators Ku, M.S et al. (2006)

TGFB1 Smooth-muscle contraction, inflammation

Kim, S.H. et al. (2007)

ADAM33 Immuno-regulation, cell proliferation Van Eerdewegh, P. et al. (2002)

Figure 1.1: Schematic diagram of the factors contributing to the heterogeneity and complexity in asthma pathogenesis and diagnosis.

The genetic susceptibility and environmental interactions which contribute to asthma as well as the key clinical features of severity such as symptoms,

exacerbations and lung function, inflammatory characteristics and their categorisation into associated phenotypes and division into endotypes are

illustrated. However, the phenotypes as well as the resulting endotypes and the interactions between them have yet to be fully understood.

Early onset ASTHMA

Symptoms

Exacerbations

Lung function

TH2 mediated Non T

H2 mediated

Phenotype A

Phenotype B

Phenotype C

Phenotype D

Endotype 1

Endotype 2

Endotype 3

Endotype 4

Endotype 5

Late onset

Genetic

Susceptibility Environment

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1.2 Recent advances in the pathophysiology of asthma

The lack of a clear understanding of the pathobiology of asthma has resulted in

relatively few new therapies over the past 4 decades. Beyond the simple ‘allergic’ and

‘inflammatory’ paradigms, the recognition that individuals with similar adaptive

immune profiles might or might not develop asthma has led to investigations of whether

intrinsic abnormalities within the respiratory airways exist. Evidence from genome wide

association studies (GWAS), such as those performed by Koppelman and colleagues,

reported a link between airway hyper-responsiveness, a fundamental feature of asthma

and the gene encoding protocadherin-1 (PCDH-1), an adhesion molecule on the airway

epithelium. In addition, various genes that are expressed on the respiratory epithelium

have also been shown to be associated with asthma (Moffatt et al. 2007; Koppelman et

al. 2009; Willis-Owen et al. 2009). As a majority of these genes observed in these

GWAS studies associated with asthma are expressed within the airway epithelium

(Moffatt et al. 2010; Zhang et al. 2012), these observations support the notion that

asthma occurs as a result of aberrant gene expression within the airway epithelium and

that epithelial cells play a pivotal role in the allergic response. Moreover, biopsy studies

in children with asthma have also demonstrated an impaired epithelium early in the

disease pathogenesis (Barbato et al. 2006; Turato et al. 2008) as well as the crucial role

in the development of immune responses within the lungs (Hammad and Lambrecht

2008). These findings further strengthen the idea that asthma occurs as a result of an

aberrant airway epithelium, rendering the epithelium vulnerable to various insults.

The potential roles of the airway epithelium in asthma have been well-documented and

characterised in numerous studies (Bucchieri et al. 2000; Holgate et al. 2000; Chakir et

al. 2001). There has been evidence to suggest that asthma is primarily a disorder of the

airway epithelium and that altered function of the epithelial barrier properties are

closely associated with the developmental onset and subsequent clinical manifestations

of asthma, rather than an allergic pathway. Moreover, an impaired epithelial barrier

function would result in the airway being susceptible to early life bacterial or viral

infections. This, in turn, acts as a stimulus to prime immature immune cells to mount an

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inflammatory response and sensitisation towards various allergens, as suggested in a

study by Medzhitov and Janeway, where the innate immune system provided the

additional signals required by the adaptive immune system to mount an appropriate

response towards pathogens rather than self or harmless environmental antigens

(Medzhitov and Janeway 2000).

Prolonged epithelial susceptibility towards various environmental insults such as

allergen, pollutant exposure and viruses, coupled with a dysregulated repair response

would lead to a chronic asthma setting involving the persistence of airway inflammation

after the removal of the insult leading to subsequent airway wall remodelling. Increased

deposition of extracellular matrix on the basement membrane and a continued physical

distortion of the epithelium due to repeated bronchoconstriction would ultimately lead

to further remodelling. An exaggerated process of airway remodelling may eventually

result in partially reversible airflow obstruction and an accelerated decline in lung

function, aspects often seen in fatal asthma (James et al. 1989; Carroll et al. 1993).

Collectively, these studies provide considerable data to support a fundamental role of

the respiratory epithelium in the onset and development of asthma. The epithelium

could be central in the disease initiation and propagation as the airway surface is the

primary contact for inhaled allergens within the lungs. Moreover, an airway epithelium

with impaired barrier function would result in increased susceptibility and passage of

airborne allergens into the sub-epithelial components of the respiratory airways such as

basement membrane, fibroblasts and endothelium, which, in turns, exacerbates the

disease state. This provides the rationale for further examination into the role of the

respiratory epithelium in the development of asthma.

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1.3 Respiratory epithelium

1.3.1 Form and function of the epithelium

The human airway epithelium is a dynamic environment composed of at least eight

morphologically different epithelial cells types which can be classified into three broad

categories based on structural, biochemical and functional properties: basal, ciliated and

secretory (Spina 1998). Basal cells are commonly observed within the conducting

airways and past studies have shown a direct correlation with the number of basal cells

to decreasing airway size (Evans and Plopper 1988; Evans et al. 1990). Moreover, the

thickness of the conducting epithelium is also directly related to the percentage of

columnar cell attachment to the basement membrane via the basal cells (Evans and

Plopper 1988). Basal cells are the only cell type within the airway epithelium to be

directly attached to the basement membrane. This is achieved by the presence of hemi-

desmosomes that expresses the α6β4 integrin necessary for the firm attachment onto the

basement membrane (Evans et al. 1989). The basal cell, akin to the epidermal cell, is

postulated to be the precursor and / or progenitor cell which differentiates into either

mucous or ciliated cells (Boers et al. 1998).

The most common of the cell types present within the conducting epithelium are the

columnar ciliated epithelial cells, found on the topmost layer of the airway epithelium.

These cells account for greater than 50% of all epithelial cells and often originate from

either basal or secretory cells (Ayers and Jeffery 1988). Normally, columnar ciliated

cells would possess around 300 cilia on each cell, thus indicating their primary role in

the unidirectional transport of secreted mucous from the lung to the throat. Secretory

cells include both goblet cells as well as Clara cells. Goblet cells or otherwise known as

mucous cells, are identified by their electron-lucent acidic-mucin granules which are

secreted into the airways to entrap foreign inhaled pathogens or environmental particles

(Jeffery 1983; Jeffery 1991). It has been suggested that an average human trachea

consists of approximately 6800 goblet cells per mm2 of surface epithelium but in

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chronic airway inflammatory diseases such as asthma or bronchitis, mucous cell

hyperplasia or metaplasia occurs, leading to excess mucous production, a pathological

finding consistent with these diseases (Lumsden et al. 1984). Goblet cells are capable of

self-renewal and may also differentiate into ciliated columnar epithelial cells following

either a mechanical or a pathogen insult (Evans and Plopper 1988).

In humans, Clara cells are located within both the bronchial as well as the bronchiolar

airways and contain electron-dense granules. Clara cells have been shown to secrete

bronchiolar surfactants and specific anti-proteases such as the secretory leukocyte

protease inhibitor in order to regulate bronchiolar epithelial integrity and immunity as

well as to prevent excessive tissue damage caused by the secretion of harmful

proteinases from neutrophils during inflammation (De Water et al. 1986; Sallenave et

al. 1994). Furthermore, Clara cells are also capable of producing oxidases that

metabolises xenobiotic compounds such as aromatic hydrocarbons found in cigarette

smoke. Evidence has also suggested that Clara cells may in fact harbour stem cell

potentials and could act as progenitor cells for either mucous or ciliated cells (Hong et

al. 2001). These cells come together to form a pseudostratified layer that lines the

conducting airways beginning at the large airway epithelium and terminating at the

alveolar epithelium (Figure 1.2). These cells perform numerous important roles such as

regulating lung fluids, removal of inhaled particles, activating the different

inflammatory cells in response to injury and the secretion of various mediators to

regulate airway smooth muscles. Moreover, cells within the airway epithelium are

constantly renewed (Crystal et al. 2008) as it is also the primary interface between

noxious external environment stimuli and the lung milieu. Hence, any damage sustained

by the epithelium has the potential to not only cause inflammation, but also contribute

to the pathogenesis of major lung diseases such as asthma and chronic obstructive

pulmonary disorder (COPD). Although traditionally regarded to be an inert barrier

against the external environment, the airway epithelium, in recent years, has been

proven to play a central role in the control and modulation of various airway function

(Holgate 1998; Holgate et al. 2000; Knight 2001). Nevertheless, the most crucial role of

the airway epithelium is still protecting the lungs and airways against external stimuli.

This is often achieved through the interaction of different protective mechanisms,

Figure 1.2: Schematic of major cell types lining the respiratory airways. Within the bronchi, the predominant cell types are basal (B), ciliated

columnar (C) and goblet (G) cells. In the bronchioles, the cell types remain relatively similar except with more Clara cell (CL) type. As the bronchioles

merge with the alveolar epithelium, Type 1 (T1) and Type 2 (T2) cells become the major cell types. The endothelium (EN) provides a division between

the epithelium (EP) and the blood stream (BS). Neutrophils (N) are shown to migrate through the blood stream to the lumen through the endothelium,

endothelial basement membrane (BM) and interstitial tissue (IN) containing both type I fibroblast (F), which are parallel to the epithelium and type II

myofibroblast that are perpendicular to the epithelium and lastly, through the epithelium via a series of ligand-receptor interactions.

Bronchi Bronchioles Alveoli

EP

IN

EN

BS

EN

B

BM

BM

F

G C

CL T 2

T 1

R N

Adapted from Tam et al, Therapeutic Advances in Respiratory Disease, 2011

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however, a physical barrier provided by various junctional complexes and supported by

an efficient and highly effective mucociliary clearance remains the most pivotal defence

against external injurious stimuli.

1.3.1.1 Mucociliary clearance

One of the critical functions of the airway epithelium is the wave-like clearance of

inhaled particles. Mucous producing goblet cells together with surfactant secreting

Clara cells and ciliated columnar epithelial cells, work in collaboration in the trapping

and removal of inhaled foreign particles from the airway lumen (Kilburn 1968). Surface

epithelial goblet cells secrete the predominant form of mucin in the human airway,

MUC5AC while MUC5B is mainly secreted by the mucous cells of the submucosal

glands (Hovenberg 1996; Wickström 1998). Regulation of mucin production can be

attributed to various factors ranging from inflammatory mediators such as

lipopolysaccharides (LPS) (Smirnova et al. 2003), growth factors such as epidermal

growth factor (EGF) or transforming growth factor – α (TGF-α) (Takeyama et al. 1999)

to environmental insults such as cigarette smoke (Shao et al. 2004). Producing the right

amount of mucin, coupled with the viscoelasticity of the mucous is critical in the

maintenance of efficient mucociliary clearance. The viscoelastic mucous layer, which

floats on the periciliary layer tethered to the apical cell surface by different mucins and

glycolipids (Sheehan et al. 2008), acts as a fluid reservoir that removes or donates liquid

to maintain homeostatic airway surface liquid (ASL) level that approximates the total

height of the cilia (Tarran et al. 2001; Leopold et al. 2009). In normal airways, the

cystic fibrosis transmembrane conductance regulator (CFTR), together with epithelial

sodium channel (ENaC) allows for the combination of chloride secretion and sodium

reabsorption to favour a healthy ion composition and ASL depth (Zeitlin 2008). This

enables proper ciliary function for an effective mucociliary clearance. However, a

dysregulation between the chloride secretion and sodium reabsorption often observed in

chronic airway diseases such as cystic fibrosis could result in a decrease of the ASL

depth, leading to mucosal glands hypertrophy and subsequently, excessive production

of mucous resulting in airflow obstruction and increased bacterial colonisation within

the respiratory airways (Zeitlin 2008). Deficient mucociliary clearance could also be a

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direct result of ciliary dysfunction. Individuals with ciliary dysfunction usually have

normal amounts of mucous production but suffer from a compromised mucociliary

clearance due to a defective ciliary beat pattern (Bush et al. 1998; Noone et al. 2004).

Despite compromised mucociliary clearance occurring in different respiratory diseases,

the airway epithelium, through the various junctional protein complexes, persevere to

provide a highly regulated and impermeable barrier in the defence of the lungs and

airways.

1.3.1.2 Junctional protein barrier

The airway epithelium, being the primary interface between the lung milieu and the

external environment, forms a complex barrier to provide the first line of physical

defence against airborne pathogens. The AECs, which adhere tightly to one another to

form an epithelial sheet lining the entire mucosal surface in contact with inhaled air,

play pivotal roles in providing a physical barrier in defending the lungs and airways.

They are cemented to each other through the formation of adhesive cell – cell contact

junctions that include tight junctions (TJs), adherens junctions (AJs), gap junctions and

desmosomes. These junctional proteins come together to form the epithelial junctional

complex (Farquhar and Palade 1963) that completely surround the cell. These junctions

play a crucial role in the formation and maintenance of epithelial barrier by mediating a

tight seal between adjacent epithelial cells (Farquhar and Palade 1963). This results in a

continuous junctional belt that interconnects with neighbouring cells which then acts as

a barrier or fence to selectively regulate the passage of ions, solutes and cells through

the paracellular space (Roche et al. 1993). Although their ultrastructure from freeze-

fracture electron microscopy suggests that these junctional complexes form stable

structures, studies have indicated that they are highly dynamic complexes capable of

various functions even in fully polarised epithelia (Nelson 2003; Irie et al. 2004;

Gumbiner 2005). These junctional complexes can be broadly defined into three

categories which consist of structural proteins needed for the initiation of the junctions,

peripheral plaque proteins associated with the actin cytoskeleton and the signalling or

polarity proteins necessary for polarisation of the epithelium. Among the myriad of

junctional complexes, TJs, which are located at the most apical point of the epithelial

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cell and are closest to the airway lumen, serve to demarcate the boundary between the

apical surface and the basolateral domains of the cell. The common model of a TJ

structure is of a stable multi-protein complex composed of integral and peripheral

membrane proteins (Tsukita et al. 2001; Aijaz et al. 2006). The major types of integral

proteins are grouped according to the number of transmembrane domains they contain,

four pass transmembrane proteins including claudins, occludin and tricellulin while

single pass transmembrane proteins include junctional adhesion molecules (JAMs) as

well as the Coxsackie and adenovirus-associated receptor (CAR) (Balda and Matter

2008).

The claudin family, which consists of at least 24 members, have been shown to be

incorporated into TJ strands when observed in cultured epithelial cells (Furuse et al.

1998). Despite studies showing their role in maintaining epithelial integrity (Tsukita and

Furuse 2002), their role in controlling transepithelial permeability remains unclear

because the functional characteristics of the majority of claudins are still unknown at

present. The identification of claudins by Furuse and colleagues advanced the

understanding of TJ structure (Furuse et al. 1998; Furuse et al. 1998). Associations of

the other integral membrane proteins with claudin further complicate the TJ structure.

Occludin, the earliest transmembrane protein to be identified, has also been implicated

in regulating the permeability properties of the TJ seal, in particular, with the regulation

of size-selective paracellular diffusion (Furuse et al. 1993; Balda et al. 1996; McCarthy

et al. 1996; Balda et al. 2000). Although TJs without occludin are relatively uncommon,

the physiological function of occludin in the TJ complex remains relatively unclear. In

epithelial tissues lacking occludin, TJ strands as well as barrier function were observed

to be present, however, in mice that were deficient in occludin expression, different

phenotypes ranging from growth retardation, deposition of minerals within the brain to

sterility seems to suggests barrier impairment to a degree (Saitou et al. 2000).

Interestingly, in studies performed using Madin-Darby canine kidney (MDCK) cells

with knocked down occludin expression, it appeared that the role of occludin within the

TJ complex remains elusive, independent of the traditional TJ barrier function that is

crucial in the communication of apoptosis to adjacent cells (Yu et al. 2005). Tricellulin,

also a tetrapass transmembrane protein, was recently identified as having structural

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similarity to occludin (Ikenouchi et al. 2005). However, in contrast to occludin and the

claudin family, tricellulin is only enriched at the tricellular TJs, where it acts to

reinforce the barrier function of the epithelium. A study by Ikenouchi et al using RNA

interference to suppress tricellulin expression resulted in barrier impairment of TJs,

which indicates the important role tricellulin has in junctional formation (Ikenouchi et

al. 2005). Underlying this membrane domain is the cytoplasmic plaque consisting of a

network of densely packed peripheral adaptor proteins which connects the integral

membrane proteins to the underlying actin cytoskeleton as well as various signalling

proteins. Within TJs, the cytoplasmic plaque functions to regulate adhesion, paracellular

permeability and the transmission of signals from cellular junctions to the interior to

control various cellular processes such as migration and gene expression. An important

and probably the most studied plaque component is the peripheral adaptor protein tight

junction protein-1 (TJP-1) or otherwise identified as zonula occludens-1 (ZO-1). Zonula

occludens-1 has a typical functional property and domain structure of scaffolding

proteins that contains multiple sites of protein-protein interaction, including 3 PDZ and

a single SH3 domain through which they bind to a number of cytoskeletal, signalling

and membrane proteins (Guillemot 2008; Fanning and Anderson 2009). Tight junction

components are also capable of engaging in interactions with proteins of other

junctional complexes and this is thought to be crucial for the proper organisation of the

integral membrane structures as well as the regulation of junction assembly, function

and signalling to the cell interior (Matter and Balda 2003; Köhler and Zahraoui 2005;

Fanning and Anderson 2009) (Figure 1.3). Located directly below the TJs are the

adherens junctions, which perform various functions including the initiation and

stabilisation of cell-cell adhesion, regulation of the actin cytoskeleton, intracellular

signalling as well as transcriptional regulation. Among the many single pass

transmembrane proteins, the most notable is E-cadherin, which belongs to the classical

cadherin family of calcium-dependent adhesion proteins. Classical cadherins have 5

characteristics extracellular cadherin domains that form trans-cadherin interactions

between adjacent cells to initiate cell-cell adhesion and formation of adherens junctions.

The cytoplasmic domain of E-cadherin binds to proteins that are responsible for the

regulation of E-cadherin endocytosis, recycling and degradation, intracellular signalling,

gene transcription and control of the actin cytoskeleton. Desmosomes, located around

Figure 1.3: Illustration of the complexity between protein-protein interactions at

the tight junction complex. Interactions between proteins such as claudins, occludins

and ZO are essential for the structural integrity of the tight junction while other

interactions between ZO and ZONAB regulate signalling pathways which originate

from the tight junction. Interactions involving cingulin potentially regulate the

transcriptional up-regulation of claudins and occludin, leading to eventual tight junction

assembly. Similarly, interactions between Par3 or Par6 and aPKC can also lead to tight

junction assembly. aPKC, atypical protein kinase C; CDK4, cell division kinase 4;

MUPP1, multi-PDZ domain protein 1; PALS1, protein associated with Lin seven 1; Par,

partitioning defective; PATJ, PALS1-associated tight junction protein; RalA, Ras-like

GTPase; Tiam1, T-lymphoma invasion and metastasis; ZO, zonula occludens; ZONAB,

ZO-1-associated nucleic acid–binding protein.

CDK4 RalA

ZO-2

ZO-1

ZO-3

Cingulin

PATJ

PALS1

ZONAB

Cdc42

MUPP1

Rac

aPKC

Par6 Par3

Tiam1

Actin cytoskeleton

Occludin

Claudins

Transmembrane proteins Peripheral proteins G-protein / regulator Kinase / Phosphatase Other signalling protein

Adapted from Shin et al, Annual Review of Cell & Developmental Biology, 2006

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the midsection of the cell, are intercellular junctions that provide robust adhesive bonds

between epithelial cells to give mechanical strength to the epithelium. Gap junctions,

which can be found at the basolateral side of the epithelial cells, are specialised cell –

cell channels that permits the diffusion of small metabolic solutes, ions and other

molecules between adjacent cells (Mese et al. 2007).

The physical barrier and fence function of the epithelium have allowed for the

association of epithelial junctional complexes as robust, rigid structures that

determinedly prevent unwanted molecular traffic and although their primary function of

providing a physical barrier remains unchanged, studies over the years have suggested

that these junctional complexes could be more dynamic in their barrier roles and

responses than previously thought (Madara 1990; Rosenblatt 2001; Matter and Balda

2003; Pilot 2005; Wang and Cheng 2007; Shen et al. 2008). Moreover, several

published reports have suggested that the composition of the junctional associated

protein complexes is flexible and is dependent on diverse factors ranging from the state

of junctional assembly to the proliferation state of the cells (Aijaz et al. 2006; Ebnet

2008). Hence, a model of TJ complexes based on a flexible network of junctional

protein strands would fit the concept of a dynamic epithelium, thereby explaining the

size selective paracellular diffusion process allowing the movement of solutes through

controlled regulation of the TJ proteins between adjacent cells. This evidence in

conjunction with the concept of a dynamic and flexible junctional complex may

suggests the importance of the airway epithelium in the regulation and modulation of

airway responses to external stimuli.

1.3.2 Airway responses in the asthmatic epithelium

Airway inflammation is increasingly gaining recognition as a key component of asthma

and represents a complex interaction of inflammatory cells and airway cells and has

been postulated that a multitude of immune cells comprising predominantly of

eosinophils, neutrophils, lymphocytes, monocytes, mast cells and basophils, with

eosinophilic infiltration being the most commonly observed (Kay 2001), are capable of

orchestrating airway inflammation. However, although there is much evidence to

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support the notion that airway inflammation drives the remodelling process within the

airway walls, there are accumulating data to support the argument that an intrinsic

abnormality of airway architecture or function might initiate or perpetuate airway

inflammation.

Airway inflammation in asthma is often restricted to the conducting airways, however,

as the disease becomes more chronic, the inflammatory infiltration extends to involve

the trachea, larynx, small airways and occasionally, the alveoli (Kraft et al. 1996).

Progression of the inflammatory process to the smaller airways only occurs as the

severity of the disease increases and becomes chronic in nature but otherwise, remains

largely restricted to the larger airways (Kraft et al. 1999). Inflammation of the

submucosa dominates in the large airway whereas in small airways, the inflammatory

response seems to be predominantly external of the airway smooth muscle (Haley et al.

1998). The exact location of airway inflammation remains controversial, however, it has

been suggested that all airway cells are capable of active participation in the

pathogenesis of airway inflammation in asthma (Springall et al. 1991; Corrigan and Kay

1992; Vignola et al. 1993; Sousa et al. 1997). Inflammatory cells, in conjunction with

mesenchymal cells, create a complex cellular network that directly regulates the

inflammatory and reparative changes within the airway (Brewster et al. 1990). These

changes are often observed in the majority of individuals with differing severity levels

of asthma and can eventuate in the progression towards airway remodelling to cause

permanent tissue alterations in both large and small airways.

Asthma is an inflammatory disease often associated with changes in the usual structure

of the airway walls. These architectural changes are collectively termed airway

remodelling. Airway remodelling in asthma usually involves changes within the airway

epithelium, with characteristic shedding of columnar epithelial cells coupled with

abnormal changes in the goblet cell, eventually resulting in mucus plugging. The most

prominent remodelling changes can usually be seen beneath this augmented epithelium,

ranging from smooth muscle hyperplasia, increased angiogenesis and innervation to the

thickening of the sub-basement membrane layer involving deposition of interstitial

collagen. Together, these changes affect the airway structure as well as its mechanical

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and functional properties and have been thought to be a major contributor to the

pathophysiology of the episodic airway dysfunction described as airway hyper-

responsiveness (AHR). Airway remodelling is a primary and consistent component of

paediatric asthma with various studies describing increased deposition of collagen and

the thickening of the lamina reticularis, angiogenesis and increased smooth muscle

(Roche et al. 1989; Aikawa et al. 1992; Kuwano et al. 1993; Li and Wilson 1997).

Although there exists a paucity of evidence of reticular basement membrane thickening

in wheezing infants, airway walls have been reported to be abnormal by several studies

in infants who subsequently develop asthma. Despite the conventional paradigm that

airway remodelling is a consequence of a chronic inflammation, recent evidence has

suggested that these remodelling processes could occur as a result of an initial stimulus

and there are cogent evidence that indicates airway remodelling occurs in early

childhood (Fedorov et al. 2005; Barbato et al. 2006). Although certain components of

airway remodelling are reversible through therapeutic interventions or spontaneously,

the more prominent abnormalities observed within the epithelium, smooth muscle,

vasculature and extracellular matrix are most likely to be sustained and refractory to

pharmacologic interventions. An elevation in airway smooth muscle (ASM) mass

caused either by hypertrophy, hyperplasia or increased deposition of extracellular

matrix is a crucial component of a remodelled wall in asthmatic airways. Despite a lack

of understanding in the precise mechanisms behind this increased mass, it has been

postulated that increased proliferation rates could potentially contribute to the

development of ASM thickening (Hirst et al. 2004).

Various models have been used to characterise airway remodelling in asthma ranging

from post-mortem examination of lung tissues to murine models. Observations of lung

tissues taken from autopsies have demonstrated profound occurrences and changes in

airway architecture in those that have died from asthma when compared to those with

asthma but died from other causes and those without asthma (Bousquet et al. 2000).

Biopsy specimens obtained from living individuals with asthma, when compared to

healthy controls, have demonstrated typical histopathological changes associated with

asthma such as reversibility of some changes with therapeutic intervention, changes in

non-inflammatory airway wall structure and alterations within the distribution of

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inflammatory cells (Van Den Toorn et al. 2001). Although these studies provide an

insight into the process of airway remodelling, they are often limited due to their highly

invasive nature to obtain a relatively small quantity of viable target tissue. Despite the

incidence of substantial epithelial damage in asthma remaining controversial, there is no

doubt that the asthmatic epithelium is intrinsically abnormal both in vivo and in vitro

(Bousquet et al. 2000; Kicic et al. 2006).

1.3.3 Intrinsic abnormalities of the asthmatic epithelium

As the initial interface between the environment and the sub-mucosa, the airway

epithelium represents the primary point of defence for the lung from the various

constituents of the environment ranging from pollutant to viruses. Injury on the airway

epithelium results in a sequence of inflammatory and cell signalling events that would

lead to either regeneration or repair. These two processes differ in that regeneration

returns the epithelium to its normal structural and functional capacity while repair

results in increased stability of the epithelium but does not confer similar structural and

functional capacity before epithelial damage. In order to comprehend the complex role

of the epithelium in asthma, studies utilising mucosal biopsies and primary cultures of

AECs obtained from donors with asthma have provided insights as well as to

demonstrate that the airway epithelium is inherently abnormal.

A seminal study by Kicic and colleagues reported that AECs obtained from children

with asthma demonstrated both biochemical and functional differences when compared

to healthy cohorts (Kicic et al. 2006). Results from their study indicate no difference in

the release of the pro-inflammatory cytokines interlukin-1β (IL-1β), interlukin-8 (IL-8)

and sICAM-1 in the different donor cohorts. These findings are in contrast to those

previously reported, which showed increased levels of ICAM-1 (Vignola et al. 1993;

Manolitsas et al. 1994), IL-1 (Borish et al. 1992; Sousa 1996) and IL-8 (Marini and

Marini 1992) production by the bronchial epithelium in donors with asthma. Kicic et al

also reported an increased in prostaglandin E2 and IL-6, which correlates with other

studies indicating the epithelial cells were not of a pro-inflammatory phenotype.

Moreover, Kicic et al demonstrated that the asthmatic epithelial cells had a greater

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proliferative capacity with greatly elevated levels of proliferating cell nuclear antigen

(PCNA) mRNA which corroborated with their reported increase in epidermal growth

factor (EGF) release despite a lack of epidermal growth factor receptor expression being

reported.

Kicic et al also showed in their study that the release of transforming growth factor β1

(TGF-β1) was markedly diminished, suggesting abnormal differentiation might occur,

which could relate to the lower expression of cytokeratin-19 expression in the asthmatic

cells. Furthermore, Gras and colleagues, in their study to investigate the feasibility of

utilising air-liquid interface (ALI) cultures of bronchial epithelium derived from

endobronchial biopsies of individuals with differing severity of asthma, demonstrated

that both inflammatory and morphological imbalances initially observed in biopsy

samples obtained from patients with mild to severe asthma continued to exist within the

reconstituted epithelial ALI cultures throughout the entire differentiation process.

Moreover, results from the study performed by Gras et al also indicated that the

epithelium of individuals with severe asthma produced significantly elevated levels of

mucin and interlukin-8 (IL-8) but diminished levels of lipoxin A4, an anti-inflammatory

factor when compared to those with mild asthma or healthy controls. Observations from

the study conducted by Gras et al indicated not only the feasibility and relevance of ex

vivo ALI cultures of bronchial epithelium obtained from endobronchial biopsies, but

also demonstrating that inherent phenotypic differences exists within the asthmatic

epithelium (Gras et al. 2012).

In another study, Stevens et al, in their study, reported that bronchial epithelial cells

obtained from donors with asthma require a significantly longer time to achieve repair

upon wounding in contrast with cells obtained from healthy donors. They also reported

that mRNA expression of the plasminogen activator inhibitor-1 (PAI-1) was greatly up-

regulated within the asthmatic cells compared to healthy ones and that protein

expression was also similarly increased. These data, taken in conjunction with a reduced

rate of proliferation in both asthmatic and healthy cells following gene silencing and

mechanical wounding demonstrate that bronchial epithelial cells obtained from donors

with asthma are inherently dysfunctional in the capability to repair wounds despite

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elevated PAI-1 levels, which have been shown to play pivotal roles in both proliferation

and repair of healthy cells (Stevens et al. 2008). Findings from a recent study by Kicic

et al showed the diminished ability of bronchial epithelial cells obtained from donors

with asthma to secrete the extracellular matrix (ECM) component of fibronectin (FN),

providing further evidence that the asthmatic epithelium have an impaired repair ability

(Kicic 2010).

Freishtat and colleagues, similarly showed that following mechanical wounding, the

rate of regeneration in the asthmatic epithelia was less efficient compared to healthy

epithelia. Moreover, the asthmatic epithelia secreted more TGF-β1, IL-1β, 10, 13 and

had markedly less mitotic cells that were more dyssynchronously distributed along the

cell cycle in contrast to healthy epithelia (Freishtat et al. 2010). These results further

support the previous findings demonstrating an impaired repair process is inherent to

bronchial epithelial cells obtained from donors with asthma and further supports the

concept that the asthmatic epithelium is intrinsically different from a healthy

epithelium. Furthermore, these results also demonstrated that commercial cell lines and

adult AECs may not always be appropriate when attempting to comprehend paediatric

airway diseases.

Although there have been significant advances over the past few decades in our

knowledge regarding the human airway epithelium, little is still known about whether

the abnormality in epithelial barrier is representative of a gene-environment interaction,

with a genetic diathesis to asthma or atopy eventually resulting in a change in epithelial

cell response during early life to potential environmental stimuli such as respiratory

viruses. In addition, respiratory viruses have been shown to be the most common trigger

for asthma exacerbations in both adults and children (Nicholson et al. 1993; Johnston et

al. 1995). However, a paucity of data on the interaction between the known intrinsic

abnormalities of the epithelium with respiratory viruses provides the rationale to further

investigate the barrier role, in particular, the tight junctional complexes of the

epithelium as a potential influence in asthma exacerbations following early life

respiratory viral infections.

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1.4 Respiratory viruses and asthma

Although airway infection during infancy and early childhood was initially perceived to

confer protection against the development of atopic asthma (Ball et al. 2000), a growing

body of evidence has resulted in a progressive shift in this concept which suggests that

respiratory viral infections as well as the resulting innate immune response can be

attributed to or associated with most acute asthma exacerbations, up to 80-85% in

children and around 60% in adults (Gama et al. 1989; Pattemore et al. 1992; Nicholson

et al. 1993; Johnston et al. 1995; Papadopoulos et al. 2003; Tan et al. 2003; Dahl et al.

2004; Sly et al. 2006). Rhinovirus (RV), coronavirus, influenza virus, adenovirus,

parainfluenza virus and respiratory syncytial virus (RSV) have been shown to be the

most common viruses to trigger wheezing within infants and cause the exacerbation of

asthma symptoms in older children and adults, however, many other factors such as age,

gender and race could determine the susceptibility towards different viral infections.

Viral infections of the respiratory tract are among the most common debilitating illness

worldwide and have been reported as a major trigger of asthma exacerbation in both

adults and in children (Lambert and Stern 1972; Minor et al. 1974; Johnston et al. 1995;

Busse et al. 2010). Early on in life, many children have wheezing episodes that are

closely related to respiratory infections, however, in most cases, wheezing episodes

associated with respiratory tract infection will diminish with age. Exceptions occur

where early life wheeze episodes can indicate the onset of asthma in certain groups of

susceptible individuals. The advancement and development of highly specific and

sensitive diagnostic methods have led to improved detection of respiratory tract viruses,

thus allowing a clear insight into the relationship between viral infections and asthma

exacerbations. Viral respiratory tract infection can have a profound effect in individuals

with established asthma and recent use of molecular diagnostic techniques such as

reverse transcription polymerase chain reaction (RT-PCR) to either supplement or

replace conventional virology techniques have reported an increase in detection of the

common viruses among adults and children (Nicholson et al. 1993; Johnston et al.

1995; Wark et al. 2002). Moreover, separate studies performed have now shown

respiratory viral infections to be associated with 80 – 85% of acute asthma

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exacerbations in children (Johnston et al. 1995) and 45% in adults (Nicholson et al.

1993). Although it has been recognised for many years that respiratory viruses can

trigger asthma exacerbations, accumulating evidence from recent studies have not only

highlighted the importance of respiratory viruses but also demonstrated that the cause of

most cases of virus induced asthma is linked to respiratory viruses such as respiratory

syncytial viruses (RSV), influenza A virus (IAV) and human rhinoviruses (HRV)

(Bizzintino et al. 2011; Fujitsuka et al. 2011; Hasegawa et al. 2011).

1.4.1 Influenza virus

Influenza viruses, classified as a RNA virus of the family orthomyxoviridae, are a major

determinant of morbidity and mortality in many worldwide epidemics or pandemics.

Influenza viruses can be classified into three immunologic types, designated A, B and

C. Each of the immunologic types of influenza viruses has one species, namely

Influenza A, B and C virus respectively. All three species are capable of infecting

humans, however, of importance is the influenza A species due to its continual

occurrence of antigenic mutation, making influenza A virus (IAV) the most virulent

human pathogen among the three influenza types capable of causing severe and often

devastating disease outbreaks. All three influenza types are similar in structure and

composition, usually observed to be spherical, between 80 – 120 nm in diameter and are

made up of a viral envelope containing two main types of glycoproteins,

haemagglutinin (HA) and neuraminidase (NA) wrapped around a central core. The

central core contains the viral genome, which is comprised of 7 – 8 pieces of single

stranded, often segmented negative sense RNA, encoding for various proteins needed

for the viral replication process.

Influenza virus infection is extremely common and is often associated with substantial

morbidity and mortality worldwide, and especially in individuals with existing asthma,

as observed by Jain et al, in their study during the 2009 H1N1 influenza A virus

pandemic (Jain et al. 2009). However, the exact mechanism in which influenza viruses

contributes to asthma exacerbations remains unclear. It has been proposed that the

AECs may play a role in the exacerbation of asthma as AECs are often the primary sites

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for most viral infections, including influenza virus infections. As demonstrated in

several studies, infection of the AEC would ultimately lead to the activation of various

signalling cascades which then initiates expression of a range of cytokines and

chemokines (Buchweitz et al. 2007; Jewell et al. 2010; Tate et al. 2011). The eventual

destruction of the AEC following infection, in conjunction with a pro-inflammatory

immune response are the major contributing factors to the influx of inflammatory cells

and AHR often associated with asthma exacerbations. A study by Park et al showed that

individuals with asthma have increased levels of IL-13, a key cytokine involved in the

formation of goblet cells leading to increased mucin production, as well as a pro-fibrotic

repair of the airway epithelium and diminished production of IFN-γ. In addition, various

studies performed which utilised AECs of healthy and individuals with asthma have

revealed that AECs obtained from individuals with asthma demonstrated augmented

expression of genes involved with airway inflammation, airway epithelial repair process

and remodelling, all of which could contribute towards viral-induced exacerbations of

asthma (Kicic et al. 2006; Stevens et al. 2008; Kicic 2010).

Airway hyper-reactivity and airway inflammation are widely thought to be orchestrated

by allergen specific TH2 cells in conjunction with basophils and eosinophils, which are

commonly present in the lungs of the majority of individuals with asthma, especially

those with allergic asthma. However, non-TH2 factors such as IFN-γ and neutrophils can

also be frequently observed within the lungs of individuals with severe or

corticosteroid-resistant asthma. It has been commonly accepted that these non-TH2

cytokines often antagonise TH2-mediated allergic diseases. In two separate studies by

Doyle et al and Román et al, they were able to demonstrate that following respiratory

infection with influenza virus eventually led to an increased production of local IFN-γ

concentrations by CD4+ and CD8+ T cells (Doyle et al. 1999; Román et al. 2002).

Furthermore, Dahl et al were able to show that infection with IAV would initiate an

intense IFN-γ response within the lungs which then leads to the development of robust,

TH1-polarising dendritic cells (DCs). The study then advanced to utilise a TH2-

dependent mouse model of allergen-induced lung inflammation to demonstrate the

ability of these robust DCs in strengthening subsequent immunity by reinforcing both

TH1 and TH2 immunoglobulins and cytokines production (Dahl et al. 2004). Moreover,

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in a recent study performed by Chang and colleagues to define the various inflammatory

cell types and processes involved in viral-induced asthma, they established and

subsequently infected an experimental murine model with IAV of subtype H3N1

(Chang et al. 2011). Their results showed that IAV infection effectively induced airway

inflammation and airway hyper-reactivity, independent of TH2-mediated or adaptive

immunity. This was achieved through the production of IL-33 in alveolar macrophages

and its corresponding receptor, ST2 in conjunction with the innate lymphoid cell

population termed “natural helper cells” (Chang et al. 2011).

Although the actual characteristics of these natural helper cells are still being delineated,

their contribution towards the development of airway hyper-reactivity and their

activation by IAV subtype H3N1 via an IL-33 dependent, fully innate pathway

demonstrates the multiple pathways influenza virus could potentially lead to the

development and exacerbation of airway inflammation. However, despite the

progression in our understanding on how viral-induced airway inflammation is

generated and the likelihood a majority of these pathways coexisting and synergistically

causing airway hyper-reactivity, inflammation and ultimately asthma, the precise

underlying mechanism remains to be established.

1.4.2 Respiratory syncytial virus (RSV)

Respiratory syncytial virus (RSV), classified as a paramyxovirus, is a ubiquitous

pathogen in all human population and is closely related to the parainfluenza virus,

measles and mumps (Cane 2001). The genome of the virus consists of a single stranded

RNA containing only 10 genes which encodes for 11 proteins, 9 of which are structural

proteins and surface glycoproteins and the remaining 2 directly involved in the

replication process upon viral entry into the host cell (Cane 2001; Hall 2001). Two

different strains of RSV, A and B, have been identified, with both being infectious but

one strain being dominant during an epidemic in a particular region. Respiratory

syncytial virus epidemics changes with the onset of different seasons, generally

occurring with either the start of autumn or winter. Due to the universal presence of

RSV within the community, virtually all children would have had an infection by the

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age of 2. Infections in infants less than two months of life are less common, however,

infections rates climb rapidly, reaching a peak during the third and fourth months of life

(Glezen et al. 1986). Repeated infections are common in all age groups and previous

infections do not confer immunity or resistance against subsequent infections even in

sequential years. No particular age group is exempt from the risk of RSV infection,

however, certain risk factors such as pre-termed neonates, individuals with cystic

fibrosis, immune-suppressed patients, low socio-economic status, crowded living

conditions, exposure to indoor air pollution and a family history of asthma or atopy

have all been implicated in a more severe disease outcome.

Respiratory syncytial virus commonly causes upper respiratory tract infections (URTI),

which are characterised by rhinitis, cough and occasionally, fever. These indicators of

URTI usually precede those of the lower respiratory tract which can be characterised by

dyspnoea and difficulty in feeding. It has been postulated that infection of the airway

epithelia by RSV initiates an inflammatory response characteristic of bronchiolitis

through the release of initial inflammatory mediators such as tumour necrosis factor-α

(TNF-α), IL-8 and eotaxin (McNamara et al. 2004; McNamara et al. 2005). Although

studies have been performed to elucidate the specific contribution of the AECs using

murine models or immortalised cell lines, none have been able to accurately reflect the

actual infection process occurring in vivo. Nonetheless, a recent study by Fonceca and

colleagues utilised AECs obtained from children with RSV infection to examine and

compare the viral replication processes and AEC responses between primary airway

epithelial and immortalised cell line cultures. They reported that viral replication

cytotoxicity as well as inflammatory mediator production was higher in primary airway

epithelial cultures compared to the immortalised cell line culture. Moreover, they also

observed that IL-8 response within the primary airway epithelial cultures were similar in

magnitude to the clinical samples obtained from the lungs of children with current RSV

bronchiolitis (Fonceca et al. 2012). Hence, their results indicate the suitability of using

primary airway epithelial cultures due to the similarities with in vivo observations.

Another study by McNamara and colleagues also utilised similar methods to investigate

the role and interaction of the airway epithelium with RSV infections. Results from

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their study showed an elevated expression of both B cell activating factor of the TNF

family (BAFF) mRNA and protein were observed in the bronchoalveolar lavage (BAL)

as well as bronchial brushings from RSV infected infants. Furthermore, BAFF mRNA

and protein expression were also seen following in vitro infection of both the primary

airway epithelium culture as well as an immortalised cell line culture. Their data

showed that BAFF is a consistent feature of airway infection and seeks to postulate a

possible role for the airway epithelium in supporting the protective immune responses

within the lung (McNamara et al. 2013).

Accumulating evidence over the years have presented a strong association between

recurrent RSV infections that require hospitalisations during early life and the

progression to asthma in later years (Noble et al. 1997; Sigurs et al. 2000; Sigurs et al.

2005; Wu et al. 2008; Sly et al. 2010). In addition, separate independent prospective

studies have documented that approximately 50% of children who experienced severe

RSV related bronchiolitis were eventually diagnosed with asthma (Pullan 1982;

Bacharier et al. 2012). Krishnamoorthy and colleagues, in their study, utilised a murine

model to investigate whether early life infection with RSV would lead to the

impairment of the regulatory T cell function and eventually lead to an increase in

susceptibility to allergic asthma. They observed that following sensitisation to

ovalbumin (OVA), repeated infection of the infant mice successfully induced allergic

airway disease characterised by airway inflammation, AHR and higher allergen-specific

IgE compared to uninfected sensitised mice. Their findings established the capability of

a viral pathogen to target an immune-regulatory mechanism during early life to initiate

an effect on the development of asthma in later life (Krishnamoorthy et al. 2012).

1.4.3 Human rhinovirus

Human rhinoviruses (HRV), members of the Picornaviridae family are commonly

classified into two distinct groups based on their receptor utilisation. Generally, HRV

are classified into HRV-A which consists of 74 serotypes and HRV-B, containing 25

serotypes. Of the 74 serotypes of HRV-A, 11 serotypes attain entry into the cells via the

low density lipoprotein (LDL) receptor family. The remaining HRV-A serotypes and all

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serotypes of HRV-B utilise the intercellular adhesion molecule-1 (ICAM-1) for

infection. Data from recent studies performed worldwide (McErlean et al. 2008; Jin et

al. 2009; Miller et al. 2009) have identified additional serotypes of HRV and based on

the analysis of their entire genome sequences, these new HRV strains has been

classified as a unique group now designated as HRV-C (Palmenberg et al. 2009).

Although the receptor molecule for HRV-C has only been recently identified, it had

already been suggested that this group of HRV was capable of causing symptomatic

responses following infection (Gern 2010).

Human rhinoviruses are single stranded RNA viruses often implicated as a trigger for

acute respiratory tract illness and upper respiratory tract infection as well as their

resulting complications including chronic bronchitis (Lambert and Stern 1972; Stanway

1994), sinusitis (Turner et al. 1992; Gwaltney et al. 1994) and bronchial asthma

(Lambert and Stern 1972; Minor et al. 1976; Stanway 1994; Gwaltney 1995; Johnston

et al. 1995). The use of molecular diagnostic techniques such as RT-PCR has identified

HRV as the most commonly found respiratory tract virus during asthma exacerbations

and are detected 65% of the time (Nicholson et al. 1993; Johnston et al. 1995; Wark et

al. 2002; Grissell et al. 2005). Moreover, in an epidemiologic study performed by

Khetsuriani and colleagues, the only virus types significantly associated with asthma

exacerbation in children between the age of 2 and 17 were rhinoviruses (Khetsuriani et

al. 2007). As in vivo studies of HRV infection can be ethically challenging at times,

studies have utilised AECs obtained from either healthy or donors with asthma to

establish cell cultures for investigating the effects of HRV infection. Wark and

colleagues, in their study, examined viral replication and the innate responses to HRV

infection in AECs from donors with asthma. They reported that viral RNA expression,

together with the release of viral particles into the supernatant, were greater in the

asthmatic cultures than in healthy controls. They also postulated that an impaired viral

induced interferon-beta (IFN-β) expression could be linked to an enhanced viral

replication within asthmatic cultures (Wark et al. 2005). In a separate study, Contoli and

colleagues also showed that a lack of interferon-lambda (IFN-λ) expression by HRV

infection highly correlated with the severity of HRV induced asthma exacerbations as

well as viral load in experimentally infected human volunteers (Contoli et al. 2006).

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Collectively, these observations of diminished responses by impaired innate immune

cells could thus be associated with increased susceptibility to HRV infection, leading to

increased viral replication, ultimately modulating airway inflammation through the

recruitment of various immune cells to cause an increase in the release of inflammatory

mediators, resulting in further asthma exacerbation.

As the airway epithelium is the initial contact point with the inhaled air, HRV infection

predominantly occurs within the airway epithelium of the upper respiratory tract and

occasionally, the lower respiratory tract. The lack of a suitable animal model of HRV

infection has led to studies utilising in vitro primary AECs from healthy and donors

with asthma to understand the mechanisms of airway inflammation and asthma

exacerbation following experimental HRV infection. These studies have examined the

production of pro-inflammatory substances, chemical mediators and adhesion molecules

in different cells of the lungs (Wark et al. 2005; Bochkov et al. 2010; Cakebread et al.

2011).

Major and minor serotypes of HRV are capable of infecting primary AEC cultures by

binding to the ICAM-1 or LDL receptor respectively. Production of pro-inflammatory

mediators such as IL-1α, IL-1β, IL-6, 8, 11, TNF-α, RANTES, GM-CSF would occur

following HRV infection. Although a study by Wegner et al have shown that the up-

regulation of the ICAM-1 receptor following chronic antigen challenge increases the

cell susceptibility towards HRV infection (Wegner et al. 1990), a study by Wark et al to

characterise the variability in the response of primary AECs to infection with different

HRV strains showed contrasting data (Wark et al. 2009). They reported that the minor

group HRV appeared to cause a more aggressive infection with intense release of

inflammatory mediators leading to a strengthened antiviral IFN-β response associated

with increased cell apoptosis and consequently, reduced viral replication (Wark et al.

2009). They also reported that primary AEC cultures from donors with asthma were less

capable of responding to a HRV infection possibly due to the impaired IFN-β response.

Their data suggests the existence of considerable diversity in the response to different

HRV strains, most notably between the major and minor groups (Wark et al. 2009).

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Viral infections have also been demonstrated to affect the barrier function of AECs. Yeo

and Jang examined the effects of HRV infection on nasal epithelial barrier function and

observed that post HRV infection, mRNA expression of TJ and adherens junction

proteins were reduced when compared to mRNA expression of the control group.

Protein expression was similarly observed to be reduced in the HRV infected cells

compared to non-infected control cells (Yeo and Jang 2010). They indicate that HRV

infection has the propensity to decrease expression of junctional protein complexes to

exert potentially detrimental effects on the nasal epithelial barrier function. However,

their study utilised cells obtained from the nasal passages and thus, might not accurately

reflect the bronchial airway setting following HRV infection. A study by Sajjan et al

demonstrated the capability of HRV infection in the disruption of barrier function of

polarised AECs obtained from tracheal trimmings of donor lungs during transplantation

(Sajjan et al. 2008). They reported a loss of the zonula occludens-1 (ZO-1) TJ protein

complexes following HRV infection and an increased in the paracellular permeability of

fluorescein isothiocynate-inulin (FITC-inulin), suggesting the ability of HRV to disrupt

epithelial barrier function in vitro. Although evident that HRV infection has the

proclivity to disrupt epithelial barrier function, there exists, fundamental gaps in our

understanding on the effects of HRV infection on the barrier function of an epithelium

that is already inherently dysregulated.

Of all observations on the aetiology of asthma exacerbations in children, HRV were by

far the most numerically important virus type, accounting for approximately 65% of all

infections detected. Hence, HRV infections can be considered as a major cause of

asthma exacerbation and therefore, the most appropriate virus type to utilise with which

to investigate the interaction with AECs and elucidate the pathogenesis of acute asthma

exacerbations. Although studies performed have suggested a close causal association

between respiratory viruses and asthma exacerbation (Johnston et al. 1995; Johnston et

al. 1996; Freymuth et al. 1999; Rakes et al. 1999; Chauhan et al. 2003) there are still

considerable gaps in our knowledge on the subsequent effects of viral infection, in

particular, HRV infection on the barrier integrity of the paediatric asthmatic epithelium

due mainly to the lack of suitable donor samples and to a lesser extent, a suitable

method of sampling from the respiratory airways.

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1.5 Assessing airway integrity

The interactions between viral infection, immunity, inflammation and remodelling has

been central to a plethora of asthma research over the years, however, attempts at

understanding the events occurring in the asthmatic epithelium, particularly in children,

have been severely hampered by the difficulty in obtaining relevant target organ tissue.

The use of animal models as a surrogate to comprehend the multifaceted aetiologies of

asthma has often been the topic of considerable debate (Pabst 2003; Corry and Irvin

2006; Shapiro 2006; Krug 2008; Shapiro 2008). Despite the controversy often

surrounding the use of animal models, they have proven to be an invaluable

experimental tool for the in-depth comprehension of disease mechanisms at a cellular

and molecular level which is otherwise, not possible in humans due to obvious ethical

reasons. Although currently constrained, ethically derived human samples through

either indirect or direct sampling of the airways for the establishment of cell cultures are

continuously gaining momentum. Sampling from the airways has major advantages as

well as its limitations. Hence, there is no optimal sampling or assessment process and

the choice to utilise one methodology over the other depends entirely upon the aims of

the study. Whatever the methodology employed, it is essential that the process is safe,

reliable, easily reproduced and with a high degree of sensitivity in the detection of

minor changes.

1.5.1 Animal models

Animal models of respiratory diseases are probably the most extensively used and

characterised in terms of the inflammatory and remodelling processes (Bartlett et al.

2008; Fattouh et al. 2008; Gueders et al. 2009). Models of acute and chronic diseases

have been widely used in investigating the molecular and cellular mechanisms

underlying the pathogenesis of various respiratory diseases such as asthma, cystic

fibrosis and chronic obstructive pulmonary disorder (COPD). Animal models are often

categorised into “small animal models”, comprising of rodents or rabbits and “large

animal models” which include cats, dogs, sheep and horses. Advantages of using small

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animal models include ease of handling, widespread availability and ability to remove

many of the immunological complexities of airway diseases through genetic

manipulations (Kips et al. 2003). Several studies have shown the feasibility of using

small animals such as murine models for the evaluation of allergic airway inflammation

(Malm-Erjefalt et al. 2001; Zhou et al. 2005; Bartlett et al. 2008) and airway hyper-

responsiveness (Hamelmann et al. 1997; Kline et al. 1998; Burchell 2009; Zosky 2009)

in asthma. Assessment of airway inflammation and hyper-responsiveness involves the

initial challenge of mice (previously sensitised to ovalbumin (OVA)) with OVA and

subsequently exposing them to either viral or allergen challenge. Airway hyper-

responsiveness can be assessed post viral or allergen challenge by using a low-

frequency forced oscillation technique to measure the partitioned components

representing the airways and lung parenchyma which involves the measurement of the

respiratory system input impedance (Zrs) in mice that have been anaesthetised,

tracheostomised, and ventilated (Zosky et al. 2004). Bronchoalveolar lavage (BAL) can

be collected following viral or allergen challenge for inflammatory cells or cytokines

quantification and antibody counts while blood leukocytes collected can be stimulated

with known mediators to observe for leukocyte responses such as degree of granulation

and granule morphology using transmission electron microscopy (TEM) (Malm-Erjefalt

et al. 2001). Mice lungs, bone marrow or nasal septum are also collected for

histopathological and immunohistochemical analysis of airway inflammation which

includes quantification of tissue or bone marrow leukocytes, TEM analysis of

intracellular distribution of eosinophil peroxidise activity as well as evaluating

ultrastructural features of eosinophil degranulation activity (Malm-Erjefalt et al. 2001;

Bartlett et al. 2008). Although the use of these murine models provide significant

insights into the synergistic interaction between viral infection and the immunological

response and are useful in the investigation of asthma pathogenesis and viral

exacerbations of asthma, their use in accessing the airway integrity currently remains

limited.

Large animal models such as sheep often provide important insights into structural,

anatomical and physiological changes which are relevant to the human airways such as

assessing changes in lung morphology and function, bronchial responsiveness and

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airway inflammation (Hein and Griebel 2003; Koumoundouros et al. 2006; Kirschvink

2008; Krug 2008). For example, allergic airway responses in sheep, either acquired

naturally or through experimental inoculation with live Ascaris suum (Abraham et al.

1983), have been used for sustained investigations of lung function decline following

repeated exposure to the allergen (Koumoundouros et al. 2006). In addition,

bronchoscopic and bronchoalveolar lavage examination in post allergen challenged

large animals have demonstrated an increase of inflammatory cells such as neutrophils,

eosinophils and macrophages in the airways (Bosse 1987; Abraham et al. 1988).

Despite the ability to conduct long-term studies allowing for concurrent intra-subject

evaluation of functional, immunological and morphological changes, large animal

models are often extremely costly and have few, or limited immunological and

molecular probes for characterising allergic airway responses (Zosky and Sly 2007).

Currently, there is no large or small animal model that completely recapitulates the

anatomical characteristics of the human asthma disease, hence, the study design and

ultimate parameter outcomes will dictate the suitability of the animal model to be

utilised. Given the current ethical and biological constraints that limit human

investigation, the use of animal models to critically comprehend the mechanisms

underlying the onset of asthma will continue to be an invaluable tool bridging the gap

between the outcomes of in vitro cellular results and the extrapolation into the human

disease setting.

1.5.2 Cell culture models

In vitro models utilising immortalised cell lines or human derived primary AEC are

often the preferred method in understanding the complex and varied functions of the

asthmatic airway epithelium. Although lung or bronchial epithelial cell lines such as

A549, BEAS-2B and 16HBE14o- are readily available as they proliferate and divide

continuously, they seldom exhibit characteristics commonly observed in the in vivo

setting. Human derived primary AECs are often obtained through various indirect or

direct sampling methods and despite their limited proliferative capacity, they retain

certain characteristics of the original in vivo setting as well as the ability to differentiate.

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Hence, the use of human derived primary AECs to establish cultures would be a more

accurate reflection of the traits exhibited by epithelial cells in vivo. It is beyond the

scope of this review to comprehensively discuss each sampling methodology

individually and refer the reader to either eloquent reviews or key original articles

where relevant.

Indirect sampling involves the measurements of airway inflammation through sputum

(Pin 1992; Brightling et al. 2000), peripheral blood (Dahl 1993) or urine (Green et al.

2004). Direct sampling includes lungs unsuitable for transplants or cadaver sources

(Hackett 2008; Hackett et al. 2009), biopsies (De Jong et al. 1993), bronchoscopy and

its associated methods such as guided or non-bronchoscopy guided cytology bronchial

brushings. Primary AECs obtained from these direct sampling methods are routinely

cultured in submerged monolayers or at air-liquid interface (ALI) if a more

physiologically similar model is required (Bayram 1998; Devalia et al. 1999; Bucchieri

2002; Lordan et al. 2002; Doherty et al. 2003). These culture models have the

advantage of flexibility, ability to alter experimental conditions and increased

opportunities for investigating cellular mechanisms, application of therapeutic

intervention and epithelial barrier integrity assessment when compared to animal

models. In addition, they also allow for the study of epithelial cell function without

interference from other cell types or tissues such as macrophages, fibroblasts and

immune cells.

Primary AECs grown at air-liquid interface cultures (Figure 1.4) utilise a specialised

defined medium to derive a differentiated phenotype. These cultures often exhibit a

pseudostratified, polarised phenotype expressing ciliated and goblet cells, together with

mucous production (Jiang et al. 2001; Chan et al. 2010). Development of high

transepithelial electrical resistance (TEER) in fully differentiated cultures provides an

indirect measurement of the formation of junctional adhesions among cells and is often

used as an indicator of epithelial layer disruption (Pedemonte 1995). Human derived

primary AEC cultures have been utilised in numerous studies to investigate the

differences between healthy and asthmatic donors. Cultured epithelial cells obtained

from asthmatic donors have displayed augmented gene expression associated with

Figure 1.4: Schematic of air-liquid interface culture process. Cells are initially seeded into culture flasks (A) and upon confluence, expansion of

submerged culture is performed where cells are seeded onto the apical surface of a semi-permeable membrane of a cell culture insert and exposed to

culture medium on both apical and basolateral side (B). Once confluence is attained in the Expansion phase, the cells are ‘air-lifted’ where the medium

is only supplied to the basolateral chamber while the apical surface of the culture insert is exposed to air (C). This culture method mimics the

conditions found within the human airway and initiates differentiation towards a muco-ciliary phenotype. Differentiated cultures which exhibit a

pseudostratified epithelium are obtained following 21- 28 days incubation at air-liquid interface and can be maintained for extended lengths of time of

up to 6 months.

Pre-expansion Expansion

(Submerged) Differentiation

Submerged cultures Air-Liquid Interface (ALI) cultures

Paediatric pAECs in culture flasks

Expansion into culture inserts

‘Air-lift’ into air-liquid interface phase

A B C

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inflammation, repair and remodelling and have been shown to have an increased

proliferative capacity but a diminished rate of repair of a mechanical wound when

compared to cells obtained from a healthy donor (Kicic et al. 2006; Stevens et al. 2008;

Kicic 2010). Asthmatic epithelial cells, when cultured at air-liquid interface, have

shown to exhibit a less differentiated phenotype expressing more basal cells with

diminished junctional complexes formation (Hackett et al. 2011; Xiao et al. 2011).

Recent published reports have shown conflicting data between normal and asthmatic

cells. In their study, Hackett and colleagues (Hackett et al. 2011) reported no significant

differences in the TEER between normal and asthmatic cultures, however, in another

study, Xiao and co-workers have shown that asthmatic cultures had a lower TEER as

well as loss of TJs (Xiao et al. 2011). These differences may be due to the age of the

donors (young adults or children in the Hackett study versus adults in the Xiao study)

and / other source from which the epithelial cells were obtained (post mortem lungs

versus bronchial brushings). Although having differing observations, these studies have

served to highlight the importance and versatility of utilising culture AECs in the

investigation of respiratory epithelial barrier integrity.

Despite these discrepancies, the current literature suggests that primary AECs from

asthma donors, when cultured either monolayer or ALI, present with an intrinsically

different phenotype from healthy donors (Kicic et al. 2006; Stevens et al. 2008; White

et al. 2008; Kicic 2010; Hackett et al. 2011; Xiao et al. 2011). This further supports the

use of human derived primary AEC cultures in asthma research. However, despite the

accumulating data demonstrating that human derived primary AEC cultures can provide

a unique insight into the asthmatic epithelium, a vast majority of studies have

predominantly utilised cells obtained from adult donors Although crucial in allowing

the characterisation of the asthmatic phenotype, data generated from these studies are

less supportive when attempting to dissect and comprehend the underlying mechanistic

changes occurring within the paediatric asthmatic epithelium. At present, no study has

been attempted to investigate epithelial barrier integrity in paediatric asthma,

furthermore, there remains a paucity of data on paediatric airway epithelial barrier

integrity following viral exacerbations and the sustained effects on epithelial function.

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1.6 Summary

Asthma is among the most common, incurable, chronic conditions worldwide, affecting

both children and young adults and is estimated to affect 300 million people worldwide.

Often refractory to treatment in many and severe in a large number of the population,

asthma has traditionally been linked to an atopy, TH2-type T-lymphocyte cell driven

process which contributes to increasing chronicity of the inflammatory response. In

addition, there has been considerable emphasis on the role of the immune system in

asthma and as atopic asthma has generally been recognised as the most common asthma

phenotype, the most widely accepted concept is one of inflammation associated with

atopy. Consequently, majority of the research directed at comprehending asthma has

focussed on the development of atopy and the association with various clinical markers

of asthma such as airway hyper-responsiveness and inflammation. However, a succinct

review of the literature have reported that the TH2 inflammation paradigm have yet to

fully account for the pathobiology of asthma, thus suggesting the possibility of other

cellular and molecular mechanisms contributing towards asthma heterogeneity.

A recent shift in research focus has emphasised how the respiratory epithelium could

contribute to asthma heterogeneity. Over the decades, data obtained have seen a change

in the understanding of the role of the respiratory epithelium, from being a static

physical barrier against foreign invading particles, to one that is dynamic, versatile and

constantly reacting to the ever changing cascades of insults from the external

environment. Although evidence has shown that an abnormal epithelium is commonly

observed in asthma, relatively few studies have addressed whether these observed

abnormalities are actually intrinsic to asthma or a consequence of airway inflammation.

Current evidence has demonstrated that the asthmatic epithelium has intrinsic properties

such as vulnerability to injury, the ability to initiate and modulate inflammatory

responses, aberrant repair as well as an increased susceptibility to environmental

pathogens, which could contribute to the pathophysiology of asthma. Regulation of the

paracellular passage in the airway epithelium is achieved via TJ complex formation and

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through the limited studies performed, evidence have demonstrated compromised

integrity, disordered assembly as well as diminished expression of TJ complexes within

the adult asthmatic epithelium. However, there remains a paucity of comprehensive

assessments of epithelial TJ expression and barrier function within the paediatric

asthmatic epithelium.

In addition, respiratory viral infections, and in particular, human rhinovirus (HRV)

infection, which accounts for the majority of acute asthma exacerbations, have also been

shown to be associated with the disruption of specific airway epithelial TJ in healthy,

adult airway epithelium, resulting in increased permeability to environmental irritants,

antigens and pathogens. Although a seminal study have shown the translocation of

bacteria across the epithelial layer at points of TJ disassembly following HRV infection,

it remains unknown whether the epithelium in asthma is more susceptible to TJ

disassembly following HRV infection, thereby facilitating sensitisation from inhaled

haptens, allergens or pathogens. Collectively, the review of the literature has identified

significant gaps in the evidence regarding the expression of epithelial TJ and barrier

function within the paediatric airway epithelium in the presence or absence of atopy and

/ or asthma and most importantly, the impact of HRV infection on epithelial TJ

expression and barrier function. This provides the justification and rationale for the

following hypotheses.

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1.7 Hypotheses and research aims

To address the gaps within the evidence, this project seeks to examine the hypotheses

that, (1) epithelial barrier function is defective in children with asthma and (2) that this

defective barrier function in asthma is independent of atopy. Moreover, this project also

investigates the hypothesis that (3) the barrier function and epithelial integrity is

compromised to a greater extent by HRV in the asthmatic airway compared to healthy

airway. Therefore, the specific aims of the experiments described in this thesis were to:

1. Compare barrier characteristics of epithelium from healthy non-atopic (HNA),

healthy atopic (HA), non-atopic asthmatic (NAA) and atopic asthmatic (AA)

paediatric subjects.

2. Determine the effects of HRV infection on barrier characteristics of epithelium

from healthy non-atopic (HNA), healthy atopic (HA) non-atopic asthmatic

(NAA) and atopic asthmatic (AA) paediatric subjects.

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CHAPTER 2: General Materials and Methods

2.1 General Materials

All general reagents and chemicals utilised in the investigation are listed with their

supplier below. Specific materials are listed in their relevant chapters.

Material name, Supplier, Supplier’s origin (City/Town; State/County, Country).

0.22 µM filter, PALL, East Hills, NY, USA

25G & 27G needles, TERUMO, Macquaire Park, NSW, Australia

2 β-Mercaptoethanol, SIGMA, St. Louis, MO, USA

2 mM deoxyribonucleotide triphosphates (dNTPs), APPLIED BIOSYSTEMS, Foster

City, CA, USA

4′,6-diamidino-2-phenylindole (DAPI), SIGMA, St. Louis, MO, USA

4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), SIGMA, St. Louis, MO,

USA

10X RT-buffer, APPLIED BIOSYSTEMS, Foster City, CA, USA

Bovine pituitary extract (BPE), SIGMA, St. Louis, MO, USA

Bovine serum albumin (BSA), SIGMA, St. Louis, MO, USA

Bronchial epithelium basal medium (BEBM), LONZA™, Basel, Switzerland

Calcium chloride (CaCl2), SIGMA, St. Louis, MO, USA

Citric acid, SIGMA, St. Louis, MO, USA

Collagen S (Type 1), BD Biosciences, Franklin Lakes, NJ, USA

Dimethyl sulfoxide (DMSO), SIGMA, St. Louis, MO, USA

DRAQ5™ stain, BIOSTATUS, Shepshed, Leicestershire, UK

Dulbecco’s Modified Eagle Medium-High Glucose (DMEM-HG), INVITROGEN,

Melbourne, VIC, Australia

Eagle’s Minimum Essential Medium (EMEM), INVITROGEN, Melbourne, VIC,

Australia

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Epidermal growth factor (EGF), SIGMA, St. Louis, MO, USA

Epinephrine, SIGMA, St. Louis, MO, USA

Ethanol, LOMB Scientific, Taren Point, NSW, Australia

Ethylene diamine tetraacetic acid (EDTA), SIGMA, St. Louis, MO, USA

Ethylene glycol tetraacetic acid (EGTA), SIGMA, St. Louis, MO, USA

Fibronectin, SIGMA, St. Louis, MO, USA

Fluorescence mounting media, DAKO, Glostrup, Denmark

Fluorescein isothiocyanate-dextran 4 (FITC-dextran 4), SIGMA, St. Louis, MO, USA

Fluorescein isothiocyanate-dextran 20 (FITC-dextran 20), SIGMA, St. Louis, MO, USA

Foetal calf serum (FCS), SIGMA, St. Louis, MO, USA

Formalin (40% aqueous solution of formaldehyde), SIGMA, St. Louis, MO, USA

Fungizone, INVITROGEN, Melbourne, VIC, Australia

Gentamicin, INVITROGEN, Melbourne, VIC, Australia

Glucose powder, SIGMA, St. Louis, MO, USA

Glycerol, SIGMA, St. Louis, MO, USA

Glycine, SIGMA, St. Louis, MO, USA

Heparin sodium, MAYNE PHARMA, Mulgrave, VIC, Australia

Hydrochloric Acid (HCl) (32%), UNIVAR, Ingleburn, NSW, Australia

Hydrocortisone, SIGMA, St. Louis, MO, USA

Insulin, SIGMA, St. Louis, MO, USA

Inulin-FITC, SIGMA, St. Louis, MO, USA

Isopropyl alcohol, SIGMA, St. Louis, MO, USA

L-glutamine, INVITROGEN, Melbourne, VIC, Australia

Magnesium chloride (MgCl2), SIGMA, St. Louis, MO, USA

Magnesium sulphate (MgSO4), SIGMA, St. Louis, MO, USA

Methanol, ANALYTICAL SCIENCES, Patumwan, Bangkok, Thailand

Minimal Essential Medium (MEM), INVITROGEN, Melbourne, VIC, Australia

MEM Non-essential amino acids, GIBCO, Melbourne, VIC, Australia

Multiscribe, APPLIED BIOSYSTEMS, Foster City, CA, USA

Nalgene 1°C Mr Frosty freezing container, WESSINGTON CRYOGENICS, Houghton-

le-Spring, Tyne & Wear, UK

Nystatin, INVITROGEN, Melbourne, VIC, Australia

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Odyssey Blocking Buffer, LI-COR Biosciences, Lincoln, NE, USA

Penicillin Streptomycin (Pen Strep), INVITROGEN, Melbourne, VIC, Australia

Potassium chloride (KCl), SIGMA, St. Louis, MO, USA

Potassium dihydrogen phosphate (KH2PO4), BDH Lab. Supplies, Poole, Dorset, UK

Protease inhibitor cocktail, SIGMA, St. Louis, MO, USA

Proteinase K, SIGMA, St. Louis, MO, USA

Random Hexamers, APPLIED BIOSYSTEMS, Foster City, CA, USA

Recombinant human epidermal growth factor (EGF), SIGMA, St. Louis, MO, USA

RNase-free DNase, QIAGEN, Hilden, Germany

RNase inhibitor, APPLIED BIOSYSTEMS, Foster City, CA, USA

RPMI-1640 media, INVITROGEN, Melbourne, VIC, Australia

Sapphire 700™ stain, LI-COR Biosciences, Lincoln, NE, USA

Sodium bicarbonate (NaHCO3), SIGMA, St. Louis, MO, USA

Sodium chloride (NaCl), SIGMA, St. Louis, MO, USA

Sodium deoxycholate, SIGMA, St. Louis, MO, USA

Sodium dihydrogen phosphate (NaH2PO4), SCHARLAU CHEMIE S.A, Barcelona,

Spain

Sodium dodecyl sulphate (SDS), SIGMA, St. Louis, MO, USA

Sodium fluoride (NaF), SIGMA, St. Louis, MO, USA

Sodium hydroxide (NaOH), SIGMA, St. Louis, MO, USA

Sodium phosphate dibasic (Na2HPO4), SIGMA, St. Louis, MO, USA

Sodium pyrophosphate, SIGMA, St. Louis, MO, USA

Sodium pyruvate, SIGMA, St. Louis, MO, USA

Sodium orthovanadate (Na3VO4), SIGMA, St. Louis, MO, USA

Sudan-Black B, SIGMA, St. Louis, MO, USA

SYBR® Green PCR Master Mix, APPLIED BIOSYSTEMS, Foster City, CA, USA

Syringe (1 ml / 5 ml / 10 ml), TERUMO, Macquaire Park, NSW, Australia

Trans-retinoic acid, SIGMA, St. Louis, MO, USA

Transferrin powder, SIGMA, St. Louis, MO, USA

Tri-iodothyronine, SIGMA, St. Louis, MO, USA

Triton-X 100, SIGMA, St. Louis, MO, USA

Trizma base, SIGMA, St. Louis, MO, USA

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Trypan-Blue, SIGMA, St. Louis, MO, USA

Trypsin, SIGMA, St. Louis, MO, USA

Trypsin/Ethylenediaminetetraacetic acid (EDTA), SIGMA, St. Louis, MO, USA

Trypsin Neutralising Solution, LONZA, Basel, Switzerland

Tween-20, ICN BIOMEDICALS, Irvine, CA, USA

Ultroser-G, PALL BIOSEPRA, Cergy-Saint Christophe, France

2.2 Antibodies

2.2.1 Primary antibodies

Rabbit Anti-human Claudin (1:200), Life Technologies, VIC, Australia

Rabbit Anti-human Occludin (1:200), Life Technologies, VIC, Australia

Polyclonal Rabbit Anti-human Zonula occluden-1 (1:200), Life Technologies, VIC,

Australia

2.2.2 Secondary antibodies

Goat Anti-Rabbit IgG FITC Conjugate (1:100), SIGMA, St. Louis, MO, USA

Goat Anti-Mouse IgG FITC Conjugate (1:100), SIGMA, St. Louis, MO, USA

Goat Anti-Mouse IRDye® 800CW (1:800), LI-COR Biosciences, Lincoln, NE, USA

Goat Anti-Rabbit IRDye® 800CW (1:800), LI-COR Biosciences, Lincoln, NE, USA

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2.3 General Equipment

2.3.1 Autoclave

Sterilisation using an Atherton autoclave (Thornbury, VIC, Australia) of equipment was

performed at 121°C for 45 min and solutions at 121°C for 40 min when required.

2.3.2 Balances

All analytical and biochemical reagents were weighed out using an Ohaus Explorer®

Balance (Derrimut, VIC, Australia).

2.3.3 Bronchoscope

Bronchoscopy guided bronchial sampling was carried out using a Pentax® FI-10RBS

portable bronchoscope.

2.3.4 Bronchial brush

All bronchial brushings were collected using Olympus® BC-25105 brushes of 10mm

length and 2mm outer diameter (Macquarie Park, NSW, Australia).

2.3.5 Centrifuges

Centrifugation was performed using either an Eppendorf 5810R refrigerated Swing

Bucket Rotor or a 5415D mini-centrifuge (Hamburg, Germany). Cytospin

centrifugation was performed using a Hettich centrifuge from Andreas Hettich GmbH

and Company KG (Tuttlingen, Baden-Württemberg, Germany).

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2.3.6 Glassware

General glassware was procured from Schott (Frenchs Forest, NSW, Australia) and

Corning (Mount Martha, VIC, Australia). All glassware was washed in detergent

overnight, rinsed three times in tap water and once in deionised water. All equipment

used for culture purposes was sterilised in an autoclave.

2.3.7 Heating devices

Heating of samples or reagents to temperatures between 37°C and 100°C was

performed using a RATEK heating block (Boronia, VIC, Australia).

2.3.8 Incubators

All established mycoplasma-free cell cultures were maintained in a Panasonic CO2

incubator (Murarrie, QLD, Australia) in an atmosphere of 5% CO2 / 95% air. All

mycoplasma-free cell line cultures were maintained in a separate, identical incubator

with the same atmospheric conditions.

2.3.9 Infrared scanner

Detection of signal on microplates from In-Cell Western™ assays was achieved using a

LI-COR Odyssey infrared scanner (Lincoln, NE, USA). Scans were performed at 700

nm and 800 nm wavelengths as required. Quantification and data analysis was

performed using the associated Odyssey v.3.0 software.

2.3.10 Laminar flow cabinets

All cell culture was performed in a National Association of Testing Authorities (NATA)

Certified Laminar Flow Cabinet from AES Environment (Balcatta, WA, Australia).

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2.3.11 Microscope

A Leica Microsystems GmbH inverted microscope (Wetzlar, Hesse, Germany) and

Nikon® Eclipse Ti inverted microscope (Coherent Scientific, Hilton, Australia) with an

attached camera was used to observe cellular morphology and cell viability. A mercury

lamp attachment on the Nikon® Eclipse Ti inverted microscope was used to observe

fluorescently stained antibody conjugates on slides.

2.3.12 pH meter

A 3310 pH meter from Jenway (Gransmore Green Felsted Dunmow, Essex, England)

was used for all pH measurements. Calibration solutions were obtained from Scharlau

(Barcelona, Catalonia, Spain).

2.3.13 Pipettes

All volumes between 1 and 25 ml were measured using a Powerpette from Jencons

(Leighton Buzzard, Bedfordshire, England) and S1 Pipet Fillers from Thermoscientific

(Wilmington, DE, USA). Gilson micropipettes (Middleton, WI, USA) were used to

measure all volumes less than 1 ml. Finnpipette® multi-channels from Thermo

Labsystems (Helsinki, South Finland, Finland) were also used for work involving 96-

well microplates.

2.3.14 Plate readers

All spectrophotometric measurements between 400 nm and 600 nm were performed

using a Thermoscientific Multiskan FC microplate photometer (Wilmington, DE, USA).

Fluorescence measurements with excitation and emission wavelengths were performed

using a PerkinElmer Enspire® multilabel plate reader (Melbourne, VIC, Australia).

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2.3.15 Real Time Quantitative PCR (RT-qPCR)

Real time quantitative PCR (RT-qPCR) was performed on an Applied Biosystems ABI

Prism® 7300 (Foster City, CA, USA). Data analysis was performed using the software

program Sequence Detection System 1.9. A PTC-100 Thermal cycler from MJ Research

was used for reverse transcription of RNA to cDNA (Boston, MA, USA).

2.3.16 Semi-dry Western Blot Transfer

The Invitrogen iBlot® Transfer System (Melbourne, VIC, Australia) was used to

perform all semi-dry transfer of proteins from eletrophoresed gels during Western blots.

Pre-made “TOP Stack” consisting of copper cathode and “BOTTOM Stack” consisting

of copper anode and PVDF membrane purchased from Invitrogen® was used to perform

the transfer as per the manufacturer’s recommendations.

2.3.17 Spectrophotometer

A Thermoscientific NanoDrop 2000C spectrophotometer was used to assess the quality

and quantity of extracted ribonucleic acid samples (Wilmington, DE, USA).

2.3.18 Stirrer, shakers and rockers

For the agitation and mixing of solutions, a stirrer (Industrial Equipment and Control

PTY LTD, Melbourne, VIC, Australia), Ratek shaker (Boronia, VIC, Australia) IKA

vortex (Petaling Jaya, Malaysia) or Stuart® rocker (Barloworld Scientific Laboratory

Group, Rochester, NY, USA) were used.

2.3.19 Tissue culture and general plastic ware

All disposable plastic culture equipment was obtained from Sarstedt (Adelaide, SA,

Australia) or BD Biosciences (San Jose, CA, USA).

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2.3.20 Water bath

When specified, certain samples or reagents were thawed or warmed using a

Thermoline water bath (Smithfield, NSW, Australia).

2.4 General Buffers and Solutions

Where appropriate, solutions were sterilised either by autoclaving for 20 minutes at

120°C at 15 pounds per square inch or passed through a 0.22 µM filter.

2.4.1 General purpose

2.4.1.1 Double deionised water (ddH2O)

Double deionised water was prepared by passing distilled water through a Milli-Q water

purification system (Millipore, North Ryde, NSW, Australia).

2.4.1.2 Ethanol (95% v/v)

To make 1000 ml of 95% (v/v final) of ethanol, 950 ml of absolute ethanol was added

to 50 ml of ddH2O. The solution was stored at room temperature (RT) until required.

2.4.1.3 Ethanol (80% v/v)


to 200 ml of ddH2O. The solution was stored at RT until required.

2.4.1.4 Ethanol (70% v/v)


to 300 ml of ddH2O. The solution was stored at RT until required.

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2.4.1.5 Hank’s Balanced Salt Solution (HBSS)

To make 1000 ml of HBSS, 0.185 g of CaCl2, 0.097 g of MgSO4, 0.4 g of KCl, 0.06 g of

KH2PO4, 8 g of NaCl, 0.047 g of Na2HPO4 and 1 g of glucose was dissolved in 900 ml

of ddH2O. Solution pH was adjusted to 7.4 and the volume was made up to 1000 ml by

adding ddH2O. The buffer was autoclaved (Refer to 2.3.1) and stored at RT until use.

2.4.1.6 (HEPES) Buffered Saline Solution

A 10X stock of 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) was

prepared by dissolving 47.6 g of HEPES, 70.7 g of NaCl, 2 g of KCl, 1.7 g of glucose

and 10.2 g of Na2HPO4 in 800 ml of ddH2O. The solution pH was adjusted to 7.4 and

the volume was made up to 1000 ml by adding ddH2O. The buffer was autoclaved as

required and stored at RT until use. The stock solution was diluted 1 part to 9 parts

ddH2O before use.

2.4.1.7 Hydrochloric acid (HCl; 10mM)

To make a 10 mM HCl solution, 10 µl of 32% HCl was diluted in 9.99 ml of ddH2O to

a final volume of 10 ml and stored at RT.

2.4.1.8 Hydrochloric acid (HCl; 4mM)

To make a 4 mM HCl solution, 17 µl of 32% HCl was diluted in 50 ml of ddH2O and

stored at RT.

2.4.1.9 Neutral Buffered Formalin (NBF)

To make 1000 ml of NBF, 900 ml of ddH2O and 100 ml of formalin were combined

with 4 g of NaH2PO4 and 6.5 g of Na2HPO4. The solution was stored at 4°C until

required.

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2.4.1.10 Phosphate Buffered Saline (PBS)

A 10X solution of PBS was initially prepared by dissolving 80 g of NaCl, 2 g of KCl,

14.4 g of Na2HPO4 and 2.4 g of KH2PO4 into 1000 ml of ddH2O. The solution was then

diluted 1 part into 9 parts ddH2O for use. The PBS solution was autoclaved (Refer to

2.3.1) to ensure solution sterility for cell culture purposes.

2.4.1.11 Sodium dodecyl sulphate (SDS) solution (20% w/v)

To make 100 ml of 20% (w/v) SDS solution, 20 g of SDS was dissolved in 100 ml of

ddH2O using a magnetic stirrer and stored at RT.

2.4.1.12 Sodium fluoride solution (NaF, 1M)

To make 10 ml of 1 M NaF solution, 0.42 g of NaF was dissolved in 10 ml of ddH2O

and stored at RT until required.

2.4.1.13 Sodium orthovanadate solution (200 mM)

To make 50 ml of 200 mM sodium orthovanadate solution, 1.85 g of sodium

orthovanadate was dissolved in 50 ml of ddH2O using a magnetic stirrer and stored at

RT.

2.4.1.14 Sodium deoxycholate (10% w/v)

To make 100 ml of 10% (w/v) sodium deoxycholate, 10 g of sodium deoxycholate was

dissolved in 100 ml of ddH2O using a magnetic stirrer and stored at RT.

2.4.1.15 Tris Buffered Saline (TBS)

A 10X solution of TBS was prepared by dissolving 80 g of NaCl, 2 g of KCl and 30 g

of Trizma Base into 1000 ml of ddH2O and the pH adjusted to 7.4. The solution was

then diluted 1 part into 9 parts ddH2O for use and stored at RT.

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2.4.1.16 Tris-Hydrochloric acid (HCl; 0.5 M)

To make 100 ml of 0.5 M Tris-HCl solution, 6 g of Trizma-Base was dissolved and

made up to a final volume of 100 ml of ddH2O. The pH was adjusted to 6.8 and stored

at 4°C until required.

2.4.1.17 Triton X-100 (10% v/v)

To make 100 ml of 10% (v/v final) Triton X-100 permeabilisation solution, 10 ml of

Triton-X stock solution was added to 90 ml of 1X PBS (refer to 2.4.1.10). The

permeabilising solution was kept at RT and away from direct sunlight until required.

2.4.1.18 Tween-20 (20% v/v)

To make 100 ml of 20% (v/v final) Tween-20 wash solution, 20 ml of Tween-20 stock

solution was added to 80 ml of 1X PBS (refer to 2.4.1.10). The wash solution was

subsequently kept at RT and away from direct sunlight until required.

2.4.1.19 Carnoy’s Fixative Solution

To make 100 ml of Carnoy’s fixative solution, 60 ml of 100% ethanol was added to 30

ml of chloroform and 10 ml of glacial acetic acid. The fixative solution was

subsequently kept at RT and away from direct sunlight until required.

2.4.2 Cell culture

2.4.2.1 Media additives

2.4.2.1.1 Bovine pituitary extract (BPE)

A 100mg/ml stock of BPE was made by dissolving 10 g of BPE powder into 100 ml of

1X HEPES buffered saline solution (refer to 2.4.1.5). The solution was centrifuged at

10000g for 30 min, the supernatant collected, re-centrifuged and filter-sterilised before

being stored at -20°C.

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2.4.2.1.2 Bovine serum albumin (BSA) stock solution (1 mg/ml)

A 1 mg/ml stock solution of BSA was prepared by dissolving 100 mg of BSA powder

into 100 ml of 1X PBS (refer to 2.4.1.10). The solution was filter-sterilised before being

stored at -20°C.

2.4.2.1.3 Epidermal growth factor (EGF) (25 µg/ml)

A 25 µg/ml stock of EGF was made by dissolving 500 µg of EGF powder into 2 ml of

BSA (refer to 2.4.2.1.2) and 18 ml of 1X HEPES buffered saline solution (refer to

2.4.1.5). The solution was then filtered-sterilised before being stored at -20°C.

2.4.2.1.4 Epinephrine (1 mg/ml)

A 1 mg/ml stock of epinephrine was made by dissolving 50 mg of epinephrine powder

into a final volume of 50 ml of 10 mM HCl solution (refer to 2.4.1.7). The solution was

then filter-sterilised before being stored at -20°C.

2.4.2.1.5 Hydrocortisone (3.6 mg/ml)

To make 3.6 mg/ml stock of hydrocortisone, 72 mg of hydrocortisone powder was

dissolved into a final volume of 20 ml of 95% (v/v) ethanol (refer to 2.4.1.2). The

solution was then filtered-sterilised before being stored at -20°C.

2.4.2.1.6 Insulin (2 mg/ml)

To make a 2 mg/ml stock of insulin, 100 mg of insulin powder was dissolved into 50 ml

of 4 mM HCl (refer to 2.4.1.8). The solution was then filter-sterilised before being

stored at -20°C.

2.4.2.1.7 Trans-retinoic acid (1 µg/ml)

A 1 µg/ml stock of trans-retinoic acid was made by dissolving 50 mg of trans-retinoic

acid powder into 5 ml of dimethyl sulfoxide (DMSO). The solution was then filter-

sterilised before being stored at -20°C.

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2.4.2.1.8 Transferrin (5 mg/ml)

A 5 mg/ml stock of transferrin was made by dissolving 100 mg of transferrin powder

and 2 ml of BSA (refer to 2.4.2.1.2) into 18 ml of 1X HEPES buffered saline solution

(refer to 2.4.1.5). The solution was then filter-sterilised before being stored at -20°C.

2.4.2.1.9 3, 3’5-Triiodo-L-thyronine sodium salt stock (6.5 µg/ml)

A 6.5 µg/ml stock solution of 3, 3’5-Triiodo-L-thyronine sodium salt stock was made

by dissolving 100 mg of 3, 3’5-Triiodo-L-thyronine sodium salt powder into 1.54 ml of

DMSO. The solution was then filter-sterilised before being stored at -20°C. The

solution was diluted 1 part to 99 parts 1X HEPES buffered saline solution (refer to

2.4.1.5) before use.

2.4.2.1.10 Ultroser-G

To reconstitute the Ultroser-G serum supplement, 20 ml of sterile ddH2O was added to

the powder and dissolved with periodic agitation for 10 min at RT under sterile

conditions. The solution was then aliquoted into 4 ml vials and stored at 4°C until

required.

2.4.2.2 Cell culture media

2.4.2.2.1 16HBE14o- cell line culture medium

The 16HBE14o- cell lines were maintained in Minimum Essential Media (MEM)

containing Earle’s Salts, FCS (10% v/v final), L-Glutamine (1% v/v final) and

penicillin/streptomycin (1% v/v final). The components were added to a final volume of

500 ml of MEM under sterile conditions and the final solution stored at 4°C until

required.

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2.4.2.2.2 A549 cell line culture medium

A549 cell lines were maintained in RPMI-1640 containing FCS (10% v/v final),

gentamicin (1% v/v final) and penicillin/streptomycin (1% v/v final). The components

were added to a final volume of 500 ml of RPMI-1640 under sterile conditions and the

final solution stored at 4°C until required.

2.4.2.2.3 Caco-2 cell line culture medium

Caco-2 cell lines were maintained in Dulbecco’s Modified Eagle Medium-High

Glucose (DMEM-HG) containing FCS (10% v/v final), MEM non-essential amino acids

(1% v/v final) and penicillin/streptomycin (1% v/v final). The components were added

to a final volume of 500 ml of DMEM-HG under sterile conditions and the final

solution stored at 4°C until required.

2.4.2.2.4 HeLa Ohio cell line culture medium

HeLa Ohio cell lines were maintained in Eagle’s Minimum Essential Medium (EMEM)

containing FCS (10% v/v final), L-glutamine (1% v/v final), non-essential amino acids

(1% v/v final), penicillin/streptomycin (0.5% v/v final), sodium pyruvate (0.2% v/v

final). The components were added to a final volume of 500 ml of EMEM under sterile

conditions and the final solution stored at 4°C until required.

2.4.2.2.5 MRC-5 cell line culture medium

MRC-5 cell lines were maintained in Eagle’s Minimum Essential Medium (EMEM)

containing FCS (5% v/v final), L-glutamine (1% v/v final), non-essential amino acids

(1% v/v final), penicillin/streptomycin (0.5% v/v final), sodium pyruvate (0.2% v/v

final). The components were added to a final volume of 500 ml of EMEM under sterile

conditions and the final solution stored at 4°C until required.

2.4.2.2.6 NuLi-1 and primary airway epithelial cell culture growth medium

NuLi-1 modified human AECs and primary AECs were maintained in a Bronchial

Epithelial Basal Media (BEBM) containing the following additives; BPE (0.05 m/ml)

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(refer to 2.4.2.1.1), EGF (0.005 µg/ml) (refer to 2.4.2.1.3), epinephrine (0.5 µg/ml)

(refer to 2.4.2.1.4), hydrocortisone (0.5 µg/ml) (refer to 2.4.2.1.5), insulin (5 µg/ml)

(refer to 2.4.2.1.6), trans-retinoic acid (0.1 ng/ml) (refer to 2.4.2.1.7), transferrin (0.01

mg/ml) (refer to 2.4.2.1.8), 3,3’5-Triiodo-L-thyronine sodium salt stock (6.5 ng/ml)

(refer to 2.4.2.1.9), gentamicin (0.05 mg/ml), penicillin/streptomycin (20 U/ml),

fungizone (0.125 µg/ml), and Ultroser-G (2% v/v final) (refer to 2.4.2.1.10). The

components were added to a final volume of 500 ml of BEBM under sterile conditions

and the final solution stored at 4°C until required.

2.4.2.3 General purpose cell culture solutions

2.4.2.3.1 Fibronectin coating buffer

To make cell culture coating buffer, 1 mg of fibronectin was diluted in 10 ml of BEBM

at 37°C for 60 min to completely dissolve the powder. To this, 1 ml of collagen S and

10 ml of BSA stock (refer to 2.4.2.1.2) were added to a final volume of 100 ml of

BEBM under sterile conditions. The solution was filter-sterilised before use and stored

at 4°C away from light until required.

2.4.2.3.2 Cryopreservation medium

To make 1 ml of cryopreservation medium, 5% (v/v final) of DMSO and 25% (v/v

final) of FCS were added to culture media in which the cells were grown (refer to

2.4.2.2.1 – 2.4.2.2.5). NuLi-1 cryopreservation medium was made up of 10% (v/v final)

of DMSO, 30% (v/v final) of FCS added to culture media in which the cells were grown

(refer to 2.4.2.2.6). This solution was only used for the freezing down and storage of

immortalised cell lines and modified primary AECs as primary AECs cannot be

successfully frozen on a routine basis.

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2.4.2.3.3 Cell protein extraction buffer (CEB)

To make 100 ml of CEB, 1 ml of 0.5 M Tris-HCl (pH 6.8, refer to 2.4.1.16), 0.59 g of

NaCl, 0.037 g of EDTA, 0.892 g of sodium pyrophosphate, 0.038 g of EGTA, 100 µl of

1 M NaF solution (refer to 2.4.1.12) and 1 ml of 200 mM sodium orthovanate solution

(refer to 2.4.1.13) was dissolved in 70 ml of ddH2O. Solution pH was adjusted to 7.4 by

the addition of HCl drop-wise. Once pH was adjusted, 1 ml of Triton X-100, 10 ml of

glycerol, 500 µl of 20% SDS solution (refer to 2.4.1.11) and 5 ml of 10% sodium

deoxycholate solution (refer to 2.4.1.14) was added and dissolved. Final volume of the

solution was made up to 100 ml with the addition of more ddH2O and then aliquoted

and stored at -20°C until required.

2.4.3 Assays and associated solutions

2.4.3.1 In-cell Western™ solutions

2.4.3.1.1 Fixative solution

To make 50 ml of fixing solution, 5 ml of formalin was added to 45 ml of 1X PBS

(refer to 2.4.1.10). This solution was made up fresh and kept at RT prior to use.

2.4.3.1.2 Triton X-100 permeabilisation solution

To make 500 ml of 0.1% (v/v final) Triton X-100 permeabilisation solution, 5 ml of an

initially prepared 10% (v/v final) Triton X-100 solution was added to 495 ml of 1X PBS

(refer to 2.4.1.10). The permeabilising solution was subsequently kept at RT and away

from direct sunlight until required.

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2.4.3.1.3 Tween-20 washing solution

To make 1000 ml of the 0.1% (v/v final) Tween-20 wash solution, 5 ml of an initially

prepared 20% (v/v final) Tween-20 solution was added to 995 ml of 1X PBS (refer to

2.4.1.10). The wash solution was subsequently kept at RT and away from direct

sunlight.

2.4.3.2 Immunocytochemistry / histochemistry solution

2.4.3.2.1 Citrate buffer

To make 1000 ml of citrate buffer, 2.1 g of citric acid and 1 g of sodium hydroxide was

dissolved in 900 ml of ddH2O and the pH adjusted to 6.0. The solution was made up to

1000 ml with ddH2O and stored at RT until required for antigen retrieval in tissue

samples during immunohistochemistry.

2.4.3.2.2 Proteinase K

Lyophilised proteinase K was reconstituted by the addition of ddH2O to a final

concentration of 10 mg/ml, aliquoted and stored at -20°C. The stock solution was

further diluted in 1X PBS (refer to 2.4.1.10) to a final concentration of 36 µg/ml before

being added to slides for antigen retrieval during immunocytochemistry.

2.4.3.2.3 Sudan Black B solution

To make 100 ml of 0.5% (w/v final) Sudan Black B, 0.5 g of Sudan Black B powder

was dissolved in 100 ml of 70% ethanol (refer to 2.4.1.4). The resulting solution was

stored at RT until use.

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2.4.3.3 Transepithelial permeability solutions

2.4.3.3.1 Fluorescein isothiocyanate-dextran (FITC-dextran)

To make 10 ml of FITC-dextran (2 mg/ml), 20 mg of FITC-dextran of either 4kDA or

20kDa molecular weight was added to HEPES-HBSS (refer to 2.4.3.3.2) in a dark room

and allowed to dissolve. The solution was then aliquoted and stored at -20°C until

required.

2.4.3.3.2 HEPES buffered Hank’s Balanced Salt Solution (HEPES-HBSS)

To make 1000 ml of HEPES-HBSS, 0.35 g of NaHCO3 and 5.96 g of 4-(2-

hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) was dissolved in 990 ml of

HBSS (refer to 2.4.1.5). Solution pH was adjusted to 7.4 and the volume was made up

to 1000 ml by adding HBSS. The buffer was autoclaved as required (refer to 2.3.1) and

stored at 4°C until required.

2.4.3.4 Western Blot buffers

2.4.3.4.1 2-(N-morpholino)ethanesulfonic acid Sodium Dodecyl Sulphate (MES

SDS) Running buffer

2-(N-morpholino)ethanesulfonic acid (MES) SDS running buffer (Life Technologies

Novex BOLT®, Melbourne, Australia) was used with all Western blot pre-cast gels

electrophoresed using the BOLT® system. 1X MES SDS running buffer was prepared

by diluting 20X BOLT® MES SDS running buffer 20-fold with the addition of ddH2O

(refer to 2.4.1.1) prior to start of Western Blot.

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2.4.3.4.2 Transfer Buffer

Transfer buffer for wet transfer in Western Blots was prepared and stored as a 10X

stock solution. Stock solution was prepared by dissolving 150 g of glycine and 30 g of

Trizma Base in 1 L of ddH2O (refer to 2.4.1.1) and stored at 4°C until required.

Working solution of 1X transfer buffer was prepared by adding 200 ml of 100%

methanol to 100 ml of 10X transfer buffer followed by 700 ml of ddH2O to make up 1

L. The 1X transfer buffer was stored at 4°C initially and transferred to -20°C freezer for

30 min prior to use.

2.5 General Methodology

2.5.1 Cell line types

All experiments conducted in this study were performed utilising primary paediatric

AECs. Due to the limited availability of the primary cells, initial optimisation

experiments were performed on immortalised cell lines or modified primary AECs and

subsequently confirmed on primary paediatric AECs. Cellular material from cell line

cultures were also used for calibration, standardisation and experimental control

purposes as described in the relevant sections.

2.5.1.1 16HBE14o- cell line

An immortalised human bronchial epithelial cell line (16HBE14o-) was obtained from

Dr Dieter Gruenert (University of California, San Francisco, USA). This cell line was

originally established from normal bronchial epithelium and transformed using a Simian

Virus 40 (SV40) large T-antigen (Cozens et al. 1994). The cell line was maintained in a

defined growth medium (refer to 2.4.2.2.1) at 37°C in an atmosphere of 5% CO2 / 95%

air. 16HBE140- RNA was also used as a calibrator for gene expression measurement by

RT-qPCR. This cell line was tested fortnightly to ensure cultures remained cleared of

mycoplasma contamination.

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2.5.1.2 A549 cell line

The adenocarcinoma human alveolar basal epithelial cell line A549 was obtained from

The Lung Institute of Western Australia at Sir Charles Gardener Hospital (Perth, WA,

Australia). A549 cells were first isolated and cultured from an explanted cancerous

tumour tissue of a 58-year old Caucasian male and shown to exhibit squamous epithelial

cell like characteristics (Giard et al. 1973). The cell line was maintained in a defined

growth medium (refer to 2.4.2.2.2) and cultured at 37°C in an atmosphere of 5% CO2 /

95% air. A549 RNA was also used as a calibrator for gene expression measurement by

RT-qPCR. This cell line was tested fortnightly to ensure cultures remained cleared of

mycoplasma contamination.

2.5.1.3 Caco-2 cell line

The human epithelial colorectal adenocarcinoma cell line was obtained from American

Type Culture Collection (ATCC) (Manassas, VA, USA). Caco-2 cells were first

isolated and cultured from a colon carcinoma and were capable of being induced to

differentiate and polarised under specific culture conditions (Fogh and Trempe 1975).

The cell line was maintained in a defined growth medium (refer to 2.4.2.2.3) and

cultured at 37°C in an atmosphere of 5% CO2 / 95% air. Caco-2 cells have been widely

used as a model to predict permeability of solutes across the physical barrier due to the

ability to become polarised and were thus used for optimisation of experimental

protocols instead of primary paediatric AECs. This cell line was tested fortnightly to

ensure cultures remained cleared of mycoplasma contamination.

2.5.1.4 HeLa Ohio cell line

The human epithelial cervix carcinoma cell line was obtained from the European

Collection of Cell Cultures (ECACC) (Porton Down, Salisbury, UK). Hela Ohio cells

were first isolated and cultured from a cervix carcinoma and have been shown to be the

most commonly utilised immortalised cell line in various scientific research. The cell

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line was maintained in a defined growth medium (refer to 2.4.2.2.4) and cultured at

37°C in an atmosphere of 5% CO2 / 95% air. HeLa Ohio cells were used for the

propagation of human rhinovirus-1B (HRV-1B). This cell line was tested fortnightly to

ensure cultures remained cleared of mycoplasma contamination.

2.5.1.5 MRC-5 cell line

The human foetal lung fibroblast cell line was obtained from American Type Culture

Collection (ATCC) (Manassas, VA, USA). MRC-5 cells were first derived from a

normal lung tissue of a 14-week old male foetus. These fibroblast cells have been

shown to be capable of 42-46 population doublings before the onset of senescence. The

cell line was maintained in a defined growth medium (refer to 2.4.2.2.5) and cultured at

37°C in an atmosphere of 5% CO2 / 95% air. MRC-5 cells were used for the titration of

crude HRV-1B. This cell line was tested fortnightly to ensure cultures remained cleared

of mycoplasma contamination.

2.5.1.6 NuLi-1 cell line

The human airway epithelial (HAE) cell, NuLi-1 was obtained from American Type

Culture Collection (ATCC) (Manassas, VA, USA). NuLi-1 cells were first derived from

a normal lung of an adult patient via dual retroviral infection with HPV-16E6/E7-LXN

(Zabner et al. 2003). Consequently, NuLi-1 cells do not undergo growth arrest when in

cell culture due to exogenous expression of the telomerase and HPV-16 E6/E7 genes

and are capable of forming polarised differentiated epithelia which exhibits

transepithelial electrical resistance as well as maintenance of ion channel physiology.

This modified primary HAE cell was maintained in defined growth medium (refer to

2.4.2.2.6) and cultured at 37°C in an atmosphere of 5% CO2 / 95% air. This modified

primary AEC was tested fortnightly to ensure cultures remained cleared of mycoplasma

contamination.

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2.5.2 Immortalised cell line culture, sub-culture and cryopreservation

Frozen stocks of immortalised cell lines were stored at -180°C in cryopreservation

medium (refer to 2.4.2.3.2). When required, cells were revived by initial thawing at

37°C, diluted in 10 ml of appropriate pre-warmed defined culture medium followed by

centrifugation at 500 g for 7 min at 4°C. Following centrifugation, supernatant was

aspirated to remove residual DMSO and the cell pellet then re-suspended using the

appropriate pre-warmed defined culture medium. Ten microliters of cell suspension was

removed and the cells counted using a haemocytometer while viability was also

assessed via trypan blue staining. The cells were then seeded into a non-coated 25 cm2

culture vessel in 5 ml of appropriate pre-warmed culture media containing additives

(refer to 2.4.2.2.1 to 2.4.2.2.5). The culture vessels were subsequently maintained in an

incubator at 37°C at an atmosphere of 5% CO2 / 95% air. In culture expansion, cells

were initially washed with HBSS (refer to 2.4.1.5) and were detached from the culture

vessels by incubating with 0.25% Trypsin (w/v) / 0.05% (w/v) EDTA solution for 4 min

at 37°C. The culture vessel was then washed with the appropriate culture media and the

resulting cell suspension collected and centrifuged at 500 g for 7 min at 4°C. The

supernatant was then aspirated and the cell pellet re-suspended in the appropriate culture

medium. A total cell count and viability stain was also performed. Cells were then

plated into new culture flasks and incubated at 37°C in an atmosphere of 5% CO2 / 95%

air in the appropriate culture medium (refer to 2.4.2.2.1 to 2.4.2.2.5).

For future use of cell lines, stocks of each cell line were frozen down and subsequently

stored in liquid nitrogen tanks. Briefly, cells were detached from flasks as described

above and re-suspended in the appropriate culture medium (refer to 2.4.2.2.1 to

2.4.2.2.5). The resulting pellet obtained was re-suspended in 1 ml of cryopreservation

medium (refer to 2.4.2.3.2) and frozen in a Mr Frosty cryopreservation vessel at -80°C

for 24 h. This provides the critical -1°C / minute cooling rate required for successful cell

cryopreservation and recovery. The frozen cells were transferred to a liquid nitrogen

storage facility for long-term storage.

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2.5.3 Ethics approval

This study was approved by both the Princess Margaret Hospital for Children’s Human

Ethics Committee and the St John of God Hospital Health Care Ethics Committee for

the recruitment of AECs from children attending theatre for non-respiratory related

elective surgery at the respective hospitals (Refer to Appendix A - C).

2.5.4 Primary paediatric airway epithelial cells

Four cohorts of paediatric airway epithelial cells (pAEC) were used in this study: pAEC

from healthy non-atopic (HNA) and atopic (HA) children with no history of asthma,

pAEC from non-atopic asthmatic (NAA) and atopic asthmatic (AA) children with mild

asthma, who did not receive any corticosteroid therapy at least 1 month prior sampling.

Samples were collected from 133 subjects who were undergoing elective surgery for

non-respiratory conditions. Asthma was defined as clinician diagnosed asthma with a

documented wheeze in the past 12 months and confirmed with positive responses to

related questions on both the International Study of Asthma and Allergies in Childhood

(ISAAC) and American Thoracic Society (ATS) questionnaires (Ferris 1978; Asher et

al. 1995). Atopy was defined by a positive radioallergosorbent (RAST) test to a panel of

common allergens (Table 2), elevated plasma IgE levels and a history of hay fever and /

or eczema.

2.5.4.1 Primary airway epithelial cell isolation

Primary AECs were collected via either trans-laryngeal, non-bronchoscopic brushings

through an endotracheal tube or a bronchoscope-guided brushing of the tracheal

mucosa. Both the non-bronchoscopic brushing and the bronchoscope-guided techniques

were established in the laboratory in which this study was performed (Lane 2005; Kicic

et al. 2006; McNamara et al. 2008). For the non-bronchoscopic brushing technique,

once each patient was anaesthetised using sevofluorane and propofol, and intubated in

theatre, an unsheathed soft nylon bronchial cytology brush was advanced through the

endotracheal tube until the tip encountered airway wall resistance usually above the

Table 2: Radioallergosorbent test (RAST) to a panel of common allergens for determination of atopy

Mixed Allergen Test Elements

EX1 (Epidermal and Animal Protein Mix 1) Cat epithelia (Felis silvestris catus)

Horse dander (Equus caballus)

Cow dander (Bos Taurus)

Dog dander (Canis lupus familiaris)

GX2 (Grass Pollen Mix 2) Couch (Cynodon dactylon)

Rye (Lolium perenne)

Timothy (Phleum pretense)

Meadow (Poa pratensis)

Johnson (Sorghum halepense)

Bahia (Paspalum notatum)

MX1 (Mould Mix 1) Penicillium notatum

Cladosporium herbarum

Aspergillus fumigatus

Alternaria alternate

HX2 (House Dust Mite Mix 2) Dermatophagoides pteronyssinus

FX5E (Food Mix 5E) Egg white

Cow’s milk

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carina. Sampling occurred using a rotational movement of the brush rather than a

probing movement. Completion of sampling would see the cytology brush being

withdrawn from the endotracheal tube, the brush tip inserted and detached into cold

sterile collection media (RPMI-1640) containing 20% (v/v final) heat-inactivated FCS.

The brushing process was repeated with a second cytology brush tip which was then

detached into the same collection media.

In the bronchoscope-guided brushing technique, the unsheathed cytology brush was

passed down the instrument port of a portable intubation bronchoscope and kept

concealed within the bronchoscope tip. The patient’s vocal cords were sprayed with

lidocaine, after which the patient was hand ventilated with a bag and mask for

approximately 2 min prior intubation. The portable bronchoscope was then passed down

through a laryngeal mask airway (LMA), directed and positioned at a suitable site for

brushing beyond the lower edge of the vocal cords. Once positioned below the vocal

cords, the cytology brush was then advanced till resistance was encountered typically

above the carina region, similar to the blind brushing technique described above.

Rotational movement was performed to sample the cells and subsequently the cytology

brush tip was retracted to just beyond the tip of the bronchoscope. Both the

bronchoscope and the concealed cytology brush were then withdrawn together. The

cytology brush would then be pushed out from the bronchoscope and the brush tip

detached into sterile collection media. This procedure was repeated once more with a

second brush tip being detached into the same collection tube.

From both techniques, the collection tube, together with the brush tips were then

immediately processed by vortexing for 15 sec to dislodge the cells off the brush tips as

well as breaking up larger cell clumps. The brush tips were then removed into another

collection tube and the process repeated. Following this, the two collection media were

pooled, centrifuged at 500 g for 7 min at 4°C to pellet the cells and subsequently

resuspended in BEBM supplemented with growth additives (refer to 2.4.2.2.6). Cell

viability and yield was determined using a haemocytometer with trypan blue exclusion.

The cell suspension was then incubated on a petri dish pre-coated with a 1:500 dilution

of CD-68 antibody for 20 minutes at 37°C in an atmosphere of 5% CO2 / 95% air to

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remove macrophages. Following that, the cell suspension was first passed through a 25-

gauge needle followed by a 27-gauge needle for the separation of cell clumps. A

fraction of the cell suspension was seeded into a culture vessel (growth area 25 cm2)

pre-coated with fibronectin coating buffer (refer to 2.4.2.3.1) and maintained in a

Panasonic CO2 incubator in an atmosphere of 5% CO2 / 95% air and at 37°C. The

remaining cell suspension was centrifuged, the supernatant aspirated and the cell pellet

dissolved in 350 µl of QIAzol lysis reagent (QIAGEN, Hilden, Germany) for

subsequent RNA extraction.

2.5.4.2 Primary airway epithelial cell sub-culture

Established primary AEC cultures were expanded over 2 passages for downstream

experimentation. In culture expansion, cells were detached from the culture vessels by

initially washing them in HBSS (refer to 2.4.1.5) and incubating with 0.25% (w/v final)

Trypsin / 0.05% (w/v final) Ethylenediaminetetraacetic acid (EDTA) solution for 4 min

at 37°C. Trypsin neutralising solution (TNS) was added to halt the action of the

Trypsin-EDTA. The resulting cell suspension was centrifuged at 500 g for 7 min at 4°C

, upon which the supernatant was aspirated and the cell pellet resuspended in BEBM

supplemented with growth additives (refer to 2.4.2.2.6). The cell suspension was seeded

onto new culture vessels (growth area 25 cm2) similarly pre-coated with fibronectin

coating buffer and incubated at 37°C in an atmosphere of 5% CO2 / 95% air as

previously described (Kicic et al. 2006; Stevens et al. 2008; Kicic 2010; Sutanto et al.

2011)

2.5.4.3 Primary airway epithelial cell culture supernatant collection

Prior to the passage or harvesting of an established primary AEC culture, the culture

media in which the cells were grown was collected and stored at -80°C for subsequent

protein analysis.

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2.5.5 Plasma and buffy coat isolation

In addition to AEC collection, 5 ml of whole blood was collected from each cohort

participant, placed into a heparin sodium collection tube, mixed and transported to the

laboratory. The whole blood was centrifuged for 10 min at 2000 g at 18°C. Plasma and

the buffy coat layer were collected into 1.5 ml micro-centrifuge tubes respectively and

stored at -80°C until required.

2.5.6 Cytospin preparation

Cell cytospins were prepared by adding 60 – 70 µl of cell suspension containing at least

50 000 cells to each slide encased in a cytospin block and centrifuged for 20 min at

1500 rpm. The resulting slides were allowed to air dry for 24 h after which they were

fixed with 4% (v/v final) neutral buffered formalin (refer to 2.4.1.9) for 10 min at room

temperature. Slides were washed three times in a bath of 1X PBS (refer to 2.4.1.10) for

10 min to remove excess NBF and allowed to air dry. Slides were then stored at -20°C

until required.

2.5.7 Human Rhinovirus

Human rhinovirus (HRV) minor serotype 1B (HRV-1B) stocks was kindly provided by

Professor Peter Wark (John Hunter Hospital, Newcastle, New South Wales, Australia).

Human rhinovirus-1B stocks obtained were generated and titrated from infected cultures

of HeLa Ohio cells as previously described (Papi and Johnston 1999). Initially

determined viral titres of 1.06 x 108 tissue culture infective dose 50% (TCID50) / ml for

HRV-1B was reconfirmed via titration in MRC-5 cells by Dr Gerry Harnett (Pathwest,

Perth, Western Australia, Australia)

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2.5.7.1 Viral propagation of human rhinovirus-1B (HRV-1B)

Viral stocks of crude HRV-1B were generated by infecting monolayer cultures of Ohio

HeLa cells until cytopathic effects were fully developed. Cells and the supernatants

were harvested, cells were disrupted by repeated freezing at -80°C and thawing and the

resultant cell debris centrifuged at 2016 g for 10 min at 4°C. The resulting supernatant

was aspirated and the cell pellet was resuspended in 1X PBS (refer to 2.4.1.10) and

stored at -80°C until required.

2.5.7.2 Titration of human rhinovirus-1B (HRV-1B)

Human rhinovirus titration was performed on the frozen aliquots by exposing confluent

monolayers of MRC-5 cells in 48-well plates to serial 10-fold dilutions of crude viral

stock. Plates were cultured for 7 days in the required growth medium (refer to 2.4.2.2.5)

and incubated at 35°C in an atmosphere of 5% CO2 / 95% air. Cytopathic effect was

assessed by visual assessment over 7 days and TCID50 / ml values were determined as

previously described (Johnston and Tyrrell 1995).

2.5.7.3 Ultra-violet (UV) light inactivation of human rhinoviral activity

To confirm that the human rhinovirus (HRV) mediated response observed were a result

of active virus, the HRV-1B serotype was UV inactivated. One millilitre of HRV-1B

crude stock virus was transferred to a petri dish with lid removed and placed 10 cm

from a UV light source in a lamina flow hood. The empty vial was also similarly placed

with lid removed to allow for maximal penetration of the UV light. The UV-inactivated

virus was exposed to UV light for at least 120 min and subsequently stored at -80°C in

multiple aliquots until required.

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2.5.7.4 Cytotoxicity assays

To determine the effects of UV-inactivated human rhinovirus-1B (HRV-1B) on primary

airway epithelial cell (pAEC) viability, cells were seeded in 96-well plates and grown to

85% confluence in BEBM containing growth additives (refer to 2.4.2.2.6). Ultra-violet

light inactivated HRV-1B was then added to the wells at a titre ranging from 2.5 – 80

TCID50 / ml and exposed for 24, 48 and 72 hours. Following exposure, supernatant were

collected and bio-banked for subsequent cytokine assessment. The CellTitre 96® Aqueous

Non-Radioactive Cell Proliferation Assay was adapted to assess the number of

metabolically active cells post viral infection and was performed as previously

described (Kicic et al. 2006; Stevens et al. 2008).

2.5.8 Immunocytochemistry

Fixed slides of epithelial cells were brought to room temperature and re-hydrated with

1X PBS (refer to 2.4.1.10) for 5 min. To improve the intensity of staining, antigen

retrieval was performed using either citrate buffer or proteinase K. For citrate buffer

antigen retrieval, slides were immersed in 0.01 M citrate buffer (refer to 2.4.3.2.1) and

heated in a microwave oven for 15 min. Slides were then removed and allowed to cool

and washed in 1X PBS (3 x 15 min / wash). For proteinase K retrieval, slides were

incubated with 36 µg/ml proteinase K solution (refer to 2.4.3.2.2) at 37°C for 30 min

and subsequently washed in 1X PBS (3 x 15 min / wash). Cells were then quenched to

minimise auto-fluorescence by incubating the slides in 0.5% (w/v final) Sudan B Black

solution (refer to 2.4.3.2.3) for 20 min at RT. After washing with 1X PBS (3 x 15 min /

wash), cells were blocked for 2 h at RT in blocking buffer (5% (w/v final) BSA, 10%

(v/v final) FBS, 0.1% (v/v final) Triton X-100 and 1% (w/v final) saponin in 1X PBS) if

cytoplasmic or nuclear proteins were to be detected. If cell surface proteins were to be

detected, the slides were incubated for 2 h at RT in saponin free blocking buffer (5%

(w/v final) BSA, 10% (v/v final) FBS, 0.1% (v/v final) Triton X-100 in 1X PBS). The

appropriate primary antibodies were diluted in the blocking buffer solution at the

desired concentration and added to the slides which were then incubated overnight at

4°C. The following day, the slides were washed in 1X TBS (refer to 2.4.1.15) with 0.1%

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(w/v) saponin (3 x 15 min / wash) for the detection of cytoplasmic markers. If cell

surface markers were to be detected, 1X TBS without saponin was used as the wash

buffer. Fluorescent secondary antibodies were prepared in blocking buffer to the

necessary concentration and added to the slides which were further incubated in the

dark, overnight at 4°C. The following day, slides were washed in 1X TBS with saponin.

Once all primary and secondary antibodies had been used to stain the cells, the nucleus

of the cells was stained with 4′, 6-diamidino-2-phenylindole (DAPI). Slides were

incubated with DAPI (1:50 000) in 1X PBS for 10 min and then washed in 1X TBS (3 x

15 min / wash). Fluorescent mounting media was used to minimise fading and slides

were visualised using a fluorescence microscope.

2.5.9 Immunohistochemistry

Paraffin embedded, formalin fixed sections were initially deparaffinised, rehydrated and

subjected to antigen retrieval by incubating sections with proteinase K for 15 minutes at

37°C. Slides were then cooled to room temperature (RT) for 10 minutes followed by 3

washes in 1X TBS (refer to 2.4.1.15) containing 0.1% (v/v) saponin. Sections were

then blocked in 5% (w/v) BSA, 10% FBS (v/v), 0.1% (v/v) TritonX-100 and 0.1% (v/v)

saponin in 1 X TBS for 1 hour at RT. After a second series of washes, sections were

incubated with the appropriate primary antibodies diluted in the blocking buffer solution

at the desired concentration and added to the slides which were then incubated

overnight at 4°C. The following day, sections were washed in 1X TBS (refer to

2.4.1.15) with 0.1% (w/v) saponin (3 x 15 min / wash) for the detection of cytoplasmic

markers. Fluorescent secondary antibodies were prepared in blocking buffer in the

necessary concentration and added to the slides which were further incubated overnight

at 4°C. The following day, slides were washed in 1X TBS with saponin. Once all

primary and secondary antibodies had been used to stain the cells, the nucleus of the

cells was stained with 4′, 6-diamidino-2-phenylindole (DAPI). Slides were incubated

with DAPI (1:50 000) in 1X PBS for 10 min and then washed in 1X TBS (3 x 15 min /

wash). Fluorescent mounting media was used to minimise fading and slides were

visualised using a fluorescence microscope.

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2.5.10 In-Cell™ Western

Primary and subsequently passaged pAECs were plated onto 96-well microplates pre-

coated with fibronectin (10 mM) at a high seeding density of 1.2 x 105 cells/cm2 and

incubated at 37°C in an atmosphere of 5% CO2 / 95% air in BEBM containing growth

supplements as described previously (refer to 2.4.2.2.6). Upon observation of confluent

cell monolayers, growth media was aspirated and cells were immediately fixed using

150 µl of 3.7% formalin (refer to 2.4.3.1.1) for 20 min at RT. The cells were then

washed with 1X PBS containing 0.1% Triton X-100 (refer to 2.4.3.1.2) (5 x 5 min /

wash) with gentle shaking if cytoplasmic or nuclear proteins were to be detected. If cell

surface proteins were to be detected, the cells were incubated for 90 min at RT using

150 µl of LI-COR Odyssey Blocking Buffer. Cells were then incubated overnight at

4°C with primary antibodies dissolved in LI-COR Odyssey Blocking Buffer at a 1:200

dilution. The following day, cells were washed with 1X PBS containing 0.1% (v/v)

Tween-20 (refer to 2.4.3.1.3) (5 x 5 min / wash) with gentle shaking and incubated with

respective IRDye® secondary antibodies, diluted in a solution of LI-COR Odyssey

Blocking Buffer (1:800) with DRAQ5 (1:10000), Sapphire700 (1:1000) and 0.2% (v/v)

Tween-20 for 60 min at RT with gentle shaking in a dark room. Cells were then washed

in 1X PBS containing 0.1% (v/v final) Tween-20 (refer to 2.4.3.1.3) (5 x 5 min / wash).

Specific antibody staining for protein expression was then immediately visualised using

an infrared imaging system at both 680 and 800 nm channels. Protein expression was

quantified using the LI-COR Odyssey v.3.0 software. The integrated intensity (I.I) of

each well at 800 nm was then normalised to the I.I of the cell densities at 680 nm in the

corresponding well.

2.5.11 Transepithelial permeability

Primary airway epithelial cells (pAECs) were seeded onto fibronectin-coated culture

inserts (Corning Incorporated Life Sciences, MA, USA) at a high seeding density of 1.2

x 105 cells/cm2 and incubated at 37°C in an atmosphere of 5% CO2 / 95% air in BEBM

containing growth supplements as described previously (refer to 2.4.2.2.6). Upon

observation of confluent cell monolayers, growth media within the apical and

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basolateral compartments was replaced with HEPES buffered Hank’s balanced salt

solution (HEPES-HBSS) (refer to 2.4.3.3.2). Two hundred microliters of FITC-dextran

of molecular weight 4kDa or 20kDa (2 mg/ml) (refer to 2.4.3.3.1) was added to the

apical chamber and 50 µl of apical solution was immediately sampled. Five hundred

microliters of HEPES-HBSS from the basolateral compartment was sampled at hourly

intervals over a period of 6 h. The volume of the basolateral compartment was

maintained by addition of 500 µl fresh buffered HBSS-HEPES. All experiments were

performed at 37°C and on a calibrated orbital shaker at 300 rpm to minimise the

unstirred buffer layer. Fluorescence of FITC-dextran was detected using a PerkinElmer

Enspire® multilabel plate reader at an excitation wavelength of 492 nm and emission

wavelength of 520 nm. The apparent permeability of the epithelial monolayer to FITC-

dextran from the apical to basolateral compartment (Papp) was then calculated following

the general equation: Papp = (dQ/dt) x (1/AC0) where dQ/dt is the steady –state flux, A is

the surface area of the membrane and C0 is the initial concentration in the donor

compartment as previously described (Stutts et al. 1981).

2.5.12 Transepithelial electrical resistance (TEER)

Upon attaining a confluent monolayer of primary AECs in the culture inserts, TEER

measurements were performed across the epithelial cell monolayer to ensure the

formation and integrity of TJs between cells. This was achieved using an epithelial

voltohmeter (Millicell-ERS voltmeter, Millipore) with silver chloride ‘chopstick’

electrodes. Triplicate measurements per well were made at 37 ± 2°C prior the start of

any transport experiments and the mean resistance calculated. The resistance obtained

from a cell-free culture insert was subtracted from the resistance measured across each

cell monolayer and corrected for the surface area of the culture insert to yield the TEER

of the epithelial cells with values expressed in Ω/cm2.

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2.5.13 Reverse Transcriptase-Polymerase Chain Reaction (RT-PCR) and Real Time

quantitative Polymerase Chain Reaction (RT-qPCR)

Primers for the genes of interest (CLDN1, OCLN, ZO1) and the house keeping gene

(PPIA) were obtained from GeneWorks (Adelaide, SA, Australia) (Appendix D). Gene

expression was analysed by two-step RT-qPCR reactions. Total cellular RNA was

extracted from cell pellets stored in either QIAzol lysis reagent or RLT buffer with 1%

(v/v) 2 β-mercaptoethanol with RNeasy mini columns (QIAGEN, Hilden, Germany)

after a DNA digest was performed with RNase-Free DNase to remove unwanted DNA.

The isolated RNA was assessed for quality and quantity by measuring

spectrophotometric absorbance at 260 and 280 nm using a NanoDrop. Reverse

transcription was performed to convert 200 ng of RNA into cDNA. A 200 ng sample of

RNA was added to a master mix containing 10X RT Buffer (2 µl), 5 mM MgCl2 (4.4

µl), 2 mM deoxyribonucleotide triphosphates (dNTPs) (1 µl), Random Hexamers (1 µl),

RNase inhibitor (0.4 µl) and Multiscribe (0.5 µl) and then made to a final volume of 20

µl with RNase free water. Samples were then placed in a thermal cycler and run on a

standard reverse transcription program of 25°C for 10 min, 48°C for 60 min and 95°C

for 5 min. The RT-qPCR reaction contained cDNA (10ng), forward and reverse primers

(0.3 µM), SYBR® GREEN PCR Master Mix (10 µl) and RNase free water to make a

final volume of 20 µl. RT-qPCR was performed on an ABI Prism® 7300 (refer to

2.3.15). Results were analysed as previously described and gene expression of all

samples was expressed relative to the expression of the house keeping gene PPIA.

2.5.14 Total cellular protein extraction

Total cellular protein from cell cultures was extracted with CEB (refer to 2.4.2.3.3)

After aspirating the culture supernatant, cell pellets were re-suspended in 900 µl of CEB

and 100 µl of protease inhibitor cocktail was added. The cell suspension was placed on

ice to prevent protein degradation. Cells were then lysed by mechanical force with a 27-

gauge needle and syringe and stored at -80°C until required.

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2.5.15 Total cellular protein quantification

Total protein concentration was determined with the micro-Bicinchoninic Acid (BCA)

Protein Assay Reagent Kit (Pierce, Rockford, IL, USA) as previously described (Kicic

et al. 2006; Stevens et al. 2008; Sutanto et al. 2011). This assay is based on the

reduction of Cu2+ to Cu1+ by protein in an alkaline medium with the highly sensitive and

selective colorimetric detection of the cuprous cation (Cu1+) by bicinchoninic acid.

Briefly, protein samples were diluted 1:5, 1:10 and 1:20 in 1X PBS (refer to 2.4.1.10)

and the BSA protein standard constructed which consisted of a concentration range

between 12.5 and 500 µg/ml. Forty microliters of sample or standard was added to each

well of a 96-well microtitre plate. Secondary kit reagents were then combined in a

50:48:2 ratios and 200 µl of the mixture added to each well and incubated for 60 min at

37°C and the absorbance of the wells read at 562 nm. The absorbance of the standards

was plotted against their known concentrations and a standard curve generated. The

concentration of the sample was then determined from the standard curve incorporating

any dilution factor utilised.

2.5.16 Western Blot

Protein was collected from cells by CEB (refer to 2.4.2.3.3), quantitated by BCA assay

(refer to 2.5.15) and stored at -80°C. Prior to Western Blot analysis, the protein samples

were thawed but placed on ice to prevent protein degradation. A 10 µg protein sample

was mixed with NUPAGE® Lithium Dodecyl Sulphate (LDS) buffer, NUPAGE®

reducing agent and ddH2O (refer to 2.4.1.1) to make up a final volume of 20 µl. The

samples were then heated for 10 min at 70°C on a heating block (refer to 2.3.7) for

optimal denaturation before being loaded into a pre-cast 4 - 12% 1.0 mm Bis-Tris Plus

polyacrylamide gel (Novex BOLT®). Samples were then electrophoresed using a Novex

BOLT® Western Blot apparatus (Life Technologies) in MES SDS running buffer (refer

to 2.4.3.4.1) at a constant 200 V for 35 min at RT. A pre-stained protein ladder was run

on all gels in addition to samples for reference purposes. After separation, proteins were

transferred to onto a PVDF membrane using a semi-dry transfer method on iBlot

(Invitrogen®) system at 200 V for 7 min or a wet transfer method at a constant 230 mA

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for 2 h at 4°C. Upon completion of protein transfer, the PVDF membrane was blocked

for non-specific staining by using the LI-COR Odyssey Blocking Buffer for 60 min at

RT. Membranes were then incubated, with gentle rocking, overnight at 4°C with the

required primary antibodies made up in LI-COR Odyssey Blocking Buffer. The

membranes were then washed 3 times (15 min per wash) in 0.2% (v/v final) Tween-20

in 1X TBS solution (refer to 2.4.1.15) at RT. After washing, membranes were incubated

in the dark with respective IRDye® secondary antibodies made up in a solution of LI-

COR Odyssey Blocking Buffer with 2% (v/v final) Tween-20 diluent for 2 h at RT with

gentle rocking. The membranes were then washed 3 times (15 min per wash) in 0.2%

(v/v final) Tween-20 in 1X TBS solution (refer to 2.4.1.15) followed by 2 times (10 min

per wash) in 1X TBS (refer to 2.4.1.15) alone. The membranes were then scanned using

the LI-COR Odyssey infrared scanner at 680 nm and 800 nm channels. Bands of protein

expression were quantified using the LI-COR Odyssey v.3.0 software. The integrated

intensity (I.I) of each band was then normalised to the I.I of the house-keeping protein,

β-actin.

2.5.17 Statistical analysis

Each experiment performed in this thesis was conducted between 6 and 8 times with at

least 2 replicates per experiment. Student’s t-test and Mann-Whitney test were used to

compare means as appropriate. Values are presented as means ± standard error of mean

(SEM) or means ± standard deviation (SD) where appropriate. All p values less than

0.05 were considered to be significant. IBM SPSS 21 software package was used to

perform all statistical analysis.

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CHAPTER 3: Optimisation of In Cell™ Western and Transepithelial

permeability assays

3.1 Introduction

Tight junctions play a role in regulating the passage of solutes through the paracellular

pathway of epithelial cells and are crucial in the establishment of various

compositionally distinct fluid compartments within the body of multicellular organisms

(Tsukita et al. 2001; Matter and Balda 2003). Moreover, for the epithelium to function

efficiently as a barrier, the inter-cellular space must remain well sealed to prevent

unwanted trafficking of injurious solutes. However, it has been postulated that there

exists, either in health or disease, along the epithelium, weak points within the TJ

barrier where solutes remain capable of traversing (Ikenouchi et al. 2005).

In order to elucidate the function of epithelial TJ complexes, the ability to accurately

assess the expression of these complexes becomes an integral requirement in various

experimental procedures. Numerous studies have been performed that have investigated

the expression of junctional complexes in either health or disease and have employed

the use of western blotting in determining junctional protein expression (West et al.

2002; Coyne et al. 2003; Xiao et al. 2011; Hardyman et al. 2013). Although considered

to be a benchmark assay for protein quantification, western blotting is an intensive

process which includes, briefly; extraction of cellular protein and denaturation, gel

electrophoresis, protein transfer onto a blotting membrane followed by blocking of non-

specific binding. The membrane is then incubated with primary followed by secondary

antibodies, upon which chemiluminescent or fluorescent detection of the target proteins

is achieved through a detector. One major limitation of western blotting is that it is only

capable of analysing a single target protein for any given sample at one time. Hence,

analysis of multiple target proteins becomes very time-consuming. This becomes more

impractical when there is a need to analyse multiple target proteins in large sample

sizes, since typically 10 – 50 µg of protein lysate is required per sample. Additionally,

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when initial sample sizes are small and repeated sampling is unavailable, the use of

traditional western blotting becomes less practical to semi-quantify protein expression.

To circumvent these limitations, a novel, quantitative immunofluorescence assay,

termed In-Cell™ Western (ICW), can be used to quantify multiple proteins

simultaneously from small sample sizes. Briefly, ICW assays involve the culture of

cells on a 96-well micro-titre plate and upon confluence, are fixed. Afterwards, cells are

blocked to prevent non-specific binding and subsequently incubated with antibodies for

specific target proteins. Cells are then incubated with a solution containing IRDye®-

labelled secondary antibodies, non-specific cell stain Sapphire700 and a DNA stain

Draq5, which acts to normalise cell numbers within each well. The micro-titre plate is

subsequently washed and analysed using a near-infrared detector, which then semi-

quantifies the expression of the target proteins, following normalisation to cell densities.

The advantage of this assay is that it is a small scale, high throughput method that

enables analysis of multiple samples and target proteins, both membrane-bound and

intracellular simultaneously. Furthermore, since cells are cultured and analysed in a

micro-titre plate set-up, it is a rapid procedure to perform as additional steps necessary

for traditional western blotting, which includes protein extraction and quantification are

no longer required.

Since this assay’s development, it has been used in a variety of applications including;

monitoring effects of drug compounds on signalling pathways (Chen et al. 2005),

timing and kinetics of cellular signal transduction (Hannoush 2008), examining

functional consequences of cell receptor mutation (Chan et al. 2009) and for the

identification of inhibitors of signalling pathways (Hoffman et al. 2010). In all

investigations mentioned, it has been demonstrated that the ICW assay provided

accurate and robust high-throughput screening for the quantitative measurements of the

target proteins. Nevertheless, the use of the ICW assay in the quantification of epithelial

junctional complex proteins remains relatively absent. Hence, in this investigation, the

ICW assay was established and optimised to elucidate the expression of multiple TJ

proteins, in AECs derived from asthmatic and non-asthmatic children.

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Ultimately, TJ expression must be translated into a functional context, especially when

trying to elucidate differences between healthy and diseased states. Transepithelial

permeability is dependent upon the functioning of various inter-cellular junctions such

as TJs, which forms a seal at the apical end of the epithelium. This acts to selectively

regulate the passage of most molecules and ions as well as restrict lateral movement of

molecules in the cell membrane. Many disease settings are often exacerbated by or

result from a loss of epithelial barrier function. As a consequence, this permits the

passage of harmful pathogens into the sub-epithelial layer, causing further

exacerbations. Hence, the ability to functionally assess the epithelial barrier integrity in

both healthy or disease states is critical in improving our understanding of the airway

epithelium.

There are several variations to the transepithelial permeability assay, however, the most

routinely utilised method involves the initial culturing of cells on a semi-permeable

culture insert (Hubatsch et al. 2007). Upon reaching confluence, cells can either be

utilised immediately for the transepithelial permeability assay or air-lifted into air-liquid

interface (ALI) cultures, whereby, cells become polarised and fully differentiated. A

small volume of fluorescently-labelled inert molecules dissolved in a buffer is then

added onto the apical layer while the basolateral layer is concurrently filled with the

same buffer without the fluorescent molecule. The culture plate, together with the insert,

is placed on a platform shaker within a humidified incubator at 37°C with 5% CO2 /

95% air for a duration of 6 hours. At regular intervals, a pre-determined volume of

buffer is collected from the basolateral layer and subsequently replenished with fresh

buffer. Following the last sample collection, sample volumes collected are then

transferred into a micro-titre plate and the absorbance read using a fluorescence counter.

Various epithelial cells in culture, including the human epithelial colorectal

adenocarcinoma cell (Caco-2), form confluent monolayers which are capable of

developing junctional complexes. Moreover, Caco-2 cells, when cultured under specific

conditions, are also capable of becoming polarised. Hence, these cells have been the

most routinely used for the in vitro assessment of transepithelial drug permeability and

absorption (Hubatsch et al. 2007). However, a more applicable model utilising human

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derived primary AECs is needed when investigating transepithelial permeability within

the airway epithelium in children.

The transepithelial permeability assay has been used in several investigations assessing

permeability following various insults (Wan et al. 1999; Dreschers et al. 2007; Xiao et

al. 2011) and have proven its reliability in providing greater insights into airway

epithelial function in adults. However, little is known on the functional consequences of

epithelial TJ complex disruptions within the paediatric population. Hence, in this

investigation, the transepithelial permeability assay was established and optimised to

allow for the functional assessment of airway epithelial barrier integrity in asthmatic

and non-asthmatic children.

3.2 In Cell™ Western assay

3.2.1 Materials

All general materials and equipment used are as described in Chapter 2 (refer to 2.1 -

2.4).

3.2.1.1 Cell lines and paediatric-derived primary airway epithelial cells

All cell lines and paediatric-derived primary AECs utilised in this investigation were

obtained and maintained as previously described (refer to 2.5.1.6 and 2.5.4).

3.2.1.2 Human Tight Junction primary antibodies

Human TJ primary antibodies to claudin-1, occludin and zonula occluden-1 were

prepared and utilised as previously described (refer to 2.5.10).

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3.2.1.3 Human Tight Junction secondary antibodies

IRDye® secondary antibodies to either rabbit or mouse primary antibodies were


3.2.2 Methods

In order to optimise this assay for primary AECs, a standard protocol derived from LI-

COR Biosciences was initially utilised as a template. As recommended by the

manufacturer, optimisation studies were performed on the following parameters, 1;

Concentration of primary antibodies, 2; Incubation temperature of primary antibodies

and 3; Concentration of secondary antibodies.

3.2.2.1 Concentration of primary antibodies

In accordance with the standard protocol, various dilutions of the different human TJ

primary antibodies between the recommended ranges of 1:50 to 1:200 were performed.

NuLi-1 cells cultured on 96 well micro-titre plates, once confluent were fixed in 3.7%

(v/v final) formalin (refer to 2.4.3.1.1) for 20 min without agitation. To determine total

protein expression, selected wells were treated with a permeabilising solution (refer to

2.4.3.1.2) while non-permeabilised wells were treated with the LI-COR odyssey

blocking buffer. The range of diluted primary antibodies was then added to the NuLi-1

cells to determine the most appropriate dilution at which optimal fluorescence occurred.

Detection of target proteins was performed by adding the recommended 1:800 dilution

of IRDye® secondary antibodies in a solution containing the cell and DNA stains

Sapphire700 and Draq5 respectively. Analysis using a near-infrared detector, which

semi-quantifies the expression of the target proteins, following normalisation to cell

densities was then performed. A similar series of experiments was then performed in

paediatric-derived primary AECs to confirm initial findings.

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3.2.2.2 Incubation temperature of primary antibodies

As recommended in the manufacturer’s protocol, incubation of primary antibodies may

be performed at either 25°C for 2.5 h or 4°C overnight. Thus, experiments were then

performed whereby NuLi-1 cells were plated and grown in 96 well micro-titre plates.

Primary antibodies were then added to wells in the dilution range recommended by the

manufacturer and plates then incubated at the two recommended temperature and

duration. Experiments were then repeated and confirmed in paediatric-derived primary

AECs to confirm initial findings.

3.2.2.3 Concentration of secondary antibodies

From the standard protocol, various dilutions of the different IRDye® secondary

antibodies between the recommended ranges of 1:200 to 1:1200 were tested. The range

of diluted secondary antibodies were added to NuLi-1 cells probed with the optimal

dilution of primary antibodies (refer to 3.2.2.1) to determine the most appropriate

secondary antibody dilution at which optimal fluorescence occurred. A similar set of

experiments were then performed in paediatric-derived primary AECs to confirm initial

findings.

3.2.3 Results / Discussion

3.2.3.1 Concentration of primary antibodies

Since a number of different primary antibodies were utilised in this investigation, the

optimal concentration of each primary antibody used was addressed. As evident, for the

semi-quantification of total protein expression, the strongest signal intensity was

observed in the 1:50 dilution of occludin, claudin-1 and ZO-1 primary antibody used

while very low signal intensity was observed in the 1:400 dilution for the same primary

antibodies (Figure 3.1 A – Total). Similar intensities were observed for membrane

bound protein expression of occludin, claudin-1 and ZO-1 (Figure 3.1 A – Membrane).

Cell normalisation staining using a combination of cell and nuclear stains Sapphire700

Figure 3.1 Effect of primary antibody concentration on NuLi-1 TJ signal intensity:

NuLi-1 cells, seeded on 96-well micro-titre plates and grown to confluence were treated

as previously mentioned (refer to 2.5.10). Incubation with primary antibodies to

occludin, claudin-1 and ZO-1 was performed at 4°C overnight followed by incubation

with IRDye™ secondary antibodies. (A) Total and membrane TJ signal intensities over

dilution range between 1:50 to 1:400 of primary antibodies observed at 800 nm channel

showed strongest intensity at 1:50 dilution while 1:400 dilution showed the weakest

intensity staining. (B) Uniform cell densities in all wells after staining with Sapphire700

and Draq5 indicates a confluent monolayer as observed at 700 nm channel. (C) Merged

image of (A) and (B) is utilised for the quantification of TJ protein expression as

normalised to cell densities. (D) Total signal intensity of occludin, claudin-1 and ZO-1

in a confluent NuLi-1 monolayer post normalisation to cell densities. Highest total TJ

signal intensity was observed in 1:50 dilution of primary antibodies, however, 1:400

dilution resulted in minimal total TJ signal intensity. (E) Similarly, membrane-bound

signal intensity of occludin, claudin-1 and ZO-1 in a confluent NuLi-1 monolayer post

normalisation to cell densities was observed to be highest in 1:50 dilution while 1:400

dilution demonstrated minimal membrane TJ signal intensity.

Dilutions

Sign

al In

tens

ity

1:50 1:100 1:200 1:4000.0

0.5

1.0

1.5

2.0

2.5occludinclaudin-1ZO-1

Dilutions

Sign

al In

tens

ity

1:50 1:100 1:200 1:4000.0

0.5

1.0

1.5

2.0


1:50 1:100 1:200 1:4000.0

0.2

0.4

0.6

D

E

A

B

C

Total Membrane

occludin claudin-1 ZO-1

Cell & Nuclear Stain

Merged

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and Draq5 respectively demonstrated uniform cell densities in all wells (Figure 3.1 B).

The target protein signal intensities were then merged with the cell normalisation stains

to then quantify target protein expression (Figure 3.1 C). When the signal intensity was

represented graphically, 1:50 dilution of the occludin, claudin-1 and ZO-1 antibodies

again demonstrated the highest total signal intensity staining while very low signal

intensity was observed in the 1:400 dilution (Figure 3.1 D). Graphical representation of

membrane signal intensities also demonstrated similar observations with highest

intensity seen at 1:50 dilutions and lowest detection seen in the 1:400 dilutions (Figure

3.1 E).

A similar series of experiments was also performed in paediatric-derived primary AEC

of healthy individuals (pAECHNA) to confirm initial observations seen in the NuLi-1

cells. As demonstrated, the strongest signal intensity was observed in the 1:50 dilution

of occludin, claudin-1 and ZO-1 primary antibody used while very low signal intensity

was observed in the 1:400 dilution for the same primary antibodies (Figure 3.2 A –

Total). Similar intensities were observed for membrane bound protein expression of

occludin, claudin-1 and ZO-1 (Figure 3.2 A – Membrane). Cell normalisation staining

using a combination of cell and nuclear stains Sapphire700 and Draq5 respectively

demonstrated uniform cell densities in all wells (Figure 3.2 B). The target protein signal

intensities were then merged with the cell normalisation stains to allow quantification of

target protein expression (Figure 3.2 C). When the signal intensity was represented

graphically, 1:50 dilution of the occludin, claudin-1 and ZO-1 antibodies again

demonstrated the highest total signal intensity staining while very low signal intensity

was observed in the 1:400 dilution (Figure 3.2 D). Graphical representation of



3.2 E). Collectively, these results indicate that the 1:50 dilutions provided the strongest

signal intensity, however, it is considered an overly strong signal which may not

provide an accurate quantification of the target antibody signal. Furthermore, data

generated showed that the 1:400 dilutions provided the lowest signal intensity for all

antibodies tested and thus could also provide an inaccurate low quantification of the

target antibody signal. As a result, a mid-range dilution of 1:200 was selected for all

Figure 3.2 Effect of primary antibody concentration on paediatric derived

pAECHNA TJ signal intensity: Paediatric derived pAECHNA cells, seeded on 96-well

micro-titre plates and grown to confluence were treated as previously mentioned (refer

to 2.5.10). Incubation with primary antibodies to occludin, claudin-1 and ZO-1 was

performed at 4°C overnight followed by incubation with IRDye™ secondary antibodies.

(A) Total and membrane TJ signal intensity over dilution range between 1:50 to 1:400

of primary antibodies observed at 800 nm channel showed strongest intensity at 1:50

dilution while 1:400 dilution showed the weakest intensity staining. (B) Uniform cell

densities in all wells after staining with Sapphire700 and Draq5 indicates a confluent

monolayer as observed at 700 nm channel. (C) Merged image of (A) and (B) is utilised

for the quantification of TJ protein expression as normalised to cell densities. (D) Signal

intensity of occludin, claudin-1 and ZO-1 in a confluent pAECHNA monolayer post

normalisation to cell densities. Highest total TJ signal intensity was observed in 1:50

dilution of primary antibodies, however, 1:400 dilution resulted in minimal total TJ

signal intensity. (E) Similarly, membrane-bound signal intensity of occludin, claudin-1

and ZO-1 in a confluent pAECHNA monolayer post normalisation to cell densities was

observed to be highest in 1:50 dilution while 1:400 dilution demonstrated minimal

membrane TJ signal intensity.

Dilutions

Sign

al In

tens

ity

1:50 1:100 1:200 1:4000

1

2

3occludinclaudin-1ZO-1

Dilutions

Sign

al In

tens

ity

1:50 1:100 1:200 1:4000

1

2


D

E

A

B

C

Total Membrane



Merged

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primary antibodies and utilised in subsequent experiments for the detection of target

protein expression.

3.2.3.2 Incubation temperature of primary antibodies

As indicated by the manufacturer and performed here, incubation of primary antibodies

were performed at either 25°C for 2.5 h or at 4°C overnight. Initial experiments

performed in NuLi-1 cells at 25°C demonstrated strongest signal intensity at 1:50

dilution of occludin, claudin-1 and ZO-1 primary antibody used while very low signal

intensity was observed in the 1:400 dilution for the same primary antibodies (Figure 3.3

A – Total). Similar intensities were observed for membrane bound protein expression of

occludin, claudin-1 and ZO-1 (Figure 3.3 A – Membrane). Cell normalisation staining

using a combination of cell and nuclear stains Sapphire700 and Draq5 respectively

demonstrated uniform cell densities in all wells (Figure 3.3 B). The target protein signal

intensities were then merged with the cell normalisation stains (Figure 3.3 C) to allow

quantification of target protein expression. When the signal intensity was represented

graphically, 1:50 dilution of the occludin, claudin-1 and ZO-1 antibodies again

demonstrated the highest total signal intensity staining while very low signal intensity

was observed in the 1:400 dilution (Figure 3.3 D). Graphical representation of



3.3 E).

However, when the NuLi-1 cells were incubated with primary antibodies at 4°C

overnight, overall signal intensities for occludin, claudin-1 and ZO-1 primary antibodies

were stronger for total protein expression (Figure 3.4 A – Total). Similarly, overall

signal intensities of occludin, claudin-1 and ZO-1 primary antibodies were also stronger

for membrane bound protein expression (Figure 3.4 A – Membrane). When the signal

intensities were represented graphically, 1:50 dilution of occludin, claudin-1 and ZO-1

primary antibodies again demonstrated highest total protein expression while minimal

or non-detectable signal intensity was observed in the 1:400 dilution (Figure 3.4 D).

Figure 3.3 Effect of incubation temperature of primary antibody at 25°C on NuLi-

1 TJ signal intensity: NuLi-1 cells, seeded on 96-well micro-titre plates and grown to

confluence were treated as previously mentioned (refer to 2.5.10). Incubation with

primary antibodies to occludin, claudin-1 and ZO-1 was performed at 25°C for 2.5 h

followed by incubation with IRDye™ secondary antibodies. (A) Total and membrane

TJ signal intensity over dilution range between 1:50 to 1:400 of primary antibodies

observed at 800 nm channel showed strongest intensity at 1:50 dilution while 1:400

dilution showed the weakest intensity staining. (B) Uniform cell densities in all wells

after staining with Sapphire700 and Draq5 indicates a confluent monolayer as observed

at 700 nm channel. (C) Merged image of (A) and (B) is utilised for the quantification of

TJ protein expression as normalised to cell densities. (D) Total signal intensity of

occludin, claudin-1 and ZO-1 in a confluent NuLi-1 monolayer post normalisation to

cell densities. Highest total TJ signal intensity was observed in 1:50 dilution of primary

antibodies, however, 1:400 dilution resulted in minimal total TJ signal intensity

staining. (E) Similarly, membrane-bound signal intensity of occludin, claudin-1 and

ZO-1 in a confluent NuLi-1 monolayer post normalisation to cell densities was

observed to be highest in 1:50 dilution while 1:400 dilution demonstrated minimal

membrane TJ signal intensity staining.

Dilutions

Sign

al In

tens

ity

1:50 1:100 1:200 1:4000.0

0.5

1.0

1.5

2.0


Dilutions

Sign

al In

tens

ity

1:50 1:100 1:200 1:4000.0

0.5

1.0

1.5

2.0


1:50 1:100 1:200 1:4000.0

0.1

0.2

0.3

0.4

D

E

A

B

C

Total Membrane



Merged

Figure 3.4 Effect of incubation temperature of primary antibody at 4°C on NuLi-1

TJ signal intensity: NuLi-1 cells, seeded on 96-well micro-titre plates and grown to

confluence were treated as previously mentioned (refer to 2.5.10). Incubation with

primary antibodies to OCLN, CLDN-1 and ZO-1 was performed at 4°C overnight

followed by incubation with IRDye™ secondary antibodies. (A) Total and membrane

TJ signal intensity over dilution range between 1:50 to 1:400 of primary antibodies

observed at 800 nm channel showed strongest intensity at 1:50 dilution while 1:400

dilution showed the weakest intensity staining. (B) Uniform cell densities in all wells

after staining with Sapphire700 and Draq5 indicates a confluent monolayer as observed

at 700 nm channel. (C) Merged image of (A) and (B) is utilised for the quantification of

TJ protein expression as normalised to cell densities. (D) Total signal intensity of

occludin, claudin-1 and ZO-1 in a confluent NuLi-1 monolayer post normalisation to

cell densities. Highest total TJ signal intensity was observed in 1:50 dilution of primary

antibodies, however, 1:400 dilution resulted in minimal total TJ signal intensity. (E)

Similarly, membrane-bound signal intensity of occludin, claudin-1 and ZO-1 in a

confluent NuLi-1 monolayer post normalisation to cell densities was observed to be

highest in 1:50 dilution while 1:400 dilution demonstrated minimal membrane TJ signal

intensity.

Dilutions

Sign

al In

tens

ity

1:50 1:100 1:200 1:4000.0

0.5

1.0

1.5

2.0


Dilutions

Sign

al In

tens

ity

1:50 1:100 1:200 1:4000.0

0.5

1.0

1.5

2.0


1:50 1:100 1:200 1:4000.0

0.2

0.4

0.6

D

E

A

B

C

Total Membrane



Merged

Looi 2015

80

Graphical representation of membrane bound protein expression also demonstrated

similar observations with highest expression seen in 1:50 dilutions and lowest or no

detection seen in the 1:400 dilutions (Figure 3.4 E).

A similar series of experiments was also performed in pAECHNA to confirm initial

observations seen in the NuLi-1 cells. As demonstrated, the strongest signal intensity

was observed in the 1:50 dilution of occludin, claudin-1 and ZO-1 primary antibody

used, while very low signal intensity was observed in the 1:400 dilution for the same

primary antibodies (Figure 3.5 A – Total). Similar intensities were observed for

membrane bound protein expression of occludin, claudin-1 and ZO-1 (Figure 3.5 A –

Membrane). Cell normalisation staining using a combination of cell and nuclear stains

Sapphire700 and Draq5 respectively confirms uniform cell densities in all wells (Figure

3.5 B). The target protein signal intensities were then merged with the cell

normalisation stains (Figure 3.5 C) to allow quantification of target protein expression.

When the signal intensity was represented graphically, 1:50 dilution of the occludin,

claudin-1 and ZO-1 antibodies again demonstrated the highest total signal intensity

staining while very low signal intensity was observed in the 1:400 dilution (Figure 3.5

D). Graphical representation of membrane signal intensities also demonstrated similar

observations with highest intensity seen at 1:50 dilutions and lowest detection seen in

the 1:400 dilutions (Figure 3.5 E).

Interestingly and of significance, when the pAECHNA were incubated with primary

antibodies at 4°C overnight, overall signal intensities for occludin, claudin-1 and ZO-1

primary antibodies were stronger for total protein expression (Figure 3.6 A – Total).

Similarly, overall signal intensities of occludin, claudin-1 and ZO-1 primary antibodies

were also stronger for membrane bound protein expression (Figure 3.6 A – Membrane).

When the signal intensities were represented graphically, 1:50 dilution of occludin,

claudin-1 and ZO-1 primary antibodies again demonstrated highest total protein

expression while minimal or non-detectable signal intensity was observed in the 1:400

dilution (Figure 3.6 D). Graphical representation of membrane bound protein expression

also demonstrated similar observations with highest expression seen in 1:50 dilutions

and lowest or no detection seen in the 1:400 dilutions (Figure 3.6 E).

Figure 3.5 Effect of incubation temperature of primary antibody at 25°C on

paediatric derived pAECHNA TJ signal intensity: Paediatric derived pAECHNA cells,

seeded on 96-well micro-titre plates and grown to confluence were treated as previously

mentioned (refer to 2.5.10). Incubation with primary antibodies to occludin, claudin-1

and ZO-1 was performed at 25°C for 2.5 h followed by incubation with IRDye™

secondary antibodies. (A) Total and membrane TJ signal intensity over dilution range

between 1:50 to 1:400 of primary antibodies observed at 800 nm channel showed

strongest intensity at 1:50 dilution while 1:400 dilution showed the weakest intensity

staining. (B) Uniform cell densities in all wells after staining with Sapphire700 and

Draq5 indicates a confluent monolayer as observed at 700 nm channel. (C) Merged


normalised to cell densities. (D) Total signal intensity of occludin, claudin-1 and ZO-1

in a confluent NuLi-1 monolayer post normalisation to cell densities. Highest total TJ


dilution resulted in minimal total TJ signal intensity staining. (E) Similarly, membrane-

bound signal intensity of occludin, claudin-1 and ZO-1 in a confluent NuLi-1

monolayer post normalisation to cell densities was observed to be highest in 1:50

dilution while 1:400 dilution demonstrated minimal membrane TJ signal intensity

staining.

Dilutions

Sign

al In

tens

ity

1:50 1:100 1:200 1:4000.0

0.5

1.0


Dilutions

Sign

al In

tens

ity

1:50 1:100 1:200 1:4000.0

0.5

1.0


1:50 1:100 1:200 1:4000.0

0.1

0.2

0.3

0.4

0.5

D

E

A

B

C

Total Membrane



Merged

Figure 3.6 Effect of incubation temperature of primary antibody at 4°C on

paediatric derived pAECHNA TJ signal intensity: Paediatric derived pAECHNA cells,

seeded on 96-well micro-titre plates and grown to confluence were treated as previously

mentioned (refer to 2.5.10). Incubation with primary antibodies to occludin, claudin-1

and ZO-1 was performed at 4°C overnight followed by incubation with IRDye™

secondary antibodies. (A) Total and membrane TJ signal intensity over dilution range

between 1:50 to 1:400 of primary antibodies observed at 800 nm channel showed

strongest intensity at 1:50 dilution while 1:400 dilution showed the weakest intensity

staining. (B) Uniform cell densities in all wells after staining with Sapphire700 and

Draq5 indicates a confluent monolayer as observed at 700 nm channel. (C) Merged


normalised to cell densities. (D) Signal intensity of occludin, claudin-1 and ZO-1 in a

confluent pAECHNA monolayer post normalisation to cell densities. Highest total TJ


dilution resulted in minimal total TJ signal intensity. (E) Similarly, membrane-bound

signal intensity of occludin, claudin-1 and ZO-1 in a confluent pAECHNA monolayer

post normalisation to cell densities was observed to be highest in 1:50 dilution while

1:400 dilution demonstrated minimal membrane TJ signal intensity staining.

Dilutions

Sign

al In

tens

ity

1:50 1:100 1:200 1:4000

1

2


Dilutions

Sign

al In

tens

ity

1:50 1:100 1:200 1:4000

1

2


D

E

A

B

C

Total Membrane



Merged

Looi 2015

81

Collectively, when comparing the different incubation temperatures tested, the results

generated demonstrate that both the length and temperatures at which primary

antibodies are incubate do affect the eventual signal intensities measured, with those

incubated at 4°C overnight having higher signal intensities. As a result, these

parameters were utilised in all subsequent incubations of primary antibodies for this

assay.

3.2.3.3 Concentration of secondary antibodies

Once the optimal dilution and incubation temperature of the different primary

antibodies was determined, the final parameter examined was the concentration of

secondary antibodies used for the detection of primary antibodies. In an initial series of

experiments using NuLi-1 cells, a range of secondary antibody dilution from 1:200 to

1:1200 was utilised. As shown for total protein expression, the strongest signal intensity

was observed in the 1:200 dilution while very low signal intensity was observed at the

1:1200 dilution of the IRDye® secondary antibodies (Figure 3.7 A – Total). Similar

intensities were observed for membrane bound protein expression (Figure 3.7 A –

Membrane). When cells were co-stained with both Sapphire700 and Draq5, uniform cell

densities were observed throughout all wells (Figure 3.7 B). Target protein signal

intensities were then merged with the cell normalisation stains (Figure 3.7 C), the

resulting images were utilised for the analysis and semi-quantification of target protein

expression. When signal intensity was represented graphically, 1:200 dilution of the

IRDye® secondary antibodies demonstrated highest total protein expression while very

low signal intensity was observed in the 1:1200 dilution (Figure 3.7 D). Graphical

representation of membrane bound protein expression also demonstrated similar

observations with highest expression seen in 1:200 dilutions and very low intensity seen

in the 1:1200 dilutions (Figure 3.7 E).

In a similar series of experiments performed in pAECHNA, the observations seen in the

NuLi-1 cells were repeated and confirmed (Figure 3.8). As demonstrated, the strongest

Figure 3.7 Effect of secondary antibody concentration on NuLi-1 TJ signal

intensity: NuLi-1 cells, seeded on 96-well micro-titre plates and grown to confluence

were treated as previously mentioned (refer to 2.5.10). Incubation with primary

antibodies to occludin, claudin-1 and ZO-1 was performed at 4°C overnight (1:200

dilution) followed by incubation with IRDye™ secondary antibodies over a dilution

range between 1:200 to 1:1200. (A) Total and membrane TJ signal intensity over the

dilution range of secondary antibodies observed at 800 nm channel showed strongest

intensity at 1:200 dilution while 1:1200 dilution showed the weakest intensity staining.

(B) Uniform cell densities in all wells after staining with Sapphire700 and Draq5

indicates a confluent monolayer as observed at 700 nm channel. (C) Merged image of

(A) and (B) is utilised for the quantification of TJ protein expression as normalised to

cell densities. (D) Total signal intensity of occludin, claudin-1 and ZO-1 in a confluent

NuLi-1 monolayer post normalisation to cell densities. Highest total TJ signal intensity

was observed in 1:200 dilution of secondary antibody, however, 1:1200 dilution

resulted in minimal total TJ signal intensity staining. (E) Similarly, membrane-bound

signal intensity of occludin, claudin-1 and ZO-1 in a confluent NuLi-1 monolayer post

normalisation to cell densities was observed to be highest in 1:200 dilution while 1:1200

dilution demonstrated minimal membrane TJ signal intensity staining.

Dilutions

Sign

al In

tens

ity

1:200 1:400 1:800 1:12000.0

0.5

1.0

1.5


Dilutions

Sign

al In

tens

ity

1:200 1:400 1:800 1:12000.0

0.5

1.0

1.5


1:200 1:400 1:800 1:12000.0

0.1

0.2

0.3

0.4

0.5

D

E

A

B

C

Total Membrane



Merged

Figure 3.8 Effect of secondary antibody concentration on paediatric derived

pAECHNA TJ signal intensity: Paediatric derived pAECHNA cells, seeded on 96-well

micro-titre plates and grown to confluence were treated as previously mentioned (refer

to 2.5.10). Incubation with primary antibodies to occludin, claudin-1 and ZO-1 was

performed at 4°C overnight (1:200 dilution) followed by incubation with IRDye™

secondary antibodies over a dilution range between 1:200 to 1:1200. (A) Total and

membrane TJ signal intensity over the dilution range of secondary antibodies observed

at 800 nm channel showed strongest intensity at 1:200 dilution while 1:1200 dilution

showed the weakest intensity staining. (B) Uniform cell densities in all wells after

staining with Sapphire700 and Draq5 indicates a confluent monolayer as observed at

700 nm channel. (C) Merged image of (A) and (B) is utilised for the quantification of TJ

protein expression as normalised to cell densities. (D) Total signal intensity of occludin,

claudin-1 and ZO-1 in a confluent pAECHNA monolayer post normalisation to cell

densities. Highest total TJ signal intensity was observed in 1:200 dilution of secondary

antibody, however, 1:1200 dilution resulted in minimal total TJ signal intensity staining.

(E) Similarly, membrane-bound signal intensity of occludin, claudin-1 and ZO-1 in a

confluent pAECHNA monolayer post normalisation to cell densities was observed to be

highest in 1:200 dilution while 1:1200 dilution demonstrated minimal membrane TJ

signal intensity staining.

Dilutions

Sign

al In

tens

ity

1:200 1:400 1:800 1:12000

1

2


Dilutions

Sign

al In

tens

ity

1:200 1:400 1:800 1:12000

1

2


1:200 1:400 1:800 1:12000.0

0.5

1.0

1.5

D

E

A

B

C

Total Membrane



Merged

Looi 2015

82

signal intensity was observed in the 1:200 dilution of occludin, claudin-1 and ZO-1

primary antibody used while very low signal intensity was observed in the 1:400

dilution for the same primary antibodies (Figure 3.8 A – Total). Similar intensities were

observed for membrane bound protein expression of occludin, claudin-1 and ZO-1

(Figure 3.8 A – Membrane). Cell normalisation staining using a combination of cell and

nuclear stains Sapphire700 and Draq5 respectively demonstrated uniform cell densities

in all wells (Figure 3.8 B). The target protein signal intensities were then merged with

the cell normalisation stains (Figure 3.8 C) to allow quantification of target protein

expression. When the signal intensity was represented graphically, 1:50 dilution of the

occludin, claudin-1 and ZO-1 antibodies again demonstrated the highest total signal

intensity staining while very low signal intensity was observed in the 1:400 dilution

(Figure 3.8 D). Graphical representation of membrane signal intensities also

demonstrated similar observations with highest intensity seen at 1:50 dilutions and

lowest detection seen at the 1:400 dilutions (Figure 3.8 E). Collectively and

comparative to primary antibody results, the data generated indicated that the 1:200

dilutions provided the strongest signal intensity and was considered an overly strong

signal which may not provide an accurate quantification of the target protein. In

addition, it was also observed that the 1:400 dilutions provided the lowest signal

intensity and thus, conversely, provides an inaccurate low quantification of the target

protein signal. As a result, a mid-range dilution of 1:800 was selected for all secondary

antibody dilutions and utilised in subsequent experiments for the detection of target

protein expression.

3.2.4 Conclusion

From the experiments conducted, an optimised and specific ICW assay protocol was

established on which all subsequent assays utilising paediatric-derived primary AECs

were based (Figure 3.9). In these assays, paediatric-derived primary AECs were plated

onto 96-well microplates pre-coated with fibronectin coating buffer (10 mM) (refer to

2.4.2.3.1) at a high seeding density of 1.2 x 105 cells/cm2 and incubated at 37°C in an

atmosphere of 5% CO2 / 95% air in BEBM containing growth supplements as described

previously (refer to 2.4.2.2.6). Upon reaching confluence, growth media was aspirated

Figure 3.9 Methodology of In-Cell™ Western (ICW) assay: A schematic

representation of the optimised ICW assay utilised in this investigation.

(Adapted from Li-Cor BioSciences)

Cells were seeded at 1.2 x 105 cells / cm2 in BEBM containing growth supplements as previously described (refer to 2.4.2.2.4).

Cells were then treated with the desired stimuli in accordance with experimental design when confluent. If no stimulus is applied, proceed onto next phase.

Fixation and permeabilisation of cells.

Incubation with primary antibody at 4°C overnight.

Incubation the cells with IRDye® secondary antibody for 1 h at RT followed by wash.

Image plate with Li-Cor Biosciences Odyssey Imager at 700 nm and 800 nm.

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and cells were immediately fixed using 150 µl of 3.7% (v/v final) formalin (refer to

2.4.3.1.1) for 20 min at RT. Cells were then washed with 1X PBS containing 0.1% (v/v

final) Triton X-100 (refer to 2.4.3.1.2) (5 x 5 min / wash at RT) with gentle shaking to

permeabilise cells. If membrane proteins were to be detected, cells were incubated for

1.5 h at RT using 150 µl of LI-COR Odyssey Blocking Buffer. Cells were then

incubated overnight at 4°C with primary antibodies diluted in LI-COR Odyssey

Blocking Buffer (1:200). The following day, cells were washed with 1X PBS containing

0.1% (v/v final) Tween-20 (refer to 2.4.3.1.3) (5 x 5 min / wash at RT) with gentle

shaking and incubated with respective IRDye secondary antibodies, made up in LI-COR

Odyssey Blocking Buffer (1:800) with DRAQ5 (1:10000), Sapphire700 (1:1000) and

0.2% (v/v final) Tween-20 for 1 h at RT with gentle shaking in a dark room. Cells were

then washed in 1X PBS containing 0.1% (v/v final) Tween-20 (refer to 2.4.3.1.3) (5 x 5

min / wash at RT). Specific antibody staining for protein expression was then

immediately visualised using an infrared imaging system at both 680 and 800 nm

channels. Protein expression was quantified using the LI-COR Odyssey v.3.0 software

whereby the integrated intensity (I.I) of each well at 800 nm was normalised to the I.I of

the cell densities at 680 nm in the corresponding well and resulting protein expression

was graphically represented in arbitrary units.

3.3 Transepithelial permeability assay

3.3.1 Materials

All general materials and equipment used are as described in Chapter 2 (refer to 2.1 -

2.4).

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3.3.1.1 Cell lines and paediatric-derived primary airway epithelial cells

All cell lines and paediatric-derived primary AECs utilised in this investigation were

obtained and maintained as previously described (refer to 2.5.1.3, 2.5.1.6 and 2.5.4).

3.3.1.2 Fluorescein isothiocynate-dextran (FITC-dextran)

Fluorescein isothiocynate-dextran of either 4kDa or 20kDa molecular weight was


3.3.1.3 HEPES buffered Hank’s Balance Salt Solution (HEPES-HBSS)

HEPES buffered Hank’s Balanced Salt Solution was prepared and utilised as previously

described (refer to 2.5.11).

3.3.2 Methods

In order to optimise this assay specific to the investigation, a standard protocol derived

from the literature and used by various groups (Hubatsch et al. 2007; Sajjan et al. 2008;

Xiao et al. 2011) was utilised. Modifications were then performed and assessed on the

following parameters, 1; the selection of the fluorescent compound of interest for flux

assay and 2; sampling period.

3.3.2.1 Fluorescein isothiocynate labelled flux compounds

Previous studies have utilised various fluorescently labelled inert compounds including

inulin and dextran for the study of airway epithelium permeability (Ehrhardt et al. 2002;

Grainger et al. 2006; Dreschers et al. 2007; Xiao et al. 2011). However, fluorescently

labelled dextran (FITC-dextran) has been consistently utilised due to its availability in a

range of molecular weights. Thus, this study chose to utilise FITC-dextran at two

different molecular weights, namely 4 and 20 kDa, at a < 3 mg/ml concentration.

Looi 2015

85

3.3.2.2 Sampling period

From past transepithelial permeability studies performed (Ehrhardt et al. 2002; Grainger

et al. 2006; Sajjan et al. 2008), the length of sampling from the receiver compartment

was found to be an essential criterion in the analysis of flux and subsequently, the

determination of apparent permeability coefficient (Papp) across the airway epithelium.

Ehrhardt et al. (2002) as well as Grainger et al. (2006), performed transepithelial

permeability experiments over a period of 4 h, whilst others performed their

transepithelial permeability experiments over 6 h (Sajjan et al. 2008). Despite the

differences in the length of sampling between these studies, it remains unknown as to

whether an increase in the sampling period would result in a more precise flux analysis

and ultimately, more accurate determination of Papp coefficients. Thus, in this

investigation, the transepithelial permeability assay was performed over a period of 6 h.

3.3.3 Results / Discussion

3.3.3.1 Fluorescein isothiocynate labelled dextran

The transepithelial permeability assay is based upon the flux of fluorescently labelled

inert compounds from the apical surface layer of cultured epithelial cells to the

basolateral layer. Therefore, one of the first criteria to be established was the selection

of a suitable fluorescently labelled inert compound. As indicated earlier, various

fluorescently labelled compound have been utilised, however, the most commonly used

compound has been fluorescein isothiocynate labelled dextran (FITC-dextran) (Balda et

al. 1996; Ehrhardt et al. 2002; Forbes and Ehrhardt 2005; Grainger et al. 2006; Xiao et

al. 2011). The advantage of FITC-dextran is the availability in a range of molecular

sizes from 4 kDa to 2000 kDa. This allows for the determination of transepithelial

permeability of the airway epithelium towards different molecular sizes and most

importantly, provides a comparative to known respiratory allergens, bacteria and viruses

of known sizes. For this study, FITC-dextran of molecular sizes 4 kDa and 20 kDa were

utilised as it allowed for the investigation of transepithelial permeability of a small and

Looi 2015

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larger sized molecules across the epithelial layer of healthy AECs. Moreover, the two

different molecular size of FITC-dextran provides the ability to add new information

about epithelial permeability as well as addressing the hypotheses and aims of this

study.

The studies utilising FITC-dextran have done so at a number of different concentrations

(Coyne et al. 2002; Grainger et al. 2006; Xiao et al. 2011). However, in most of them,

the concentration is always greater than 1 mg / ml (w/v final). Thus, in this

investigation, a concentration of 2 mg / ml (w/v final) was selected in order to provide a

higher concentration of FITC-dextran within the apical surface to maintain the sink

conditions in the receiver chamber within the culture well.

3.3.3.2 Sampling period

A variety of studies utilising the transepithelial permeability assay have been performed

over different lengths of time. Ehrhardt et al. (2002) as well as Grainger et al. (2006), in

their studies, performed the transepithelial permeability assay over a period of 4 h while

in other studies by Hubatsch et al (2007) and Sajjan et al. (2008), the transepithelial

permeability assay was performed over 6 h. In this investigation, a series of experiments

utilising different cell types were initially conducted over 4 h and subsequently, 6 h to

ascertain whether the length of the sampling period had any effect on absorbance and

ultimately, permeability coefficient levels. As demonstrated, an increase in absorbance

was observed over 4 h of sampling time in Caco-2, NuLi-1 and pAECHNA cells (Figures

3.10 A, B, and C respectively). However, when the experiments were extended over 6 h,

a further increase in absorbance was observed at the 5 h sampling interval. Absorbance

values continue to increase within the Caco-2 cells (Figure 3.10 A) while a plateau in

absorbance values was attained by the 6 h sampling interval in NuLi-1 and pAECHNA

cells (Figures 3.10 B & C). Collectively, the data generated suggest that maximum

absorbance is achieved at 6 h which then corresponds to a maximum calculated

permeability coefficient. As a result, a sampling period of 6 h was then utilised in all

subsequent transepithelial permeability assays.

Figure 3.10 Effect of sampling time on FITC-dextran molecules across cell

monolayers: Cells were cultured on Corning transwell inserts and grown to confluence

and transepithelial permeability assay performed as previously mentioned (refer to

2.5.11). An increase in FITC-dextran molecules of both 4kDa and 20kDa in the

basolateral compartment is observed over 4 h in confluent monolayers of (A) Caco-2

cells, (B) NuLi-1 cells and (C) pAECHNA cells. (A) When sampling time was increased

over 6 h, a further increase in both FITC-dextran molecules was observed within the

basolateral compartment in confluent monolayers of Caco-2 cells. (B and C) An

increase in both FITC-dextran molecules was similarly observed within the basolateral

compartment in confluent monolayers of NuLi-1 cells and pAECHNA cells at the 5 h

time point before a plateau was attained at the 6 h time point for both cell types. Results

are the mean ± SEM of five samples.

Time (h)

Abso

rban

ce a

t 520

nm

0 1 2 3 40.0

0.5

1.0

1.5

5 6

4kDa20kDa

Time (h)

Abso

rban

ce a

t 520

nm

0 1 2 3 40

50

100

150

5 6

20kDa4kDa

Time (h)

Abso

rban

ce a

t 520

nm

0 1 2 3 40

10

20

30

40

50

5 6

20kDa4kDa

A

B

C

Looi 2015

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3.3.4 Conclusion

From the experiments conducted in this chapter, a standard transepithelial permeability

assay protocol was established on which all subsequent assays utilising paediatric-

derived primary AECs of this thesis were based (Figure 3.11). Here, primary AECs

were seeded onto culture inserts (Corning Incorporated Life Sciences, MA, USA) pre-

coated with fibronectin coating buffer at a density of 1.2 x 105 cells/cm2. Culture inserts

were then incubated at 37°C in an atmosphere of 5% CO2 / 95% air in BEBM

containing growth supplements as described previously (refer to 2.4.2.2.6). Growth

media within the apical and basolateral compartments was replaced with HEPES

buffered Hank’s balanced salt solution (HEPES-HBSS) (refer to 2.4.3.3.2). FITC-

dextran of molecular weight 4kDa or 20kDa (2 mg/ml w/v final) (refer to 2.4.3.3.1) was

added to the apical chamber and 50 µl of apical solution was immediately sampled. Five

hundred microliters of HEPES-HBSS from the basolateral compartment was sampled at

hourly intervals over a period of 6 h. The volume of the basolateral compartment was

maintained by addition of 500 µl fresh buffered HBSS-HEPES. All experiments were

performed at 37°C and on a calibrated orbital shaker at 100 rpm to minimise the

unstirred buffer layer. Fluorescence of FITC-dextran was detected using a PerkinElmer

Enspire® multilabel plate reader at an excitation wavelength of 492 nm and emission

wavelength of 520 nm. The apparent permeability of the epithelial monolayer to FITC-

dextran from the apical to basolateral compartment (Papp) was then calculated following

the general equation: Papp = (dQ/dt) x (1/AC0) where dQ/dt is the steady –state flux, A is

the surface area of the membrane and C0 is the initial concentration in the donor

compartment as previously described (Stutts et al. 1981).

Figure 3.11 Methodology of Transepithelial permeability assay: A schematic

representation of the optimised transepithelial permeability assay utilised in this

investigation.

Cells were seeded at 1.2 x 105 cells / cm2 in BEBM containing growth supplements as previously described (refer to 2.4.2.2.6).

Cells were then treated with the desired stimuli in accordance with experimental design when confluent. If no stimulus is applied, proceed onto next phase.

Growth media replaced with HEPES-HBSS in basolateral chamber. FITC-dextran (2mg/ml w/v final) in HEPES-HBSS added to apical chamber.

Over 6 h, 500µl of HEPES-HBSS sampled from basolateral chamber at determined time point and subsequently replaced with fresh HEPES-HBSS.

Sampled HEPES-HBSS then transferred to 96 well micro-titre plate.

Fluorescence detected via PerkinElmer Enspire® multilabel plate reader at excitation wavelength of 492 nm and emission wavelength of 520 nm.

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CHAPTER 4: Effect of human rhinovirus infection on tight junction

disassembly and the subsequent changes to barrier function

4.1 Introduction

A pseudostratified mucosal barrier consisting of a multitude of cell types function as the

initial protective interface between the internal milieu of the lung and the external

environment of micro-organisms, aeroallergens and noxious gases within the human

respiratory tract. A range of junctional complexes from tight junctions (TJ) to connexins

constitutes this protective barrier. Tight junctions that are located at the apical borders

of adjacent epithelial cells play a major role in the maintenance of epithelial barrier

function. They serve to regulate the movement of ions and solutes as well as to prevent

unwanted migration of pathogens and their products to the sub-epithelial space. Thus, a

breach in the epithelial barrier function, such as disruption in the TJ protein, zonula

occludens protein-1 (ZO-1) (Sajjan et al. 2008) may increase the paracellular traffic of

pathogenic molecules into the interstitium, resulting in the release of pro-inflammatory

cytokines such as interlukin-8 (IL-8), eotaxin, regulated on activation, normal T-cell

expressed and secreted (RANTES), interferon-inducible protein-10 (IP-10) and

macrophage inflammatory protein 1α (Wark et al. 2007).

Compromisation of the airway barrier integrity occurs after exposure to injurious

stimuli including airborne pollutants, aeroallergens, bacteria and respiratory viruses.

Respiratory viruses such as human rhinovirus (HRV) are common insults of the airway

epithelium and despite evidence suggesting that a significant reduction in TJ protein

expression and a disassembly of the apical junction proteins is associated with an

attenuated barrier function (Sajjan et al. 2008; Rezaee et al. 2011; Xiao et al. 2011), the

molecular mechanisms involved in the regulation of airway epithelial TJ proteins

following live HRV infection and the effects on barrier function are still not well

understood. Hence, this investigation was conducted to establish a direct correlation

between HRV infection, altered TJ expression and transepithelial permeability by

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utilising a modified human AEC, NuLi-1 with the hypothesis that TJ gene expression

would be altered following HRV infection which translates into a disassembly of TJ

complexes, resulting in a change in epithelial permeability. Therefore, using healthy

AECs, the specific aims of this chapter was to assess mRNA and protein of TJ

expression changes following HRV-1B infection using qPCR focussed arrays and in-

cell westerns. Furthermore, effects of HRV-1B infection on barrier function were

directly correlated using the optimised transepithelial permeability assay.

4.2 Materials and Methods

The general materials and methods used in this part of the investigation are listed in

detail in Chapter 2.

4.2.1 Cell culture

The modified human AEC, NuLi-1 and the sub-culture methodology used in this

investigation has been described in detail in Chapter 2 (refer to 2.4.2.2.6, 2.5.1.6, 2.5.2)

4.2.2 Human rhinovirus and titrations

Human rhinovirus (HRV) minor serotype 1B (HRV-1B) utilised in this investigation

has been described in detail in Chapter 2 (refer to 2.5.7).

4.2.3 Human tight junction Polymerase Chain Reaction (PCR) arrays

To assess for genes associated with TJ disassembly and altered barrier function, a

focused PCR array consisting of 84 key genes encoding for proteins forming the

epithelial barrier was utilised. Briefly, total cellular RNA was initially extracted as

previously described (refer to 2.5.13) and RNA quality and quantity assessed using a

NanoDrop. Reverse transcription was performed according to the manufacturer’s

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guidelines using the provided RT2 First Strand Kit which converts 1000 ng of RNA

into cDNA. A 1000 ng sample of RNA was added to a reverse transcription mix

containing 5X Buffer BC3 (4 µl), Control P2 (1 µl), RE3 Reverse Transcription Mix (2

µl) and made to a final volume of 10 µl with RNase free water. Samples were then

placed in a thermal cycler and run on a RT2 Profiler™ PCR array reverse transcription

program of 95°C for 10 min, 95°C for 15 sec and 60°C for 1 min. RT-qPCR was

performed on an ABI Prism® 7300 (refer to 2.3.15).

4.2.4 Infection of cell cultures

The modified human AEC, NuLi-1, grown on culture plates or inserts until confluence,

were infected with HRV-1B and incubated for 24 h. Briefly, growth media was replaced

with fresh basal media and subsequently infected with HRV-1B at a 50% Tissue Culture

Infectivity Dose (TCID50) of 2.5, 10, 20, 40 or 80 x 104 TCID50/ml. After each

incubation period, media was collected and stored at -80°C and infected cells were

utilised for various downstream assays.

4.2.5 In Cell™ Western assay

The optimised In Cell™ Western assay, as described in detail (refer to Chapter 3) was

utilised for the determination of TJ membrane protein expression prior and following

human rhinovirus infection.

4.2.6 MTS cell viability assay

For the assessment of NuLi-1 cell viability, a CellTitre 96® AQueous Non-Radioactive

Cell Proliferation Assay (Promega, Madison, WI, USA) was utilised. This is a

colorimetric assay based upon the conversion of a tetrazolium salt into a coloured

compound by dehydrogenase enzymes found only in metabolically active cells. The

assay was performed in accordance with the manufacturer’s instructions and

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measurements were recorded at 0, 24, 48 and 72 h post HRV infection to determine the

percentage of viable cells.

4.2.7 Quantification of Human Rhinovirus viral copy number

Viral copy number of all HRV samples was determined quantitatively via two-step RT-

PCR reactions using a HRV-1B advanced kit and HRV standard kit (both

PrimerDesign Ltd, UK) in combination with MultiScribe™ Reverse Transcriptase

and Taqman Universal Master Mix (Applied Biosystems, USA) as previously

described (Sutanto et al. 2011). Briefly, 200 ng total RNA was reverse transcribed in a

20 l total reaction volume containing 1X RT buffer, 5.5 mM MgCl2, 0.5 mM of each

of the dNTPs, 1 µl HRV-1B/ACTB primer mix, 0.4 U/l RNase inhibitor, 0.5 U/l

MultiScribe reverse transcriptase and RNase-DNase free water. The reactions were

carried out as follow: initial primer incubation step at 25C for 10 minutes followed by

1 h incubation at 48C and ended by heating at 95C for 5 minutes. The cDNA was then

used in a final PCR reaction volume of 20 l containing 1X Taqman Universal Master

Mix, 1 l HRV-1B primer/probe mix and 5 l of cDNA which has been diluted 5-fold.

The PCR conditions were as described by manufacturer: 50C for 2 minutes, 95C for

10 minutes followed by 40 cycles of 15 seconds at 95C and 1 minute at 60C. A copy

number was determined from a set of standards ranging from 2 copy number/mL to

2x107 copy number/ml that was included in each run.

4.2.8 Real-Time Quantitative Polymerase Chain Reaction (RT-qPCR)

Extraction and quantification of RNA, as well as the methodology for RT-qPCR has

been described in detail in Chapter 2 (2.5.13).

4.2.9 Single-stranded DNA (ssDNA) apoptosis assay

To determine the percentage of cells that underwent apoptosis during HRV infection, a

ssDNA apoptosis ELISA kit was utilised. This procedure is based upon the selective

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denaturation of DNA in apoptotic cells by formamide and the detection of denatured

DNA with a specific monoclonal antibody for ssDNA. The assay was performed in

accordance with the manufacturer’s instructions as previously described (Sutanto et al.

2011). Briefly, cells were plated onto 96-well microplates pre-coated with fibronectin

(10 mM) at a seeding density of 6 x 104 cells/cm2 and incubated at 37°C in an

atmosphere of 5% CO2 / 95% air in BEBM containing growth supplements as described

previously (refer to 2.4.2.2.6). Cells were then infected with HRV-1B at three viral

concentrations: 2.5 x 104 TCID50/ml, 10 x 104 TCID50/ml and 80 x 104 TCID50/ml for

24 h. The plates were then centrifuged at 200 g for 5 minutes and the media collected

and replaced with 200 µl of 80% (v/v final) methanol fixative and incubated at RT for

30 minutes. The fixative was removed from the cell monolayers and the plates dried at

RT for 1-2 h to allow for permanent attachment of cells to the plate. Once fully dry, 50

µl of formamide solution was added to each well and incubated at RT for 10 minutes.

To denature the DNA in apoptotic cells, plates were heated to 75°C for 10 minutes in an

oven, cooled in a refrigerator for 5 minutes and finally the formamide removed. Wells

were then rinsed 3 times with 1X PBS and blocked with 200 µl of 3% (w/v final) non-

fat milk solution for 1 h at 37°C. The blocking solution was removed and replaced with

100 µl of supplied antibody mixture to each well followed by a 30 minutes incubation at

RT. Plates were then washed a further 3 times with 250 µl of 1X wash solution and 100

µl of supplied ABTS solution added to each well and incubated for 30 minutes at RT.

The reaction was stopped by the addition of 100 µl of stop solution and resulting

absorbance read at 405 nm.

4.2.10 Transepithelial permeability assay

The optimised transepithelial permeability assay, as described in detail (refer to Chapter

3) were utilised for the functional determination of epithelial TJ membrane integrity

prior and following human rhinovirus infection.

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Before statistical evaluation, all results were tested for population normality and

homogeneity of variance, and where applicable, a Student t test was performed.

Experiments were performed in at least in triplicates and all data were non-parametric,

with analysis performed using Mann-Whitney test. Values were presented as mean ±

SD. All p values less than 0.05 were considered to be significant.

4.3 Results

4.3.1 Effect of human rhinoviral infection on NuLi-1 cell viability

Although HRV has been reported to have a cytotoxic effect on pAECs (Stevens 2009),

the effects on epithelial TJ and subsequently barrier function, remains largely

unanswered. Hence, to determine the effects of HRV infection on epithelial barrier

integrity and function in a modified human AEC, NuLi-1, initial validation of cellular

viability response upon infection by HRV-1B was conducted. Human rhinovirus

cytotoxicity assays were performed to determine the effects of HRV-1B exposure on

NuLi-1 cellular viability. The assays were performed using viral titres from 2.5 to

80x104 TCID50/ml HRV-1B and infection times of 24 to 72 h to assess both dose

response and time course effects.

Infection of NuLi-1 cells with HRV-1B affected culture viability in both a time and

dose dependent manner. Following infection with a viral titre of 2.5x104 TCID50/ml of

HRV-1B, despite significant loss of cell viability at 24 h (93.2% ± 2.2) and 48 h (93.6%

± 1.2) but no significant loss at 72 h (95.2% ± 5.2), infection with the low viral titre

demonstrated the least effect on cellular viability at all infection times (Figure 4.1 -

2.5x104 TCID50/ml; p<0.05). Interestingly, no significant loss of viability was observed

when infected with viral titre of 10x104 TCID50/ml of HRV-1B at 24 h (100% ± 2.8)

and 48 h (93.7% ± 4.1) but significance was observed when infection was extended to

72 h (79.6% ± 3.1) (Figure 4.1 - 10x104 TCID50/ml; p<0.05). However, when infected

Figure 4.1 Effect of HRV-1B on cellular viability in NuLi-1 over time: NuLi-1 cells, seeded on 96-well micro-titre plates were grown to confluence

subsequently infected with a range of HRV-1B titres (2.5 – 80x104 TCID50/ml) and cell viability assessed at 24, 48 and 72 h post infection via a

colorimetric MTS assay as described (refer to 4.2.6). Results were presented as whisker box plots (mean, Min – Max) in percentages from at least three

different experiments with each data assayed in triplicate and normalised to non-infected control cells (---). Infection with HRV-1B caused a time and

dose-dependent cytotoxic effect on NuLi-1 cells. *Statistical significance relative to control (p < 0.05).

Viral titre (x104 TCID50/ml)

Cel

l Via

blity

(% r

elat

ive

to c

ontr

ol)

Control 80 40 20 10 2.5 80 40 20 10 2.5 80 40 20 10 2.5

0

25

50

75

100

12572h48h24h

*

*

*

** **

* *

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with viral titre of 20x104 TCID50/ml of HRV-1B, increasing significant loss of viability

was observed at 24 (88.8% ± 3.7), 48 (91.1% ± 1.2) and 72 h (60.8% ± 4.8) (Figure 4.1

- 20x104 TCID50/ml; p<0.05). When infected with viral titre of 40x104 TCID50/ml of

HRV-1B, no significant loss of viability was observed at 24 (82.3% ± 10) and 48 h

(79.2% ± 8.3) but significance was observed when infection was extended to 72 h

(32.1% ± 4.8) (Figure 4.1 - 40x104 TCID50/ml; p<0.05). Infection with the maximal

viral titre of 80x104 TCID50/ml of HRV-1B demonstrated significantly increased loss of

cellular viability at 24 (91.8% ± 0.2), 48 h (81.4% ± 6.3) with the greatest loss of

viability observed at 72 h (25.9% ± 0.6) (Figure 4.1 - 80x104 TCID50/ml; p<0.05).

4.3.2 Apoptotic response and viral replication following infection with HRV-1B

Apoptotic responses in virally infected cells are key protective mechanisms to prevent

and minimise viral replication and release. Previous studies have demonstrated an

apoptotic response in pAECs following HRV-1B infection (Stevens 2009; Sutanto et al.

2011), however, prior utilising NuLi-1 cells for the determination of tight junctional

expression and response to HRV infection, initial validation of cellular apoptotic

response to HRV-1B infection was conducted. Hence, a ssDNA apoptosis ELISA was

performed to determine the extent of apoptosis in NuLi-1 cells following infection with

HRV-1B.

Infection with HRV-1B induced significant apoptotic response at all viral titres,

however, the level of apoptosis induced was dependent on the viral titre, with a 50%

increase in apoptosis at a low viral titre of 2.5x104 TCID50/ml of HRV-1B compared to

non-infected controls (Figure 4.2A; p<0.05). Furthermore, determination of viral copy

number in HRV infected samples after 24 h infection with viral titres of 2.5 to 40x104

TCID50/ml of HRV-1B demonstrated significant increase in viral copy numbers,

indicative of increased viral replication, which was concomitant with increasing viral

titres (Figure 4.2B; p<0.05).

Figure 4.2 Effect of HRV-1B on apoptosis and viral replication in NuLi-1: (A) NuLi-1 cells, seeded on 96-well micro-titre plates and grown to

80% confluence were infected with a range of HRV-1B titres (2.5 – 40x104 TCID50/ml) for 24 h and apoptotic response of the cells determined using a

colorimetric assay ssDNA apoptosis kit (refer to 4.2.9). Following infection with HRV-1B, overall apoptotic response increases with significance

observed at all viral titres (p<0.05). Data were presented as mean ± SD percentage apoptosis relative to control from at least three different experiments

with each data assayed in triplicate. (B) NuLi-1 cells were established and infected with a range of HRV-1B titres (2.5 – 40x104 TCID50/ml) for 24 h.

Cells were harvested to extract RNA and HRV-1B RNA measured using qPCR. Viral copy number increased with increasing viral titre and was

significant for all viral titres used (p<0.05). Data were normalised to microgram RNA and presented as mean ± SD; n = 3 individual experiments each

performed in duplicates. *Statistical significance relative to control (p<0.05). **Statistical significance relative to control (p<0.01).

A B

Control 2.5 10 20 400

50

100

150

200

250 **


Incr

ease

in a

popt

osis

(% r

elat

ive

to c

ontr

ol)

****

*

Control 2.5 10 20 400

10

20

30

40

50


Vira

l cop

y #/

g R

NA

*

*

**

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4.3.3 Effect of HRV-1B infection on mRNA expression of tight junction complexes

In order to directly correlate HRV infection with altered expression in tight junctional

and associated proteins, a focussed qPCR array was performed. Briefly, RNA was

collected from non-infected and infected cultures, isolated, purified and transferred onto

a focussed array which contained primers for 84 key genes encoding for proteins that

form impermeable barriers between epithelial cells to regulate polarity, proliferation and

differentiation.

Results demonstrated that following infection with viral titre of 20x104 TCID50/ml of

HRV-1B for 9 h, down-regulation of mRNA was observed for 58 key genes encoding

for epithelial barrier junction proteins with the most down-regulated being CLDN8

(14.2-fold) and up-regulation of 26 key genes with the most up-regulated being Crb3

(1.9-fold). Interestingly, from the data, significant down-regulation was only observed

for CLDN8 (14.2-fold), PTEN (3.5-fold), CLDN12 (2.3-fold), ASH1L (1.9-fold), and

ZO-1 (1.3-fold) (Figure 4.3 – CLDN8, PTEN, CLDN12, ASH1L, ZO-1; p<0.05).

However, of relevance to the barrier integrity of the respiratory epithelium, the

ubiquitously expressed claudin-1, occludin and ZO-1 were analysed following HRV-1B

infection. The data generated demonstrated down-regulation of mRNA for claudin-1

(1.1-fold), occludin (1.2-fold) and ZO-1 (1.3-fold), however, significance was only

observed for ZO-1 (Figure 4.3 – ZO-1; p<0.05).

4.3.4 Effect of human rhinovirus infection on membrane tight junction disassembly

At present, there remains a paucity of data on the effects HRV infection has on other

tight junctional complexes despite evidence from a seminal study demonstrating

disassembly of ZO-1 protein (Sajjan et al. 2008). Hence, an In-Cell™ Western assay

(refer to Chapter 3) was utilised to corroborate membrane protein disassembly of

claudin-1, occludin and ZO-1 with their respective down-regulated mRNA expression

following HRV-1B infection.

Figure 4.3 Effect of HRV-1B on mRNA expression of TJ in NuLi-1: NuLi-1 cells, seeded on 12-well plates and grown to confluence were infected

with a viral titre of 20x104 TCID50/ml of HRV-1B for 9 h. Cells were harvested to extract RNA and human tight junction gene expression assessed

using a focused RT-qPCR array (refer to 4.2.3). Down-regulation of mRNA was observed in 58 key genes with significance observed in CLDN8,

PTEN, CLDN12, ASH1L and ZO-1 following HRV-1B infection. Down-regulation was also observed for the identified genes of interest (Blue) with

significance observed for ZO-1. Data were presented as fold up- or down-regulation from at least three different experiments relative to non-infected

control cells. *Statistical significance relative to non-infected control (p<0.05).

Human TJ genes

mR

NA

expr

essi

on(fo

ld u

p- o

r do

wn-

regu

latio

n re

lativ

e to

con

trol

)

CLD

N8

PEC

AM

1SP

TA1

PTEN

CLD

N19

MA

GI2

IGSF

5ES

AM

CLD

N12

ICA

M2

AR

HG

EF2

ASH

1LZA

KC

LDN

14C

LDN

3SP

TBM

LLT4

CLD

N9

CR

B1

VAPA

PAR

D3

TJP3

CLD

N15

CLD

N5

CSN

K2A

2C

TNN

A3

PAR

D6B

JAM

2A

CTN

2M

AR

K2

EPB

41C

ASK

F11R

TJP1

GN

AI1

CD

99M

PDZ

MA

GI1

OC

LNJA

M3

RA

C1

LLG

L2C

GN

CLD

N1

RH

OA

CTT

NC

LDN

18TJ

AP1

INA

DL

CTN

NA

1SY

MPK

MPP

6A

CTN

4LL

GL1

CLD

N6

AC

TN1

ILK

SMU

RF1

AC

TN3

ICA

M1

CLD

N7

CD

K4

CTN

NB

1A

MO

TL1

CLD

N10

CTN

NA

2TJ

P2C

LDN

11PR

KC

IC

SDA

MPP

5H

CLS

1C

LDN

17SP

TAN

1C

SNK

2A1

CLD

N2

PRK

CZ

TIA

M1

CD

C42

CLD

N16

CLD

N4

CSN

K2B

PAR

D6A

CR

B3

-20

-15

-10

-5

0

5

*

*

** *

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The data demonstrated high basal membrane occludin expression (0.038AU ± 0.007)

within NuLi-1 cells, while, basal membrane claudin-1 and ZO-1, which demonstrated

similar levels of expression were lower (0.015AU ± 0.002 and 0.016AU ± 0.002

respectively) when compared to membrane occludin expression (Figure 4.4 - Control).

Following infection with a viral titre of 2.5x104 TCID50/ml of HRV-1B for 24 h, a

decrease in membrane occludin and ZO-1 expression was observed (0.02AU ± 0.001

and 0.012AU ± 0.0004 respectively), although this was not statistically significant.

However, a significant decrease in membrane claudin-1 expression (0.007AU ± 0.0002)

was observed at the same viral titre (Figure 4.4 - 2.5x104 TCID50/ml; p<0.05). When

NuLi-1 cells were infected with a high viral titre of 20x104 TCID50/ml of HRV-1B,

significant decreases in all membrane protein expressions of claudin-1 (0.007AU ±

0.0001), occludin (0.016AU ± 0.002) and ZO-1 (0.007AU ± 0.00009) were observed in

contrast to non-infected controls (Figure 4.4 – 20x104 TCID50/ml; p<0.05).

4.3.5 Effect of human rhinovirus infection on transepithelial permeability

Epithelial paracellular permeability is a key functional indicator of cellular junctional

integrity. Hence, to correlate HRV-1B induced disassembly of membrane TJ with

barrier function, a transepithelial permeability assay was performed after 24 h infection

with a low (2.5x104 TCID50/ml) and high viral titre (20x104 TCID50/ml) of HRV-1B.

From the data generated following infection with a low viral titre (2.5x104 TCID50/ml)

of HRV-1B for 24 h, a significant increase in permeability to FITC-dextran of 4 kDa

(292.6 x 10-4 cm/sec ± 2.4) and 20 kDa (249.8 x 10-4 cm/sec ± 10.33) was observed

when compared to non-infected controls (221.9 x 10-4 cm/sec ± 8.5 and 115 x 10-4

cm/sec ± 4.5 respectively) (Figure 4.5 - 2.5x104 TCID50/ml; p<0.05). Moreover,

permeability to FITC-dextran 4 kDa was observed to be significantly higher (292.6 x

10-4 cm/sec ± 2.4) when compared to 20 kDa (249.8 x 10-4 cm/sec ± 10.33) following

infection with the low viral titre of HRV-1B. Interestingly, when NuLi-1 cells were

infected with a high viral titre (20x104 TCID50/ml) of HRV-1B for 24 h, significant

increase in transepithelial permeability of FITC-dextran 4 kDa (357.9 x 10-4 cm/sec ±

12.2) and 20 kDa (302.2 x 10-4 cm/sec ± 7.2) was similarly observed compared to non-

Figure 4.4 Effect of HRV-1B infection on membrane TJ protein expression in

NuLi-1: NuLi-1 cells, seeded on 96-well micro-titre plates and grown to confluence

were treated as previously mentioned (refer to 2.5.10). Cells were exposed to two

different HRV-1B titres (2.5 and 20x104 TCID50/ml) for 24 h and membrane TJ protein

expression assessed via a previously optimised In-Cell™ Western assay (refer to 4.2.5).

As observed, overall membrane claudin-1, occludin and ZO-1 protein expression

decreased following exposure to HRV-1B at both viral titres. Significance was observed

for claudin-1 at the lower titre while significance was observed for all three TJ protein

at the higher viral titre (p<0.05). Data were normalised to cell numbers and presented as

mean ± SD; n = 5 individual experiments each performed in duplicates. *Statistical

significance relative to control (p<0.05).


Mem

bran

e pr

otei

n ex

pres

sion

(Arb

itary

uni

ts)

Control 2.5 200.00

0.01

0.02

0.03

0.04

0.05

claudin-1occludinZO-1

* *

*

*

Figure 4.5 Effect of HRV-1B infection on transepithelial permeability in NuLi-1:

NuLi-1 cells, seeded onto Corning transwell inserts and grown to confluence were

infected with two different HRV-1B titres (2.5 and 20x104 TCID50/ml) for 24 h and

epithelial permeability assessed via a transepithelial permeability assay (refer to 4.2.10).

As observed, overall transepithelial permeability to FITC-dextran 4 kDa and 20 kDa

was significantly increased at both viral titres (p<0.05). Results are presented as mean ±

SD; n = 5 individual experiments each performed in duplicates. *Statistical significance

relative to control (p<0.05). #Statistical significance relative to 20 kDa (p<0.05).

Control 2.5 200

100

200

300

4004 kDa20 kDa

*#

*

*


Papp

coe

ffici

ent (

cm/s

ec) x

10-4

*#

#

Looi 2015

97

infected controls (221.9 x 10-4 cm/sec ± 8.5 and 115 x 10-4 cm/sec ± 4.5 respectively)

(Figure 4.5 - 20x104 TCID50/ml; p<0.05). In addition, permeability to FITC-dextran 4

kDa was observed to be significantly higher (357.9 x 10-4 cm/sec ± 12.2) when

compared to 20 kDa (302.2 x 10-4 cm/sec ± 7.2) following infection with the low viral

titre of HRV-1B. Interestingly, when comparing transepithelial permeability between

the viral titres, permeability to FITC-dextran of 4 and 20 kDa was observed to be

significantly higher following infection with the high viral titre in contrast to epithelial

permeability post infection with the low viral titre.

4.4 Discussion

The associations between HRV infection, expression and regulation of TJ genes and

subsequent disassembly of TJ proteins resulting in impairment of epithelial barrier

functions are poorly understood. Various studies have used cell lines or pAECs for

understanding the effects of viral infection on membrane TJ dissociation alone or

disassembly in membrane TJ proteins leading to an observed increase in epithelial

permeability (Sajjan et al. 2008; Comstock et al. 2011; Xiao et al. 2011). However, no

study has yet incorporated both and at present, this is the first study to examine a direct

correlation between live-viral infection and the effects on TJ gene expression,

disassembly of membrane TJ proteins and eventual impairment of barrier function

resulting in increased transepithelial permeability.

Airway epithelial cells are of vital importance during viral infections as they serve as

the host cell for viral replications as well as initiating the innate and adaptive immune

responses. Past studies have demonstrated that pAECs are susceptible to HRV

infections and are able to successfully replicate resulting in a cytotoxic effect (Subauste

et al. 1995; Papadopoulos et al. 2000; Sutanto et al. 2011; Cakebread et al. 2014).

However, limited access to pAECs as well as variability between different donor

epithelia often restricts and confounds the experimental design and subsequent

interpretation of data. Hence, to answer the main objective of this study, this

investigation utilised modified human AECs, NuLi-1 for assessing the associations

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between HRV infection with changes in TJ gene, protein expression as well as

alterations in transepithelial permeability.

Initial validation results demonstrated a typical viral titre and time dependant effect on

NuLi-1 viability following HRV-1B infection. Similarly, elevated cellular apoptosis as

well as increased viral replication was also observed when infected with increasing viral

titres of HRV-1B. Collectively, these observations parallel those observed by others in

pAECs (Stevens 2009; Sutanto et al. 2011), where they similarly reported reduced

cellular viability, increased levels of apoptosis as well as increased viral replication

following infection with HRV-1B, thus validating the utilisation of NuLi-1 cells in this

investigation.

Tight junction complexes which are the most apical of the junctional complex family

not only serve to provide intercellular adhesion but also form a continuous permeability

barrier which regulates the trafficking of molecules across the epithelial layer. A myriad

of junctional complex interact to provide structural support for the regulation and

integrity of airway epithelium. Perturbations in these integral TJ complexes could

possibly result in increased epithelial permeability, facilitate trafficking of aeroallergens

or pathogens as well as increased exposure of basolateral receptors to the inhaled air.

Through a focussed qPCR array, this study was able to demonstrate a down-regulation

of 58 key genes with an up-regulation of 26 key genes encoding for epithelial barrier

junction proteins following infection with HRV-1B. In particular, claudin-8 mRNA

(CLDN8), which is responsible for a TJ seal, was found to be significantly down-

regulated following infection, while the pore-forming claudin-2 mRNA (CLDN2), was

found to be up-regulated. However, past evidence have demonstrated a wide disparity in

claudin expression among different tissue types (Morita et al. 1999) and in a study

conducted by Coyne and colleagues, they provide further evidence of respiratory airway

specific expression of claudins (Coyne et al. 2003). Based on these evidence, the

present study focussed on the key gene encoding for claudin-1, occludin and ZO-1 and

from the focussed qPCR array analysis, the data obtained mirror those reported by Yeo

and colleague (Yeo and Jang 2010), where they demonstrated significant decreases in

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mRNA levels of claudin-1, occludin and ZO-1, hence, indicating the effects of HRV-1B

on modulating TJ gene expression.

Furthermore, corroborating the gene expression findings, data obtained from this study

similarly demonstrated significant disassembly of membrane claudin-1, occludin and

ZO-1 proteins following infection with HRV-1B. Furthermore, disassembly of these TJ

protein complexes was shown to result in increased transepithelial permeability of

macromolecules. Collectively, these observations, in addition to mirroring those

reported by Sajjan and colleagues (2008), where they also demonstrated the effects of

HRV-1B infection on the dissociation of membrane ZO-1 proteins, also extends those

observations with the investigation of expression of membrane claudin-1 and occludin

proteins. Observations from this study suggest that increase epithelial permeability

could possibly result in increased exposure of basolateral receptors thereby facilitating

increased viral entry and eventual dissemination throughout the respiratory system.

Moreover, this increased transepithelial permeability would also result in co-migration

of bacteria into the sub-epithelial space, an observation reported by Sajjan and co-

workers (2008). Elevated entry of pathogens could potentiate the inflammatory

response, which could exacerbate existing conditions especially in airway diseases such

as cystic fibrosis and asthma.

4.5 Conclusion

This study demonstrated the direct associations between alterations in a number of TJ

genes, protein expression and the impaired functionality of the epithelium in regulating

passage of macromolecules following HRV infection. Data demonstrated that HRV

infection was capable of modulating the expression of TJ genes encoding for claudin-1,

occludin and ZO-1 as well as causing a reduction of membrane protein expression. The

diminished membrane protein expression resulted in increased transepithelial

permeability of macromolecules and when interpreted collectively, strongly suggests

the capacity of HRV in disrupting epithelial integrity and attenuating barrier function.

However, this study utilised a non-asthmatic modified primary human AEC type, thus,

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to corroborate these findings in unmodified primary human AECs, further studies

utilising human-derived pAECs are required to determine the association of HRV

infection on modulating TJ gene, protein expression and barrier functionality.

Moreover, this study only investigated the expression and functionality of tight

junctional complexes within a non-asthmatic setting. Given that HRV infection is the

most common trigger of acute asthma in children, further studies to determine whether

there is an intrinsic vulnerability of the epithelium in asthma following HRV infection is

required.

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CHAPTER 5: Epithelial barrier integrity and function in paediatric

asthma

5.1 Introduction

The respiratory airway epithelium is the primary interface with the external

environment and under normal circumstances forms a physical barrier complemented

with the mucociliary escalator clearance to form the first line of defence. In addition, it

also represents a dynamic system of innate host defence mechanisms (Diamond et al.

2000; Holgate et al. 2000). During normal function, the epithelium forms a highly

regulated and impermeable barrier through the formation of tight junctions (TJ) at the

apical end of the ciliated columnar cells. In conjunction with TJ, adherens junctions

(AJ), hemidesmosomes and desmosomes form a continuous junctional belt (Farquhar

and Palade 1963) which interconnects with neighbouring cells to enable inter-cell

communication as well as selectively regulating intercellular trafficking of ions, solutes

and cells through the paracellular space (Roche et al. 1993).

Impairment of the epithelial barrier function and dysregulation of the junctional

complex proteins can often result in the development or exacerbation of numerous

diseases. For example, filaggrin is a pivotal protein involved in epidermal

differentiation and the maintenance of an intact skin barrier function. However, a

functional mutation in the gene which encodes for the protein can result in increased

predisposition towards atopic dermatitis (Palmer et al. 2006). Furthermore, a recent

study by De Benedetto and colleagues (2011) has implicated claudin-1 as a novel

susceptibility gene for atopic dermatitis, where it might be involved in barrier

dysfunction and TH2 polarisation. With relevance to airway diseases, due to the

traditional difficulties in accessing tissue from appropriate human populations, the

majority of investigations have focussed on TJ disruptions in response to exogenous

stimuli through immortalised cell lines or animal models. These have included; house

dust mite allergen Dermatophagoides pteronyssinus 1 (Der p1) exposure on

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immortalised AEC lines (Wan et al. 2000; Friedlander and Busse 2005; Johnston 2005)

as well as ovalbumin exposure on sensitised mice (Evans et al. 2002; Tillie-Leblond et

al. 2007). Although these studies provide critical insight for the response of TJ to

external stimuli, due to the experimental models used in these studies, little can be

extrapolated to establish TJ profile patterns and investigate how these differ between

healthy and disease states. Surprisingly, despite advancements in the ability to obtain

primary airway cellular tissue (Doherty et al. 2003; Looi et al. 2011), there remains

limited studies that have attempted to profile TJs in healthy and disease states (de Boer

et al. 2008; Xiao et al. 2011). In addition, due to the lack of appropriate controls for

atopic asthmatic cohorts in de Boer and colleagues’ study, the observations from the

study failed to account for the potential impact atopy might exert on TJ profile

expression. Furthermore, the studies by de Boer et al (2008) and Xiao et al (2011)

predominantly utilised adult-derived primary AECs. Hence, there remain limited studies

on basal tight junctional complex gene and protein expression within the paediatric

epithelium. This provides the rationale for assessing whether the abnormality of

dysregulated TJ gene and protein expression is intrinsic or extrinsic to the paediatric

asthmatic epithelium.

Although a previous seminal study has identified intrinsic differences between healthy

and asthmatic epithelium (Kicic et al. 2006), a paucity of data exists on whether these

intrinsic differences extend to the expression of epithelial junctional proteins. Thus, this

study tested the hypotheses that epithelial barrier integrity and function is defective in

children with asthma and that a defective barrier function in asthma is independent of

atopy. Utilising cell cultures obtained from paediatric cohorts, this study was able to

assess basal levels of multiple membrane TJ gene and protein expression in the presence

or absence of atopy. Moreover, basal levels of barrier function were also assessed to

determine basal epithelial permeability of solutes across the epithelial layer.

Collectively, this study attempts to characterise the basal tight junctional complex

profiles in non-asthmatic and asthmatic cohorts in the presence or absence of atopy and

establishes the premise upon which future studies involving injurious stimuli can be

peformed.

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detail in Chapter 2

5.2.1 Patient Demographics

As previously described (refer to 2.5.4), four cohorts were used in this study. For this

section of the investigation, samples were obtained from 56 healthy non-atopic (HNA;

23 female, 33 male), 39 healthy atopic (HA; 14 female, 25 male), 13 non-atopic

asthmatic (NAA; 4 female, 9 male) and 25 atopic asthmatic (AA; 10 female, 15 male)

children who were not currently receiving any corticosteroid therapy. Patient

demographics are summarised in Table 5.1.

5.2.2 Cell culture

Maintenance and subsequent sub-culture of paediatric derived pAECs used in this

investigation has been described in detail in Chapter 2 (refer to 2.4.2.2.6 and 2.5.4).

5.2.3 In Cell™ Western

The optimised In Cell™ Western assay, as described in detail within Chapter 3 was

utilised for the determination of basal TJ membrane protein expression.

5.2.4 Immunocytochemistry

Immunocytochemistry of ex vivo pAECs prepared on cytospins has been described in

detail in Chapter 2 (refer to 2.5.6, 2.5.8).

Table 5.1 Demographic of patient cohort categorised according to atopy

HNA – Healthy non-atopic; HA – Healthy atopic; NAA – Non-atopic asthmatic;

AA – Atopic asthmatic; F – Female; M - Male

Phenotype Gender Average Age (yr)

Age Range (yr) Number Total (n)

HNA F 7.2 1.9 – 18.4 23 56

M 5.3 1.4 – 10.9 33

HA F 8.4 2.5 – 15.5 14 39

M 6.9 2.2 – 16.4 25

NAA F 5.7 4.3 – 6.9 4 13

M 5.2 2.2 – 11 9

AA F 8.1 2.9 – 14.7 10 25

M 7.4 3.9 – 13.5 15

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5.2.5 Real-Time Quantitative Polymerase Chain Reaction (RT-qPCR)

Extraction and quantification of RNA, as well as the methodology for RT-qPCR has

been described in detail in Chapter 2 (refer to 2.5.13).


The optimised transepithelial permeability assay, as described in detail within Chapter 3

was utilised to elucidate functionality of epithelial TJ membrane expression at basal

level.


Before statistical evaluation, all results were tested for population normality and

homogeneity of variance, and where applicable, a Student t test was performed.

Experiments were performed in at least duplicates using a minimum of three patients of

each cohort per experiment. Statistical analyses were performed using a Mann-Whitney

non-parametric test and values presented are mean ± SD. P values less than 0.05 were

considered to be significant.

5.3 Results

5.3.1 Basal tight junction gene expression

Knowing that basal membrane TJ proteins in cultured human AEC lines are expressed

in different levels, basal gene expression of three TJs, claudin-1, occludin, and ZO-1 in

non-asthmatic and asthmatic pAECs were assessed immediately ex vivo following non-

bronchoscope brushing of the airways.

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5.3.1.1 Comparison between pAECs of non-asthmatic and asthmatic cohorts

Data generated showed that when mRNA expression was compared between both

cohorts, the asthmatic cohorts demonstrated significantly higher levels of basal TJ

mRNA expression of claudin-1 (1.4-fold) and occludin (2.6-fold) (Figure 5.1 A – B

respectively; p<0.05). However, there was no statistically significant difference in ZO-1

mRNA expression between non-asthmatic and asthmatic cohorts (Figure 5.1 C).

Furthermore, when comparing between the three TJ, mRNA expression of claudin-1

was the most highly expressed in both non-asthmatic (7.86AU ± 0.95) and asthmatic

(11.28AU ± 1.03) cohorts, followed by occludin (non-asthmatic, 2.26AU ± 0.18 ;

asthmatic, 6.04AU ± 0.34) and with mRNA expression of ZO-1 being the least

expressed (non-asthmatic, 1.07AU ± 0.12 ; asthmatic, 1.16AU ± 0.1). (Figure 5.1 A –

C). The difference in mRNA expression of each TJ within the cohorts was observed to

be significant (Figure 5.1 A – C; Appendix E 1 & 2; p<0.05).

5.3.1.2 Comparison between pAECs of non-asthmatic and asthmatic cohorts based on

atopic status

When the non-asthmatic and asthmatic cohorts were further classified according to

atopy, data generated demonstrated significantly lower levels of ex vivo claudin-1

mRNA expression in the pAECHA cohorts (4.2AU ± 0.5) compared to pAECHNA

(10.9AU ± 1.1) (Figure 5.2 A; p<0.05). Interestingly, no significant difference in

claudin-1 mRNA expression was observed between pAECHNA, pAECNAA and pAECAA

cohorts. However, when pAECHA was compared to pAECNAA and pAECAA,

significantly higher levels of claudin-1 gene expression was observed in both pAECNAA

(11.1AU ± 1.5) and pAECAA (14.5AU ± 3.3) cohorts (Figure 5.2 A; p<0.05). Claudin-1

mRNA expression was not observed to be significantly different between pAECHA and

pAECAA.

In contrast, ex vivo occludin mRNA expression was observed to be significantly higher

in pAECHA (2.6AU ± 0.2), pAECNAA (5.9AU ± 0.6) and pAECAA (6.2AU ± 0.3) when

compared to pAECHNA (1.8AU ± 0.2). Similarly, occludin mRNA expression was also

Figure 5.1 Ex vivo mRNA expression of TJs from pAECs of non-asthmatic and

asthmatic cohorts: mRNA expression of claudin-1 (green), occludin (blue) and ZO-1

(red) in pAECs of non-asthmatic and asthmatic cohorts was quantified by RT-qPCR as

described (refer to 5.2.4). (A) mRNA expression of claudin-1 was significantly higher

in the asthmatic cohort compared to the non-asthmatic counterpart. (B) mRNA

expression of occludin was observed to be significantly higher in the asthmatic cohort.

(C) No significant differences were observed between the two cohorts for expression of

ZO-1. (A – C) mRNA expression of claudin-1 was most abundantly expressed

compared to occludin and ZO-1. *Statistical significance relative to non-asthmatic

cohort (p < 0.05). Statistical significance between tight junctions (Appendix E; p<0.05).

Gen

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Non-asthmatic Asthmatic0

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A

B

C

Figure 5.2 Ex vivo mRNA expression of TJs in pAECS of non-asthmatic and

asthmatic cohorts with each cohort further categorised based on atopy: mRNA

expression of claudin-1 (green), occludin (blue) and ZO-1 (red) in each phenotypic

cohort was quantified by RT-qPCR as described (refer to 5.2.4). (A) mRNA expression

of claudin-1 was significantly lower in pAECHA compared to pAECHNA. There was no

significant difference between pAECHNA, pAECNAA and pAECAA. However, both

pAECNAA and pAECAA demonstrated significantly higher expression levels when

compared to pAECHA while no significant difference was observed between pAECNAA

and pAECAA. (B) In contrast, mRNA expression of occludin was significantly higher in

pAECHA, pAECNAA and pAECAA compared to pAECHNA. Moreover, pAECNAA and

pAECAA demonstrated significantly higher mRNA expression of occludin compared to

pAECHA. There was no significant difference in mRNA expression between pAECNAA

and pAECAA. (C) mRNA expression of ZO-1 was observed to be higher in pAECHA,

pAECNAA and pAECAA compared to pAECHNA. When compared to pAECHA, lower ZO-

1 expression was observed in pAECNAA while pAECAA showed higher expression.

Similarly, mRNA expression of ZO-1 was also higher in pAECAA compared to

pAECNAA. However, statistical analysis did not demonstrate any significant differences

in mRNA expression of ZO-1 between the phenotypic cohorts. (A – C) mRNA

expression of claudin-1 was significantly most abundantly expressed in all phenotypic

cohorts compared to occludin and ZO-1. *Statistical significance (p<0.05). Statistical

significance between tight junctions (Appendix F; p<0.05).

Gen

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HNA HA NAA AA0

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observed to be significantly higher in pAECNAA and pAECAA in comparison to pAECHA

(Figure 5.2 B; p<0.05). No significant statistical difference was shown between

occludin mRNA expression in pAECNAA and pAECAA cohorts.

When ZO-1 mRNA expression was assessed in all four cohorts, higher levels of ZO-1

mRNA expression were observed in pAECHA (1.3AU ± 0.25), pAECNAA (1.1AU ±

0.13) and pAECAA (1.5AU ± 0.32) cohorts compared to pAECHNA (1AU ± 0.19).

However, the data obtained did not demonstrate any statistical significance between all

phenotypic cohorts (Figure 5.2 C).

When assessing the level of mRNA expression of each TJ within the individual

phenotypic cohort, mRNA expression of claudin-1 was observed to be the highest,

followed by occludin and ZO-1. The same expression profile was observed in all 4

phenotypic cohorts and was of significance (Figure 5.2 A – C; Appendix F 1 – 4;

p<0.05).

5.3.2 Basal tight junction protein expression

Having shown differences in ex vivo TJ gene expression, ex vivo protein levels of the

same three TJ were assessed to determine basal expression levels. Membrane claudin-1,

occludin and ZO-1 TJ protein expression were determined via immunocytochemistry

staining of cells obtained ex vivo.


Immunocytochemical staining of pAECs from the non-asthmatic cohort demonstrated

stronger intensities of membrane claudin-1, occludin and ZO-1 TJ protein in ex vivo

cytospin samples in comparison with pAECs from the asthmatic cohort (Figure 5.3).

When comparing between the TJs within each cohort, strongest intensity of membrane

claudin-1 was observed followed by occludin with membrane ZO-1 showing the lowest

intensity in pAECs of the non-asthmatic cohort (Figure 5.3 – Non-asthmatic).

Figure 5.3 Ex vivo membrane protein expression of TJs from pAECs of non-asthmatic and asthmatic cohorts: Cytospins were prepared from

cells obtained from non-asthmatic and asthmatic cohorts as previously described (refer to 2.5.6). Briefly, slides were incubated with primary antibodies

to claudin-1 (CLDN-1), occludin (OCLN) and zonula occluden-1 (ZO-1) for 24 h at 4°C followed by fluorescently conjugated secondary antibodies

(FITC in green) for a similar period. The slides were counterstained with DAPI, which illuminates cellular nuclear material (blue).

Immunocytochemical staining of claudin-1, occludin and ZO-1 respectively in a representative sample of pAECs from non-asthmatic cohort

demonstrates strong membrane protein expression of each tight junction protein. In contrast, immunocytochemical staining of claudin-1, occludin and

ZO-1 respectively within the asthmatic cohort demonstrates a marked decrease in membrane protein expression of each tight junction. Images are

representative of n = 3 (Total magnification 100x).

Non-asthmatic Asthmatic

CLDN-1

OCLN

ZO-1

CLDN-1

OCLN

ZO-1

DAPI

DAPI

DAPI

DAPI

DAPI

DAPI

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Interestingly, in pAECs of the asthmatic cohort, strongest intensity was observed for

membrane occludin, while claudin-1 and ZO-1 showed similar diminished intensities

(Figure 5.3 – Asthmatic).


atopic status

When pAECs from the non-asthmatic and asthmatic cohorts were further classified

according to atopy, immunocytochemical staining of cytospins demonstrated strongest

intensity for membrane claudin-1 within the pAECHNA cohorts, followed by similar

intermediate intensities in both pAECHA and pAECAA cohorts, while markedly

diminished intensity was observed for pAECNAA cohorts (Figure 5.4 – CLDN-1). A

similar profile of intensity staining was observed for membrane occludin (Figure 5.4 –

OCLN). Comparable strong intensity was also observed for membrane ZO-1 in

pAECHNA cohorts, however, pAECHA cohort showed a marked decrease in intensity

followed by pAECAA cohort with pAECNAA cohort showing minimal intensity for

membrane ZO-1 TJ protein (Figure 5.4 – ZO-1).

When assessing the level of intensities of the TJ protein within each phenotypic cohort,

the results demonstrated strongest intensity of membrane claudin-1, followed by

occludin and subsequently ZO-1 within the pAECHNA cohort (Figure 5.4 – HNA).

However, in the pAECHA cohort, similar levels of intensity were observed for both

membrane occludin and ZO-1 with the lowest intensity observed in membrane claudin-

1 (Figure 5.4 – HA). Interestingly, strongest intensity was observed for membrane

occludin followed by membrane claudin-1 with membrane ZO-1 showing markedly

decreased intensity in the pAECAA cohort (Figure 5.4 – AA). However, within the

pAECNAA cohorts, strongest intensity was observed for membrane claudin-1 while both

membrane occludin and ZO-1 demonstrated similar markedly diminished levels of

intensity (Figure 5.4 – NAA).

Figure 5.4 Ex vivo membrane protein expression of TJs of pAECs from non-

asthmatic and asthmatic cohorts with each cohort further categorised based on

atopy: Cytospins were prepared from cells obtained from each phenotypic cohorts as

previously described (refer to 2.5.6). Briefly, slides were incubated with primary

antibodies to claudin-1 (CLDN-1), occludin (OCLN) and zonula occluden-1 (ZO-1) for

24 h at 4°C followed by fluorescently conjugated secondary antibodies (FITC in green)

for a similar period (refer to 5.2.4). Immunocytochemical staining of claudin-1,

occludin and ZO-1 respectively in a representative sample of pAECHNA demonstrates

the strongest membrane protein expression of each tight junction protein while

intermediate levels of membrane claudin-1, occludin and ZO-1 expression of each tight

junction protein were observed in representative samples of pAECHA and pAECAA

respectively. Immunocytochemical staining of membrane claudin-1, occludin and ZO-1

respectively in a representative sample of pAECNAA demonstrates the lowest expression

of each tight junction protein among the four phenotypic cohorts. Images are

representative of n = 3 (Total magnification 100x).

I

HNA HNA HNA

HA HA HA

AA AA AA

NAA NAA NAA

CLDN-1 OCLN ZO-1

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5.3.3 In vitro tight junction protein expression

Differences in membrane TJ protein expression between pAECs of non-asthmatic and

asthmatic cohorts have been previously demonstrated via immunocytochemical

staining. However, to validate this observation as well as to semi-quantify expression

levels of the three membrane TJ proteins, in vitro monolayer cultures of non-asthmatic

and asthmatic pAECs, in conjunction with an In-Cell™ Western assay was utilised for

this assessment.


Data obtained demonstrated that membrane claudin-1 expression in pAECs of the non-

asthmatic cohort was significantly higher (0.09AU ± 0.04) in contrast to pAECs of the

asthmatic cohort (0.01AU ± 0.005) (Figure 5.5 A; p<0.05). Similarly, membrane

occludin expression was observed to be significantly higher (0.09AU ± 0.02) in pAECs

of the non-asthmatic cohort compared to their asthmatic counterpart (0.03AU ± 0.006)

(Figure 5.5 B; p<0.05). Significantly higher levels of membrane ZO-1 was observed in

pAECs from the non-asthmatic cohort (0.02AU ± 0.005) in comparison to the asthmatic

cohort (0.01AU ± 0.002) (Figure 5.5 C; p<0.05).

When assessing membrane TJ protein expression of pAECs within the non-asthmatic

cohort, results demonstrated similar levels of membrane claudin-1 (0.09AU ± 0.04) and

occludin (0.09AU ± 0.02) expression with membrane ZO-1 showing the lowest level of

expression (0.02AU ± 0.005). No significant difference in membrane claudin-1 and

occludin expression was observed in pAECs of the non-asthmatic cohort. However,

significance was observed for the difference in membrane TJ protein expression

between occludin and ZO-1 (Figure 5.5 B – C, Non-asthmatic; Appendix G1; p<0.05).

In contrast, pAECs from the asthmatic cohort demonstrated significantly lower levels of

membrane claudin-1 expression (0.01AU ± 0.005) when compared to occludin (0.03AU

± 0.006) (Figure 5.5 A – B, Asthmatic; Appendix G2; p<0.05). No significant

difference in membrane claudin-1 and ZO-1 expression was observed in pAECs of the

asthmatic cohort. However, membrane occludin expression (0.03AU ± 0.006) was

Figure 5.5 Basal membrane TJ protein expression in pAECs from non-asthmatic

and asthmatic cohorts: pAECs seeded on 96-well micro-titre plates and grown to

confluence were treated as previously mentioned (refer to 2.5.10) and membrane TJ

protein expression assessed via a previously optimised In-Cell™ Western assay as

described (refer to 3.2.4). (A) pAECs from asthmatic cohort expressed significantly

lower levels of membrane TJ claudin-1. (B) Membrane occludin was also significantly

lower in pAECs of asthmatic cohort compared to non-asthmatic. (C) Similarly, pAECs

from asthmatic cohort also showed lower levels of membrane ZO-1 expression when

compared to pAECs of non-asthmatic cohort. (A-C) Similar levels of membrane protein

expression were observed for claudin-1 and occludin in pAECs of non-asthmatic cohort

and was significantly elevated compared to membrane ZO-1. Interestingly, pAECs of

asthmatic cohort demonstrated significantly elevated level of basal membrane occludin

protein expression compared to claudin-1 and ZO-1. No significant difference in

membrane protein expression was observed between claudin-1 and ZO-1 in pAECs of

asthmatic cohort. Data were normalised to cell numbers and presented as mean ± SD.

*Statistical significance relative to non-asthmatic cohort (p<0.05). Statistical

significance between TJ protein in each cohort (Appendix G; p<0.05).

Non-asthmatic Asthmatic0.00

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observed to be significantly higher compared to ZO-1 expression (0.01AU ± 0.002) in

pAECs of the asthmatic cohort (Figure 5.5 B – C, Asthmatic; Appendix G2; p<0.05).


atopic status


atopy, data demonstrated lower membrane claudin-1 protein expression in the pAECHA

(0.014AU ± 0.002), pAECNAA (0.006AU ± 0.003) and pAECAA (0.011AU ± 0.003)

cohorts when compared to pAECHNA (0.033AU ± 0.005) cohort. Significance was

observed in both pAECHA and pAECAA compared to pAECHNA (Figure 5.6 A; p<0.05).

In addition, membrane claudin-1 expression was observed to be lower in both the

pAECNAA (2.3-fold) and the pAECAA (1.3-fold) cohorts in comparison with the pAECHA

cohort. Furthermore, a lower level of membrane claudin-1 expression was observed in

the pAECNAA (1.8-fold) cohort in contrast with the pAECAA cohort. However, due to the

limited availability of pAECNAA, statistical analysis could not be performed for claudin-

1 expression to determine the level of significance.

Similarly, occludin expression was observed to be significantly lower in the pAECHA

(0.053AU ± 0.009), pAECNAA (0.023AU ± 0.013) and pAECAA (0.04AU ± 0.007)

cohorts in comparison with pAECHNA (0.148AU ± 0.038) cohort (Figure 5.6 B; p<0.05).

Furthermore, membrane occludin expression was observed to be lower in both

pAECNAA (2.3-fold) and pAECAA (1.3-fold) cohorts in comparison with the pAECHA

cohort. In addition, a lower level of membrane occludin expression was observed in the

pAECNAA (1.7-fold) cohort in contrast with the pAECAA cohort. However, these

differences in membrane expression were not significant.

When ZO-1 expression was assessed, results generated were similar to claudin-1 and

occludin, with lower expression in the pAECHA (0.017AU ± 0.002), pAECNAA

(0.011AU ± 0.004) and pAECAA (0.008AU ± 0.002) cohorts in contrast to the pAECHNA

(0.04AU ± 0.01) cohort (Figure 5.6 C; p<0.05). Comparison of ZO-1 protein expression

Figure 5.6 Basal membrane TJ protein expression in pAECs from non-asthmatic

and asthmatic cohorts with each cohort further categorised based on atopy: pAECs

from each phenotypic cohort, seeded on 96-well micro-titre plates and grown to

confluence were treated as previously mentioned (refer to 2.5.10) and membrane TJ

protein expression assessed via a In-Cell™ Western assay as described (refer to 3.2.4).

Due to limited numbers of pAECNAA, the cohort was excluded from statistical analysis

for claudin-1 and ZO-1 membrane expression. (A) As observed, membrane claudin-1

expression was highest within the pAECHNA cohorts and significant differences were

shown when compared to pAECHA and pAECAA cohorts. However, there was no

significant difference in membrane protein expression between pAECHA and pAECAA.

(B) Similarly, membrane occludin expression was highest within the pAECHNA cohort

and was significantly elevated when compared to pAECHA, pAECNAA and pAECAA

cohorts. No significant differences were observed between pAECHA, pAECNAA and

pAECAA cohorts. (C) Membrane claudin-1 expression was similarly observed to be

elevated in pAECHNA compared to pAECHA, pAECNAA and pAECAA. However this was

only significant for pAECHA and pAECAA. Interestingly, membrane ZO-1 expression

was significantly higher in pAECHA compared to pAECAA cohorts. (A – C) Membrane

occludin was observed to be most expressed in all phenotypic cohorts compared to

claudin-1 and ZO-1 and this was shown to be significant for pAECHNA, pAECHA and

pAECAA cohorts. No significant difference in membrane expression was observed for

all phenotypic cohorts between membrane claudin-1 and ZO-1 proteins. Data were

normalised to cell numbers and presented as mean ± SD. *Statistical significance

relative to pAECHNA (p<0.05). Statistical significance between TJ protein in each

phenotypic cohort (Appendix H; p<0.05).

Mem

bran

e pr

otei

n ex

pres

sion

(Arb

itary

uni

ts)

HNA HA NAA AA0.00

0.05

0.10

0.15

0.20

n=8 n=5n=2n=10

**

Mem

bran

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otei

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pres

sion

(Arb

itary

uni

ts)

HNA HA NAA AA0.00

0.05

0.10

0.15

0.20

n=10 n=7n=4n=10

**

*

Mem

bran

e pr

otei

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pres

sion

(Arb

itary

uni

ts)

HNA HA NAA AA0.00

0.05

0.10

0.15

0.20

n=10 n=5n=2n=10

***

A

B

C

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between the pAECHA, pAECNAA and pAECAA cohorts demonstrated lower expression in

both pAECNAA (1.5-fold) and pAECAA (2.1-fold) cohorts and this was observed to be

significant for the pAECAA cohort. Interestingly, in contrast to membrane claudin-1 and

occludin expression, membrane ZO-1 expression was observed to be lower in the

pAECAA (1.4-fold) cohort compared to the pAECNAA. However, due to the limited

availability of pAECNAA, statistical analysis could not be performed for ZO-1

expression to determine the level of significance.

5.3.4 In vitro transepithelial permeability

Knowing that epithelial paracellular permeability is an indicator of a functional barrier,

transepithelial permeability towards two different sizes of inert macromolecules was

also performed at baseline. The flux of the fluorescently labelled inert macromolecule

through the epithelial monolayer allowed for barrier functionality to be determined.


Results generated demonstrates increased levels of basal transepithelial permeability to

both FITC-dextran of both 4 (344.9 x 10-4 cm/sec ± 37.3) and 20 kDa (130.2 x 10-4

cm/sec ± 41.9) for the asthmatic cohort compared to the non-asthmatic cohort (302.9 x

10-4 cm/sec ± 20.5 and 96.5 x 10-4 cm/sec ± 26.8 respectively) (Figure 5.7 A). However,

no statistical significance between the two cohorts was observed. When assessing

epithelial permeability to the different sized inert macromolecule within each cohort, a

significantly higher level of transepithelial permeability for FITC-dextran of 4 kDa was

observed in the non-asthmatic (302.9 x 10-4 cm/sec ± 20.5) and asthmatic (344.9 x 10-4

cm/sec ± 37.3) cohort in comparison with FITC-dextran of 20 kDa (96.5 x 10-4 cm/sec ±

26.8 and 130.2 x 10-4 cm/sec ± 41.9 respectively) (Figure 5.7 A; p<0.05).

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atopic status


atopy, interestingly, the results generated demonstrated lower permeability to FITC-

dextran of 4 kDa in the pAECHA (1.3-fold, 272.5 x 10-4 cm/sec ± 14.6), pAECNAA (79.4

x 10-4 cm/sec ± 0.9) and pAECAA (344.9 x 10-4 cm/sec ± 37.3) cohorts compared to the

pAECHNA (370.3 x 10-4 cm/sec ± 13.2) cohort and this was observed to be significant

for pAECHA (Figure 5.7 B – HA; p<0.05). Transepithelial permeability of pAECNAA to

FITC-dextran 4 kDa was observed to be lower (3.4-fold) than pAECHA. However,

pAECAA demonstrated slightly higher permeability (1.3-fold) when compared to

pAECHA. Moreover, when comparing permeability between pAECNAA and pAECAA

cohort, lower epithelial permeability was observed within the pAECNAA cohort (4.3-

fold). Despite these observations, due to the limited availability of pAECNAA, statistical

analysis could not be performed to determine the level of significance.

Transepithelial permeability to FITC-dextran 20 kDa was observed to be similarly

lower in pAECHA (179.7 x 10-4 cm/sec ± 53.9), pAECNAA (24.4 x 10-4 cm/sec ± 6.2) and

pAECAA (183.6 x 10-4 cm/sec ± 44.1) cohorts when compared to the pAECHNA cohort

(201.7 x 10-4 cm/sec ± 34.1) (Figure 5.7 B). However, this was not observed to be

significant. Transepithelial permeability of pAECNAA to FITC-dextran 20 kDa was

observed to be lower (7.3-fold) than pAECHA. However, pAECAA demonstrated similar

levels of permeability when compared to pAECHA. Moreover, when comparing

permeability between pAECNAA and pAECAA cohorts, lower epithelial permeability was

observed for the pAECNAA cohort (7.5-fold). However, statistical analysis did not

demonstrate any significant differences in transepithelial permeability to FITC-dextran

20 kDa between the cohorts. Due to limited numbers of pAECNAA, this cohort was

precluded from analysis.

Figure 5.7 Basal transepithelial permeability in pAECs from non-asthmatic and

asthmatic cohorts and with each cohort further categorised based on atopy: pAECs

from non-asthmatic and asthmatic cohorts, seeded onto Corning transwell inserts and

grown to confluence were treated as previously mentioned (refer to 2.5.11). A

transepithelial permeability assay as described was performed to determine basal

epithelial permeability to FITC-dextran 4 kDa and 20 kDa (refer to 3.3.4). (A) pAECs

from asthmatic cohort demonstrated higher basal transepithelial permeability to both

FITC-dextran 4 kDa and 20 kDa but no significance was observed when compared to

the non-asthmatic cohort. Permeability to FITC-dextran 4 kDa was observed to be

significantly greater in both non-asthmatic and asthmatic cohorts when compared to

FITC-dextran 20 kDa. (B) pAECs further categorised based on atopy demonstrate lower

transepithelial permeability to FITC-dextran 4 kDa in pAECHA, pAECNAA and pAECAA

cohorts and this was shown to be significant for pAECHA and pAECAA. Due to the low

sample size of pAECNAA, this cohort was excluded from statistical analysis. Lower

transepithelial permeability to FITC-dextran 20 kDa was observed in pAECHA,

pAECNAA and pAECAA cohorts compared to pAECHNA however, this was not shown to

be significant. Permeability to FITC-dextran 4 kDa was observed to be greater in all

phenotypic cohorts compared to 20 kDa, however, significance was only observed for

pAECHNA, pAECHA and pAECAA cohorts. *Statistical significance relative to pAECHNA

(p<0.05). # Statistical significance between FITC-dextran 4 and 20 kDa (p<0.05).


100

200

300

400

5004kDa20kDa

Papp

coe

ffici

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cm/s

ec) x

10-4

n=10 n=7

#

#

HNA HA NAA AA0

100

200

300

400

5004kDa20kDa

Papp

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cm/s

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n=5 n=5 n=2 n=5

#

*#

#

A

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5.4 Discussion

Changes in transepithelial permeability are often a feature of airway inflammation,

however, the exact molecular mechanisms involved in the regulation of epithelial

permeability remain poorly understood. A pivotal component of the conductive airway

epithelium is the apico-lateral junctional complexes consisting of tight and adherens

junctions, which contributes significantly to the barrier function as well as providing a

link to the cellular cytoskeleton via various adaptor proteins. Although the presence of

TJs between epithelial cells has long been recognised, there still remains a lack of

understanding of the regulation of junctional complex assembly and disassembly and

the resulting changes in epithelial permeability in respiratory diseases such as asthma.

Previous studies have shown the effects of TJ gene and protein expression following

various insults (Sajjan et al. 2008; Comstock et al. 2011; Xiao et al. 2011) in both non-

asthmatic and asthmatic epithelium, however, these studies were performed in either

established epithelial cell lines or adult-derived primary AECs. Currently, there exists a

paucity of data on the intrinsic gene and protein expression of epithelial tight junctional

complexes particularly within the paediatric asthmatic population. This investigation

attempted to provide new insights into the presently limited understanding on baseline

expression of epithelial membrane TJ complexes in paediatric asthma.

Results from this parallel ex vivo and in vitro study demonstrated significant increases

in ex vivo mRNA expression of claudin-1 and occludin in pAECs of the asthmatic

cohorts in contrast to the non-asthmatic counterparts, while, ZO-1 gene expression was

not found to be significantly different. These findings extend those of Xiao and

colleagues (2011), who have previously reported an increase in mRNA expression of

occludin but not ZO-1 in adult individuals with asthma. Interestingly, when these

cohorts were sub-categorised based upon atopy, mRNA expression of claudin-1 was

observed to be significantly higher in the atopic asthmatic cohort in contrast to the

healthy atopic phenotype. The mRNA expression of occludin was also significantly

higher in the asthmatic cohort, irrespective of atopy, when compared to the non-

asthmatic cohort. This suggests that the presence of asthma might be characterised by an

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altered expression of specific TJ complex genes. Furthermore, when comparing mRNA

expression between non-atopic and atopic cohorts, claudin-1 was revealed to be

significantly lower in the atopic compared to the non-atopic phenotype within the non-

asthmatic cohort. This is consistent with an earlier work in individuals with atopic

dermatitis conducted by De Benedetto and colleagues (2011), where they showed

reduced mRNA expression levels of claudin-1 and claudin-23 compared to healthy

subjects.

Elevated mRNA expression of occludin in non-asthmatic atopic patients from this study

contrasts that of De Benedetto and colleagues, highlighting that the possible difference

in expression could be attributed to the different epithelial sites assessed, the epidermal

layer of the skin in De Benedetto’s study in contrast with the respiratory bronchial

epithelium in the present study. In addition, differences in age range as De Benedetto’s

study utilised an adult cohort in contrast to this study, which utilised a paediatric cohort.

The dichotomy in occludin mRNA expression could also be partially explained by the

complex interactions between the myriad of junctional proteins. The current lack of data

provides the rationale for the assessment of basal TJ mRNA expression in the AECs of

non-asthmatic or asthmatic paediatric cohorts with or without atopy.

Within the asthmatic cohort, mRNA expression of claudin-1, occludin and ZO-1 was

higher in the atopic asthmatic phenotype in comparison to non-atopic asthmatic. This

suggests that although atopy might be implicated in the significant augmentation of TJ

protein expression, as demonstrated by De Benedetto and colleagues in non-asthmatic

individuals, the precise role of atopy in altering TJ complex expression, whether causal,

co-contributing or a consequence of asthma especially within asthmatic cohorts

warrants further investigation.

Interestingly, when membrane TJ protein expression was assessed, ex vivo expression of

claudin-1, occludin and ZO-1 proteins were markedly lower within the asthmatic

compared to the non-asthmatic cohort. This observation was further validated with in

vitro analysis of membrane TJ proteins in submerged paediatric derived pAECs cultures

from non-asthmatic and asthmatic cohorts, which is consistent to those observed by

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Xiao and colleagues (2011) However, this study is unique with regards to several

factors. The children with asthma are all mild asthmatics who have not taken either

inhaled β2-agonists or corticosteroids in the past six months prior to sampling. This

enables the investigation of intrinsic TJ protein expression to be as similar to unaltered

conditions as possible. In addition, classification based on atopic status within non-

asthmatic and asthmatic cohorts, provides further insights into the potential differences

in intrinsic TJ protein expression. However, one drawback to this approach is the

limited availability of children who are non-atopic asthmatic. Nonetheless, despite low

numbers of non-atopic asthmatic subjects, data from this study demonstrated that

membrane TJ protein expression of claudin-1, occludin and ZO-1 progressively

diminishes with the involvement of either atopy, asthma or a combination of both, an

observation validated with in vitro analysis of membrane TJ proteins in submerged

paediatric derived pAECs cultures from each phenotypic cohort. When comparing

between non-atopic and atopic phenotypes within the non-asthmatic cohort, lower

membrane claudin-1 expression in the atopic phenotype corroborates the observation of

a decreased mRNA expression of claudin-1, an observation in line with those reported

by De Benedetto and colleagues showing reduced expression of claudin-1 protein in

patients with atopic dermatitis compared to non-atopic individuals.

A decrease in membrane occludin expression was observed in the non-asthmatic atopic

phenotype which contrasts observations of an elevated mRNA expression of occludin,

suggesting that the effect on membrane occludin expression involves a post-

transcriptional mechanism, an observation which mirrors those of Xiao and co-workers

(2011). Collectively, this disparity in mRNA and protein expression could suggest

probable post-transcriptional regulation by transcription factors such as Snail (Ohkubo

and Ozawa 2004) or other factors such as cytokines IL-4, IL-13 and IFN-γ (Ahdieh et

al. 2001; Bruewer et al. 2005), resulting in either increased TJ protein internalisation

and redistribution or disruption in their interaction with the scaffolding proteins. In

addition, host microRNAs have also been identified to play a regulatory role. For

example, miR-122a, which when overexpressed due to increased TNF-α levels, has

been demonstrated to result in degradation of occludin, thereby contributing to

increased intestinal epithelial permeability (Ying et al. 1991; Bradding et al. 1994; Ye

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et al. 2011). In another recent study, Yang and colleagues (2014) showed that

repression of claudin-1 expression post-transcriptionally by miR-155 resulted in

decreased epithelial barrier functionality (Yang et al. 2014). Although microRNAs

functions have been studied in human disease, their roles in the control of signalling

pathways in epithelial cells, their impact on the development and phenotypic stability of

immune cells as well as their regulation of inflammation in allergic diseases such as

asthma have only recently been uncovered (Makeyev and Maniatis 2008; Djuranovic et

al. 2011). Moreover, due to the numerous biological functions microRNAs regulate,

modulating their expression could potentially provide rationale for new therapeutic

regimen. MicroRNAs have already been shown to limit allergic airway inflammation

mouse models (Mattes et al. 2009; Collison et al. 2011; Collison et al. 2011; Qin et al.

2012), inhibition of hepatitis C virus (HCV) replication in primates (Lanford et al.

2010) or suppression of tumour metastasis in mouse models (Liu et al. 2011; Yang et al.

2012). Furthermore, the dichotomy in TJ mRNA and protein expression could also be

explained by a probable compensatory effect via other epithelial junctional complexes.

This has been demonstrated in occludin knock-out studies showing that occludin was

not required for the formation of morphologically intact TJs and strongly suggested that

other junctional complexes were capable of compensating for the lack of occludin

expression (Saitou et al. 1998; Saitou et al. 2000). Strikingly, unlike membrane claudin-

1 and occludin protein expression, which were observed to be lower in the non-atopic

asthmatic phenotype, membrane ZO-1 expression was observed to be lower in the

atopic asthmatic phenotype, an observation which parallels that of de Boer and

colleagues (2008). However, due to the limited availability of non-atopic asthmatic

participants, the elevated expression of membrane claudin-1 and occludin protein as

well as the diminished expression of membrane ZO-1 protein expression in atopic

asthmatic cannot be fully attributed to atopy rather than asthma, a limitation which

needs to be addressed in future.

When these differences in basal membrane TJ protein expression were translated into a

functional context, data from this study demonstrated high permeability to

macromolecules in submerged monolayer pAECs cultures of both non-asthmatic and

asthmatic cohort. However, in both cohorts, permeability towards the 20 kDa dextran

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was lower, compared to 4 kDa, suggesting that for aeroallergens, environmental

particles or respiratory viruses to breach the epithelial barrier, these pathogens or their

fragments must have relatively small molecular weights. Interestingly, when the non-

asthmatic cohort was sub-categorised based on atopy, significantly diminished

permeability to 4 kDa dextran pAECHA compared to pAECHNA cohort was observed.

This contrasts with observations reported by De Benedetto and colleagues (2011),

where they reported increased permeability of FITC-conjugated albumin in the

epidermal cells of adult individuals with atopic dermatitis. Furthermore, De Benedetto

and colleagues also reported lower transepithelial electrical resistance measurements

within the same sample population, an observation consistent with the increase in

transepithelial permeability. The contrasting observations presented in this study

suggest potential differences in membrane tight junctional complex permeability

between different epithelium despite similar mRNA and protein expression profiles

reported. In addition, samples obtained from a paediatric population in this study in

contrast with an adult population indicate that a less environmentally exposed

epithelium may translate to a less immunologically active epithelium hence, reduced

impact on epithelial TJ dissociation thereby maintaining epithelial permeability.

When comparing pAECHNA and the pAECNAA cohorts, the pAECNAA cohorts

demonstrated less epithelial permeability to both sizes of the inert macromolecule.

Although pAECAA cohorts demonstrates increased levels of epithelial permeability

compared to the pAECNAA cohorts, indicating a possibility of asthma in altering

epithelial tight junctional complex mRNA, protein expression as well as function, the

present limited number of pAECNAA samples highlights the limitation of this study as

well as the need for attention when attempting to elucidate the effects of asthma on in

vitro epithelial TJ complex protein expression and function.

Despite current limitations of this investigation such as the use of submerged monolayer

culture for the assessment of epithelial permeability, preliminary observations have

provided the rationale as well as various interesting avenues for future studies. This

would involve the assessment of not only additional epithelial junctional complexes to

profile basal expression levels of both mRNA and protein, but also the extent of atopy

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as a contributing factor and its association with epithelial junctional complex

expression. This could provide further insights into understanding potential

compensatory responses by other junctional complexes to maintain a protective barrier

against further environmental insult.

5.5 Conclusion

This study was able to distinguish intrinsic differences in basal expression of epithelial

TJ mRNA and protein between non-asthmatic and asthmatic cohorts that might partially

explain the increased susceptibility to aeroallergen sensitisation and pathogenic

challenges in children with asthma. Furthermore, the results demonstrate differences

between non-atopic and atopic phenotypes in the non-asthmatic cohort, suggesting that

the presence of atopy might be a contributor to an increased predisposition towards

membrane TJ protein disassembly. However, due to limited availability of non-atopic

asthmatic subjects, the relationship between atopy and asthma, whether solely or co-

contributing to membrane TJ protein disassembly certainly requires further

investigation. Furthermore, when alterations in membrane TJ protein were translated

into a functional aspect, although non-significant, differences in transepithelial

permeability between non-asthmatic and asthmatic cohorts and subsequently, individual

phenotypes, were observed. However, in all phenotypes, transepithelial permeability

towards small-sized macromolecules was significantly higher, which would indicate the

increased susceptibility for various injurious stimuli such as aeroallergens, haptens or

pathogens to traverse the epithelium based on molecular size. Additional samples would

provide greater determination of the differences between individual phenotypes and

enable future investigations into TJ protein expression and barrier functionality in

response to injurious stimuli and addressing the limitations within the current

knowledge in epithelial tight junctional complex biology.

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CHAPTER 6: Effects of human rhinovirus on epithelial barrier

integrity and function in vitro and its role in paediatric asthma

6.1 Introduction

Respiratory viruses have been identified as causal agents of epithelial barrier

compromisation, altered innate immunity defences and modulation of cell membranes

(Jacoby et al. 1988; Kristjansson et al. 2005; Sajjan et al. 2008). Human rhinovirus in

particular has been identified as a primary trigger in the majority of asthma

exacerbations in children (Johnston et al. 1995; Friedlander and Busse 2005; Johnston

2005). Although HRV typically affects the upper respiratory airways, there are evidence

to suggest the association between lower airway infection and subsequent increased

bronchial responsiveness in patients with asthma (Fraenkel et al. 1995; Grunberg et al.

1997). In vitro studies have also shown that viral replication is elevated in the bronchial

epithelial cells of asthmatic patients due to reduced antiviral protein responses as

compared to bronchial epithelial cells from healthy participants (Wark et al. 2005;

Contoli et al. 2006).

Human rhinovirus infections have also been previously demonstrated to affect the

barrier integrity and function of healthy AECs. Yeo and Jang (2010) examined the

effects of HRV infection on nasal epithelial barrier function and observed that post

HRV infection, mRNA expression of TJ and adherens junction proteins were reduced

compared to the non-infected healthy control group. In the same study, protein

expression was similarly observed to be reduced in the HRV infected cells compared to

non-infected controls. Findings from this study indicate that HRV infection has the

propensity to decrease expression of junctional protein complexes in order to exert

potentially detrimental effects on the nasal epithelial barrier function. However, their

study utilised cells obtained from the nasal passages of non-asthmatic, non-atopic

individuals and thus, do not reflect the bronchial airway setting following HRV

infection, especially in asthma. A seminal study by Sajjan and colleagues (2008)

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demonstrated the capability of HRV infection to disrupt the barrier function of polarised

AECs obtained from tracheal trimmings of donor lungs during transplantation.

Observations from this study reported a loss of the ZO-1 TJ protein complexes

following HRV infection and an increased in the paracellular permeability of FITC-

inulin, suggesting the ability of HRV to disrupt epithelial barrier function in vitro.

Although evident that HRV infection has the proclivity to disrupt epithelial barrier

function, there currently exists a paucity of data in the understanding of the effects of

HRV infection on barrier integrity and function of a paediatric epithelium that is already

inherently dysregulated.

Along with the known association between HRV infection and disruption of the

epithelial barrier from previous in vitro studies (Sajjan et al. 2008; Yeo and Jang 2010;

Comstock et al. 2011), this study hypothesised that HRV infection within asthmatic

airways results in greater compromisation of epithelial barrier integrity and function

compared to healthy airways. Utilising cell cultures obtained from a paediatric cohort,

this study was able to assess membrane expression levels of multiple membrane TJ

proteins prior and following HRV infection through an In-Cell Western® assay.

Moreover, through a transepithelial permeability assay adapted for pAECs, this study

was also able to functionally assess transepithelial permeability of inert solutes of two

different molecular weights across the epithelial layer.



detail in Chapter 2



section of the investigation, samples were obtained from 16 healthy non-atopic (HNA; 8

female, 8 male), 17 healthy atopic (HA; 7 female, 10 male), 5 non-atopic asthmatic

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(NAA; 2 female, 3 male) and 6 atopic asthmatic (AA; 2 female, 4 male) children who

did not previously receive any corticosteroid therapy. Patient demographics specific for

this chapter are summarised in Table 6.1.

6.2.2 Cell culture




The source, propagation and viral titre determination of the HRV-1B utilised in this

investigation has been described in detail in Chapter 2 (refer to 2.5.7, 2.5.7.1 and

2.5.7.2).


Paediatric derived pAECs grown on culture plates or inserts, were infected with HRV-

1B and incubated for 24 h. Briefly, growth media was replaced with fresh basal media

and subsequently infected with HRV-1B at a 50% Tissue Culture Infectivity Dose

(TCID50) of 2.5 x 104 TCID50/ml and 10 or 20 x 104 TCID50/ml to mimic an in vivo

chronic and acute viral infection. After each incubation period, media was collected and

stored at -80°C and infected cells were utilised for various downstream assays.

6.2.5 In Cell™ Western

The optimised In Cell™ Western assay, as described in detail within Chapter 3 was

utilised for the determination of TJ membrane protein expression prior and following

HRV-1B infection.






HNA F 7.1 3.0 – 15.1 8 16

M 4.1 1.4 – 6.2 8

HA F 5.9 3.3 – 15.4 7 17

M 5.9 2.2 – 16.4 10

NAA F 6.9 6.9 – 7 2 5

M 3.9 2.2 – 6.1 3

AA F 8.1 2.9 – 14.7 2 6

M 6.6 5.3 – 8.1 4

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was utilised for the functional determination of epithelial TJ membrane integrity prior

and following HRV infection.


Data were tested for population normality and homogeneity of variance, and where

applicable, a Student t test was performed. Experiments were performed in at least

duplicates and using a minimum of three patients of each cohort per experiment.

Statistical analyses were performed using Mann-Whitney non-parametric test and

values presented are mean ± SD. P values less than 0.05 were considered to be

significant.

6.3 Results


expression after 24 and 48 h

Having previously demonstrated that the basal membrane TJ protein expression of

claudin-1, occludin and ZO-1 in paediatric derived pAECs differed between non-

asthmatic and asthmatic cohorts, this chapter focused on the effects of HRV-1B

infection on TJ expression. Hence, this investigation characterised the changes to

epithelial TJ proteins following HRV-1B infection in non-asthmatic and asthmatic

cohorts using an In-Cell™ Western assay to assess barrier integrity and a transepithelial

permeability assay for the assessment of barrier functionality. A low viral titre of 2.5 x

104 TCID50/ml HRV-1B and a high viral titre of 20 x 104 TCID50/ml HRV-1B was

utilised in this study to mimic a chronic and acute viral infection respectively.

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When assessing membrane claudin-1 expression between non-asthmatic and asthmatic

cohorts, results obtained in this section demonstrated significantly lower basal

membrane claudin-1 (11.08%) within the asthmatic cohort compared to the non-

asthmatic counterpart (100%) (Figure 6.1 A – 0 h; p<0.05), corroborating basal data

from the previous chapter. Following 24 h infection with a low viral titre of 2.5 x 104

TCID50/ml of HRV-1B, a significant decrease in membrane claudin-1 expression

(35.5% ± 0.002) was observed in comparison with non-infected controls in pAECs of

non-asthmatic cohort (Figure 6.1 A – Non-asthmatic, 24 h; p<0.05). However, at 48 h

post infection with the same viral titre, a significant increase back to baseline values in

membrane claudin-1 expression (105.8% ± 0.002) was shown within pAECs from the

non-asthmatic cohort (Figure 6.1 A – Non-asthmatic, 48 h; p<0.05). When infected

with a high viral titre of 20 x 104 TCID50/ml of HRV-1B for 24 h, a significant decrease

in membrane claudin-1 expression (25.3% ± 0.001) was observed in comparison with

non-infected controls (Figure 6.1 A – Non-asthmatic, 24 h; p<0.05). Interestingly, at 48

h post infection, a significant increase towards baseline values in membrane claudin-1

expression (82.2% ± 0.002) was similarly observed (Figure 6.1 A – Non-asthmatic, 48

h; p<0.05).

In addition, 24 h post infection with the higher viral titre resulted in a significant

decrease in membrane claudin-1 expression (25.3% ± 0.001) within pAECs of the non-

asthmatic cohort compared to the lower viral titre (35.5% ± 0.002) (Figure 6.1 A – Non-

asthmatic, 24 h; p<0.05). Furthermore, 48 h post infection, infection with the higher

viral titre continued to demonstrate significantly lower levels of membrane claudin-1

expression (82.2% ± 0.002) compared to expression levels post infection with the lower

viral titre (105.8% ± 0.002) (Figure 6.1 A – Non-asthmatic, 48 h; p<0.05).

When membrane claudin-1 expression of pAECs from the asthmatic cohort were

assessed following infection with viral titre of 2.5 x 104 TCID50/ml of HRV-1B, the data

demonstrated, at 24 h post infection, significant differences in membrane claudin-1

expression in pAECs of the asthmatic cohort (8.53% ± 0.0002) was observed in

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comparison to non-infected controls (Figure 6.1 A – Asthmatic, 24 h; p<0.05). In

addition, 48 h post infection, a further significant decrease in membrane claudin-1

expression was observed (3.63% ± 0.001) compared to non-infected controls (Figure 6.1

A – Asthmatic, 48 h; p<0.05). Similarly, following 24 h infection with a high viral titre

of 20 x 104 TCID50/ml of HRV-1B, the data demonstrated a significant decrease

(12.02% ± 0.002) in membrane claudin-1 expression in pAECs of the asthmatic cohort

compared to non-infected controls (Figure 6.1 A – Asthmatic, 24 h; p<0.05). At 48 h

post infection, a further decrease in membrane claudin-1 expression was observed

(3.73% ± 0.002) compared to non-infected controls (Figure 6.1 A – Asthmatic, 48 h;

p<0.05).

Interestingly, 24 h post infection, infection with the high viral titre resulted in increased

levels of membrane claudin-1 expression (12.02% ± 0.002) in contrast to decreased

expression levels post infection with the low viral titre (8.53% ± 0.0002). The

differences in expression was observed to be significant (Figure 6.1 A – Asthmatic, 24

h; p<0.05). Moreover, 48 h post infection, significant differences in membrane claudin-

1 expression was observed between both viral titres (Figure 6.1 A – Asthmatic, 48 h;

p<0.05).

When assessing membrane occludin expression between non-asthmatic and asthmatic

cohorts, results corroborated data from the previous chapters, demonstrating

significantly lower basal membrane occludin (40.3%) within the asthmatic cohort

compared to the non-asthmatic counterpart (100%) (Figure 6.1 B – 0 h; p<0.05).

Following 24 h infection with low viral titre of 2.5 x 104 TCID50/ml of HRV-1B, a

significant decrease in membrane occludin expression (57.8% ± 0.002) was observed

compared to non-infected controls in pAECs of non-asthmatic cohort (Figure 6.1 B –

Non-asthmatic, 24 h; p<0.05). Interestingly, a significant increase in membrane

occludin expression (83.5% ± 0.002) was shown at 48 h post infection (Figure 6.1 B –

Non-asthmatic, 48 h; p<0.05). After 24 h infection with a high viral titre of 20 x 104

TCID50/ml of HRV-1B, a significant decrease in membrane occludin expression (28.3%

± 0.001) was similarly observed compared to non-infected controls in pAECs of the

non-asthmatic cohort (Figure 6.1 B – Non-asthmatic, 24 h; p<0.05). In contrast, a

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significant increase towards baseline values in membrane occludin expression (71.4% ±

0.002) was observed 48 h post infection (Figure 6.1 B – Non-asthmatic, 48 h; p<0.05).

In addition, 24 h post infection with a higher viral titre resulted in a significant decrease

in membrane occludin expression (28.3% ± 0.001) within pAECs of the non-asthmatic

cohort in contrast to expression levels following infection with the lower viral titre

(57.8% ± 0.002) (Figure 6.1 B – Non-asthmatic, 24 h; p<0.05). Furthermore, 48 h post

infection, despite an increase in membrane occludin expression in pAECs of the non-

asthmatic cohort, infection with the higher viral titre continued to demonstrate

significantly lower level of membrane occludin expression (71.4% ± 0.002) in contrast

to expression levels post infection with the lower viral titre (83.5% ± 0.002) (Figure 6.1

B – Non-asthmatic, 48 h; p<0.05).

When membrane occludin expression of pAECs from the asthmatic cohort were

assessed following infection with a low viral titre of 2.5 x 104 TCID50/ml of HRV-1B,

the data demonstrated, at 24 h post infection, a significant decrease in membrane

occludin expression in pAECs of the asthmatic cohort (7.5% ± 0.003) compared to non-

infected controls (Figure 6.1 B – Asthmatic, 24 h; p<0.05). Furthermore, a continued

decrease in membrane occludin expression was observed (3.56% ± 0.005) 48 h post

infection (Figure 6.1 B – Asthmatic, 48 h; p<0.05). Following infection with a high

viral titre of 20 x 104 TCID50/ml of HRV-1B, results obtained demonstrated a

significant decrease in membrane occludin expression (3.1% ± 0.002) 24 h post

infection compared to non-infected controls (Figure 6.1 B – Asthmatic, 24 h; p<0.05).

Moreover, a sustained decrease in membrane occludin expression (3.3% ± 0.005) was

observed at 48 h post infection compared to non-infected controls(Figure 6.1 B –

Asthmatic, 48 h; p<0.05).

When assessing effects of viral titres on membrane occludin expression, 24 h post

infection with the higher viral titre resulted in significantly lower levels of membrane

occludin expression (3.1% ± 0.002) in contrast to expression levels following infection

with the lower viral titre (7.5% ± 0.003) (Figure 6.1 B – Asthmatic, 24 h; p<0.05).

Interestingly, 48 h post infection, data demonstrated a further decrease in membrane

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occludin expression following infection with the lower viral titre (3.56% ± 0.005) while

infection with higher viral titre showed a sustained low level of membrane occludin

expression (3.3% ± 0.005), however, there was no significant difference in membrane

occludin expression between both viral titres (Figure 6.1 B – Asthmatic, 48 h).

Assessment of membrane ZO-1 expression between non-asthmatic and asthmatic

cohorts demonstrated significantly lower basal membrane ZO-1 (57.4%) expression

within the asthmatic cohort compared to the non-asthmatic counterpart (100%),

corroborating earlier observations from the previous chapter (Figure 6.1 C – 0 h;

p<0.05). Following 24 h infection with a low viral titre of 2.5 x 104 TCID50/ml of HRV-

1B, a significant decrease in membrane ZO-1 expression (66.5% ± 0.0007) was

observed in comparison with non-infected controls in pAECs of non-asthmatic cohort

(Figure 6.1 C – Non-asthmatic, 24 h; p<0.05). A significant increase towards baseline in

membrane ZO-1 expression (95.1% ± 0.001) was observed at 48 h post infection

(Figure 6.1 C – Non-asthmatic, 48 h; p<0.05). When infected with a high viral titre of

20 x 104 TCID50/ml of HRV-1B, a significant decrease in membrane ZO-1 expression

(54.2% ± 0.0005) was similarly observed compared to non-infected controls at 24 h post

infection (Figure 6.1 C – Non-asthmatic, 24 h; p<0.05). A significant increase towards

baseline in membrane ZO-1 expression (76.5% ± 0.001) was equally observed at 48 h

post infection (Figure 6.1 C – Non-asthmatic, 48 h; p<0.05).

In addition, 24 h post infection with the higher viral titre resulted in a significant

decrease in membrane ZO-1 expression (54.2% ± 0.0005) within pAECs of the non-

asthmatic cohort in contrast to the expression levels following infection with the lower

viral titre (66.5% ± 0.0007) (Figure 6.1 C – Non-asthmatic, 24 h; p<0.05). Furthermore,

48 h post infection, despite an increase in membrane ZO-1 expression in pAECs of the

non-asthmatic cohort, infection with the higher viral titre continued to demonstrate

significantly lower levels of membrane ZO-1 expression (76.5%± 0.001) in contrast to

expression levels post infection with the lower viral titre (95.1% ± 0.001) (Figure 6.1C

– Non-asthmatic, 48 h; p<0.05).

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When membrane ZO-1 expression of pAECs from the asthmatic cohort was assessed

following infection with a low viral titre of 2.5 x 104 TCID50/ml of HRV-1B, the data

demonstrated a significant decrease in membrane ZO-1 expression (2.5% ± 0.0005) at

24 h post infection in comparison to non-infected controls (Figure 6.1 C – Asthmatic,

24 h; p<0.05). A further significant decrease in membrane ZO-1 expression was

observed (0.53% ± 0.001) at 48 h post infection compared to non-infected controls

(Figure 6.1 C – Asthmatic, 48 h; p<0.05). Data obtained following infection with a high

viral titre of 20 x 104 TCID50/ml of HRV-1B demonstrated a significant decrease in

membrane ZO-1 expression (1.9% ± 0.0006) at 24 h post infection compared to non-

infected controls (Figure 6.1 C – Asthmatic, 24 h; p<0.05). A further significant

decrease in membrane ZO-1 expression was observed (0.29% ± 0.0005) at 48 h post

infection with the high viral titre compared to non-infected controls (Figure 6.1 C –

Asthmatic, 48 h; p<0.05).

When assessing effects of viral titres on membrane ZO-1 expression, 24 h post infection

with the higher viral titre resulted in significantly lower levels of membrane ZO-1

expression (1.9% ± 0.0006) in contrast to expression levels following infection with the

lower viral titre (2.5% ± 0.0005) (Figure 6.1 C – Asthmatic, 24 h; p<0.05).

Furthermore, 48 h post infection, data demonstrated a significantly greater decrease in

membrane ZO-1 expression following infection with the higher viral titre (6.9-fold,

0.29% ± 0.0005) while infection with the lower viral titre showed a similar decrease in

membrane ZO-1 expression but of a smaller magnitude (4.8-fold, 0.53% ± 0.001)

(Figure 6.1 C – Asthmatic, 48 h; p<0.05).

When assessing membrane TJ protein expression of pAECs within the non-asthmatic

cohort following infection, results showed similar profiles of decreased membrane

protein expression at 24 h post infection with low viral titre, demonstrating the lowest

expression in membrane claudin-1 (35.5% ± 0.002), followed by occludin (57.8% ±

0.002) and ZO-1 (66.5% ± 0.0007) (Figure 6.1 A – C, Non-asthmatic, 24 h; p<0.05).

Interestingly, the greatest increase towards baseline expression values was observed in

membrane claudin-1 (105.8% ± 0.002), followed by ZO-1 (95.1% ± 0.001) and

occludin (83.5% ± 0.002) at 48 h post infection with the low viral titre (Figure 6.1 A –

Figure 6.1 Membrane TJ protein expression over time in pAECs from non-

asthmatic and asthmatic cohorts following viral infection: pAECs seeded on 96-well

micro-titre plates and grown to confluence were infected with two different HRV-1B

titres (2.5 and 20 x 104 TCID50/ml) over 48 h and membrane TJ protein expression

assessed via an optimised In-Cell™ Western assay as previously described (refer to

3.2.4). (A) Membrane claudin-1 protein expression decreased following infection with

HRV-1B at both viral titres at 24 h in the non-asthmatic cohort. However, at 48 h post

infection, a significant increase in membrane claudin-1 expression was observed in this

cohort. Significant difference in membrane claudin-1 expression was observed in

pAECs of asthmatic cohort at 24 h post infection compared to 0 h. Further significant

decrease in expression was observed at 48 h following infection compared to 0 h. (B)

Membrane occludin protein expression decreased following infection with HRV-1B at

both viral titres at 24 h in the non-asthmatic cohort. However, at 48 h post infection, a

significant increase in membrane occludin expression was observed within this cohort.

Significant decrease in membrane occludin expression was observed in pAECs of

asthmatic cohort at 24 h post infection with both viral titres compared to 0 h. Further

significant decrease in expression was similarly observed at 48 h following infection

compared to 0 h. (C) A significant decrease in membrane ZO-1 protein expression was

observed following infection with HRV-1B at both viral titres at 24 h in the non-

asthmatic cohort. However, at 48 h post infection, a significant increase in membrane

ZO-1 expression was observed in this cohort. Significant decrease in membrane ZO-1

expression was observed in pAECs of asthmatic cohort at 24 h post infection with both

viral titres compared to 0 h. Further significant decrease in expression was also

observed at 48 h following infection compared to 0 h. Data were normalised to cell

numbers and presented as mean ± SD percentage relative to control. *Statistical

significance relative to non-infected controls (p<0.05).

Time (h)

Mem

bran

e cl

audi

n ex

pres

sion

(% r

elat

ive

to c

ontr

ol)

0 24 480

25

50

75

100

125

150

2.5 x 104 TCID50/ml20 x 104 TCID50/ml

Non-asthmatic

2.5 x 104 TCID50/ml20 x 104 TCID50/ml

Asthmatic

* *

**

**

** *

Time (h)

Mem

bran

e oc

clud

in e

xpre

ssio

n(%

rel

ativ

e to

con

trol

)

0 24 480

25

50

75

100

125

150

2.5 x 104 TCID50/ml20 x 104 TCID50/ml

2.5 x 104 TCID50/ml20 x 104 TCID50/ml

Non-asthmatic

Asthmatic

* *

**

*

*

* *

Time (h)

Mem

bran

e ZO

1 ex

pres

sion

(% r

elat

ive

to c

ontr

ol)

0 24 480

25

50

75

100

125

150

2.5 x 104 TCID50/ml20 x 104 TCID50/ml

Non-asthmatic

2.5 x 104 TCID50/ml20 x 104 TCID50/ml

Asthmatic

* *

*

**

*

* *

A

B

C

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C, Non-asthmatic, 48 h; p<0.05). Following 24 h infection with a high viral titre, the

lowest expression was observed in membrane claudin-1 expression (25.3% ± 0.001),

followed by occludin (and 28.3% ± 0.001) and ZO-1 (54.2% ± 0.0005) (Figure 6.1 A –

C, Non-asthmatic, 24 h; p<0.05). A greatest increase towards baseline expression

values was similarly observed in membrane claudin-1 (82.2% ± 0.002), followed by

ZO-1 (76.5% ± 0.001) and occludin (71.4% ± 0.002) at 48 h post infection (Figure 6.1

A – C, Non-asthmatic, 48 h; p<0.05).

Similarly, when assessing membrane TJ protein expression of pAECs within the

asthmatic cohort following 24 h infection with a low viral titre, the lowest expression

was observed in membrane ZO-1 (2.5% ± 0.0005), followed by occludin (7.5% ± 0.003)

and claudin-1 (8.53% ± 0.0002) (Figure 6.1 A – C, Asthmatic, 24 h; p<0.05).

Surprisingly, similar levels of expression was observed between membrane claudin-1

(3.63% ± 0.001) and occludin (3.56% ± 0.005) with lowest expressed being membrane

ZO-1 (0.53% ± 0.001) at 48 h post infection (Figure 6.1 A – C, Asthmatic, 48 h;

p<0.05). Infection with a high viral titre demonstrated the greatest decrease in

membrane ZO-1 (1.9% ± 0.0006), followed by occludin (3.1% ± 0.002) and claudin-1

(12.02% ± 0.002) at 24 h post infection (Figure 6.1 A – C, Asthmatic, 24 h; p<0.05).

Interestingly, at 48 h post infection, similar levels of expression were observed between

membrane claudin-1 (3.73% ± 0.002) and occludin (3.3% ± 0.005) with membrane ZO-

1 being the lowest (0.29% ± 0.0005) (Figure 6.1 A – C, Asthmatic, 48 h; p<0.05).


atopic status


atopy, the results generated corroborated data from the previous chapter demonstrating

lower basal expression of membrane claudin-1 in pAECHA (10.2%), pAECNAA (5.7%)

and pAECAA (6.1%) when compared to pAECHNA (100%). In addition, following 24 h

infection with a low viral titre of 2.5x104 TCID50/ml of HRV-1B, results showed a

greater magnitude in the decrease of membrane claudin-1 expression in the pAECHNA

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cohort (33.2% ± 0.01) while expression of membrane claudin-1 in pAECHA was

maintained (10.3% ± 0.005) and a decrease in membrane claudin-1 protein expression

was observed in the pAECNAA (5.4%) and pAECAA (5.2% ± 0.001) cohorts when

compared to non-infected controls (Figure 6.2 A – 24 h; p<0.05). Interestingly, 48 h

post infection, significant return to baseline values for membrane claudin-1 expression

was observed in the pAECHNA (114% ± 0.002) and pAECHA (12.3% ± 0.02) cohorts,

while a decreased expression was observed in the pAECNAA (0.8%) and pAECAA (2.9%

± 0.005) cohorts in comparison to non-infected controls (Figure 6.2 A – 48 h; p<0.05).

When the effects of a low viral titre infection on membrane claudin-1 protein expression

at 24 h post infection was assessed, membrane expression was found to be lower in

pAECHA (10.3% ± 0.005), pAECNAA (5.4%) and pAECAA (5.2% ± 0.001) in contrast to

pAECHNA (33.2% ± 0.01). This was observed to be significant for the pAECHA and

pAECAA cohorts. Expression of membrane claudin-1 protein was also lower in

pAECNAA (5.4%) and pAECAA (5.2% ± 0.001) in comparison with pAECHA (10.3% ±

0.005) and this was observed to be of significance for the pAECAA cohort. Although

there was no observable difference in membrane claudin-1 expression between the

pAECNAA and pAECAA cohorts, due to the limited availability of pAECNAA, statistical

analysis could not be performed (Figure 6.2 A – 24 h; p<0.05). At 48 h post infection,

membrane expression of claudin-1 was similarly lower in the pAECHA (12.3% ± 0.02),

pAECNAA (0.8%) and pAECAA (2.9% ± 0.005) in contrast to pAECHNA (114% ±0.002).

This was observed to be significant for the pAECHA and pAECAA cohorts. Expression of

membrane claudin-1 protein was also lower in pAECNAA (0.8%) and pAECAA (2.9% ±

0.005) in comparison with pAECHA (12.3% ± 0.02) and this was observed to be of

significance for the pAECAA cohort. Moreover, membrane claudin-1 expression was

observed to be lower in pAECNAA (0.8%) compared to pAECAA (12.3% ± 0.02).

However, due to the limited availability of pAECNAA, statistical analysis could not be

performed (Figure 6.2 A – 48 h; p<0.05) for this cohort.

Following 24 h infection with a high viral titre of 20 x 104 TCID50/ml of HRV-1B, a

decrease in membrane claudin-1 protein expression was demonstrated in the pAECHNA

(21.6% ± 0.003), pAECHA (8.8% ± 0.005) and pAECNAA (5.4%) cohorts while an

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increase in expression was observed in the pAECAA (8.1% ± 0.008) cohort. These were

observed to be significant in the pAECHNA, pAECHA and pAECAA cohorts when

compared to non-infected controls (Figure 6.2 D – 24 h; p<0.05). Interestingly, despite

an increase in membrane claudin-1 expression at 48 h post infection compared to 24 h

post infection, membrane claudin-1 expression remained significantly lower than non-

infected controls in the pAECHNA (76.5% ± 0.003) and pAECHA (9.9% ± 0.02) cohorts.

In contrast, a decrease in membrane claudin-1 expression was observed in pAECNAA

(0.8%) and pAECAA (3.1% ± 0.005) cohorts when compared to non-infected controls.

This was observed to be significant in the pAECAA cohort (Figure 6.2 D – 48 h;

p<0.05). Due to the limited availability of pAECNAA, statistical analysis could not be

performed for this cohort.

In the assessment of the effects of a high viral titre infection on membrane claudin-1

protein expression at 24 h post infection, membrane expression of claudin-1 was lower

in pAECHA (8.8% ± 0.005), pAECNAA (5.4%) and pAECAA (8.1% ± 0.008) in contrast to


pAECAA cohorts. Expression of membrane claudin-1 protein was also lower in the

pAECNAA (5.4%) and pAECAA (8.1% ± 0.001) in comparison with pAECHA (8.8% ±

0.005) and this was observed to be of significance for the pAECAA cohort (Figure 6.2 D

– 24 h; p<0.05). Although membrane claudin-1 expression was observed to be lower in

pAECNAA compared to the pAECAA cohorts, statistical analysis could not be performed

due to limited availability of pAECNAA. At 48 h post infection, membrane expression of

claudin-1 was similarly lower in pAECHA (9.9% ± 0.02), pAECNAA (0.8%) and pAECAA

(3.1% ± 0.005) in contrast to pAECHNA (76.5% ± 0.003). This was observed to be

significant for the pAECHA and pAECAA cohorts. Expression of membrane claudin-1

protein was also lower in pAECNAA (0.8%) and pAECAA (3.1% ± 0.005) in comparison

to pAECHA (9.9% ± 0.02) and this was observed to be of significance for the pAECAA

cohort. Moreover, membrane claudin-1 expression was observed to be lower in

pAECNAA (0.8%) compared to pAECAA (3.1% ± 0.005). However, due to the limited

availability of pAECNAA, statistical analysis could not be performed (Figure 6.2 D – 48

h; p<0.05).

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When membrane occludin expression was assessed, the results also corroborated data

from the previous chapter demonstrating lower basal expression of membrane occludin

in pAECHA (35.9%), pAECNAA (27.2%) and pAECAA (27.5%) when compared to

pAECHNA (100%). In addition, following 24 h infection with a low viral titre, data

showed a greater decrease of membrane occludin expression in the pAECHNA cohort

(46.4% ± 0.03) while expression in pAECHA was maintained (34.5% ± 0.02) and a

decrease in membrane occludin protein expression was observed for both the pAECNAA

(20.2% ± 0.03) and pAECAA (20.2% ± 0.01) cohorts (Figure 6.2 B – 24 h; p<0.05).

Interestingly, 48 h post infection, although an increase in membrane occludin

expression was observed compared to expression levels at 24 h post infection,

expression was still significantly lower in pAECHNA (84.7% ±0.02) when compared to

non-infected controls. Further decrease in membrane occludin expression was observed

in the pAECHA (20% ± 0.02), pAECNAA (3.8%) and pAECAA (12.6% ± 0.02) cohorts

when compared to non-infected controls and this was significant for both the pAECHA

and pAECAA cohorts (Figure 6.2 B – 48 h; p<0.05). Due to the limited availability of

pAECNAA, statistical analysis could not be performed.

In assessing the effects of a low viral titre infection on membrane occludin protein

expression at 24 h post infection, membrane expression of occludin was significantly

lower in pAECHA (34.5% ± 0.02), pAECNAA (20.2% ± 0.03) and pAECAA (20.2% ±

0.01) in contrast to pAECHNA (46.4% ± 0.03). Expression of membrane occludin was

also significantly lower in pAECNAA (20.2% ± 0.03) and pAECAA (20.2% ± 0.01) in

comparison with pAECHA (34.5% ± 0.02). In addition, there was no observable

differences in membrane occludin expression between the pAECNAA and pAECAA

cohorts (Figure 6.2 B – 24 h; p<0.05). At 48 h post infection, membrane expression of

occludin was similarly lower in pAECHA (20% ± 0.02), pAECNAA (3.8%) and pAECAA

(12.6% ± 0.02) compared to pAECHNA (84.7% ±0.02). This was observed to be

significant for the pAECHA and pAECAA cohorts. Expression of membrane occludin was

also lower in pAECNAA (3.8%) and pAECAA (12.6% ± 0.02) in comparison with

pAECHA (20% ± 0.02) and this was observed to be of significance for the pAECAA

cohort. Moreover, membrane occludin expression was observed to be lower in


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availability of pAECNAA, statistical analysis could not be performed to determine

significance (Figure 6.2 B – 48 h; p<0.05).


significant decrease in membrane occludin protein expression was demonstrated in the

pAECHNA (16.2% ± 0.007), pAECHA (21% ± 0.01), pAECNAA (3.7% ± 0.001) and

pAECAA (14.7% ± 0.01) cohorts when compared to non-infected controls (Figure 6.2 E

– 24 h; p<0.05). Interestingly, despite an increase in membrane occludin expression at

48 h post infection compared to expression levels at 24 h, membrane occludin

expression remained significantly lower than non-infected controls in the pAECHNA

(68.7% ± 0.01) cohort. In contrast, a further decrease in membrane occludin expression

was observed in pAECHA (17.7% ± 0.01), pAECNAA (3.6%) and pAECAA (11.8% ±

0.02) cohorts when compared to expression levels both at 24 h post infection as well as

in non-infected controls. This was observed to be significant in both the pAECHA and

pAECAA cohorts (Figure 6.2 E – 48 h; p<0.05). Due to the limited availability of

pAECNAA, statistical analysis could not be performed to determine significance.

When the effects of a high viral titre infection on membrane occludin protein expression

at 24 h post infection were assessed, membrane expression of occludin was significantly

higher in pAECHA (21% ± 0.01) compared to pAECHNA (16.2% ± 0.01) while membrane

occludin expression was lower in pAECNAA (3.7% ± 0.001) and pAECAA (14.7% ±

0.01) compared to pAECHNA (16.2% ± 0.01). Expression of membrane occludin protein

was also significantly lower in pAECNAA (3.7% ± 0.001) and pAECAA (14.7% ± 0.01)

in comparison with pAECHA (21% ± 0.01) (Figure 6.2 E – 24 h; p<0.05). Membrane

occludin expression was observed to be lower in pAECNAA (3.7% ± 0.001) compared to

pAECAA cohorts (14.7% ± 0.01). At 48 h post infection, membrane expression of

occludin was similarly lower in pAECHA (17.7% ± 0.01), pAECNAA (3.6%) and pAECAA

(11.8% ± 0.02) in contrast to pAECHNA (68.7% ± 0.01). This was observed to be

significant for pAECHA and pAECAA cohorts. Expression of membrane occludin protein

was also lower in pAECNAA (3.6%) and pAECAA (11.8% ± 0.02) in comparison with

pAECHA (17.7% ± 0.01) and this was observed to be of significance for the pAECAA

cohort. Moreover, membrane occludin expression was observed to be lower in

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availability of pAECNAA, statistical analysis could not be performed to determine

significance (Figure 6.2 E – 48 h; p<0.05).

Similarly, when membrane ZO-1 protein expression was assessed, the data generated

corroborated results from the previous chapter demonstrating lower basal expression of

membrane ZO-1 in pAECHA (49.2%), pAECNAA (40%) and pAECAA (45%) when

compared to pAECHNA (100%). In addition, following 24 h infection with a low viral

titre, the obtained data showed the greatest magnitude in decrease of membrane ZO-1

expression in the pAECHNA cohort (59.1% ± 0.01). A decrease in expression of

membrane ZO-1 was similarly observed in pAECHA (43.8% ± 0.005), pAECNAA

(38.5%) and pAECAA (36.3% ± 0.002) cohorts when compared to non-infected controls

(Figure 6.2 C – 24 h; p<0.05). Interestingly, , although an increase in membrane ZO-1

expression was observed at 48 h post infection compared to expression levels at 24 h,

membrane ZO-1 expression was still significantly lower in pAECHNA (94.3% ±0.004) in

contrast to non-infected controls. A further decrease in membrane ZO-1 expression was

observed in the pAECHA (27.3% ± 0.001), pAECNAA (11.8%) and pAECAA (3.8%)

cohorts compared to non-infected controls and this was significant for the pAECHA

cohort (Figure 6.2 C – 48 h; p<0.05). Due to the limited availability of pAECNAA and

pAECAA, statistical analysis could not be performed to determine the level of

significance.

In the assessment of the effects of a low viral titre infection on membrane ZO-1 protein

expression at 24 h post infection, membrane expression of ZO-1 was lower in pAECHA

(43.8% ± 0.005), pAECNAA (38.5%) and pAECAA (36.3% ± 0.002) in contrast to


pAECAA cohorts. Expression of membrane ZO-1 protein was also lower in pAECNAA

(38.5%) and pAECAA (36.3% ± 0.002) compared to pAECHA (43.8% ± 0.005) and this

was observed to be of significance for the pAECAA cohort. Moreover, there was no

observable difference in membrane ZO-1 expression between pAECNAA and pAECAA

cohorts and since there was limited availability of pAECNAA, statistical analysis could

not be performed (Figure 6.2 C – 24 h; p<0.05). At 48 h post infection, membrane

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expression of ZO-1 was similarly lower in pAECHA (27.3% ± 0.01), pAECNAA (11.8%)

and pAECAA (3.8%) compared to pAECHNA (94.3% ±0.004). This was observed to be

significant for the pAECHA cohort. Expression of membrane ZO-1 protein was also

lower in pAECNAA (11.8%) and pAECAA (3.8%) in comparison to pAECHA (27.3% ±

0.01). Moreover, membrane ZO-1 expression was observed to be lower in pAECNAA

(0.8%) compared to pAECAA (12.3% ± 0.02), however, due to the limited availability of

pAECNAA and pAECAA, statistical analysis could not be performed to determine

significance (Figure 6.2 C – 48 h).


significant decrease in membrane ZO-1 protein expression was demonstrated in the

pAECHNA (54% ± 0.003), pAECHA (30.9% ± 0.003), pAECNAA (37.7%) and pAECAA

(25.7% ± 0.0004) cohorts when compared to non-infected controls (Figure 6.2 F – 24 h;

p<0.05). Interestingly, despite an increase in membrane ZO-1 expression at 48 h post

infection compared to expression levels at 24 h, membrane ZO-1 expression remained

significantly lower than non-infected controls in the pAECHNA (63.1% ± 0.005) cohort.

In contrast, a further decrease in membrane ZO-1 expression was observed in the

pAECHA (23.5% ± 0.01), pAECNAA (7.3%) and pAECAA (2.7% ± 0.001) cohorts when

compared to expression levels at 24 h post infection as well as in non-infected controls.

This was observed to be significant in pAECHA and pAECAA cohorts (Figure 6.2 F – 48

h; p<0.05). Due to the limited availability of pAECNAA, statistical analysis could not be

performed to determine significance at this time point.

When assessing the effects of a high viral titre infection on membrane ZO-1 protein

expression at 24 h post infection, membrane expression of ZO-1 was lower in pAECHA

(30.9% ± 0.003), pAECNAA (37.7%) and pAECAA (25.7% ± 0.0004) in contrast to

pAECHNA (54% ± 0.003). This was observed to be significant for the pAECHA and

pAECAA cohorts (Figure 6.2 F – 24 h; p<0.05). Expression of membrane ZO-1 protein

was also lower in pAECNAA (37.7%) and pAECAA (25.7% ± 0.0004) in comparison with

pAECHA (30.9% ± 0.003). This was observed to be of significance in pAECAA cohort

(Figure 6.2 F – 24 h; p<0.05). Membrane ZO-1 expression was also observed to be

lower in pAECNAA (37.7%) compared to pAECAA cohorts (25.7% ± 0.0004), however,

Figure 6.2 Membrane TJ protein expression over time in pAECs from non-asthmatic and asthmatic cohorts following viral infection with each

cohort further categorised based on atopy: pAECs seeded on 96-well micro-titre plates and grown to confluence were infected with two different

HRV-1B titres (2.5 and 20 x 104 TCID50/ml) for 24 and 48 h and membrane TJ protein expression assessed via an optimised In-Cell™ Western assay as

previously described (refer to 3.2.4). (A - C) Following infection with HRV-1B at viral titre of 2.5 x 104 TCID50/ml ( ), a significant decrease in

membrane claudin-1 (green), occludin (blue) and ZO-1 (red) protein respectively was observed in pAECHNA cohort at 24 h. However, at 48 h post

infection, membrane protein expression for all 3 tight junctions was significantly higher. A significant decrease in all 3 membrane tight junction

protein expression was observed in pAECHA, pAECNAA and pAECAA cohorts at 24 h post infection. In all 3 phenotypic cohorts at 48 h following

infection, significantly sustained decreased membrane protein expression was observed for claudin-1 while a significantly further decrease was

observed for both occludin and ZO-1. (D – F) Following infection with HRV-1B at viral titre of 20 x 104 TCID50/ml ( ), a significant decrease in all

3 membrane tight junction protein was observed in pAECHNA cohort at 24 h. Although membrane protein expression for all 3 tight junctions were

significantly higher at 48 h compared to 24 h, expression levels remain significantly lower compared to non-infected control. A significant decrease in

all 3 membrane tight junction protein expression was observed in pAECHA, pAECNAA and pAECAA cohorts at 24 h post infection. In all 3 phenotypic

cohorts at 48 h following infection, significantly sustained decreased membrane protein expression was observed for claudin-1 and occludin while a

significantly further decrease was observed for ZO-1. Data were normalised to cell numbers and presented as percentage mean ± SD; n = 5 individual

experiments each performed in duplicates with the exception of pAECNAA phenotype (n=2). *Statistical significance relative to non-infected control

(p<0.05).

0 24 480

25

50

75

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*

*

* *0 24 48

0

25

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*

*

**

0 24 480

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50

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*

*

**

0 24 480

25

50

75

100

125

150

*

*

* *0 24 48

0

25

50

75

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125

150

*

* *

0 24 480

25

50

75

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150

* *

**

A

D

B C

E F

Me

mb

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rote

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xpre

ssio

n (

% r

elat

ive

to c

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tro

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Time (h)

2.5 x 104 TCID50/ml 20 x 104 TCID50/ml

0 24 480

25

50

75

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125

150HNA C

HNA HA NAA AA

0 24 480

25

50

75

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125

150HNA C

HNA HA NAA AA

0 24 480

25

50

75

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125

150HNA C

HNA HA NAA AA

0 24 480

25

50

75

100

125

150HNA C

HNA HA NAA AA

0 24 480

25

50

75

100

125

150HNA C

HNA HA NAA AA

0 24 480

25

50

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150HNA C

HNA HA NAA AA

0 24 480

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150HNA C

HNA HA NAA AA

Looi 2015

134

due to limited availability of pAECNAA, statistical analysis could not be performed to

determine significance. At 48 h post infection, membrane expression of ZO-1 was

similarly lower in pAECHA (23.5% ± 0.01), pAECNAA (7.3%) and pAECAA (2.7% ±

0.001) in contrast to pAECHNA (63.1% ± 0.005). This was observed to be significant for

the pAECHA and pAECAA cohorts. The expression of membrane ZO-1 protein was also

lower in pAECNAA (7.3%) and pAECAA (2.7% ± 0.001) compared to pAECHA (23.5% ±

0.01) and this was observed to be of significance for the pAECAA cohort. Moreover,

membrane ZO-1 expression was observed to be lower in pAECNAA (7.3%) compared to

pAECAA (2.7% ± 0.001). However, due to the limited availability of pAECNAA,

statistical analysis could not be performed (Figure 6.2 F – 48 h; p<0.05).

6.3.2 Effect of human rhinovirus infection on in vitro transepithelial permeability

In order to correlate HRV-1B induced disassembly of membrane TJ with barrier

function, a transepithelial permeability assay was performed after 24 h infection with a

viral titre of 10x104 TCID50/ml of HRV-1B. This viral titre has been previously

demonstrated (Kicic, unpublished data) to result in a significant loss of cellular viability

but no increase in cellular apoptosis in pAECs of non-asthmatic and asthmatic cohorts.

Hence, any change in epithelial permeability would be implicated with an alteration of

membrane TJ protein expression.


Non-asthmatic pAECs demonstrated an increase in transepithelial permeability of

FITC-dextran 4 (427 x 10-4 cm/sec ± 17.8) and 20 (187.9 x 10-4 cm/sec ± 27.1) kDa

following 24 h infection with HRV-1B when compared to non-infected controls (302.9

x 10-4 cm/sec ± 20.5 and 99.1 x 10-4 cm/sec ± 32.7 respectively). This was observed to

be significant for FITC-dextran 4 kDa (Figure 6.3 A; p<0.05). However, a significantly

higher level of transepithelial permeability for FITC-dextran of 4 kDa (427 x 10-4

cm/sec ± 17.8) was observed when compared to epithelial permeability towards FITC-

dextran 20 kDa (187.9 x 10-4 cm/sec ± 27.1) (Figure 6.3 A – Infected; p<0.05).

Figure 6.3 Transepithelial permeability in pAECs from non-asthmatic and asthmatic cohorts following viral infection: pAECs from non-

asthmatic and asthmatic cohorts, seeded onto Corning transwell inserts and grown to confluence were treated as previously mentioned (refer to 2.5.11).

Following 24 h infection with a viral titre of 10 x 104 TCID50/ml of HRV-1B, an optimised transepithelial permeability assay as previously described

was performed to determine epithelial permeability to FITC-dextran 4 kDa (blue) and 20 kDa (grey) (refer to 3.3.4). (A) Increase in transepithelial

permeability towards both FITC-dextran 4 and 20 kDa was observed in pAECs of non-asthmatic cohorts following infection, however, this was only

observed to be significant for FITC-dextran 4 kDa. Epithelial permeability towards FITC-dextran 4 kDa was significantly higher when compared to

FITC-dextran 20 kDa in non-infected and infected pAECs of non-asthmatic cohort. (B) No significant difference in transepithelial permeability

towards both FITC-dextran 4 and 20 kDa was observed in pAECs of asthmatic cohorts following infection. However, permeability towards FITC-

dextran 4 kDa was significantly higher when compared to FITC-dextran 20 kDa in non-infected and infected pAECs of asthmatic cohort. Results are

presented as mean ± SD. *Statistical significance relative to non-infected control (p<0.05). # Statistical significance relative to FITC-dextran 20 kDa

(p<0.05).

Non-infected Infected0

100

200

300

400

5004kDa20kDa

*#

Pap

p co

effic

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(cm

/sec

) x10

-4

n=10 n=10

#


100

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Pap

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n=7 n=7

##

A B

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135

Similarly, in pAECs of the asthmatic cohort following infection with HRV-1B, data

demonstrated an increase in transepithelial permeability of FITC-dextran 4 and 20 kDa

compared to non-infected controls but this was not found to be statistically significant

(Figure 6.3 B). However, a significantly higher level of transepithelial permeability for

FITC-dextran of 4 kDa (371.1 x 10-4 cm/sec ± 17.8) was observed when compared to

epithelial permeability for FITC-dextran 20 kDa (219.1 x 10-4 cm/sec ± 27.1) (Figure

6.3 B – Infected; p<0.05).


atopic status


atopy, the results demonstrated a significant increase in transepithelial permeability for

both FITC-dextran 4 kDa (442.4 x 10-4 cm/sec ± 6.8) and 20 kDa (288.9 x 10-4 cm/sec ±

6.9) within the pAECHNA cohorts following infection compared to non-infected controls

(370.3 x 10-4 cm/sec ± 13.3 and 201.8 x 10-4 cm/sec ± 34.2 respectively) (Figure 6.4 A;

p<0.05). In addition, a significantly higher level of transepithelial permeability towards

FITC-dextran of 4 kDa (442.4 x 10-4 cm/sec ± 6.8) was observed in comparison to

epithelial permeability towards FITC-dextran 20 kDa (288.9 x 10-4 cm/sec ± 6.9)

(Figure 6.4 A – Infected; p<0.05).

A significant increase in permeability to FITC-dextran 4 (461.8 x 10-4 cm/sec ± 13.9)

and 20 kDa (276.9 x 10-4 cm/sec ± 2.9) was observed following similar exposure to

HRV-1B in the pAECHA cohorts compared to their non-infected controls (272.6 x 10-4

cm/sec ± 14.6 and 188.3 x 10-4 cm/sec ± 34.8 respectively) (Figure 6.4 B; p<0.05). In

addition, a significantly higher level of transepithelial permeability towards FITC-

dextran of 4 kDa (461.8 x 10-4 cm/sec ± 13.9) was observed when compared to

epithelial permeability towards FITC-dextran 20 kDa (276.9 x 10-4 cm/sec ± 2.9)

(Figure 6.4 B – Infected; p<0.05).

Figure 6.4 Transepithelial permeability in pAECs from non-asthmatic and asthmatic cohorts following viral infection with each cohort

further categorised based on atopy: pAECs from non-asthmatic and asthmatic cohorts, seeded onto Corning transwell inserts and grown to

confluence were treated as previously mentioned (refer to 2.5.11). Following 24 h infection with a viral titre of 10 x 104 TCID50/ml of HRV-1B, an

optimised transepithelial permeability assay as previously described was performed to determine epithelial permeability to FITC-dextran 4 kDa (blue)

and 20 kDa (grey) (refer to 3.3.4). (A) Significant increase in transepithelial permeability was observed for both FITC-dextran 4 and 20 kDa following

infection in the pAECHNA cohort. Epithelial permeability to FITC-dextran 4 kDa was also significantly higher compared to FITC-dextran 20 kDa in

both non-infected and infected pAECHNA. (B) Significant increase in transepithelial permeability was observed for both FITC-dextran 4 and 20 kDa

following infection in the pAECHA cohort. Epithelial permeability to FITC-dextran 4 kDa was also significantly higher compared to FITC-dextran 20

kDa in both non-infected and infected pAECHA. (C) No observable difference in epithelial permeability was shown in pAECNAA cohort and due to the

low sample size of pAECNAA, statistical analysis could not be performed to determine significance and was excluded from comparative statistical

analysis. (D) Significant increase in transepithelial permeability was only observed for FITC-dextran 4 kDa following infection in the pAECAA cohort.

Epithelial permeability to FITC-dextran 4 kDa was also significantly higher compared to FITC-dextran 20 kDa in only the infected pAECAA. Results

are presented as mean ± SD. *Statistical significance relative to non-infected control (p<0.05). # Statistical significance relative to FITC-dextran 20

kDa (p<0.05).


100

200

300

400

500*#

*

n=5 n=5

#


100

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n=5 n=5

#

*#


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A B

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*

*

Papp

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oef

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10

-4

Looi 2015

136

An increase in permeability to FITC-dextran 4 (91.6 x 10-4 cm/sec) and 20 kDa (37.7 x

10-4 cm/sec) was observed following similar exposure to HRV-1B in the pAECNAA

cohorts when compared to their non-infected controls (78.6 x 10-4 cm/sec and 30.6 x 10-

4 cm/sec) (Figure 6.4 C). Furthermore, a higher level of transepithelial permeability to

FITC-dextran of 4 kDa (91.6 x 10-4 cm/sec) was observed in comparison to epithelial

permeability to FITC-dextran 20 kDa (37.7 x 10-4 cm/sec) (Figure 6.4 C – Infected).

However, due to the limited availability of pAECNAA cultures, statistical analysis could

not be performed to determine significance.

Similarly, following infection with HRV-1B in the pAECAA cohort, an increase in

permeability to FITC-dextran 4 (442.2 x 10-4 cm/sec ± 17.2; p<0.05) and 20 kDa (268.1

x 10-4 cm/sec ± 16.3) was observed compared to non-infected controls (320.5 x 10-4

cm/sec ± 39.9 and 221.7 x 10-4 cm/sec ± 28.9) (Figure 6.4 D). In addition, a

significantly higher level of transepithelial permeability towards FITC-dextran of 4 kDa

(442.2 x 10-4 cm/sec ± 17.2) was observed compared to the epithelial permeability

towards FITC-dextran 20 kDa (268.1 x 10-4 cm/sec ± 16.3) (Figure 6.4 D – Infected;

p<0.05).

Interestingly, no significant differences in epithelial permeability was observed (Figures

6.4 A - D, Infected) when assessing epithelial permeability to FITC-dextran 4 and 20

kDa following viral infection between pAECHNA, pAECHA, pAECNAA and pAECAA

cohorts.

6.4 Discussion

A seminal study by Sajjan and colleagues (2008) has previously shown that HRV

infection has the capacity to disrupt epithelial barrier integrity disruption in non-

asthmatic AECs. In a separate study, Kicic and colleagues (2006) have previously

demonstrated that there are intrinsic biochemical and functional differences between the

epithelium of children with and without asthma and that the paediatric asthmatic

epithelium exhibits an inability for complete repair after injury (Stevens et al. 2008).

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Hence, when viewed collectively, these studies provide the rationale and aims of the

current study, which aimed to assess tight junctional expression and barrier

functionality in response to HRV infection in paediatric AECs of non-asthmatic and

asthmatic cohorts.

The airway epithelium is constantly exposed to a myriad of potentially injurious

physical, chemical and biological agents. Some of these agents are capable of causing

inflammatory responses which can also result in further exacerbations of any underlying

respiratory conditions such as asthma, chronic obstructive pulmonary disease (COPD)

or other serious respiratory diseases. Human rhinoviruses have been shown to be the

most common precipitants of acute exacerbations of asthma (Johnston et al. 1995;

Corne et al. 2002; Chauhan et al. 2003; Grissell et al. 2005; Papi et al. 2006) and that

different serotypes of HRV could induce different responses in AECs (Nakagome et al

2014). However, the lack of a suitable animal model of asthma and HRV infection has

led to numerous studies utilising in vitro primary AECs from both non-asthmatic and

asthmatic individuals to better understand the airway biology following experimental

HRV infection in different cells of the lungs (Wark et al. 2005; Bochkov et al. 2010;

Cakebread et al. 2011).

Although Yeo and Jang were able to demonstrate an association between HRV infection

and decreased expression of epithelial junctional complexes as well as impaired barrier

function through measurement of transepithelial electrical resistance measurements, the

study was performed strictly on nasal derived AECs. A previous study by Lopez-Souza

and co-workers (2009) have demonstrated differences in resistance to HRV infection

between nasal and bronchial epithelial cells while others have demonstrated increased

susceptibility of bronchial epithelial cells to HRV infection (Jakiela et al. 2008).

Furthermore, evidence from past studies have demonstrated increased viral replication

in bronchial epithelial cells of asthmatic patients compared to healthy individuals,

suggesting that HRV infection could be an important cause of lower respiratory airway

disease (Papadopoulos et al. 2000; Contoli et al. 2006). Collectively, this emphasises

the limitation of utilising nasal AECs to study junctional complex expression in

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bronchial asthma as well as the need for caution when extrapolating data to bronchial

AECs.

A body of evidence has shown that HRV infections not only constitute the most

common cause of acute illness and wheezing during infancy, but has also been

implicated in the possible development and subsequent exacerbations of asthma later in

life (Kotaniemi-Syrjänen et al. 2003; Hyvärinen et al. 2005; Lemanske et al. 2005;

Kusel et al. 2007; Lee et al. 2007; Jackson et al. 2008; Kusel et al. 2012). Despite this

evidence which have been pivotal in demonstrating dissociation of specific TJ

complexes following HRV infection or exposure to cigarette smoke extract resulting in

a change in epithelial permeability, present studies have continued to utilise non-

paediatric cohorts for assessing epithelial barrier integrity. Hence, there remains a

paucity of data within the paediatric population and thus, findings from this study would

provide a glimpse of epithelial TJ expression profiles and function following HRV

infection in paediatric asthma. This complements those observed by Xiao and

colleagues in providing a snap-shot overview of epithelial TJ expression profiles and

function in early life for comparison to adulthood.

Basal expression data in this chapter is corroborated by those observed in Chapter 5,

demonstrating differences in basal membrane TJ protein expression within asthmatic

pAECs compared to non-asthmatic controls. Furthermore, when infected with HRV-1B

for 24 h, a loss in membrane TJ expression as observed in both non-asthmatic and

asthmatic cohorts could indicate an increased propensity for the movement of

pathogenic molecules or aeroallergens into the basement membrane. Interestingly,

despite the loss in both occludin and ZO-1 expression following infection at 24 h within

the asthmatic cohort, no change in membrane claudin-1 expression was observed. This

suggests a possible compensatory effect by the epithelial cells to maintain barrier

integrity, as shown by Saitou and colleagues (1998; 2000) in their occludin knock-out

studies demonstrating that occludin was not required for the formation of

morphologically intact TJs and strongly suggested that other junctional complexes were

capable of compensating for the lack of occludin expression.

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Interestingly, when HRV infection was extended over 48 h, a restoration of membrane

TJ expression of claudin-1, occludin and ZO-1 was observed within the non-asthmatic

cohort, suggesting that, in individuals without asthma, the epithelial cells might attempt

to re-assemble membrane TJ complexes in order to regain barrier integrity. In contrast,

membrane TJ protein expression within the asthmatic cohort was not restored. This

complements previous findings that the asthmatic epithelium is intrinsically different

and dysregulated (Kicic et al. 2006; Stevens et al. 2008; Kicic 2010) and suggests that

infection with HRV could result in a further loss of restorative capability in re-forming

membrane TJ complexes over extended periods.

When the data obtained in this study was further categorised according to atopy, it was

observed that the pAECHA, pAECNAA and pAECAA cohorts demonstrated similar

diminished membrane TJ protein expression profiles at a basal level in contrast to

pAECHNA. When infection was extended over 48 h, similar decrease in membrane TJ

protein expression profiles were observed for the pAECHA, pAECNAA and pAECAA

cohorts compared to pAECHNA, which demonstrated a restoration of membrane TJ

protein expression. This suggests the possible contribution of atopy in influencing TJ

protein expression, as demonstrated in studies investigating the effects of atopic

dermatitis on epidermal barrier integrity and function (De Benedetto et al. 2011; Kuo et

al. 2012) as well as the potential interaction with asthma in influencing eventual

membrane TJ protein expression.

Epithelial permeability towards various sized molecules is often a defining hallmark of

barrier integrity as demonstrated in numerous studies (Wan et al. 1999; Dreschers et al.

2007; Sajjan et al. 2008; Xiao et al. 2011). In the current study, when the observed post

HRV infection differences in membrane TJ protein expression were assessed

functionally, an increase in permeability, especially towards smaller sized

macromolecules, was observed in pAECs from the non-asthmatic cohort. A separate

study by Stevens and colleagues (2009) demonstrated that the viral titre utilised in this

study did not result in cellular apoptosis. Hence any change in epithelial permeability

could very likely be attributed to the disassembly of membrane TJ proteins following

HRV infection. Although different cell types were utilised, results from this study is in

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140

accordance as well as extend those from Sajjan and colleagues (2008), where they

demonstrated a significant increase in epithelial permeability following HRV infection

in polarised immortalised bronchial epithelial cell lines when compared to non-infected

or sham-infected controls.

Data from this chapter corroborated earlier observations of low basal membrane protein

expression levels in pAECAA and went on to demonstrate that although infection with

HRV causes further decreases in specific membrane TJ proteins, this does not result in

significantly increased epithelial permeability. Collectively, these results suggest that,

with the current monolayer asthmatic culture model, HRV infection has no additional

impact on an epithelium which is already inherently dysregulated and impaired.

When the data was further classified according to atopy, despite significant increases in

epithelial permeability towards the small sized macromolecule following HRV infection

in the pAECHNA, pAECHA, pAECNAA and pAECAA cohorts, there were no observable

differences between the cohorts. However, a drawback of the approach to assessing

barrier functionality in the present study is the use of a monolayer culture model, which

might not be the most reflective of an in vivo environment. Xiao and colleagues (2011),

in their study, utilised differentiated adult-derived pAECs of both non-asthmatic and

asthmatic cohorts to demonstrate significant differences in baseline epithelial

permeability of macromolecules of two different sizes. Interestingly, when basal

epithelial permeability of pAECs from non-asthmatic and asthmatic cohorts in this

present study were compared to the observations by Xiao and colleagues, results from

this study demonstrated significantly higher levels of baseline permeability. This

suggests that a lack of differentiation within the pAECs could partially explain the lack

of observable difference between the phenotypic cohorts following HRV infection

despite significant differences in permeability at baseline. This emphasises the potential

limitation of using a monolayer culture model in the assessment of transepithelial

permeability and provides the rationale for utilising a well-differentiated culture model

of paediatric derived pAECs to elucidate airway epithelium biology in response to HRV

infections.

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6.5 Conclusion

Human rhinovirus infections have been shown to be a major trigger in asthma

exacerbations (Nicholson et al. 1993; Johnston et al. 1995; Wark et al. 2002; Grissell et

al. 2005; Bizzintino et al. 2011; Cox et al. 2013) and in this chapter, data obtained has

demonstrated that disassembly of specific TJ complexes ensues in response to HRV

infection in both pAECs of non-asthmatic and asthmatic cohorts. However, restoration

of membrane TJ complexes occurs within pAECs of the non-asthmatic cohort while

sustained disassembly of TJ complexes were observed in pAECs of the asthmatic

cohort. Increases in epithelial permeability following HRV infection in the non-

asthmatic cohort further confirm disassembly of TJ complexes. However, a non-

significant difference in permeability post infection in the asthmatic cohort indicates a

lack of observable impact of HRV infection on increasing epithelial permeability. This

is observed despite an apparent decline in membrane TJ complexes expression. In

addition, although results from the different phenotypic cohorts are indicative of an

increase in epithelial permeability following HRV infection, the non-significant

differences between non-atopic and atopic phenotypes in each cohort is contrary to the

significant differences observed in TJ protein expression within the cohorts. This

suggests and establishes the premise for the utilisation of a physiologically

representative culture model in the assessment of transepithelial permeability.

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CHAPTER 7: Effects of human rhinovirus on epithelial barrier

integrity and function in well-differentiated air-liquid interface

cultures

7.1 Introduction

In healthy individuals, the respiratory epithelium is observed to be a pseudostratified

columnar layer consisting of ciliated, goblet and basal cells. These cells are involved in

a continual process of epithelial repair in response to cell senescence and injury,

characterised by cell proliferation and differentiation to maintain a homeostatic

epithelial layer (Bishop 2004; Rawlins and Hogan 2006; Crystal et al. 2008). In airway

diseases such as asthma, there are data that indicate that the respiratory epithelium has

an important role in the progression and pathogenesis of disease (Knight 2003; Holgate

2011). Moreover, studies have also shown that in asthma, the disease process often

originates in early life (Van Den Toorn et al. 2001; Evans et al. 2002; Stevens 2009).

Hence, the rationale for examining the respiratory epithelium in early life in order to

better comprehend the origins of this chronic disease.

Cell culture models utilising immortalised cell lines or human derived primary AEC are

often the preferred method for studying the complex and varied functions of the

respiratory epithelium (Grainger et al. 2006) . Moreover, the ability of human derived

primary AEC to differentiate in vitro whilst retaining their in vivo characteristics

provides a physiologically more representative culture model to assess cellular

characteristics, mechanisms and potential therapeutic interventions (Grainger et al.

2006; Lin et al. 2007; Kesimer et al. 2009; Pezzulo et al. 2011). Human derived

differentiated primary AEC cultures have been utilised in several studies to investigate

the differences in the respiratory epithelium between healthy and asthmatic individuals,

however, most of these studies were performed utilising adult-derived primary AECs

(Hackett et al. 2011; Xiao et al. 2011). Despite a previous study utilising paediatric

derived differentiated primary AECs to illustrate differences between healthy and

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asthmatic paediatric individuals (Parker 2010), there is a paucity of data on respiratory

epithelial barrier integrity and function in children and, in particular, children with

asthma. Furthermore, at present, there have been limited attempts to investigate the

airway epithelial integrity and function in paediatric asthmatic population following

exposure to viral exacerbations and the effects on epithelial function.

Hence, this study sought to test the hypothesis that HRV disrupts the epithelial barrier

integrity through disassembly of TJ proteins and that infection with HRV would result

in elevated transepithelial permeability in asthmatic cohorts. The initial aim of this

investigation was to generate reliable, robust air-liquid interface cultures (ALI) of

ciliated, pseudostratified pAECs from non-asthmatic and asthmatic children.

Subsequently, this investigation then compares and contrasts the epithelial barrier

integrity at basal level and following exposure to human rhinovirus between non-

asthmatic and asthmatic children. Utilising cell cultures obtained from paediatric

cohorts, this study was able to generate well-differentiated ALI cultures. Together with

TEER measurements and a permeability assay adapted for pAECs, this study was able

to functionally assess transepithelial permeability of inert macromolecules of two

different molecular weights following HRV infection between non-asthmatic and

asthmatic cohorts in the presence or absence of atopy.



detail in Chapter 2.



section of the investigation, samples were obtained from 6 healthy non-atopic (HNA; 5

female, 1 male), 4 healthy atopic (HA; 1 female, 3 male), 2 non-atopic asthmatic (NAA;

0 female, 2 male) and 4 atopic asthmatic (AA; 3 female, 1 male) children who did not

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previously receive any corticosteroid therapy. Patient demographics are summarised in

Table 7.1.

7.2.2 Cell culture



7.2.3 Establishment of air-liquid interface (ALI) cultures

Primary AECs were initially grown on 6.5-mm Transwell-Clear inserts 0.4 µm pore size

pre-coated with human placental collagen type IV, which has been previously

demonstrated to support AEC growth (Tillie-Leblond et al. 2007). Cells were grown

under submerged conditions in Bronchial-Air Liquid Interface (B-ALI™, Lonza) growth

media until confluent. To differentiate into ciliated pseudostratified AECs, media was

removed from the apical side. This was considered Day 0 of ALI culture and the start of

the experimental period. Cells were then grown in B-ALI™ differentiation media, added

to the basolateral side every alternate day, and the apical side washed with tissue-culture

sterile 1X PBS (refer to 2.4.1.10) weekly. Cultures were grown for 28 days at ALI to

ensure maximal differentiation as assessed by the presence of beating cilia as well as

mucus production, as evident by mucus build-up on the apical side of the cultures.

7.2.4 Immunohistochemistry for visualisation of ciliated and goblet cells

Selected non-asthmatic and asthmatic cultures grown at ALI were washed with tissue-

culture sterile 1X PBS, fixed in Carnoy’s fixative solution (refer to 2.4.1.19) for 24 h,

dehydrated in 100% ethanol and paraffin embedded. Five-micrometre sections were

obtained according to standard procedure and subsequently stained with haematoxylin

and eosin as well as periodic acid-Schiff (PAS).






HNA F 3.1 1.5 – 4.2 5 6

M 2.2 2.2 1

HA F 3.4 3.4 1 4

M 4.3 3.1 – 5.5 3

NAA F – – – 2

M 7 2.4 – 11.5 2

AA F 4 3.4 – 4.5 3 4

M 16.7 16.7 1

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7.2.5 Immunofluorescence and confocal microscopy

Cells were fixed using ice cold methanol and acetone (1:1 dilution) for occludin or ZO-

1 stains. Following fixation, cells were washed in 1X TBS (3 x 5 min / wash) (refer to

2.4.1.15). Samples were blocked for non-specific binding in diluent containing 10%

normal goat serum (v/v final) (Gibco, USA), 10% FCS (v/v final) and 1% BSA (w/v

final) (refer to 2.4.2.1.2) in 1X TBS for 30 min at room temperature. Cells were

incubated with primary antibodies to occludin and ZO-1 for 1 h in diluent, followed by

washing with 1X TBS, (6 x 10 min / wash). Secondary antibodies in diluent were then

added to the cells for 1 h in the dark at room temperature. Cells were washed with 1X

TBS for 1 h and counterstained with Hoechst 33342 (Sigma, USA) for 5 min. All

samples were mounted with fluorescent mounting medium containing DABCO (Sigma,

USA). Control samples were included where the primary antibody was omitted to

eliminate any non-specific secondary antibody binding from analysis. Due to limited

availability of samples as well as the time limitation of this pilot study, staining of

membrane claudin-1 was not performed.

Samples were imaged using a Nikon A1 inverted confocal microscope (Nikon, Japan),

with a Nikon Plan Apo VC 60x NA 1.4 oil immersion objective (Nikon, Japan) and

NIS-AR Elements software (v4.2.22, Nikon, Japan), with 6 random fields acquired per

sample. Individual channels were captured sequentially, where a 405 nm laser was used

for Hoechst 33342 with collection through a 450/50 bandpass filter, whilst AF488 and

AF568 were imaged with 488 nm and 561 nm lasers, and 525/50 and 585/50 bandpass

filters respectively. Z-stack images with step size of 0.5 µm were collected with a

pinhole of 35.8 µm (1.2 A.U. for 488 nm laser), where the top and bottom of the stacks

were determined visually due to the uneven nature of the membrane.

7.2.6 Stereological analysis and quantification of tight junction expression

Quantification of TJ expression was performed on Maximum Intensity Projections

(ImageJ, NIH, USA) using standard stereology techniques (Schmitz et al. 1999; Matter

and Balda 2003; Aijaz et al. 2007). Briefly, grid overlays of 1000 µm2 (Grid Plugin,

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ImageJ, NIH, USA) were applied to images obtained from 6 random fields of each

sample in an unbiased manner. Where the TJs form a complete junction with a

contiguous membrane on both sides of the grid line, they were counted and analysed as

previously described (Rønn et al. 2000). However, where the connections between cells

are interrupted, the junction was excluded from the analyses. The number of junctions

were then tabulated and expressed as an average of the sample.


The source, propagation and viral titre determination of the human rhinovirus minor

serotype 1B (HRV-1B) utilised in this investigation has been described in detail in

Chapter 2 (refer to 2.5.7, 2.5.7.1 and 2.5.7.2).


Paediatric-derived pAECs cultured on inserts, were infected with HRV-1B and

incubated for 24 h. Briefly, 50 µl of HRV-1B at a 50% Tissue Culture Infectivity Dose

(TCID50) of 10 x 104 TCID50/ml was added apically to the culture inserts for 5 h, after

which HRV-1B was removed and the culture inserts allowed to incubate for an

additional 19 h at 33°C in an atmosphere of 5% CO2 / 95% air. Infection with this viral

titre has been previously demonstrated to elicit cellular responses such as inflammation

but no significant increase in cell apoptosis (Stevens 2009). After each incubation

period, basolateral media was collected and stored at -80°C and infected cells were

utilised for various downstream assays.

7.2.9 Transepithelial electrical resistance measurement

Transepithelial electrical resistance (TEER) measurement were performed on days 7,

14, 21 and 28 to ensure the formation and integrity of epithelial TJ between cells using

an epithelial voltohmeter with silver chloride ‘chopstick’ electrodes. Prior to

measurement, the apical layer of the ALI cultures were washed with tissue-culture

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sterile 1X PBS (refer to 2.4.1.10) and HEPES buffered Hank’s Balanced Salt Solution

(HEPES-HBSS) (refer to 2.4.3.3.2) added to both apical and basolateral sides to

equilibrate the cultures. Triplicate measurements per well were made and the mean

resistance calculated. The resistance obtained from a cell-free culture insert was

subtracted from the resistance measured across each cell monolayer and corrected for

the surface area of the culture insert to yield the TEER of the epithelial cells with values

expressed in Ω/cm2.



was utilised for the functional determination of epithelial TJ membrane integrity prior

and following human rhinovirus infection.


Where applicable, data were tested for population normality and homogeneity of

variance, and a Student t test was performed. Experiments were performed in at least in

duplicates and using a minimum of three patients of each cohort per experiment.

Statistical analyses were performed, where applicable, using Mann-Whitney non-

parametric test and values presented are mean ± SD and P values less than 0.05 were

considered to be significant.

7.3 Results

7.3.1 Establishment of air-liquid interface cultures

When visualised under a light microscope at a low magnification (10X), the confluent

monolayer of cells demonstrated a typical ‘cobblestone’ appearance that was

characteristic of epithelial cells in culture. The confluent cell monolayer was then ‘air-

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lifted’ according to the supplied protocol. At ~20 days post ‘air-lifting’, mucus was

visible on the apical layer and this was evident until the termination of the experiment.

When visualised under high magnification (40X), ciliary movement was also observed

at ~ 20 days post ‘air-lift’ until experiment was performed, at ~ 31 days post ‘air-lift’.

7.3.2 Physical properties of pAEC derived ALI cell cultures of non-asthmatic and

asthmatic cohorts

When viewed under high magnification (40X Oil), the presence of a number of layers of

cells was observed, including ciliated cells on the apical surface following haematoxylin

and eosin (H&E) stains. Furthermore, observations showed well-differentiated, multi-

layered ALI cultures of 3 – 4 cell layers in vitro, with ciliated cells on the apical surface

of the epithelial cell layer (Figure 7.1 A and B respectively). To further assess

differentiation and confirm the cellular identity of pAEC ALI culture, periodic acid-

Schiff (PAS) staining which was performed, revealed the presence of goblet cells

associated with mucus secretion at the apical surface (Figure 7.1 C and D respectively).

The observed presence of a mucus-producing, multi-layered epithelium containing

ciliated and goblet cells strongly indicated the complete differentiation of pAECs into a

well-differentiated and functional epithelium.

When pAEC ALI cultures of both non-asthmatic and asthmatic cohort were further

classified according to atopy, well-differentiated, multi-layered ALI cultures of 2 – 4

cell layers, with ciliated cells observed on the apical surface of the epithelial cell layer

following H&E stains (Figure 7.2 A - D) was observed within each phenotype while

PAS staining revealed the presence of a mucus layer and goblet cells in all phenotypes

(Figure 7.2 E - H).

Figure 7.1 Generation of ALI cultures from pAECs of non-asthmatic and

asthmatic cohorts: Air-liquid interface cultures of airway epithelial cells derived from

non-asthmatic (A and C) and asthmatic (B and D) paediatric donors were fixed,

embedded and sectioned for subsequent analysis of morphology and cilia formation

(black arrows) by haematoxylin-and-eosin (H&E) and mucin production (red arrows) by

periodic-acid-schiff (PAS) stained sections respectively. Images are representative of

n=3 (Total magnification 40X Oil).

A C

D B

H&E PAS

Figure 7.2 Generation of ALI cultures from pAECs of non-asthmatic and

asthmatic cohorts with each cohort further categorised based on atopy: Air-liquid

interface cultures of airway epithelial cells derived from healthy non-atopic (A and E);

healthy atopic (B and F); non-atopic asthmatic (C and G) and atopic asthmatic (D and

H) paediatric cohorts were fixed, embedded and sectioned for subsequent analysis of

morphology and cilia formation (black arrows) by haematoxylin-and-eosin (H&E) and

mucin production (red arrows) by periodic-acid-schiff (PAS) stained sections

respectively. Images are representative of n=3 (Total magnification 40X Oil).

A

B

C

D

E

F

G

H

H&E PAS

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expression in pAEC derived well differentiated air-liquid interface (ALI) cultures

Having demonstrated differences in in vitro TJ protein expression on monolayer

cultures following HRV infection, protein expression of the same TJs were assessed in

well-differentiated pAEC ALI cultures of non-asthmatic and asthmatic cohorts

following infection. Due to the limited availability of samples in this pilot study, the

analysis of membrane claudin-1 expression was not performed. Membrane occludin and

ZO-1 TJ protein expression were determined via confocal microscopy of well-

differentiated ALI cultures on semi-permeable insert membranes.


Under confocal microscopy, the presence of a continuous, uninterrupted membrane

protein expression of both occludin and ZO-1 TJs proteins were observed in the apical

regions of the in vitro differentiated ALI cultures of both non-asthmatic (Figure 7.3 A -

D) and asthmatic (Figure 7.4 A – D) cohorts prior viral infection. A less continuous and

more punctate membrane protein expression of both TJs was observed in the apical

regions of both non-asthmatic (Figure 7.3 E – H) and asthmatic (Figure 7.4 E - H)

cohorts following viral infection. A more disrupted membrane protein expression in the

apical region was observed in the asthmatic ALI (Figure 7.4 H) cultures compared to

the non-asthmatic counterpart (Figure 7.3 H).

When assessing membrane occludin expression between non-asthmatic and asthmatic

cohorts, results obtained demonstrated significantly lower basal membrane occludin

within the asthmatic cohort compared to the non-asthmatic counterpart (Figure 7.5 A –

Non-infected). Following 24 h infection with viral titre of 10 x 104 TCID50/ml of HRV-

1B, results demonstrated minimal change in membrane occludin expression (276.6AU ±

17.9) compared to their non-infected controls (278.6AU ± 11.7) in the non-asthmatic

cohort. When membrane occludin expression from the asthmatic ALI cultures were

assessed following infection, the data demonstrated significantly diminished membrane

occludin expression (37.3AU ± 5.9) in the asthmatic ALI cultures in comparison to non-

Figure 7.3 Membrane TJ protein expression in ALI cultures generated from

pAECs of non-asthmatic cohorts following viral infection: Air-liquid interface

cultures of airway epithelial cells derived from non-asthmatic paediatric donors were

fixed and incubated with primary antibodies to zonula occluden-1 (ZO-1) (green; A and

E) and occludin (red; B and F) for 1 h at room temperature followed by secondary

antibodies for 1 h in the dark at room temperature. Cultures were then counterstained

with Hoechst 33342, which illuminates cellular nuclear material (blue). (C and G)

Merged image of (A) and (B), (E) and (F) respectively demonstrates the areas of tight

junction expression. (D) Confocal imaging of occludin and ZO-1 in representative ALI

samples of pAECs from non-infected non-asthmatic cohort demonstrates continuous

uninterrupted membrane protein expression of both TJ proteins at the apical region of

the differentiated culture (white arrows). (H) Confocal imaging of occludin and ZO-1 in

representative ALI samples of pAECs from infected non-asthmatic cohort demonstrates

loss in expression of membrane protein expression of both TJ proteins at the apical

region of the differentiated culture (red arrows). Images are representative of n=3 (Total

magnification 60X Oil).

A

B

C

D

E

F

G

H


pAECs of asthmatic cohorts following viral infection: Air-liquid interface cultures of

airway epithelial cells derived from asthmatic paediatric donors were fixed and

incubated with primary antibodies to zonula occluden-1 (ZO-1) (green; A and E) and

occludin (red; B and F) for 1 h at room temperature followed by secondary antibodies

for 1 h in the dark at room temperature. Cultures were then counterstained with Hoechst

33342, which illuminates cellular nuclear material (blue). (C and G) Merged image of

(A) and (B), (E) and (F) respectively demonstrates the areas of tight junction

expression. (D) Confocal imaging of occludin and ZO-1 in representative ALI samples

of pAECs from non-infected asthmatic cohort demonstrates continuous uninterrupted

membrane protein expression of both TJ proteins at the apical region of the

differentiated culture (white arrows). (H) Confocal imaging of occludin and ZO-1 in

representative ALI samples of pAECs from infected asthmatic cohort demonstrates loss

in expression of membrane protein expression of both TJ proteins at the apical region of

the differentiated culture (red arrows). Images are representative of n=3 (Total


A

B

C

D

E

F

G

H

50 µm

Figure 7.5 Expression of membrane TJ protein in ALI cultures from pAECs of non-asthmatic and asthmatic cohorts following viral infection:

pAECs seeded on culture inserts and grown to confluence were treated as previously mentioned (refer to 7.2.3). Cultures were infected with HRV-1B

at 10 x 104 TCID50/ml for 24 h and membrane protein expression assessed via confocal microscopy and standard stereology techniques (refer to 7.2.5).

(A) Minimal decrease in membrane occludin expression following infection with HRV-1B was observed in ALI cultures of the non-asthmatic cohort.

Significant decrease in membrane occludin expression was observed in ALI cultures of the asthmatic cohort following infection. Significant difference

in membrane protein expression was also observed between ALI cultures of asthmatic and non-asthmatic cohorts. (B) A significant decrease in

membrane ZO-1 expression was observed in ALI cultures of non-asthmatic cohort following HRV-1B infection. Similarly, a significant decrease in

membrane ZO-1 expression was also observed in ALI cultures of asthmatic cohort. Difference in membrane ZO-1 expression between ALI cultures of

non-asthmatic and asthmatic cohorts was observed to be of significance. Data were presented as mean ± SD relative to control. *Statistical significance

relative to non-infected controls (p<0.05) # Statistical significance relative to non-asthmatic cohort (p<0.05).


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infected controls (165.7AU ± 10.8). Interestingly, following 24 h infection, membrane

occludin expression was observed to be significantly lower (37.3AU ± 5.9) in the

asthmatic cultures in comparison to the non-asthmatic cultures (276.5AU ± 17.9)

(Figure 7.5 A; p<0.05)

Similarly, when ZO-1 expression was compared between non-asthmatic and asthmatic

cohorts, results obtained demonstrated significantly lower basal ZO-1 expression in

asthmatic ALI pAECs (325.2AU ± 5.2) compared to non-asthmatic ALI pAECs (338.4

± 6.7) (Figure 7.5 B, Non-infected; p<0.05). After 24 h infection, results showed a

significantly lower membrane ZO-1 expression (270.6AU ± 8.5) in contrast to non-

infected, non-asthmatic controls (338.4AU ± 6.7) (Figure 7.5 B; p<0.05). When

membrane ZO-1 expression of the asthmatic cultures were analysed, results

demonstrated significantly lower ZO-1 expression (239.1AU ± 13.9) in comparison to

non-infected controls (325.2AU ± 5.2). In addition, following 24 h infection, membrane

ZO-1 expression was observed to be significantly lower (239.1AU ± 13.9) in the

asthmatic cohort in comparison with the non-asthmatic cohort (270.6AU ± 8.5) (Figure

7.5 B; p<0.05).


atopic status


atopy, the presence of a continuous, uninterrupted membrane protein expression of both

TJs was observed in the apical regions of the in vitro differentiated ALI cultures of non-

asthmatic non-atopic (Figure 7.6 A - D), non-asthmatic atopic (Figure 7.7 A - D) and

asthmatic atopic (Figure 7.8 A - D) cohorts prior viral infection. In contrast, a less

continuous and more punctate membrane protein expression of both TJs was observed

in the apical regions of all three phenotypic cohorts following viral infection (Figure 7.6

– 7.8 E - H respectively). Due to limited availability of pAECNAA and the time

constraints of this preliminary investigation, the pAECNAA cohort was excluded from all

statistical and comparative analysis.


pAECHNA cohorts following viral infection: Air-liquid interface cultures of airway

epithelial cells derived from non-asthmatic, non-atopic paediatric donors were fixed and







of pAECs from non-infected non-asthmatic cohort demonstrates continuous







A

B

C

D

E

F

G

H


pAECHA cohorts following viral infection: Air-liquid interface cultures of airway

epithelial cells derived from atopic, non-asthmatic paediatric donors were fixed and







of pAECs from non-infected non-asthmatic cohort demonstrates continuous







A

B

C

D

E

F

G

H

50 µm 50 µm


pAECAA cohorts following viral infection: Air-liquid interface cultures of airway

epithelial cells derived from atopic, asthmatic paediatric donors were fixed and







of pAECs from non-infected asthmatic cohort demonstrates continuous uninterrupted

membrane protein expression of both TJ proteins at the apical region of the

differentiated culture (white arrows). (H) Confocal imaging of occludin and ZO-1 in

representative ALI samples of pAECs from infected asthmatic cohort demonstrates loss

in expression of membrane protein expression of both TJ proteins at the apical region of

the differentiated culture (red arrows). Images are representative of n=3 (Total


A

B

C

D

E

F

G

H

50 µm

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In the assessment of membrane protein expression, data demonstrated higher basal

expression of membrane occludin in pAECHA (300.1AU ± 15.4) when compared to

pAECHNA (257.3AU ± 15.1). In contrast, basal membrane occludin expression was

observed to be significantly lower in pAECAA (165.7AU ± 10.8) when compared to both

pAECHNA and pAECHA ALI cultures (Figure 7.9 A, Non-infected; p<0.05). In addition,

following 24 h infection with a viral titre of 10x104 TCID50/ml of HRV-1B, the

obtained data showed a significant decrease in membrane occludin expression within

the pAECHNA and pAECAA cohorts (201.8AU ± 17.5 and 37.3AU ± 5.9 respectively)

while expression of membrane occludin in pAECHA cohort was significantly elevated

(351.1AU ± 5.1) (Figure 7.9 A, Infected; p<0.05).

When assessing the effects of the viral infection on membrane occludin protein

expression at 24 h post infection, membrane expression of occludin was significantly

higher in pAECHA (351.1AU ± 5.1) in contrast to pAECHNA (201.8AU ± 17.5).

Expression of membrane occludin was also significantly lower in pAECAA (37.3AU ±

5.9) in contrast to pAECHNA (201.8AU ± 17.5) and pAECHA (351.1AU ± 5.1).

Similarly, when membrane ZO-1 protein expression was assessed, data generated

demonstrated significantly lower basal expression of membrane ZO-1 in pAECHA

(343.8AU ± 6.3) when compared to pAECHNA (364.4AU ± 6.3) (Figure 7.9 B, Non-

infected; p<0.05). Interestingly, basal expression of membrane ZO-1 was observed to

be higher in the pAECAA cohort compared to pAECHNA. However, this was found to be

not significant (Figure 7.9 B, Non-infected). In addition, following 24 h infection with a

viral titre of 10x104 TCID50/ml of HRV-1B, the obtained data showed a significant

decrease in membrane ZO-1 expression within the pAECHNA (228.1AU ± 9.6), pAECHA

(303.6AU ± 13.1) and pAECAA (204.2AU ± 10.7) when compared to non-infected

controls (Figure 7.9 B, Infected; p<0.05).

When assessing the effects of the viral infection on membrane ZO-1 protein expression

at 24 h post infection, membrane expression of ZO-1 was lower pAECAA (204.2AU ±

10.7) in contrast to pAECHNA (228.1AU ± 9.6). Membrane expression of ZO-1 was

significantly higher in pAECHA (303.6AU ± 13.1) when compared to pAECHNA (Figure

Figure 7.9 Expression of membrane TJ protein in ALI cultures from pAECs of non-asthmatic and asthmatic cohorts with each cohort further

categorised based on atopy: pAECs seeded on culture inserts and grown to confluence were treated as previously mentioned (refer to 7.2.3). Cultures

were infected with HRV-1B at 10 x 104 TCID50/ml for 24 h and membrane protein expression assessed via confocal microscopy and standard

stereology techniques (refer to 7.2.5). (A) Following infection with HRV-1B at viral titre of 10 x 104 TCID50/ml, a significant decrease in membrane

occludin expression was observed in pAECHNA and pAECAA ALI cultures compared to non-infected controls. However, a significant increase in

membrane occludin expression was observed in pAECHA ALI cultures in contrast to non-infected controls. (B) A significant decrease in membrane

ZO-1 expression was observed in pAECHNA, pAECHA and pAECAA ALI cultures following infection compared to non-infected controls. Data were

presented as mean ± SD; n = 4 individual experiments each performed in duplicates. *Statistical significance relative to non-infected control (p<0.05).


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7.9 B, Infected; p<0.05). Expression of membrane ZO-1 was also significantly lower in

pAECAA (204.2AU ± 10.7) in comparison with pAECHA (303.6AU ± 13.1) (Figure 7.9

B, Infected; p<0.05).

7.3.4 Effect of human rhinovirus infection on transepithelial electrical resistance

(TEER) and permeability in pAEC derived well differentiated air-liquid interface

(ALI) cultures

Having demonstrated a change in TJ expression, transepithelial permeability towards

the same inert macromolecule sizes of 4 and 20 kDa was then assessed across the

differentiated ALI cultures following HRV infection.


Results generated demonstrated higher transepithelial resistance (RT) of non-infected

controls (851.4 Ω/cm2 ± 156.6) and following HRV infection, a significant decrease in

RT (402.8 Ω/cm2 ± 54.6) was observed (Figure 7.10 A - TEER; p<0.05). Consistent with

the decrease in RT, infected non-asthmatic ALI cultures demonstrated an increase in

levels of transepithelial permeability to FITC-dextran 4 (22.4 x 10-4 cm/sec ± 8.4) and

20 (3.1 x 10-4 cm/sec ± 0.5) kDa following 24 h infection in contrast with non-infected

controls (9.2 x 10-4 cm/sec ± 2.4 and 2.6 x 10-4 cm/sec ± 0.4 respectively) (Figure 7.10

A). This was observed to be significant for FITC-dextran 4 kDa (Figure 7.10 A – 4 kDa;

p<0.05). Moreover, when assessing epithelial permeability to the different sized inert

macromolecule within the non-infected non-asthmatic cultures, significantly higher

level of transepithelial permeability towards FITC-dextran of 4 kDa (9.2 x 10-4 cm/sec ±

2.4) was observed in comparison to FITC-dextran 20 kDa (2.6 x 10-4 cm/sec ± 0.4)

(Figure 7.10 A – Non-infected; p<0.05). Similar observations of significantly higher

level of transepithelial permeability to FITC-dextran of 4 kDa (22.4 x 10-4 cm/sec ± 8.4)

was observed in comparison to FITC-dextran 20 kDa (3.1 x 10-4 cm/sec ± 0.5) in the

infected non-asthmatic cultures (Figure 7.10 A - Infected; p<0.05).

Figure 7.10 Transepithelial electrical resistance (RT) and permeability in ALI cultures from pAECs of non-asthmatic and asthmatic cohorts:

pAECs from non-asthmatic and asthmatic cohorts, seeded onto Corning transwell inserts and grown to confluence were cultured at ALI conditions

until differentiation and mucus generation occurred. Following 24 h infection with a viral titre of 10 x 104 TCID50/ml of HRV-1B, an optimised

transepithelial permeability assay as previously described was performed to determine epithelial permeability to FITC-dextran 4 kDa (blue) and 20

kDa (grey) (refer to 3.3.4). (A) Corroborating the reduced RT, an increase in transepithelial permeability towards both FITC-dextran 4 and 20 kDa was

observed in pAEC ALI cultures of non-asthmatic cohorts following infection . Epithelial permeability towards FITC-dextran 4 kDa was significantly

higher when compared to FITC-dextran 20 kDa in non-infected and infected pAEC ALI cultures of non-asthmatic cohort. (B) Significant reduction in

RT was concomitant with significant difference in transepithelial permeability towards FITC-dextran 4 kDa but not 20 kDa in pAEC ALI cultures of

asthmatic cohorts following infection. However, permeability towards FITC-dextran 4 kDa was significantly higher when compared to FITC-dextran

20 kDa in non-infected and infected pAEC ALI cultures of asthmatic cohort. Results are presented as mean ± SD; n = 8 individual experiments each

performed in duplicates with the exception of asthmatic cohort (n=6). *Statistical significance relative to non-infected control (p < 0.05). # Statistical

significance relative to FITC-dextran 20 kDa (p<0.05).


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Similarly, in pAECs of the asthmatic cohort prior to infection with HRV, data

demonstrated higher RT in asthmatic pAEC ALI cultures (611.5 Ω/cm2 ± 350.7) and

following infection, a significant decrease (183.7 Ω/cm2 ± 112.8) was observed (Figure

7.10 B - TEER; p<0.05). In addition, consistent with the decrease in RT, a significant

increase in transepithelial permeability of FITC-dextran 4 kDa (31.4 x 10-4 cm/sec ±

5.5) (Figure 7.10 B; p<0.05) was observed in infected cultures in contrast to non-

infected controls, however, permeability to FITC-dextran 20 kDa, although higher, was

found to be non-significant. Nonetheless, when assessing epithelial permeability to the

different sized inert macromolecule in the non-infected asthmatic cultures, significantly

higher level of transepithelial permeability towards FITC-dextran of 4 kDa (15.4 x 10-4

cm/sec ± 3.2) was observed in comparison to FITC-dextran 20 kDa (6.1 x 10-4 cm/sec ±

1.8) (Figure 7.10 B – Non-infected; p<0.05). A significantly higher level of

transepithelial permeability to FITC-dextran of 4 kDa (31.4 x 10-4 cm/sec ± 5.5) was

observed in comparison to epithelial permeability towards FITC-dextran 20 kDa (6.2 x

10-4 cm/sec ± 2.7) in the infected asthmatic cultures (Figure 7.10 B - Infected; p<0.05).


atopic status


atopy, results generated demonstrated a higher RT in the pAECHNA ALI cohort (994.4

Ω/cm2 ± 1.9) and post HRV-1B infection, RT in pAECHNA ALI cohort was significantly

lower (452.7 Ω/cm2 ± 2.8) (Figure 7.11 A – TEER; p<0.05). Consistent with the

decrease in RT, infected pAECHNA ALI cultures demonstrated an increase in levels of

transepithelial permeability of FITC-dextran 4 (16.9 x 10-4 cm/sec ± 1.3) and 20 (3.1 x

10-4 cm/sec ± 0.8) kDa following infection compared to non-infected controls (7.2 x 10-

4 cm/sec ± 1.8 and 1.8 x 10-4 cm/sec ± 0.4 respectively). This was observed to be

significant for FITC-dextran 4 kDa (Figure 7.11 A – 4 kDa; p<0.05). In addition,

transepithelial permeability to FITC-dextran of 4 kDa (16.9 x 10-4 cm/sec ± 1.3) was

significantly higher in comparison to FITC-dextran 20 kDa (3.1 x 10-4 cm/sec ± 0.8)

(Figure 7.11 A – Infected; p<0.05).

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In comparison to non-infected controls (708.4 Ω/cm2 ± 1.9), a significant decrease in RT

was demonstrated following HRV-1B infection in the pAECHA ALI cohort (352.9

Ω/cm2 ± 2.5) (Figure 7.11 B – TEER; p<0.05). In conjunction with the decrease in RT,

infected pAECHA ALI cultures similarly demonstrated increase in levels of

transepithelial permeability of FITC-dextran 4 (49.3 x 10-4 cm/sec ± 18.8) and 20 (3 x

10-4 cm/sec ± 0.3) kDa following infection with compared to non-infected controls (5.6

x 10-4 cm/sec ± 1.4 and 2.1 x 10-4 cm/sec ± 0.2 respectively). This was observed to be

non-significant for FITC-dextran 4 kDa (Figure 7.11 A – 4 kDa; p = 0.055).

Furthermore, when assessing epithelial permeability to the different sized inert

macromolecule following infection, transepithelial permeability to FITC-dextran of 4

kDa (49.3 x 10-4 cm/sec ± 18.8) was significantly higher in comparison to epithelial

permeability to FITC-dextran 20 kDa (3 x 10-4 cm/sec ± 0.3) (Figure 7.11 B – Infected;

p<0.05).

In contrast to non-infected controls (291.4 Ω/cm2 ± 1.3), a significant decrease in RT

was similarly observed following HRV-1B infection in pAECNAA ALI cohort (81.1

Ω/cm2 ± 2) (Figure 7.11 C – TEER; p<0.05). Together with the reduction in RT,

infected pAECNAA ALI cultures demonstrated an increase in levels of transepithelial

permeability to FITC-dextran 4 (27.7 x 10-4 cm/sec ± 13.6) and 20 (4.2 x 10-4 cm/sec ±

2.6) kDa following infection which was in contrast to non-infected controls (15.7 x 10-4

cm/sec ± 3.5 and 4 x 10-4 cm/sec ± 1.2 respectively) (Figure 7.11 C). Moreover,

epithelial permeability to the different sized inert macromolecules following infection

was observed to be higher for FITC-dextran of 4 kDa (27.7 x 10-4 cm/sec ± 13.6) in

comparison to FITC-dextran 20 kDa (4.2 x 10-4 cm/sec ± 2.6) (Figure 7.11 C -

Infected). However, due to the limited availability of pAECNAA ALI, statistical analysis

could not be performed to determine the level of significance.

When compared to non-infected controls (931.7 Ω/cm2 ± 1.9), significant decrease in R-

T was observed in pAECAA ALI cohort (286.5 Ω/cm2 ± 13.2) following HRV infection

(Figure 7.11 D – TEER; p<0.05). Concurrent with the decrease in RT, infected pAECAA

ALI cultures also showed significant increase in level of transepithelial permeability of

FITC-dextran 4 (33.9 x 10-4 cm/sec ± 7.1) but not FITC-dextran 20 kDa (6.8 x 10-4

Figure 7.11 Transepithelial electrical resistance (RT) and permeability in ALI cultures from pAECs of non-asthmatic and asthmatic cohorts

with each cohort further categorised based on atopy: pAECs from non-asthmatic and asthmatic cohorts, seeded onto Corning transwell inserts and

grown to confluence were cultured at ALI conditions until differentiation and mucus generation occurred. Following 24 h infection with a viral titre of

10 x 104 TCID50/ml of HRV-1B, an optimised transepithelial permeability assay as previously described was performed to determine epithelial

permeability to FITC-dextran 4 kDa (blue) and 20 kDa (grey) (refer to 3.3.4). (A) Concomitant with a significantly reduced RT, significant increase in

transepithelial permeability was observed for FITC-dextran 4 following infection in the pAECHNA cohort. Epithelial permeability to FITC-dextran 4

kDa was also significantly higher compared to FITC-dextran 20 kDa in both non-infected and infected pAECHNA. (B) Associated with a significant

decreased RT, an increase in transepithelial permeability was observed for FITC-dextran 4 following infection in the pAECHA cohort, however, this was

observed to be just not significant. Epithelial permeability to FITC-dextran 4 kDa was also significantly higher compared to FITC-dextran 20 kDa in

both non-infected and infected pAECHA. (C) Concomitant with a significantly reduced RT, an increase in transepithelial permeability was observed for

FITC-dextran 4 following infection in the pAECNAA cohort, however, due to low sample size of pAECNAA, statistical analysis could not be performed

to determine significance and was excluded from comparative statistical analysis. (D) Associated with a significant decreased RT, an increase in

transepithelial permeability was observed for FITC-dextran 4 following infection in the pAECAA cohort, increase in transepithelial permeability was

observed for both FITC-dextran 4 and 20 kDa following infection in the pAECAA cohort. Epithelial permeability to FITC-dextran 4 kDa was also

significantly higher compared to FITC-dextran 20 kDa in only the infected pAECAA. Results are presented as mean ± SD; n = 4 individual experiments

each performed in duplicates with the exception of pAECNAA phenotype (n=2). *Statistical significance relative to non-infected control (p<0.05). #Statistical significance relative to FITC-dextran 20 kDa (p<0.05).


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cm/sec ± 3.8) following infection with HRV-1B in comparison to non-infected controls

(11.7 x 10-4 cm/sec ± 7.2 and 5.1 x 10-4 cm/sec ± 2.1 respectively). Furthermore,

epithelial permeability to the different sized inert macromolecule was not significantly

different in non-infected pAECAA cultures (Figure 7.11 D – Non-infected). However,

following infection, significantly higher epithelial permeability to FITC-dextran of 4

kDa (33.9 x 10-4 cm/sec ± 7.1) was observed in comparison to FITC-dextran 20 kDa

(6.8 x 10-4 cm/sec ± 3.8) (Figure 7.11 D – Infected; p<0.05).

When RT between the phenotypic cohorts were assessed, pAECHA, pAECNAA and

pAECAA ALI cultures demonstrated significantly lower basal and post infection RT

values compared to pAECHNA. Furthermore, when assessing the magnitude of decrease

in RT following HRV-1B infection, pAECNAA and pAECAA ALI cohorts demonstrated a

greater magnitude of decrease (3.6-fold and 3.3-fold respectively) when compared to

pAECHNA and pAECHA (2.2-fold and 2-fold respectively). Similar observations were

shown when data obtained from the ALI cultures were further analysed based on atopy.

Interestingly, when assessing epithelial permeability to FITC-dextran 4 and 20 kDa

following infection, permeability to FITC-dextran 4 kDa was just significantly higher in

pAECHA (49.3 x 10-4 cm/sec ± 18.8) ALI cohort compared to pAECHNA (16.9 x 10-4

cm/sec ± 1.3), while there was no significant difference in pAECNAA and pAECAA

cultures when compared to pAECHNA ALI cultures. Further comparative analysis

between pAECHA, pAECNAA and pAECAA did not demonstrate any significant

difference in epithelial permeability towards FITC-dextran 4 kDa following HRV-1B

infection.

7.4 Discussion

Air-liquid interface (ALI) cultures are an immensely powerful tool for the investigation

of the human respiratory epithelium in vitro and are used to model airway epithelial

differentiation, injury and repair; to assess the function of specific genes and biological

pathways as well as for the assessment of gene transfers (Wu et al. 1986; Whitcutt et al.

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1988; Gruenert et al. 1990; De Jong et al. 1993; Pickles et al. 1998; Fulcher et al. 2005;

Zabner et al. 2005; Pierrou et al. 2007; Ross et al. 2007). Hence, this study aimed to

recapitulate the observations of the previous chapters by providing additional insights

on tight junctional complex expression through the utilisation of a well-differentiated

culture model which better mimics the in vivo tracheo-bronchial epithelium compared to

submerged monolayer cultures. This would offer a better understanding of the paediatric

asthmatic epithelium and the effects on barrier compromisation following HRV

infection.

Initial observations of ALI cultures in this study are consistent with other studies that

have demonstrated that ALI cultures consists of various cells types including ciliated,

mucus and basal cells (Parker 2010; Hackett et al. 2011). Despite the small samples

sizes tested in this preliminary in vitro study of well-differentiated non-asthmatic and

asthmatic epithelium, results from confocal microscopy stereological analysis

corroborated earlier findings demonstrating lower basal membrane TJ protein

expression within the asthmatic epithelium compared to non-asthmatic counterpart.

Basal transepithelial permeability was not significantly different between non-asthmatic

and asthmatic ALI cultures, corroborating earlier findings. However, lower basal RT

values were observed in the asthmatic ALI cultures in contrast to their non-asthmatic

counterpart, which strongly suggests that the asthmatic epithelial ALI cultures could be

less differentiated and retaining a more basal characteristic, an observation in line with

that demonstrated by Hackett and colleagues (2011) showing increased expression of

cytokeratin-5, a cytoskeleton protein expressed in basal cells. Moreover, the lack of a

significant difference in epithelial permeability could possibly be attributed to the

severity of asthma as this study utilised samples obtained from paediatric individuals

who had mild, stable asthma and were not on corticosteroid therapy for at least 3

months. Future studies involving samples obtained from paediatric individuals with

moderate to severe asthma could certainly anticipate a difference in epithelial

permeability, as previously demonstrated by Xiao and colleagues despite the study

utilising adult derived asthmatic airway epithelial cells.

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Interestingly, when the data were further categorised according to atopy, it was

observed that the pAECHA ALI cultures demonstrated significantly higher levels of

basal membrane occludin expression and no significant difference in basal membrane

ZO-1 expression compared to pAECHNA and pAECAA ALI cultures, which contrasts

earlier findings demonstrating significantly lower membrane occludin and ZO-1

expression within the pAECHA cultures. This suggests the possible role of atopy in

regulating membrane TJ protein expression, as demonstrated by De Benedetto and

colleagues (2011) in non-asthmatic individuals. Interestingly, basal epithelial

permeability was not significantly different between pAECHNA, pAECHA and pAECAA

ALI cultures, suggesting that allergic sensitisation of non-asthmatic or asthmatic airway

epithelium does not occur through trafficking of aeroallergens through the epithelial

layer into the sub-epithelial space but rather, through the interaction with dendritic cells,

as demonstrated in a study by Jahnsen and colleagues (2001).

However, as most studies which have investigated the contributing role of atopy in

affecting TJ complexes have utilised the epidermal layer (Weidinger et al. 2006; De

Benedetto et al. 2011; Kuo et al. 2012), there remains a lack of studies examining the

impact of atopy on TJ complex expression within the respiratory airways. As such, the

contrariety of the data could possibly be attributed to the type and severity of the atopic

status, wherein an increased allergic reaction to a particular allergen could result in

compensatory effects by different epithelial TJs not assessed in this study, to maintain

barrier functionality. In addition, the dichotomy in the data could also be possibly

attributed to the structure of the ALI cultures, being well-differentiated and stratified

compared to submerged monolayer cultures or the use of confocal microscopy and

stereological analysis compared to the In-Cell™ Western assay in providing TJ

membrane analysis.

When membrane occludin and ZO-1 was assessed following infection, the data showed

disassembly of membrane TJ protein interaction at the apical regions in pAECHNA and

pAECAA ALI cultures, with an exaggerated disassembly in pAECAA ALI cultures. In

contrast, membrane TJ protein was elevated in the pAECHA ALI cultures but

interestingly, a reduction in RT with an increase in epithelial permeability was observed

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in pAECHA ALI cultures following infection. The contrast in data could be attributed to

a probable dissociation of other junctional proteins that not assessed in the current

study, hence resulting in increased permeability. In pAECHNA and pAECAA ALI cultures

following infection, a disassembly of membrane TJ protein was concomitant with a

reduction in RT and a corresponding increase in epithelial permeability in both ALI

cultures with a greater increase observed in the pAECAA ALI cultures. Despite the

counter intuitiveness as well as the demonstrated impaired apoptotic response to

infection (Wark et al. 2005), infection with HRV could induce the disassembly of TJ

proteins through the initiation of apoptosis, albeit delayed, eventually leading to the

dissociation of infected cells from the neighbouring non-infected cells. Consequently,

TJ disassembly and dissociation from adjacent cells could ultimately result in increased

epithelial permeability. This increase in permeability could either facilitate the passage

of inhaled pathogenic agents from the airway lumen to the sub-epithelial space or

conversely, provide the opportunity for cells associated with the innate immune system

to migrate out from the submucosa into the airway lumen to sample aeroallergens,

bacteria, fine particulate matter or respiratory viruses resulting in allergic sensitisation,

infection and inflammation in each process. Although a restoration of TJ proteins was

observed in pAECHNA cultures in contrast to a sustained loss of TJs in pAECHA,

pAECNAA and pAECAA cultures following extended HRV infection in earlier chapters,

whether barrier functionality changes with extended periods or recurrent viral infections

in ALI cultures require further investigation.

Collectively, the results emphasises the importance of using ALI cultures in the

assessment of barrier functionality. More importantly, the results indicate that HRV

infection would lead to an increase in epithelial permeability in ALI cultures, with a

greater magnitude in increase observed in asthmatic ALI cultures. This would result in

elevated passage of pathogenic challenges or migration of defence cells, eventually

leading to increased sensitisation, infection, and further asthma exacerbations.

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7.5 Conclusion

Human AECs cultured in vitro at air-liquid interface (ALI), are capable of forming a

pseudostratified epithelium with functioning TJs, cilia and mucin production. There are

limited data regarding the effects of HRV infection on epithelial TJs and the subsequent

effects on barrier function in paediatric cohorts. Data obtained in this investigation,

which corroborated findings from earlier chapters, demonstrated diminished basal TJ

expression in asthmatic epithelial cells and an exaggerated disassembly of TJ proteins

following HRV infection concomitant with increased epithelial permeability. However,

the dichotomy in the data with regards to membrane TJ protein expression and epithelial

permeability in submerged monolayer cultures and ALI cultures is attributed to the

difference in structure between the two culture models. The decision to utilise

submerged monolayer cultures or ALI cultures is dependent on the experimental

questions raised and the assessment methodologies used, as data from this study has

clearly demonstrated and supported the need for ALI cultures in contrast to data from

submerged monolayer culture when assessing barrier functionality due to the ability of

ALI cultures in establishing barrier integrity and function. However, in a previous study

by Hackett and colleagues (2011), they demonstrated significantly higher levels of the

basal cell marker, cytokeratin-5 in asthmatic epithelial cells, indicating that asthmatic

cells are more basal-like. Hence, in this circumstance, the use of submerged monolayer

cultures with only a single cell type for the assessment of membrane TJ protein

expression prior and post HRV infection would also be suitable, in addition to the use of

more sophisticated methods such as confocal microscopy and stereological assessments.

Collectively, this study has demonstrated, in addition to the applicable use of confocal

microscopy and stereology for the assessment of TJ protein expression, the importance

of ALI cultures in the assessment of barrier functionality at basal levels or in response

to HRV infection to demonstrate the impact of infection in causing greater impairment

of barrier function within the asthmatic epithelium.

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Chapter 8: General Discussion

Asthma has often been regarded as a variety of disorders initiated at different phases

throughout life by a spectrum of environmental factors interacting with a susceptible

genetic background. Paediatric asthma is among the most common chronic conditions

worldwide, resulting in a substantial burden on the health care system, society as well as

families. Despite decades of asthma research, there have been no significant advances in

treatment, emphasising the need to investigate possible alternative cellular and

molecular mechanisms which could contribute to the heterogeneity of asthma. There is

increasing consensus that the respiratory epithelium as well as being a physical barrier

between the external environment and internal parenchyma, also plays important roles

in actively responding to noxious irritants, allergens and pathogens.

A number of studies have suggested that the epithelium is abnormal in asthma (Holgate

2007), however is unclear whether these abnormalities are intrinsic to asthma because

atopic status, a common association with asthma, has not been adequately accounted for

(Laberge et al. 1999; Wark et al. 2005; Contoli et al. 2006; Kicic et al. 2006; Qiu et al.

2007). Furthermore, most evidence has been derived from adult cohorts and is therefore

temporally dissociated from early disease initiating pathogenic factors. Thus, there is a

need for more direct information to determine whether observations regarding the

epithelium can be attributed to intrinsic abnormalities, a consequence of chronic

inflammation, related to an atopic predisposition. These gaps in the literature provide

the rationale for the current hypotheses that epithelial barrier integrity and function is

defective in children with asthma and that the defective barrier integrity and function in

asthma is independent of atopy. Finally, infection with respiratory viruses, in particular,

HRV, account for an estimated 90% of acute asthma exacerbations in children and have

been shown to be implicated in the development of asthma (Sly et al. 2006; Kusel et al.

2007; Kusel et al. 2012). Hence, this study also tested the hypothesis that epithelial

barrier integrity and function is compromised to a greater extent following HRV

infection in the asthmatic compared to healthy airway.

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This study firstly characterised the basal tight junction (TJ) expression of claudin-1,

occludin and ZO-1 within paediatric derived pAECs of non-asthmatic and asthmatic

children. The data demonstrated basal differences in TJ gene and protein expression

between non-asthmatic and asthmatic children, indicative of an intrinsic alteration in

epithelial barrier integrity within the asthmatic AECs. The rationale for including these

three TJ proteins stems from their crucial roles in regulating epithelial permeability, as

previously demonstrated (Farquhar and Palade 1963; Furuse et al. 1993; Roche et al.

1993; Nusrat et al. 1995; Balda et al. 1996; McCarthy et al. 1996; Balda et al. 2000;

Coyne et al. 2003; Wang et al. 2003). Although studies have identified some

differences in epithelial barrier integrity in asthma (de Boer et al. 2008; Swindle 2009;

Xiao et al. 2011), there remains a lack of comprehensive analyses of the roles airway

epithelial TJ proteins play in supporting the epithelial barrier infrastructure and

regulating epithelial permeability. Moreover, a paucity of data within the paediatric

asthmatic population further fuels the need for paediatric junctional complex analyses as

studies have shown the effects of early life environmental challenges resulting in the

development of asthma (Sly et al. 2006; Kusel et al. 2007). This study addresses the

current gaps within the literature and from the observed data, the intrinsically altered

expression in asthmatic AECs could translate to an inherently impaired barrier function.

This suggests an increased propensity for allergic sensitisation or the induction of

inflammatory responses following pathogenic challenge. Furthermore, intrinsic

differences between the non-asthmatic and asthmatic epithelial cells implicate the

airway epithelium as a potential therapeutic target to reduce predisposition or to

ameliorate disease severity in asthma through improvement of barrier integrity and

function or limiting viral replication.

Interestingly, the data indicated a discordance between the ex vivo gene and protein

expression within asthmatic AECs. This could suggest direct transcriptional repression

or post-transcriptional regulation by various factors including the pleotropic

transcription repressor Snail, which act to down-regulate expression of TJ gene

components required for junctional complex formation or decrease translation of

junctional proteins (Ohkubo and Ozawa 2004). T-helper type-2 cytokines have also

been implicated such as IL-4 and IL-13 which has been shown to inhibit cellular

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migration and wound healing respectively (Ahdieh et al. 2001), This, in turn leads to a

diminished epithelial barrier integrity and consequently, a possible decreased barrier

function. Other factors that might account for the discordance in the observations could

be host microRNAs such as miR-122a, which are involved in the degradation of

occludin as well as the involvement of miR-155 in post-transcriptional repression of

claudin-1 expression, resulting in decrease barrier functionality. Further studies are

required in this area to identify novel molecules that could be translated into a

personalised therapeutic regimen for the improvement of impaired barrier function in

asthma.

Although a previous study by Xiao and colleagues (2011) demonstrated increased

airway epithelial permeability in adults with moderate and severe asthma, little is

known about airway epithelial permeability in paediatric asthma, whether it is intrinsic

or a consequence of airway inflammation. In the current study, basal barrier function of

ALI cultures established from paediatric individuals with mild, stable asthma and not on

corticosteroid treatment for at least 3 months showed no significant difference in the

levels of epithelial permeability compared to non-asthmatic counterpart However, future

studies involving the assessment of barrier functionality in paediatric individuals with

moderate to severe asthma would anticipate an increase in epithelial permeability.

Moreover, when assessing epithelial permeability towards various sized inert molecules,

data demonstrated significantly increased permeability towards smaller sized molecules

in both non-asthmatic and asthmatic epithelial cells. These observations suggest that the

asthmatic airway epithelium has increased susceptibility towards small molecular sized

allergens or pathogens including house dust mites, fine particulate matter in diesel

exhaust, cigarette smoke extract or respiratory viruses. With the increased facilitation of

haptens, allergens or pathogens across the airway epithelium, this may translate to an

increased predisposition towards allergic sensitisation and / or viral infection. Repeated

exposure to allergen and / or viral infection could subsequently result in co-infection

with other micro-organisms such as bacteria, as demonstrated by Sajjan and colleagues

(2008). Having shown differences in basal epithelial permeability between non-

asthmatic and asthmatic paediatric airway epithelium, the next focus will be

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determining the pathways behind these differences, through the integration of genomic

or mRNA profiles to advance the current understanding of tight junctional complexes.

Collectively, these observations are to the first to implicate an association between

basally dysregulated barrier integrity and impaired barrier function towards small-sized

particles in paediatric asthma.

Data from the second approach demonstrated differential expression of basal TJ

complexes in non-asthmatic and asthmatic AECs that were atopic, suggestive of the

involvement of atopy in altering TJ expression. Despite ongoing controversy about the

relationship between atopy and asthma, there is substantial evidence supporting the role

of atopy in asthma development and progression (Shirakawa et al. 2000; de Boer et al.

2008; Holgate et al. 2009; Scott et al. 2010; Spergel 2010; Baraldo et al. 2012;

Lambrecht and Hammad 2012). However, most studies that have investigated the

contributing role of atopy to the disruption of TJ complexes, have utilised the epidermal

layer (Palmer et al. 2006; Weidinger et al. 2006; De Benedetto et al. 2011; Kuo et al.

2012) and few have examined the impact of atopy on TJ complex expression within the

respiratory airways, let alone, the interaction of atopy with asthma in altering TJ

complex expression. Furthermore, the only study that have attempted this, utilised non-

atopic non-asthmatic and atopic asthmatic subjects, raising the issues of appropriate

controls and whether the reported data adequately addressed the significance of atopy in

altering junctional complex expression in asthma (de Boer et al. 2008). Thus, this study

established baseline values for tight junctional complex expression of non-asthmatic

and asthmatic AECs with or without atopy. Interestingly, basal TJ complex expression

profiles of pAECHA were observed to be significantly different from pAECHNA and

resembled those of pAECAA. This indicates the contribution of atopy in the alteration of

tight junctional complex expression and consequently, a probable increase in epithelial

permeability towards aeroallergen sensitisation or viral infection.

In the assessment of barrier integrity in the presence of atopy, the only study to have

been performed have been in the assessment of atopic dermatitis (De Benedetto et al.

2011). At present, there remains a paucity of data on the influence of atopy on tight

junctional function of the respiratory airways. Pilot data from this study demonstrated

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pAECHA had no significant difference in basal epithelial permeability compared to

pAECHNA. These findings are in contrast with those observed by De Benedetto and

colleagues (2011) which could likely be attributed to the different epithelial sites

sampled. An alternative is the suggestion that pAECHA could in fact, be a ‘pre-

asthmatic’ phenotype and as data obtained showed, pAECHA ALI cultures had

significantly higher basal permeability towards small sized molecules, which could

indicate increased susceptibility towards fragments of aeroallergens or respiratory

viruses, thereby facilitating sensitisation or infection, eventually leading to the induction

of asthma.

Although this study has shown differences in epithelial barrier integrity and function

between non-atopic and atopic AECs, the major limitation of the study was the

availability of non-atopic asthmatic AECs and the inability to perform longitudinal

follow-up assessment. Nonetheless, future work to further define the relationship

between atopy and barrier integrity and function should categorise the type and severity

of atopy and correlate this with the level of impairment in barrier integrity and barrier

functionality could then be subsequently assessed. Focussed PCR arrays could also be

performed to determine the magnitude difference in gene expression of TJ complexes

between atopic and non-atopic AECs. Furthermore, genomic and proteomic pathway

analysis would elucidate the effects of atopy on TJ disassembly and epithelial

permeability. Collectively, these observations highlight the significance of atopy in the

influence of tight junctional complex formation and the contribution towards altering

epithelial barrier function in health and disease.

In a seminal study, respiratory viral infections were shown to cause TJ complex

disorganisation and altered epithelial permeability AECs from non-asthmatic (Sajjan et

al. 2008). Hence, very little is known about TJ complex expression and function in

asthmatic AECs following viral insult. Therefore, the hypothesis that HRV infection

differentially disrupts TJs and barrier function in asthmatic compared to non-asthmatic

AECs was tested in the final set of experiments.

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Data demonstrated significant disassembly of TJ complex following 24 h HRV-1B

infection in both non-asthmatic and asthmatic AECs but 48 h post infection, a

restoration towards non-infected levels of expression was observed for non-asthmatic

cultures, while, diminished TJ expression levels was sustained in asthmatic cultures.

When barrier function was assessed at 24 h, data demonstrated significantly reduced

basal RT in asthmatic compared to non-asthmatic AECs and was concomitant with an

increased epithelial permeability. Although not performed in the current study, future

experiments to assess epithelial permeability at 48 h would anticipate a restitution of

barrier functionality in non-asthmatic epithelial cells concomitant with the observed

restoration of membrane TJ protein levels to non-infected levels. However, whether

decreased barrier functionality in asthmatic epithelial cells would be sustained remains

to be answered

When the data was stratified according to atopy, pilot data demonstrated significant

decrease in TJ expression following 24 h HRV-1B infection in pAECHNA cultures with

restoration to non-infected levels 48 h post infection. Interestingly, the decrease in TJ

expression in pAECHA cultures was sustained at 48 h after infection. This profile of

sustained decrease was similar to both pAECNAA and pAECAA cultures. The evidence

presented suggests that HRV-1B infection in conjunction with the presence of either

atopy and / or asthma could contribute to the sustained disorganisation and disassembly

of TJ complexes. This would have serious implications, especially for individuals with

asthma as continued disassembly of TJ complexes could potentially translate to a

prolonged compromised barrier state. Hence, the likelihood of allergens, haptens or

pathogens traversing the epithelial layer into the sub-epithelial and endothelial layers

would be significantly increased. The consequence of such an increased epithelial

permeability could enhance airway inflammation in a manner which leads to disease

exacerbations through generation of a variety of pro-inflammatory cytokines including

IL-1β, IL-6, GM-CSF and IL-11 (Einarsson et al. 1996; Terajima et al. 1997; Sanders et

al. 1998; Sanders et al. 2001) as well as chemokines such as IL-8, IP-10 and RANTES

(Subauste et al. 1995; Schroth et al. 1999; Donninger et al. 2003; Spurrell et al. 2005),

ultimately resulting in chronic airway inflammation and eventually, structural

remodelling (Kumar et al. 2002; Johnson et al. 2004; Grainge et al. 2011).

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As this study is the first to investigate the relationship between atopy and HRV

infection on tight junctional complex expression, there remains a lack of comparative

literature from which to draw significant conclusions. Hence, further investigations

involving genomic and pathway analyses for TJs of asthmatic AECs in the presence or

absence of atopy could provide candidate TJ genes of interest for this compensatory

effect. In addition, matched investigations which further categorises the type and

severity of atopy with the level of dysregulation in barrier integrity and impairment in

barrier function are warranted to elucidate the contributions of atopy on regulating TJ

complexes during HRV infection.

Current findings have highlighted the emergence of HRV-C as a predominant trigger of

asthma exacerbations in children (Evans et al. 2002; Stevens 2009; Baraldo et al. 2012;

Cakebread et al. 2014). Furthermore, recent studies have successfully propagated HRV-

C in vitro through the use of well-differentiated air-liquid interface cultures (Ahdieh et

al. 2001; Ohkubo and Ozawa 2004; Tillie-Leblond et al. 2007; Schneider et al. 2012).

The present data has demonstrated a sustained decreased of TJ complex expression in

pAECHA, pAECNAA and pAECAA except pAECHNA at extended infection periods.

However, limited availability of paediatric AEC cultures and time constraints has

prevented the present investigation from determining the effects of extended HRV

infection as well as repeated viral infection on barrier functionality. The next phase of

the study may utilise HRV-C to assess the effects on TJ complex disassembly and

barrier function impairment in non-asthmatic and asthmatic AECs in the presence and

absence of atopy as well as the effect of time. Genomic and proteomic pathway analysis

could be performed in conjunction with focussed qPCR arrays to elucidate the potential

pathways in which HRV-C infection is capable of altering tight junctional complex

expression. Furthermore, in vivo experiments utilising appropriate mouse models which

simulate the different facets of asthma could be performed to provide additional insights

on the effects of HRV infection on the airway epithelium.

The initial studies focussed on TJ proteins as past investigations have implicated TJ in

the regulation of epithelial permeability. Based on these preliminary findings, future

studies should also examine the global expression and function of tight junctional

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167

complexes such as junctional adhesion molecules (JAMs), tricellulin, ZO-2, ZO-3 as

well as the other members of the claudin family as well as their relationship with barrier

functionality. The present findings highlight the potential relationship between atopy

and asthma in altering tight junctional complex formation. However, future

investigations with increased numbers of non-atopic asthmatic participants would better

define this relationship and the effects on barrier integrity and permeability. In addition,

the current study has also demonstrated the effects of HRV infection on altering

epithelial permeability. The increase in epithelial permeability would result in increased

susceptibility to additional insults from aeroallergens, haptens or pathogens, resulting in

airway inflammation and / or asthma exacerbations. Conversely, innate immune cells

such as dendritic cells (DCs), which have been shown to express TJ proteins (Sung et

al. 2006), often interact directly with TJ proteins to sample the airway lumen without

disruption of the epithelial barrier (Takano et al. 2006; Blank et al. 2011; Veres et al

2011). However, in the presence of HRV infection, as shown in the current findings, the

presence and interaction of certain subsets of DCs with epithelial cells could potentially

result in further perturbation of epithelial integrity and permeability through increased

exposure surface to haptens, allergens or pathogens, resulting in further allergic

sensitisation or infection of the airways (Jahnsen et al. 2001; Jahnsen et al. 2006). This

would have serious consequences in the initiation and perpetuation of asthma

exacerbations. Future studies should examine the interaction between DCs or other cells

of innate immunity such as neutrophils with airway epithelial cells and the resulting

effect on bronchial epithelial integrity and permeability in non-asthmatic and asthmatic

epithelial cells with or without atopy.

In summary, epithelial barrier integrity is intrinsically altered in asthmatic compared to

non-asthmatic AECs and that this intrinsic characteristic translates to impaired epithelial

barrier function towards small sized molecules. Furthermore, this investigation has

provided pilot data on the possible contribution of atopy to the dysregulation of barrier

integrity and altering barrier function. These characteristics were further examined by

exposing AECs to HRV. A greater impact of HRV on barrier integrity in asthmatic

compared to non-asthmatic AECs was observed. In addition to demonstrating that

asthmatic AECs have an intrinsically different tight junctional complex expression and

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168

function, this study has also raised the questions of the impact of early life sensitisation

and infection in causing dysregulation of barrier integrity and function. Finally, this

investigation has provided new insight regarding important intrinsic epithelial

vulnerability in asthma and has provided further rationale for investigating epithelium-

centred asthma therapies in young children with asthma.

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169

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Appendix A

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Appendix B

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Appendix C

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Appendix D

CCATTATGGCGTGTAAAGTCA

TGAGCACTGGAGAGAAAGGA PPIA

Reverse

Forward

CTGGGTAAAAAGAGTAGGCTGGC Reverse

AGGCCTGATGAATTCAAACCG Forward Occludin

TCTTCTGCACCTCATCGTCTT Reverse

GGCAGATCCAGTGCAAAGTC Forward Claudin

CCGCCCGCTCCTCACGCCACAG Reverse

GCCCGTGCCCCGCTCGCTCTC Forward ZO-1

Sequence Primer Gene

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Appendix E

1:

pAECNon-asthmatic Claudin-1 Occludin ZO-1

Claudin-1 * *

Occludin * *

ZO-1 * *

2:

pAECAsthmatic Claudin-1 Occludin ZO-1

Claudin-1 * *

Occludin * *

ZO-1 * *

*, p<0.05

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Appendix F

1:

pAECHNA Claudin-1 Occludin ZO-1

Claudin-1 * *

Occludin * *

ZO-1 * *

2:

pAECHA Claudin-1 Occludin ZO-1

Claudin-1 * *

Occludin * *

ZO-1 * *

3:

pAECNAA Claudin-1 Occludin ZO-1

Claudin-1 * *

Occludin * *

ZO-1 * *

4:

pAECAA Claudin-1 Occludin ZO-1

Claudin-1 * *

Occludin * *

ZO-1 * *

*, p<0.05

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Appendix G

1:

pAECNon-asthmatic Claudin-1 Occludin ZO-1

Claudin-1 N.S *

Occludin N.S *

ZO-1 * *

2:

pAECAsthmatic Claudin-1 Occludin ZO-1

Claudin-1 * N.S

Occludin * *

ZO-1 N.S *

*, p<0.05

N.S, Non-significant

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Appendix H

1:

pAECHNA Claudin-1 Occludin ZO-1

Claudin-1 * N.S

Occludin * *

ZO-1 N.S *

2:

pAECHA Claudin-1 Occludin ZO-1

Claudin-1 * N.S

Occludin * *

ZO-1 N.S *

3:

pAECAA Claudin-1 Occludin ZO-1

Claudin-1 * N.S

Occludin * *

ZO-1 N.S *

*, p<0.05

N.S, Non-significant

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